Recombinant Protein Production Potential of South African Microalgae

Recombinant protein production potential of South African microalgae

Julanie Stapelberg

Submitted in partial fulfilment of the requirements for the degree of

MAGISTER SCIENTIAE: PLANT BIOTECHNOLOGY

In the Faculty of Natural and Agricultural Sciences

University of Pretoria

Pretoria

May 2019

Supervisor: Dr Bridget Crampton

Co-supervisors: Dr Robyn Roth and Dr Michael Crampton DECLARATION

I, Julanie Stapelberg declare that the thesis, which I hereby submit for the degree of Magister Scientiae at the University of Pretoria, is my own work and has not previously been submitted by me for a degree at this or any other tertiary institution.

SIGNATURE:

DATE:

Recombinant protein production potential of South African microalgae

Julanie Stapelberg

Supervisor: Dr Bridget Crampton

Co-supervisor: Dr Robyn Roth

Co-supervisor: Dr Michael Crampton

Department: Plant and Soil Sciences

Degree: Magister Scientiae

SUMMARY

Therapeutic recombinant proteins, including complex glycoproteins, antibodies and other small molecules, have many current and future applications in biopharmaceuticals, biomaterials, nutraceuticals, agriculture, animal health and cosmeceutical industries (Rasala & Mayfield, 2015). Existing expression systems used to produce these proteins include mammalian, plant, insect and microbial culture systems, with each portraying their own strengths and weaknesses (Andersen & Krummen, 2002). For mammalian cell cultures, proteins, including those for human and animal function, can be manipulated and produced in their active form but the system is costly. Microbial systems are less expensive but may not have the necessary transcription, translation and post- translational tools to produce viable proteins without further alterations (Rasala & Mayfield, 2015). Therefore, the current expression systems available to produce recombinant proteins lack either in their ability to produce viable proteins, have low protein yields, and are not easily manipulated. Some of these systems also incur high a cost of processing. The benefit of a novel expression system, through algal culture, could overcome many challenges by offering low production costs, fast growth rates, easy culture, simple transgenic manipulation, and modified abilities of transcription and translation (Feng et al., 2014).

The most advanced genetic work that has been done on green algae is that of Chlamydomonas reinhardtii. The nuclear, chloroplast and mitochondrial genomes of C. reinhardtii have been sequenced with optimised transformation vectors and protocols created for each (Merchant et al., 2007). The nuclear genome has already been engineered to successfully express an assortment of soluble proteins, enzymes or immunotoxins of therapeutic or industrial value. While C. reinhardtii is the model microalgae currently used in research, it may not be the best strain for large scale production of heterologous proteins (Rasala & Mayfield, 2015). Other microalgae strains might have faster growth rates and tolerate commercial culturing conditions better (Taunt et al., 2018).

Microalgae represent a rich collection of species that offer numerous advantages for both biotechnology and biomedical industries (Doron et al., 2016). Many South African microalgae species have not been identified, let alone characterised for their biotechnology potential. The aim of this project was to establish and optimise the nuclear transformation protocols of C. reinhardtii within our current South African laboratory conditions. A previously successful vector system

harbouring a C. reinhardtii (Cr) codon optimised green fluorescent protein (CrGFP) was used along with endogenous algal regulatory elements for an optimised molecular toolkit. South African microalgae isolated across the country were then cultured, screened and identified by genetic sequencing. From a database of over 500 indigenous microalgae isolates, nine isolates indicated potential as promising protein expression platforms. This project lays down the foundation for initiating a heterologous microalgae protein expression system within South Africa.

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PREFACE

This Thesis is divided in four chapters as set out below:

Chapter 1: Introduction to microalgae and their application as an expression system for recombinant protein production. This Chapter provides detail on green microalgae, on their structure, reproduction, habitats, environment and growth requirements. The role microalgae may play in the economy and the benefit of microalgal expression systems over current recombinant protein production platforms is discussed. Further details are then provided on the transformation methods, including nuclear and chloroplast transformation comparisons, and the molecular toolkit that has been explored for microalgal protein expression.

Chapter 2: Transformation of two strains of the model green microalgae C. reinhardtii is described in this Chapter. A gene construct containing the C. reinhardtii codon optimised Crgfp transgene driven by the Hsp70-rbSc2 promoter with she ble antibiotic resistance gene as a selectable marker was electroporated into the nuclear genome. The CrGFP production was then quantified in live microalgae cells by fluorescence.

Chapter 3: In this Chapter selected South African microalgae isolates, obtained from the microalgal culture collection of South Africa (MiCCSA), were subjected to growth curve analysis in comparison to C. reinhardtii. Growth conditions were on both mixotrophic and freshwater media with a variety of light intensities. The microalgae culturing conditions were also optimised. The South African microalgae with the fastest growth rates were identified based on compound microscopy and 18S rRNA Sanger sequencing.

Chapter 4: This Chapter outlines the general conclusion of this study, proposes recommendations for future work, as well as exploring the broad potential of microalgae for South Africa’s bio-economy.

Aspects of this study were presented at the following conferences:

The 8th International Conference on Algal Biomass, Biofuels and Bioproducts. Seattle, USA. Stapelberg J, Roth RL, Crampton MC, Crampton BG. June 2018. Development of a heterologous expression platform in indigenous South African microalgae. (Poster Presentation)

The 31st Congress of the Phycological Society of Southern Africa. Hartbeespoort, South Africa. Stapelberg J, Roth RL, Crampton MC, Crampton BG. July 2018. The recombinant protein production potential of South African microalgae. (Poster Presentation)

South African Society for Bioinformatics Conference, Golden Gate, South Africa. Stapelberg J, Roth RL, Crampton MC, Crampton BG. October 2018. Why we should focus on microalgal biotechnology in South Africa. (Oral Presentation)

South African Genetics Society Conference, Golden Gate, South Africa. Stapelberg J, Roth RL, Crampton MC, Crampton BG. October 2018. The identification, growth analysis and biotechnological potential of South African microalgae. (Poster Presentation)

Chapter 3 has been written up as a journal article and will be submitted to a peer review journal for publication.

ACKNOWLEDGEMENTS

There are many people whom I would like to thank; this MSc was not simply a project for a degree but has taught me so many fundamental skills and has developed my passion for microalgae research. I truly believe that microalgae are invaluable to making our planet sustainable, with endless potential in several industries. (There is currently an entire building powered by algae in Germany with plans for an entire sustainable algae city in Sweden). I am therefore most grateful to everyone who allowed me the opportunity to pursue this passion.

Thank you to my supervisor and role model, Dr Bridget Crampton, who knew I was interested in a microalgae protein expression and who opened the door to this MSc project. To Dr Michael Crampton as the initiator at the CSIR and to Dr Robyn Roth who was a constant support and who understood that I work in the laboratory at night and therefore granted me the freedom to experiment and learn from all my failures. To Monique Smit for tolerating the constant presence of my microalgae cultures in the Phycology room and for informing me when the laboratory moved locations. Thank you to the kind staff at CSIR who allowed me to use their equipment, and to all my colleagues, lab mates and my friends for the mutual motivation. Thank you to my funding bodies, the University of Pretoria and National Research Foundation for providing the resources and the CSIR for supporting project costs.

A hearty thanks also to all the members, staff and friends at the Forestry and Agricultural Biotechnology Institute (FABI for allowing me to work there so that I may support my master’s studies and for allowing me to present my work- despite it being a somewhat off-topic from foci- Eucalyptus trees, avocadoes or fungi. Thank you especially to Dr Bernard Slippers for always challenging my love for microalgae and urging me to actively research its pros compared to fungi.

A sincere thank you to the SANPCC community who allowed me the opportunity to attend their conference and get further inspired by the wonderful work South Africa is doing on algae.

And finally, I owe a huge debt of gratitude to my family for their love and encouragement and continued support of my research passions. You mean more to me than you would ever know. Lastly to John Rogers, for his innate understanding, patience and reassurance.

I am grateful for everyone on this journey, it has been an absolute privilege!

Table of Contents CHAPTER 1

1.1 General Introduction to Algae…………...... 1 1.2 Algal Culturing Techniques ...... 6 1.3 Recombinant Protein Production ...... 8 1.4 Algae in Biotechnology...... 12 1.4 Aims and Objectives ...... 20 1.6 References ...... 21

CHAPTER 2

Abstract ...... 29

2.1 Introduction ...... 30

2.2 Materials and Methods ...... 33

2.3 Results ...... 40

2.4 Discussion ...... 50

2.5 Conclusion ...... 57

2.6 References...... 58

CHAPTER 3

Abstract ...... 62

3.1 Introduction ...... 63

3.2 Materials and Methods ...... 67

3.3 Results ...... 73

3.4 Discussion ...... 84

3.5 Conclusion ...... 91

3.6 References…………………………………………………………………………………………………………………………..92

CHAPTER 4

4.1 Introduction ...... 99

4.2 Final Conclusion ...... 105

4.3 References...... 106

APPENDICES

Appendix A ...... 110

Appendix B ...... 112

Appendix C ...... 113

Appendix D ...... 114

Appendix E ...... 116

Appendix F ...... 117

Appendix G ...... 118

Appendix H ...... 119

Appendix I ...... 131

Appendix J ...... 137

References ...... 138

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List of Tables and Figures

TABLES

Table 1.1. Description of typical Chlorophyte cellular structures………………………………………..….………….4 Table 1.2. Summary of the advantages of a microalgae expression platform compared to aspects of other expression systems…………………………………………………………………………………………………....………….11 Table 1.3. Current successes in microalgal recombinant protein production………………………...………..12 Table 1.4. Comparison of transgene expression in the nucleus and chloroplast of microalgae………..17 Table 1.5. Selectable markers used for screening for positive microalgae transformants………..….…..18 Table 1.6. Promoters and terminators for transgene expression within microalgae……………………..…20 Table 2.1. The detailed genotypes and fundamental cell properties of two C. reinhardtii strains….…33 Table 2.2. Primers used in this study for amplification of the Crgfp transgene………..…….………………..34 Table 2.3. The rGFP dilutions utilised to generate the fluorescent standard curve in this study……...39 Table 2.4. Influence of Crgfp-pChlamy4 linearized plasmid volume on electroporation arcing and number of colonies produced on a plate………………………………………………………………….……………………..45 Table 2.5. Transformation data for the C. reinhardtii strains CC-125 and CC-400….………………..……….46 Table 3.1. The 18S rRNA primers used in this study for the classification of microalgae………………….70 Table 3.2. List of additional microalgae GenBank accession numbers used in this study.…………………71 Table 3.3. The growth rate, maximum OD cell density and estimated cell concentration of each microalgae isolate grown on either AF6 or TAP media…………………………………………………….………………76 Table 3.4. Identification results from the BLASTn (NCBI) analysis for each South African microalgal isolate……………………………………………………………………………………………………………….…………………………….78

FIGURES

Figure 1.1. The broad classification of algae into their different divisions, with Chlorophyta further classified into classes. Image redesigned from (Kumar, 2014)…………………………………………………………..1 Figure 1.2. The common photosynthetic algae in relation to the visible colour spectrum and water depth in an aquatic environment (Bolduc, 2015)………………………………………………………………………………2 Figure 1.3. Structure range of the Chlorophytes. From the microscopic unicellular green microalgae Chlamydomonas reinhardtii (Library, 2017), to coenobium Volvox carteri (von Der Heyde et al., 2015), filamentous Rhizoclonium ramosum (Zhi–Juan et al., 2016), and multicellular marine Ulva rotundata (Goff, 2011). Diagram (Stapelberg, 2018)……………………………………………………………..………….3 Figure 1.4. Diagram indicating the structure of unicellular Chlamydomonas Chlorophyte containing general microalgal features. Adapted (Adams, 2011)…………………………………………………………………..……3 Figure 1.5. Diagrammatic representation of the sexual and asexual reproduction of the microalgae, Chlamydomonas (Gastineau et al., 2014)………………………………………………….……………………………………..5 Figure 2.1. KAPA HiFi PCR amplification of the Crgfp gene from the pCrGFP plasmid ………………..……40 Figure 2.2. A colony PCR amplification of the Crgfp region in Crgfp-pBSK containing E. coli colonies……………………………………………………………………………………………………………………………………………41 Figure 2.3. Restriction enzyme double digest of recombinant plasmid Crgfp-pBSK …………………………42 Figure 2.4. Restriction enzyme double digest of plasmid pChlamy4 with BamHI and XbaI……………...42 Figure 2.5. A schematic diagram showing the Crgfp transgene cloned into the pChlamy4 vector…...43 Figure 2.6. PCR amplification of the Crgfp region of Crgfp-pChlamy4 plasmids……………………….……….43

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Figure 2.7. Confirmation of the correct Crgfp-pChlamy4 vector by Sanger Sequencing alignment…..44 Figure 2.8. Restriction enzyme digest of recombinant plasmid Crgfp-pChlamy4 with ScaI………………44 Figure 2.9. A zeocin kill curve of C. reinhardtii strains CC-125 and CC-400 ………………….…………………..45 Figure 2.10. Colonies after electroporation transformation of the CC-125 and CC-400 strains………..46 Figure 2.11. Growth curves of the untransformed and Crgfp transformed C. reinhardtii strains …….47 Figure 2.12. The C. reinhardtii CC-125 and CC-400 transformed cells seen with (A) light microscopy and fluorescence microscopy of (B) transformed and (C) untransformed cells………..………………………48 Figure 2.13. Quantiﬁcation of the relative CrGFP ﬂuorescence of four selected (A) CC-400 and (B) CC- 125 transformed whole cell transformants…………….……………………………………………………………………….49 Figure 2.14. The C. reinhardtii (A) CC-125 and (B) CC-400 colony PCR screen for Crgfp-pChlamy4 transformants …………………………………………………………………………………………………………………………………49 Figure 2.15. The exponential decay waveform as a measure of T pulse which is the rate of which the voltage decays to 1/e (~37%) of the peak value (BioRad, 1988)………………………………………………….……51 Figure 3.1. The five phases associated with the growth of a closed system microalgal culture……….63 Figure 3.2. The comparison of nitrogen, magnesium, sulphur, phosphorous, chlorine and calcium nutrient concentrations levels utilised in AF6 and TAP medium………………………………………………………65 Figure 3.3. The respective locations of over 500 Indigenous freshwater microalgae isolated across South Africa as part of the MiCCSA database………………………….……………………………………………………….67 Figure 3.4. Schematic diagram of the 6 well microplates used in the growth analysis …………………….68 Figure 3.5. Stock cultures of the selected 40 South African microalgae grown on AF6 media………….74 Figure 3.6. Growth curve analysis of 11 microalgae isolates with 8-hour light (250 μmol m-2.s-1)/ 16- hour dark phase cycles in AF6 medium.……………………………………..………………………………………….………..74 Figure 3.7. Growth curve analysis of 11 microalgae isolates with 8-hour light (250 μmol m-2.s-1)/ 16 hour dark phase cycles in TAP medium…………………………………………………….……………………………………..75 Figure 3.8. The biomass production for each microalgae isolate on AF6 and TAP media after a period of 14 days………………………………………………………………..……………………………………………………………….……..75 Figure 3.9. Maximum likelihood molecular phylogenetic analysis of 18s rRNA microalgae isolates………………………………………………………………………………………………………………………………….…………80 Figure 3.10. Contamination of TAP medium microalgae stock cultures ………………………….………….……81 Figure 3.11. A zeocin kill curve of the South Africa micoalgal isolates……………………………………………..82 Figure 3.12. Microalgae rejuvenation growth in liquid TAP media after cryopreservation……….….…..83 Figure 4.1. Closed systems for cultivating microalgae………………………….…………..……………………………101 Figure 4.2. The input for microalgae biomass may produce a range of products which have an inverse relationship between the volume and cost of products produced……..……………..………………104

LIST OF ABBREVIATIONS

6× His Polyhistidine tag AD Anno domini ANOVA Analysis of variance Bp Base pairs BC Before the common era CaMV 35S Cauliflower mosaicvirus 35S CFSE Carboxyﬂuorescein diacetate succinimidylester CFU Colony forming units

CO2 Carbon dioxide Cox1 Cyclooxygenase gene 1 CrGFP C. reinhardtii codon optimised green fluorescent protein CRISPR Clustered Regularly Interspaced Short Palindromic Repeats CSIR Council for Scientific and Industrial Research CTAB Cetyl trimethylammonium bromide ddH20 Double distilled Millipore water DNA Deoxyribonucleic acid dNTPs Deoxyribonucleotide triphosphate DFP Dual fluorescent protein EDTA Ethylene diamine triacetic acid ELISA Enzyme-linked immunosorbent assay EtBr Ethidium bromide FDA US food and drug administration g Gravitational force GAPDH Glyceraldehyde-3-phosphate dehydrogenase GFP Green fluorescent protein GMO Genetically modified organism GRAS Generally regarded as safe hGH Human growth hormone HSM High salt medium Hsp70A Heat shock protein 70A IPTG Isopropyl β-D-1-thiogalactopyranosid ITS Internal transcriber spacer region Kbp Kilobase pairs

kDA Kilo dalton LB Lysogeny broth MAT Mating type MCL Maximum composite likelihood MCS Multiple cloning site MiCCSA Microalgal culture collection of South Africa mRNA Messenger RNA mt- Minus mating type mt+ Plus mating type NADPH Nicotinamide adenine dinucleotide phosphate NCBI National centre for biotechnology nos Nopaline synthase promoter NPQ Non-photochemical quenching OD Optical density PCR Polymerase chain reaction PPFD Photosynthetic photon flux density psaD Photosystem I protein D R2 Regression coefficient rbcS2 Ribulose bisphosphate carboxylase small chain 2 RFU Relative fluorescent units rGFP Recombinant GFP ROS Reactive oxygen species RT-qPCR Reverse transcription polymerase chain reaction RuBisCO Ribulose-1,5-bisphosphate carboxylase/oxygenase SANPCC South African national phycology culture collection SOC Super optimal broth TAE Tris-acetate-EDTA TAG Triacylglycerols TAP Tris-acetate-phosphate TEV Tobacco etch virus Tm Melting temperature UTR Untranslated region UV Ultraviolet X-Gal 5-bromo-4-chloro-3indolyl-β-d-galactoside

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Chapter 1 Introduction to microalgae and their application as an expression system for recombinant protein production

Chapter 1

1.1 General Introduction to Algae

Algae is the name given to a large group of oxygenic, predominantly aquatic organisms within the Domain Eukarya and Kingdom Protista. The primary establishment of algae is that they are photosynthetic and are able to convert carbon dioxide (CO2) and water into carbohydrates, fats, and proteins through the utilisation of sunlight (Silva, 2016). Despite the photosynthetic similarity, algae lack complex structures associated with land plants such as roots, stems, leaves or vascular systems (Barsanti & Gualtieri, 2014). Eukaryotic algae are polyphyletic -they evolved from multiple ancestors - and therefore are extremely diverse. Based on their morphological structural, reproductive and molecular features, algae have been classified into 27 distinct phyla (Kumar, 2014). These phyla are branched across the tree of life where algal species can be found within the five main supergroups of eukaryotic organisms: Excavata, Chromalveolata, Rhizaria, Archaeplastida and Unikonta (Barsanti & Gualtieri, 2014). The classification of algae, over these five supergroups, has been described as six divisions (Figure 1.1). The first division is the Chlorophyta, known as green algae; the second Phaenophyta, referred to as brown algae; then Pyrrhophyta containing the dinoflagellates; Chrysophycophyta consisting of diatoms; Rhodophyta known as red algae and finally Euglenophyta containing the euglenoids (Kumar, 2014). These divisions are then further classified into classes, orders, family, genus and species level depending on the structure and organization of the cell, the presence of organelles, and the pattern of reproduction (Gerber, 2006).

Figure 1.1: The broad classification of algae into their different divisions, with Chlorophyta further classified into classes. Image redesigned from (Kumar, 2014).

1.1.1 Green microalgae Most algae are photosynthetic with pigments like those of terrestrial plants, namely chlorophyll a and chlorophyll b. They differ in their photoreceptors and accessory pigments which absorb light of different wavelengths (Figure 1.2). In algae, chlorophyll a is the predominant pigment, also found within plants and bacteria. Unique to algae however, is that they also have other forms of chlorophyll and accessory pigments such as carotenoids, xanthophylls, phycobillins (Barsanti & Gualtieri, 2014).

Chapter 1

Figure 1.2: The common photosynthetic algae in relation to the visible colour spectrum and water depth in an aquatic environment. Image redesigned from (Bolduc, 2015).

Further phototrophic niche differentiation occurs due to the absorption spectra of water, the main habitat of algae. This allows algae to utilise light across the colour spectrum consisting of red, far- red, orange, green and blue light which infiltrates the water less as the water depth increases (Rockwella et al., 2015). Algae in shallow water of approximately 50 m or closer to the water surface, would contain accessory pigments which absorb the longer wavelength of red light. These accessory pigments are green and consist of chlorophyll a and b, resulting in the common name of green algae (Bolduc, 2015). The same can be said for brown algae, which contain a brown accessory pigment - fucoxanthin, which absorbs orange light at a deeper water level of approximately 100 m (Kumar, 2014). As the water becomes deeper, less light can penetrate. As a result, at approximately 200 m only blue-green light is able to be absorbed by red accessory pigments - phycobiliproteins, from the red algal species (Metting, 1996).

The availability of light at the correct wavelength and intensity, is crucial for the growth of photosynthetic algae. Algae within the Chlorophyte division require the least water to cultivate and therefore they are desired for industrial purposes. To detect changes in the colour, intensity, and quality of light, most algae contain phytochromes which allow the algae to react and adapt structurally and physiologically to their environment (Barsanti & Gualtieri, 2014, Rockwella et al., 2015).

Green algae structure Structurally the size of Chlorophyta algae can range from tiny microscopic microalgae, less than 1 μm in diameter to larger macroalgae, such as seaweeds and giant kelp which can reach up to 200 m in length (Bolduc, 2015). Microscopic algae can be organized in different conformations and can be unicellular, colonial (filamentous / coenocytic) or multicellular (Figure 1.3). They can be singular unicellular organisms or colonial - where the single-celled organisms come together and are surrounded by a gelatinous sphere. The colonial groups are either microscopic or macroscopic. Filamentous colonial groups are joined end to end and can be either branched or unbranched arrangements (Kumar, 2014).

Chapter 1

The colonial algae can even be complex, multicellular structures such as seaweed. Most multicellular algae are marine species which have a shaped vegetative body referred to as a thallus (Bolduc, 2015). Microalgae may also be motile or nonmotile, free floating or benthic (attached to a surface such as rocks or even other organisms). Most chlorophytes are unicellular, and microalgae are easier to cultivate, with faster growth rates and easier transformation (Richmond & Hu, 2013).

Figure 1.3: Structure range of the Chlorophytes. From the microscopic unicellular green microalgae Chlamydomonas reinhardtii (Library, 2017), to coenobium Volvox carteri (von Der Heyde et al., 2015), filamentous Rhizoclonium ramosum (Zhi–Juan et al., 2016), and multicellular marine Ulva rotundata (Goff, 2011). Diagram (Stapelberg, 2018).

Though structural characteristics may separate algae, like all living eukaryotes, microalgae contain the typical cellular organelles such as a mitochondria, a nucleus, an endoplasmic reticulum, a golgi apparatus, a plasma membrane and vacuoles (Silva, 2016). The unicellular microalgae also share unique Chlorophyte characteristics considering their flagella, stigma, pyrenoid, chloroplast and cell walls as indicated in the diagram of a Chlamydomonas cell (Figure 1.4) with further cellular descriptions (Table 1.1).

Figure 1.4: Diagram indicating the structure of unicellular Chlamydomonas Chlorophyte containing general microalgal features. Image redesigned from (Adams, 2011).

Chapter 1

Table 1.1: Description of typical Chlorophyte cellular structures

Microalgae Function Reference structure Flagella Single or multiple whip-like flagella allow the microalgae gliding (Tomaselli, or swimming motility. Movement may also occur by cellular 2004) protrusion where water enters through a pore one side of the cell and is expelled through a pore on the opposite side. Stigma/ Found within motile cells it senses light direction and intensity. (Barsanti & Eyespot Gualtieri, 2014) Pyrenoid Forms part of the algal carbon concentrating mechanism by (Meyer et al., fixing CO2 and maintaining a carbon dioxide rich environment 2017) for the enzyme, Ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO), as well as storing starch. Chloroplast For photosynthesis, the chloroplasts contain chlorophylls a and (Turmel et al., b, may be cup-shaped, tubular or even form elaborate 2006) networks. The number of chloroplasts per cell varies significantly between algae but is usually fixed in each algal species. In the algae C. reinhardtii there is only one prominent cup-shaped chloroplast per cell whereas Chara vulgaris can contain about 100 chloroplasts per cell. Cell walls Cell walls are often thin, rigid and made from different (Atkinson et carbohydrates, with cellulose as one of the major cell wall al., 1972) components. There are a few exceptions where cellulose is absent, and the cell wall is formed by other glycoproteins such as non-cellulosic polysaccharides in the case of Chlorella spp. and Volvox spp., or xylan and mannan in the case of Caulerpa spp.

Green microalgae ecology Structural adaptations have allowed Chlorophyte microalgae to populate a range of environmental conditions with differences in habitat, temperature, salinity, CO2 concentrations and other external factors (Tomaselli, 2004). Most green microalgae are primarily aquatic organisms with 10% in marine and 90% growing within freshwater; several algal species have even adapted to survive in brackish or wastewater (Tragin et al., 2016). Their distribution continues to be diverse as some green microalgae are terrestrial and are known to grow on land, trees, rocks and in moist soil.

Algae can exist as independent growth forms, or they can form symbiotic relationships with other organisms. Symbiosis proves mutualistic as the algae provide oxygen and nutrients whilst obtaining protection from predators. One common example is the plant-like composite organism, lichen, which consists of a symbiotic fungi surrounding the photosynthesising, green algae (Barsanti & Gualtieri, 2014).

Algae can grow in a range of temperatures, whilst the majority prefer temperatures between 14 and 30°C (Andersen, 2005). Some extremophiles inhabit abnormal environments, such as hot springs, deserts, frigid marine water or ice. The cryophilic algae Koliella Antarctica grows within ice (Vona et al., 2004), whilst the thermophilic algae, Cyanidium caldarium inhabits acidic hot springs with an upper temperature limit of 56°C (Doemel & Brock, 1971).

Chapter 1

Extremophiles are often able to endure stresses, for example: the marine Dunaliella spp. can tolerate temperatures up to 40°C and can grow with high salt concentrations up to 0.8 M (Walker et al., 2005). The majority of algae require CO2 to produce oxygen, but concentrations that are too high become toxic (Seckbach, 2015). Few acidophilic algae can tolerate high CO2 with pH values as low as 0.05 (Gross, 2000). Other physical parameters within an environment include the turbidity, nutrient and metal concentrations with different habitats leading to the adaption of diverse microalgae life strategies (Barsanti & Gualtieri, 2014). Some microalgae may adapt to a range of conditions, such as the single species Stichococcus sp. which can inhabit various environments from soil, glacier surfaces or fresh water (Latala, 1991).

Green microalgae reproduction strategies As the habitats of algae differ, so may their reproduction strategies. Many microalgae vary in their life cycle and reproduction, from simple asexual cell division in a stable environment to sexual reproduction -which may adapt to environmental changes, or a combination of both (Figure 1.5). Asexual reproduction is the division of a single parent cell to form genetically identical offspring (Peng et al., 2012). Division occurs through either binary fission where a cell splits in two, or by vegetative budding where the parental algae fragments into pieces and these cells may form a new colony (Barsanti & Gualtieri, 2014). In sexual reproduction, two parental cells each undergo individual meiosis to form gametes, which will form a zygote when united (Barsanti & Gualtieri, 2014). Sexual reproduction is generally controlled by environmental events and exchange of genetic material may lead to stronger and more environmentally adapted microalgae offspring (Gastineau et al., 2014). The reproduction of some species may be more complex with multi-generational cycles which alternate from asexual (haploid cells) to sexual (diploid cells) (Gastineau et al., 2014). It is of fundamental necessity to understand algae structure, physiology and reproduction for commercial cultivation.

Figure 1.5: Diagrammatic representation of the sexual and asexual reproduction of the microalgae, Chlamydomonas. Image adapted from (Gastineau et al., 2014).

Chapter 1

1.2 Algal Culturing Techniques The growth requirements of green algae are those associated with photosynthesis; namely light, water and CO2, as well as nutrients. Aspects influencing the growth kinetics of algal cultures may include an even greater variety of factors, such as the salinity (ionic strength and composition), temperatures, and pH values of the medium. Although algal species will differ in their optimal growth conditions, many green microalgae would be able to grow under the general growth requirements. Those that thrive would be considered the simplest and most cost-effective for commercial scale-ups.

1.2.1 General culture parameters Macronutrients Nutrients are elements required for growth and maintenance of life, and those for green microalgal growth are like those of higher plants. The majority of algae (95%) consists of hydrogen, carbon and oxygen provided naturally by the environment (Rasala & Mayfield, 2015), and those added to the medium can be categorized into essential macronutrients and some selected micronutrients. In the medium, macronutrients are required in a high concentrations of grams per litre whereas micronutrients are considered trace elements required in much smaller amounts of micrograms per litre (Procházková et al., 2014). Although algal species and strain specific responses have been identified in biomass culturing, the effect of these nutrients on microalgal physiology may follow a predominant pattern (Fields et al., 2014).

One of the most fundamental nutrients in algal culture is nitrogen (N), as it is required for the synthesis of proteins and nucleic acids - the building blocks of cells. In nature, nitrogen can be - + supplied in many forms including nitrate (NO3 ), nitrogen dioxide (NO2), ammonium (NH4 ), urea

((NH2)2CO) or other organic compounds (Mandal et al., 2018). Although nitrate is the most commonly supplied nitrogen source in algal cultures, ammonium is preferred as it can be directly incorporated into organic compounds and does not require the enzyme nitrate reductase to reduce from nitrate to nitrite, to ammonium (Procházková et al., 2014). The first instance of nitrogen depletion acclimatisation requires a variety of metabolic changes within the cells (Glibert et al., 2016). Microalgae start to synthesize nitrogen-scavenging enzymes for protein degradation, causing proteins such as RuBisCO, to be recycled into amino acids and new ribosomes (Sreedharan et al., 2018). A decrease in the photosynthetic activity of microalgae sees storage compounds being converted into those without nitrogen, such as triacylglycerides (TAGs) and starch (Procházková et al., 2014). If the cells do not acclimatise, then the lack of nitrogen would trigger the vegetative cells to undergo gametogenesis and form mating type gametes for zygote formation, which halts the growth process (Abe et al., 2004). A nutrient rich medium would therefore guarantee a high growth rate of microalgal cells (Glibert et al., 2016).

Another macronutrient essential for algal growth is phosphorous. Although microalgal biomass contains less than 1% of phosphorus it is absolutely crucial for the biosynthesis of phospholipids and nucleic acids in DNA and RNA (Deng et al., 2011), for the modification of protein function, and for energy transfer (Procházková et al., 2014). Algal cultures predominantly utilise inorganic phosphate 2- such as hydrogen phosphate (HPO4 ) and the uptake of phosphorus can be increased when exposed to high irradiance (Procházková et al., 2014, Andersen, 2005). A phosphorous deficiency in microalgae culture causes a reduction in the protein and chlorophyll content, causing defective cell division, lowers the rate of respiration and photosynthetic carbon fixation, and subsequently

Chapter 1 reduces growth rates (Sizova et al., 2001). Microalgal growth can be further affected by the sulphur content, a key macronutrient for cellular processes. Sulphur is required for the synthesis of many proteins, cell walls, membrane sulpholipids, thiol compounds that participate in the stress response, vitamins and the formation of disulphide bonds (Andersen, 2005). In microalgae medium, the most 2- cost-effective source of sulphur is the inorganic sulphate (SO4 ) and when there is a clear sulphur 2- deprivation, the gene expression is altered to ensure that SO4 is maintained by intracellular protein degradation (Procházková et al., 2014). To prevent the degradation of intracellular proteins, which will undoubtebly affect microalgal physiology, the amount of sulphur present in the medium should be sufficient (Andersen, 2005).

Micronutrients Required in a much smaller scale, most micronutrients can already be present in the medium without further addition and can include the following: iron, manganese, zinc, nickel, boron, vanadium, cobalt, copper, molybdenum and selenium, among others (Andersen, 2005). Most of these micronutrients are divalent metal cations and function in enzymes or as cofactors in many different metabolic pathways vital for microalgae to grow (Zhang, 2016). The essential macronutrients, resources such as light and CO2, and conditions such as pH can also influence the micronutrient requirements and uptake. For example, increased light intensity requires a higher concentration of iron and manganese for electron transport in photosynthesis (Gordon & Cook, 2016). Most algae absorb nutrients from the water over their entire cell surface through physical diffusion, membrane transport or chemical dissociation (Morales-Sánchez et al., 2015). The environment may therefore impact their ability to absorb nutrients with external factors able to further influence microalgae growth.

Light As photosynthetic organisms, green microalgae would attain a higher cell division rate, thus greater biomass would be produced with increasing light intensity. Morphological studies have also shown that the increase in light may lead to larger cells (Latala, 1991).

Temperature One of the most significant environmental conditions for microalgal growth is temperature. For most Chlorophytes, the optimal growth temperature is between 20-30°C (Singhania et al., 2017), but laboratory cultures may be grown from between 5-38°C (Andersen, 2005). Temperature has a defining effect on cell morphology with species geographically located in warmer conditions being correlated with a smaller cell size (Latala, 1991). The higher temperature allows for a greater metabolic rate which decreases the cell volume requirements (Latala, 1991).

Salinity The salt concentrations in water may affect the microalgae osmotic potential and may also influence gas solubility, especially that of CO2, water density and viscosity (Andersen, 2005). The water salinity is a determining factor for the metabolism rate and thus the photosynthesis and respiration intensities of microalgae. If the salt concentration is too low for the particular species, such as marine microalgae, then the metabolism will decrease and the dwarfism phenomenon may be observed (Latala, 1991). However if the salt concentrations exceeds a species’ requirements then cell division and daughter cell formation may be inhibited (Latala, 1991). The optimum salinity for a given species is usually the same as in its natural habitat and should therefore be cultured under

Chapter 1 such conditions (Andersen, 2005). Some microalgae, such as the Chlorella species, show great adaptability and may be cultured in both fresh, and saltwater conditions (Latala, 1991).

Halophilic cultures such as the Dunaliella species, have many technical difficulties associated with transformation. High salt concentrations significantly reduce the efficiency of many antibiotics and herbicides, thereby limiting the range of selectable markers for transformation (Walker et al., 2005). Transformation efficiency may also be hindered as DNA degradation is apparent during introduction into the cell (Walker et al., 2005).

Therefore, the inputs for microalgae growth are simple namely, CO2, sunlight, water, and some nutrients. Their ability to photosynthesize solar energy into chemical energy, sugar, and oxygen makes them primary producers -the fundamental role player of any ecosystem.

1.2.2 Role of algae in global society Chlorophytes are considered ancestors to land plants and therefore they have had very important role in many ecosystems. They fix 40% of the world’s carbon and deliver 70% percent of the earth’s atmospheric oxygen (Gordon & Cook, 2016). Microalgae are the foundation for the aquatic food chains, with microalgae nourishing fish and crustaceans, which in turn feed larger species and continue up the food chain to larger predators and even humans (Barsanti & Gualtieri, 2014). Historically algae have been a part of the human diet across the globe; from 14 000 before the common era (BC) archaeological evidence indicates that algae have been eaten by the natives in Chile, with written evidence from anno Domini (AD) 300 supporting their consumption in China and from AD 600 in Ireland (Wells et al., 2017).

Today the consumption of microalgae has commercialised Chlorella species for human and animal nutrition due to its high protein content, as a source of nutrients and high-value compounds such as pigments, fatty acids and anti-oxidants (Vigani et al., 2015). The uses for microalgae are becoming more widespread, to include pharmaceuticals, nutraceuticals, fertilizer, bioplastics and even environmental biosensors (Ho et al., 2016). Several microalgal species naturally accumulate high levels of oil in their dry mass which can be converted to biodiesel, gasoline, ethanol, methane, and other biofuels (Gordon & Cook, 2016).

The microalgae fundamental characteristics as phototroph, unicellular cells has opened a new application for the bio-manufacturing of microalgae as a sustainable manufacturing platform to produce a range of higher-value products. With their ease of transformation, higher value products can include recombinant proteins of therapeutic value, glycoproteins, antibodies, and many other industrially valuable proteins (Wannathong et al., 2016).

1.3 Recombinant Protein Production Protein production is the synthesis, modification and regulation of proteins from within living organisms. It is the central dogma of molecular biology which explains how the coded genetic elements from DNA is transcribed into messenger RNA, then translated or used to synthesize a protein which can undergo post-translational modifications. With recombinant organisms, a DNA transgene is inserted into the host organism and it is the host organism’s transcription, translation and post-translational machinery that is then utilised to produce the recombinant protein. Therefore in vivo expression systems include living cells and their cellular machinery which use supplied genetic templates to build and construct proteins (Andersen & Krummen, 2002).

Chapter 1

The commercial requirements for proteins began as early as 1796, when the cow-pox vaccine was produced by Edward Jenner (Cook, 1996). Later in 1982, the very first recombinant protein product available to consumers was recombinant insulin (Johnson, 1983), and the production of recombinant proteins has grown ever since. Over the last 60 years, biomanufacturing has developed significantly due to advances in genetic engineering and biotechnology (Rasala & Mayfield, 2015). Many of these proteins are therapeutic, and include complex glycoproteins, antibodies, subunit vaccines, immunotoxins, metabolites, hormones, industrial enzymes and other small molecules. The production of such proteins are greatly desired since they have many current and future applications in biopharmaceuticals, biomaterials, nutraceuticals, agriculture, animal health and cosmeceutical industries (Rasala & Mayfield, 2015). The global protein therapeutic market alone is expected to grow at a compounded annual growth rate of around 8.6% with expectations to reach $315.90 billion by 2025 (Wood, 2016 ).

1.3.1 Current expression systems for recombinant proteins Currently recombinant proteins can be expressed in transgenic cell cultures of bacteria, yeasts, fungi, mammals, plants, insects, or via transgenic animals (Demain & Vaishnav, 2009). None of these expression systems are alike and each must be assessed based on their operation costs, ease of use, protein quality, functionality, production speed, yield and their post-translational modification profiles.

Mammalian and insect expression systems The main advantage of mammalian expression systems is that they can produce recombinant proteins, of mammalian origin, with complete biological activity. Their eukaryotic system allows for proper protein folding, post-translational modifications including acetylation (Decottignies et al., 1995), methylation (Doerfler, 1981), S-thiolation, glycosylation (Mathieu-Rivet et al., 2014) and product assembly (Khan, 2013). The most widely used host mammalian cells for heterologous protein expression are the Chinese hamster ovary cells and mouse myeloma cells. These cultures contain live cells, and are therefore at risk for contamination with animal viruses (Khan, 2013). Another disadvantage of mammalian expression system is that the culturing equipment, complex technology, and cells themselves are extremely expensive (Khan, 2013). Therefore, products derived from this platform result in higher costs unfeasible for poorer and developing countries.

Insect cell cultures would also be able to post-translate proteins through eukaryotic machinery. Unlike mammalian cells, no large and complex glycans are attached to the expressed proteins and therefore crystallization should not be inhibited (He et al., 2014). This makes insect cultures the expression of choice for membrane proteins; however, the use of insect cells is also constrained by the expensive operational and technical costs (van Oers et al., 2015).

Bacterial expression systems As a more cost-effective strategy, inexpensive microbial expression systems have been utilised. Their quick replication, inexpensive cost, time-saving procedure, mature and easy genetic manipulations makes them sought after operationally (Sanchez-Garcia et al., 2016). However, as prokaryotes they may lack the molecular chaperones, and post-translational modifications that are required for the correct folding and functioning of eukaryotic proteins. To facilitate protein folding, the expression of post-translational machineries has been incorporated with the heterologous protein, however the

Chapter 1 eukaryotic proteins may still lack proper biological function (He et al., 2014). One of the most widely used hosts for heterologous protein production is the bacterium, Escherichia coli.

Toxicity in E. coli cell cultures due to endotoxin production is a problem resulting in the inclusion of complex downstream processing steps before product application (Demain & Vaishnav, 2009). The bacteria, Bacillus spp. is often preferred since it does not produce harmful exo- or endotoxins. Microbial cell cultures are economically feasible, but limited to producing small and structurally simple proteins (Demain & Vaishnav, 2009, Sanchez-Garcia et al., 2016).

Yeast expression systems A eukaryotic expression system that can be cultured in the same manner as prokaryotic microbial cells, is yeast. Yeast is easy to manipulate genetically, it can produce high yields of heterologous protein, may be cost-effective and the eukaryotic machinery allows for post-translational modifications (Kim et al., 2015). Although yeast can handle disulphide rich protein expression, even the most cultured yeast, Saccharomyces cerevisiae tends to hyper-glycosylate heterologous proteins, which may produce an immunogenic response in humans (Demain & Vaishnav, 2009). Other drawbacks of such cultures is their susceptibility to contamination and the hard cell walls which make it difficult to disrupt and purify the recombinant protein (He et al., 2014).

Plant expression systems Another eukaryotic expression system includes both transgenic whole plants and transgenic plant cell cultures. The use of whole plants for the synthesis of recombinant proteins can be a more economical and scalable option than microbial and mammalian production systems (He et al., 2014). Plants are also not susceptible to animal or human viruses, making them a safer option. Other key advantages include rapid production and the ability to produce unique protein glycol forms (Sack et al., 2015). One limitation within plants is that proteolysis impairs the quality of the recombinant proteins produced and although inhibition of specific proteases is currently being developed, their high cost and impact on protein quality make it unfeasible (Sack et al., 2015). Their biggest limitation is perhaps that plants require more time and space to grow than alternate expression systems (Mayfield & Franklin, 2005).

The choice of an appropriate expression system would depend on the practicality of implementation, on the equipment available and on the biochemical and biological properties of the recombinant proteins (Demain & Vaishnav, 2009). It is clear however, that the current expression systems available to produce recombinant proteins remain limited, since they lack either in their ability to produce viable proteins, sufficient yield and viability, cost of processing, risk of contamination or their ease of transgenic manipulations. A very promising alternative protein expression system, which can overcome many of these limitations, is that of green microalgae (Mayfield & Franklin, 2005).

Microalgal expression systems Microalgae offer advantages from both prokaryotic and eukaryotic expression systems. As unicellular organisms, culturing microalgae is similar to that of prokaryotes - inexpensive with easy scalability (Andersen, 2005). Within a few weeks, a single cell can expand to cell biomass of thousands of litres, and within a matter of months this may increase to hectares of microalgae pond production (Rasala & Mayfield, 2015). The unicellular nature of most microalgae circumvents environmental distresses and their haploid vegetative state would allow for easier transformation

Chapter 1 where recessive mutants immediately show a phenotype (Jinkerson & Jonikas, 2015). The unicellular, asexual and sexual reproduction of microalgae enables easier genetic modification through transformation, and simple breeding implementations (Rasala & Mayfield, 2015).

Most microalgae are Generally Regarded As Safe (GRAS) and do not sequester any animal or human viruses (Taunt et al., 2018). Furthermore, microalgae contain eukaryotic machinery and would be able to express, post-translationally modify, and secrete intricate proteins (Taunt et al., 2018). Microalgae can properly subject proteins to glycosylation, acetylation, S-thiolation, methylation and the production of mammalian like N-glycans for biologically functional heterologous proteins (Jinkerson & Jonikas, 2015). Their production systems are flexible to both prokaryotic and eukaryotic proteins as the chloroplast contains prokaryotic transcriptional and translational machinery (Taunt et al., 2018). Therefore, based on the numerous advantages microalgae holds great potential as a protein expression system and should be explored further.

1.3.2 Expression of recombinant proteins in green microalgae The advantage of using microalgae as an expression host has already allowed several proteins of economic value to be produced (Table 1.3). Recombinant proteins include nutritional supplements, industrial enzymes, enhanced animal feeds, antibodies, immunotoxins, subunit vaccines as well as novel anti-cancer therapeutics (Walsh, 2018, Rasala & Mayfield, 2015). Even the yield of these heterologous proteins can be high; heterologous protein levels may surpass 0.25% of the total soluble protein with expression up to 21% with chloroplast transformation (Rasala & Mayfield, 2015).

Table 1.2: Current successes in microalgal recombinant protein production

Recombinant Application Genome Expression Remark Reference protein engineered microalgae Human Treatment to Chloroplast C. reinhardtii TSP accumulation (Wang et al., glutamate prevent 0.3%, fully 2008) decarboxylase diabetes 1 functional 2 Antihypertens Blood Chloroplast C. reinhardtii Decreased blood (Carrizalez- ive pressure pressure in López et al., polypeptide regulation hypertensive rats 2018) Human Treatment of Nuclear C. reinhardtii / First study (Eichler- Erythropoietin anaemia Dictyosphaeriu accumulated to Stahlberg et protein m pulchellum 100 μg/L; second al., 2009); study to 500 μg/L (Bashir et al., 2018) Human Treatment Chloroplast C. reinhardtii / Fully functional (Franklin & immunoglobul against herpes P. tricornutum Mayfield, in A/ B single- simplex virus 2005); chain (Vanier et al. antibody 2015) Monoclonal Diagnostics Nuclear Phaeodactylu Antibodies (Hempel et immunoglobul and therapy m tricornutum excreted into salt al., 2017) in against water medium IgG antibodies Marburg virus were relatively pure

Chapter 1

Sexual stage Antibodies for Chloroplast C. reinhardtii Parasites were (Gregory et antigen of a vaccine prevented from al., 2012); Plasmodium inhibiting developing within (Patra et falciparum malaria the mosquito al.,2015) transmission vector E7 protein of Antibody for Chloroplast C. reinhardtii TSP accumulation (Demurtas et human human 0.12%, induces al., 2013) papillomaviru papillomavirus specific tumour s type 16 -16 cancer protection vaccine Synthetic Nutritional Chloroplast C. reinhardtii Success when (Tevatia et taurine animal feed taurine precursors al., 2019) were transformed as separate genes as opposed to a fusion protein Human Nutritional Nuclear C. reinhardtii Human (Hou et al., selenoprotein selenium selenoprotein 2013) 15 supplements accumulated to detectible levels Flounder Increased Nuclear Chlorella Up to 400 μg/L (Kim et al., growth growth rate of ellipsoidea accumulated in the 2002) hormone aquaculture algal culture, and when fed to flounder they exhibited a 25 % greater increase in body size after 30 days β-1,4- Paper and Nuclear C. reinhardtii Increased (Rasala et al., endoxylanase food bleaching recombinant 2012) protein yield by fusing transgene to she ble antibiotic resistant marker Xylanase, α- Feed additive Chloroplast Dunaliella First (Georgianna galactosidase, salina transformation of et al., 2013) phytase Dunaliella chloroplast Rabbit Anti-microbial Nuclear C. ellipsoidea Fully functional (Chen et al., neutrophil activity 2001) peptide-1

1.4 Algae in Biotechnology Despite the clear commercial demand, the methodologies for recombinant algal biotechnology are only partially developed (Díaz‐Santos et al., 2013). Complete genomes have only been published for a few unicellular green algal organisms. The world’s smallest free-living eukaryotic organism, Ostreococcus tauri, has been characterised as a model organism for eukaryotic genome evolution (Derelle et al., 2006). As a model for viral/algal interactions and adaption to photo symbiosis the

Chapter 1 algae, Chlorella variabilis NC64A was completely sequenced (Blanc et al., 2010) with several genetic transformations systems established in green algae, C. reinhardtii and the multicellular V. carteri (Walker et al., 2005).

The most studied algal organism is C. reinhardtii, a unicellular green microalgae which has been used for the past 60 years to research fundamental biological processes including photosynthesis, flagella assembly, metabolic pathways, sexual reproduction and the circadian rhythm (López‐Paz et al., 2017). Functional genome annotation furthered our understanding of the biological, physiological, developmental, cellular, and molecular processes when the C. reinhardtii haploid genome was sequenced and made available in 2007. Research on this organism is continually updating our understanding of algae (Harris, 2009, Merchant et al., 2007).

The complete C. reinhardtii genome project sequenced the nuclear, chloroplast and mitochondrial genomes of this organism (Shrager et al. 2003). In total, C. reinhardti is predicted to contain approximately 15,000 genes of which 741 are found in the nuclear genome, 99 genes in the 203 kb chloroplast genome (Merchant et al., 2007, Jinkerson & Jonikas, 2015) and eight genes in the 16 kb mitochondrial genome (Walker et al., 2005). To date, all three of its genomes have been successfully transformed (Rasala et al., 2010).

Classified as the “green yeast”, C. reinhardtii is the most characterised microalgal expression system (Shrager et al. 2003). Ironically, the first C. reinhardtii transformation was with yeast DNA (Rochaix & Van Dillewijn, 1982). Several years later the first stable transformation of the C. reinhardtii nuclear genome restored the nitrate reductase function in a mutant strain (Kindle et al., 1989). The transformation technique of particle bombardment was utilised and since then methods to transform microalgae have been improved.

1.4.1 Algae transformation techniques Many different transformation methods have been successfully employed for C. reinhardtii namely: particle bombardment (Stevens & Purton, 1997, Kindle et al., 1989), electroporation (Shimogawara et al., 1998), the glass-bead method (Purton, 2007, Kindle et al., 1991), lithium acetate/polyethylene glycol (PEG)-mediated method (Feng et al., 2014), and Agrobacterium tumefaciens-mediated transformation (Úbeda-Mínguez et al., 2015). Each technique has its own advantages and disadvantages with respect to the transformation rate, controllability, repeatability and requirements of laboratory apparatus (Feng et al., 2014). Additional techniques for the transformation of algae also include carbide whiskers (Dunahay, 1993) and nanoparticles (Wang et al., 2016), but these are very technical and not reputable yet (Jinkerson & Jonikas, 2015).

For nuclear transformations with the model C. reinhardtii, the first successful transformation method was a biolistic particle delivery system developed in the late 1980’s (Kindle et al., 1989). With this method metal microparticles, such as gold or tungsten, are coated with DNA which gets projected at a high velocity, into the algal cells. However this method requires specialised, expensive equipment, is complex, damages most cells, and produces very few transformants (Jinkerson & Jonikas, 2015). One of the first successful transformations included the nit1 gene (encodes nitrate reductase) into C. reinhardtii (Sodeinde & Kindle, 1993).

Chapter 1

The glass bead method is a more cost-effective approach which requires no specialised equipment (Kindle et al., 1991). The algal cells are simply vortexed with the exogenous DNA in a tube containing glass beads (Lumbreras & Purton, 1998). This glass bead method was also used to successfully transform the nit1 gene into C. reinhardtii, but with fewer transformants obtained (Sodeinde & Kindle, 1993). However, transformation requires that there is no cell wall, as a cell wall can inhibit the foreign DNA from entering the cells. The removal of the cell wall is required through enzymatic treatment, or through the transformation of cell wall deficient strains (Purton, 2007). The cell wall degrading enzyme can interfere with transformation (D’Souza et al., 2018) and cell wall deficient strains would not be available for novel microalgae isolates.

A more efficient method than the glass bead method is electroporation, which can yield up to 100 times more transformants with the same quantity of exogenous DNA (Coll, 2006, Jinkerson & Jonikas, 2015). Electroporation introduces DNA into the algal cells through the application of an electric pulse (Biorad, 1988). Currently three of the six microalgae phyla that have been electrotransformed include: the Chlorophyta lineage with C. reinhardtii, C. ellipsoidea, Chlorella vulgaris, D. salina and Scenedesmus obliquus; the Heterokontophyta with Nannochloropsis sp. and Phaeodactylum tricornutum as well as the Rhodophyta phylum with Cyanidioschyzon merolae (Kotnik et al., 2015). Electroporation has successfully transformed the hepatitis B surface antigen (HBsAg) gene into the D. salina (Geng et al., 2003).

Recently, the plant pathogen A. tumefaciens has been applied as a method for gene delivery and has been implemented as a transformation method for a few microalgal species namely: Porphyra sp., Haematococcus pluvialis, Dunaliella bardawil, C. vulgaris, Schyzochitrium sp., Tetraselmis chuii and C. reinhardtii (Úbeda-Mínguez et al., 2015). Agrobacterium-meditated transformation offers several advantages over other methods including a high transformation efficiency, the ability to introduce larger DNA segments, less likelihood of rearrangement and higher likelihood of transgene integration into transcriptionally active regions; this leads to a low copy number which may reduce the silencing tendency (Úbeda-Mínguez et al., 2015).

The first successful Agrobacterium-meditated transfer of T-DNA carrying the genes coding for Green Fluorescent Protein (GFP) to the nuclear genome of C. reinhardtii was observed in 2004 (Kumar et al., 2004). There was a 50-fold higher transformation frequency than that of glass bead transformations. However after 14 years of optimisation, Agrobacterium-mediated transformation efficiencies were still 2.5 to 60-fold lower with respect to electroporation (Mini et al., 2018). This method was re-evaluated in 2018 in C. reinhardtii, and Mini and co-workers concluded that there was no real benefit to Agrobacterium transformation over electroporation, and that it was merely a more labour-intensive procedure (Mini et al., 2018).

The expression of heterologous proteins can be achieved by introducing the target gene into the nuclear genome, into the chloroplast genome or into the nuclear genome with a targeting signal that directs the transgene RNA into the chloroplast for expression (Doron et al., 2016). Comparable technologies can be used for both chloroplast and nuclear transformations of microalgae species with both having their own sets of advantages depending on the heterologous protein (Doron et al., 2016).

Chapter 1

Nuclear transformation The advantages of nuclear transformation for a microalgal expression system are that proteins can be correctly expressed, folded as they would be within their eukaryotic origin, and signal peptides can be attached onto the transgenes to ensure the heterologous protein is targeted to the endoplasmic reticulum and secreted into the extracellular media (Rasala et al., 2014). This type of automatic secretion cannot be achieved in chloroplast expression and the heterologous proteins would remain in the plastid, requiring added downstream purification processing steps such as cell lysis (Wittkopp, 2018).

Unfortunately, transgene silencing can occur at both the transcriptional and post-transcriptional level within the nuclear genome of C. reinhardtii. Silencing results if the transgene is methylated by DNA or histone methylation, and is induced if the construct was inserted multiple times within the nuclear genome (Jinkerson & Jonikas, 2015). Epigenetic mechanisms which could circumvent silencing and improve transgene expression include: codon optimisation, inserting native algal introns in the transcript, coupling the transgene with resistance to a selecting agent and using smaller constructs with a lower likelihood to be fragmented by endonucleases (Jinkerson & Jonikas, 2015).

Chloroplast transformation The chloroplast genome of higher plants and algae consists of a double stranded circular genome that can range from 50- 295 kb in size of which, 20- 30 kb are usually inverted repeat sequences. The protein synthesis machinery is prokaryotic in nature and therefore lacks many eukaryotic post- translational modifications (Jinkerson & Jonikas, 2015). Unlike the single nucleus, there can be up to 100 chloroplasts within a mesophyll cell with 100 genome copies per chloroplast. Therefore a transgene inserted within the chloroplast could have 10 000 genome copies being expressed within a singular cell (Doron et al., 2016). However, this would be subject to the microalgae species, as popular species such as Chlamydomonas spp. or Chlorella spp. only have one chloroplast per cell.

Chloroplast transformation offers several advantages for protein expression when compared to nuclear transformation and heterologous protein yields can be 20 times greater through chloroplast transformation (Jinkerson & Jonikas, 2015). Due to multigene operons, genes encoding several enzymes from a complex pathway could be introduced in a single transformation event and be simultaneously expressed (Doron et al., 2016). Targeting the foreign DNA for specific integration into the genome is possible and positional effects and gene silencing would be reduced (Walker et al., 2005). The formation of disulphide bonds, required for quaternary structure and correct protein folding, occurs readily in the benign chloroplast without the occurrence of undesirable glycosylations (Wannathong et al., 2016). Since introduction of foreign genes is not as random as with nuclear transformations, less variability exists between the different transgenic clones and less extensive screening would be required (Doron et al., 2016). Comparison of nuclear and chloroplast transformation in algae is summarised in Table 1.4.

Chapter 1

Table 1.3: Comparison of transgene expression in the nucleus and chloroplast of microalgae

Nucleus Chloroplast Reference Protein synthesis machinery is Protein synthesis machinery is (Day & eukaryotic in nature with inducible prokaryotic gene expression with Goldschmidt‐ gene expression eukaryotic chaperones Clermont, 2011) Eukaryotic post-translational Can properly fold and assemble (Wittkopp, modifications include glycolysation, disulphide-bonded proteins, 2018) acetylation, S-thiolation, phosphorylation methylation and even the but cannot perform glycosylation of production of mammalian like N- proteins glycans Detectable levels of protein High levels of total soluble proteins (Wannathong et expression al., 2016) Random integration can lead to Specific integration minimises positional (Kindle et al., positional effects, gene silencing or effects and gene silencing 1991) low transgene expression Transgene size limit Can accept large DNA inserts with (Day & multiple genes and repetitive elements Goldschmidt‐ Clermont, 2011) Suited for recombinant proteins that Suited for immunotoxins, antibodies (Doerfler, 1981) require targeting to specific cellular and oral vaccines locations or extracellular secretion

Chloroplast transformation is more challenging than nuclear transformation since it requires more time, extensive optimisation and elaborate resources (Jinkerson & Jonikas, 2015). To express a transgene of interest in the nuclear genome, regulatory genes are required.

1.4.2 Expression of heterologous protein products Transformed into the host organism, an expression vector includes the gene cassette that contains a transgene as well as necessary elements to ensure the expression and regulation of an inserted gene. The molecular toolkit for manipulating gene expression in C. reinhardtii is well established and refined with the development of selectable markers, reporter genes and regulatory genes which can be extended to other microalgae (Walker et al. 2005).

Selectable Markers After transformation, the positive microalgae colonies must be distinguished from the untransformed ones; selectable markers aid this screening process and help maintain the transformed state (Sizova et al., 2001). Transformation markers that are often used in microalgae nuclear transformation include antibiotic, herbicide and auxotrophic markers (Table 1.5).

An antibiotic resistance marker is a gene which produces a protein that allows the transformed cell to easily tolerate an antibiotic. After transformation, the cells are cultured with this antibiotic at a concentration that kills wild type cells and therefore only the transformed cells are selected for (Chankova et al., 2007). This is considered the most frequently used method with the greatest success for selecting stable microalgae transformants (Garcia-Echauri & Cardineau, 2015). The same principle applies for herbicide resistant genes, and the type of herbicide resistance genes expressed in microalgae are similar to those expressed in higher plants (Jiang et al., 2014).

Chapter 1

To avoid the use of antibiotics or herbicides, under the principle of photosynthetic competency, light may be used as a selecting agent. Microalgal cells have been mutated so that their photosynthetic machinery is disrupted, only the microalgae that are transformed would contain a gene to restore their ability to photosynthesize and grow under light (Day & Goldschmidt‐Clermont, 2011). This selection method targets the chloroplast and is not relevant for nuclear transformation. However, the same principle may be followed with metabolic competency, where the microalgae is mutated to be dysfunctional in a primary growth gene, leaving only the transformed microalgae to grow under the nutrient conditions supplied (Lumbreras & Purton, 1998). For example, the mutant strain, C. reinhardtii arg9-2 lacks a functional N-acetyl ornithine aminotransferase and requires arginine supplemented in the media to grow. The only way to grow without the addition of arginine is to rectify the ability to synthesize a functional enzyme, by transforming the arg9 gene into the mutant strain (Remacle et al., 2009). The application of these markers are limited to the respective mutant strains and the external metabolites required might complicate selection (Jinkerson & Jonikas, 2015).

Table 1.4: Selectable markers used for screening for positive microalgae transformants.

Gene Protein expressed Selection agent Expression Reference microalgae Antibiotic selectable markers Sh ble Phleomycin/ bleomycin Zeocin Nannochloropsis (Stevens et al., resistance protein salina, 1996) C. reinhardtii Cry1 Ribosomal protein S14 Emetine C. reinhardtii (Nelson et al., 1994) aphVIII Aminoglycoside Paromomycin C. reinhardtii (Sizova et al., phosphotransferase VIII 2001) aph70 Aminoglycoside Hygromycin Neochloris (Berthold et phosphotransferase oleoabundans al., 2002) aadA Adenlytransferase Spectinomycin C. reinhardtii (Cerutti et al., 1997) tetx NADP-requiring Tetracycline C. reinhardtii (Garcia- oxidoreductase hydroxylates Echauri & enzyme Cardineau, 2015) nptII Aminoglycoside G418 sulphate C. vulgaris (Yang et al., 3'phosphotransferase 2015) enzyme

Herbicide selectable marker bar Phosphinothricin N- Basta Symbiodinium (Ortiz- acetyltransferase spp. Matamoros et al., 2015) gat Glyphosate Glyphosate C. reinhardtii (Jiang et al., acetyltransferase 2014)

Auxotrophic selectable marker arg7 Argininosuccinate lyase Arginine media C. reinhardtii (Debuchy et al., 1989)

Chapter 1

nit1 Nitrate reductase Ammonium C. reinhardtii (Kindle et al., media + nitrate 1989) media nic7 Quinolinate synthetase Nicotinamide C. reinhardtii (Ferris, 1995) media thi10 Hydroxyethylthiazole kinase Thiamine media C. reinhardtii (Ferris, 1995)

Promotors and terminators Regulatory elements, promotors and terminators are required to drive the expression of both the transgene and selectable marker within the microalgae genome (Table 1.6). In most cases the transgene and selectable marker require a promotor and terminator gene cassette of their own, but multi-gene expression systems are possible in microalgae (Kilian et al., 2012, Noor-Mohammadi et al., 2014).

Research has shown that exogenous promoters, such as the Cauliflower Mosaic Virus 35S (CaMV 35S) (Doron et al., 2016) and the Agrobacterium nopaline synthase (nos) (Díaz‐Santos et al., 2013), promoters, often used in higher plant transformations, can drive transgene expression within algae. However much greater transgene expression is obtained when endogenous algal promoters are used (Feng et al., 2014). Native promoters are thought to contain enhancer elements, decreasing the chance of transgene silencing and contributing to high transcription levels (Jinkerson & Jonikas, 2015). The most successful constitutive promoters for microalgae are the photosystem I protein D (psaD) (Fischer & Rochaix, 2001) and the ribulose bisphosphate carboxylase (rbcS2) (Ferrante et al., 2008, Rasala et al., 2013).

In addition to promoters, the type of terminator within the gene cassette may also enhance transgene expression within microalgae. Just as endogenous promoters yield greater protein expression, native terminators were found to yield greater translation efficiencies by regulating mRNA stability and polyadenylation signals (Díaz‐Santos et al., 2013). The most efficient microalgal terminator has been the 3’ UTR of the rbcS2 gene (Schroda et al., 2000).

Heterologous proteins are normally expressed under constitutive promoters for a high protein yield for algae. However, for any potentially toxic gene products, expression should only occur after reaching a specific cell density and inducible promoters are required instead (Rasala & Mayfield, 2015). With C. reinhardtii inducible promoters include the nit1 promoter, induced by ammonium starvation, the ca1 promoter, induced when the CO2 concentrations drop and the cyc6 promoter which is induced when copper is depleted (Schroda et al., 2000).

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Table 1.5: Summary of the promoters and terminators used for transgene microalgae over the past decade

Promoter Terminator Expression microalgae Expression levels Reference rbcS2 rbcS2 C. reinhardtii Highest expression (Díaz‐Santos levels obtained for C. et al., 2013, reinhardtii Scranton et al., 2016) PsaD PsaD C. reinhardtii Efficient promoter (Ferrante et al., 2008) CaMV 35S nos C. reinhardtii Low expression levels (Díaz‐Santos et al., 2013) nos nos C. reinhardtii Expression levels are (Díaz‐Santos low, but 5-fold et al., 2013) increase compared to CaMV 35S rbcs2 CaMV 35S D. salina Efficient expression (Simon et al., 2016) 33kDa 3’ UTR C. ellipsoidea Expression level equal (Park & Park, peptide gene to CaMV 35S 2004) of a Chlorella sp. viral promoter vcp2 vcp2 3′ UTR Nannochloropsis spp. Bidirectional and able (Kilian et al., to express two 2012) proteins at once

Algae offer the therapeutic protein production industry many advantages over existing expression systems. While C. reinhardtii is an excellent model, faster-growing and more robust native species of green algae are better suited to intensive commercial cultivation (Taunt et al., 2018). Microalgae biodiversity is enormous. It has been predicted that approximately 200,000-800,000 species in many different genera exist of which only 50,000 species are defined and fewer than 20 have been genetically transformed (Úbeda-Mínguez et al., 2015). Most of these are decent laboratory models but are not appropriate for commercial cultivation. Many uncharacterised microalgae exist in South African freshwaters with potential for species to surpass the industrial microalgae biotechnology requirements, cultivation and recombinant protein production of C. reinhardtii.

The development of genetic transformation methods for newly identified industrially promising microalgal species is thus important. The aim of this study was to explore the feasibility of microalgae biotechnology within a South African context. The practicality of transgene expression was tested within the C. reinhardtii nuclear genome, and South African microalgae isolates that had biotechnology potential were identified with the ambition of developing future microalgal heterologous protein production systems.

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1.5 Aims and Objectives The study aimed to develop a heterologous protein expression platform in C. reinhardtii under the current laboratory conditions, as well as analyse how the growth of C. reinhardtii may compare to indigenous South African green microalgae. To achieve this, the following objectives were undertaken:

• To obtain and culture model C. reinhardtii strains. • To clone a C. reinhardtii (Cr) codon optimised Crgfp gene into the pChlamy4 vector. • To transform the recombinant Crgfp-pChlamy4 vector into the nuclear region of the model green microalgae C. reinhardtii through electroporation. • To analyse the DNA integration of the Crgfp transgene by PCR and to quantify the CrGFP protein production by fluorescent spectrophotometry. • To screen cultured indigenous freshwater South African microalgal species based on their growth rate potential. • To identify the green microalgal isolates chosen to genus level by compound microscopy and Sanger sequencing of the conserved 18S rRNA region. • To test all microalgal isolates for tolerance to the selection antibiotic, zeocin. • To optimise South African microalgae culturing procedures to current South African laboratory conditions. • To cryopreserve the biotechnologically important microalgae identified in this study. • To engage with the public to highlight the importance of microalgae for the socio-economy, bio-economy, health and sustainability in South Africa.

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Wang X, Brandsma M, Tremblay R, Maxwell D, Jevnikar AM, Huner N, Ma S, 2008. A novel expression platform for the production of diabetes-associated autoantigen human glutamic acid decarboxylase (hGAD65). BMC Biotechnology 8, 87. Wannathong T, Waterhouse JC, Young RE, Economou CK, Purton S, 2016. New tools for chloroplast genetic engineering allow the synthesis of human growth hormone in the green alga Chlamydomonas reinhardtii. Applied Microbiology and Biotechnology 100, 5467-77. Wells ML, Potin P, Craigie JS, Raven JA, Merchant SS, Helliwell KE, Smith AG, Camire ME, Brawley SH, 2017. Algae as nutritional and functional food sources: revisiting our understanding. Journal of Applied Phycology 29, 949-82. Wittkopp TM, 2018. Nuclear Transformation of Chlamydomonas reinhardtii by electroporation. The Salk Institute for Biological Studies. https://bio-protocol.org/pdf/bio-protocol2837.pdf. Wood L, 2016. Protein therapeutics market analysis and trends - therapeutic proteins, application, function - forecast to 2025. Research and Markets. Wurm, F.M., 2004. Production of recombinant protein therapeutics in cultivated mammalian cells. Nature Biotechnology 22, 1393. Yang B, Liu J, Liu B, Sun P, Ma X, Jiang Y, Wei D, Chen F, 2015. Development of a stable genetic system for Chlorella vulgaris—A promising green alga for CO2 biomitigation. Algal Research 12, 134-41. Zhang ESB, 2016. Enhanced microalgae growth and lipid production: a study of cytostatic inhibitors and glycerol assimilation. Thesis. Institute of Molecular Bioscience, The University of Queensland. Zhi–Juan Z, Huan Z, Guo–Xiang L, Zheng–Yu H, 2016. Rhizoclonium ramosum sp. nov.(Cladophorales, Chlorophyta), a new fresh-water algal species from China. Fottea Olomouc 16, 12–21.

Chapter 2 Nuclear transformation of Chlamydomonas reinhardtii

Chapter 2

Abstract The green microalgae species, Chlamydomonas reinhardtii has been extensively used as the model for microalgae recombinant protein production. It was therefore the prime target to investigate the feasibility of establishing microalgal biotechnology under the available laboratory environment at the CSIR. In this study, a gene encoding the Chlamydomonas codon optimised green fluorescent protein (CrGFP) was cloned into a pChlamy4 vector under the control of the chimeric Hsp70A-rbcS2 promoter and selected for with a bleomycin resistant gene, she ble. The verified plasmid was then electro-transformed into the nuclear region of two C. reinhardtii strains: CC-125, a wild type which contains a cell wall; and CC-400, a cell wall deficient mutant. For successful transformation the transformation procedure required optimisation with regards to the cell concentrations, the voltage, the resistance, the DNA concentrations and the temperature utilised. The microalgae transformants were assessed at the DNA level by PCR and further at the CrGFP protein production level by fluorescence spectrophotometry. Results indicated stable insertion of the Crgfp transgene with detectable levels of CrGFP fluorescence above autofluorescence for all transformed C. reinhardtii colonies. The CC-400 strain indicated higher expression than the CC-125 strain and different levels of expression were noted for each transformation event.

Keywords: microalgae, Chlamydomonas, nuclear transformation, endogenous algal promoters, selection marker, green fluorescent protein

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2.1 Introduction Microalgae are currently considered ideal recombinant protein expression systems due to their rapid growth rates, lower production costs and easy, safe culturing (Ramos‐Martinez et al., 2017). The microalga C. reinhardtii was subject to nuclear transformation with the pChlamy4 vector, which contains the most ideal genetic elements for transgene expression. The green algae C. reinhardtii is the model organism for microalgae transformation and recombinant protein expression research (Fischer et al. 2006; Ledford et al., 2007; Peers et al., 2009).

Chlamydomonas spp. may reproduce either asexually, under favourable growth conditions; or sexually, under environmental stress or a combination of both (Gastineau et al., 2014). This infers that the microalgae may undergo haploid (single set of chromosomes) and diploid (double set of chromosomes) stages with their genetic mating type material categorised as either the plus mating type (mt+) or the minus mating type (mt-) (Harris, 2009). It is due to its sexual reproduction and use as a laboratory model that many different strains and mutants of C. reinhardtii have come to existence (Pröschold et al., 2005, Procházková et al., 2014, Flowers et al., 2015). The initial laboratory strain of C. reinhardtii was isolated by Smith in 1945 and through subsequent cultivation the cell wall mutant, CC-400 was created by Davies in 1971. In 1968 Gillham cultivated a C. reinhardtii strain that lost its nitrate reductase function, and through successive cultivations the CC-125 strain emerged from the Chlamydomonas Resource Centre in 1980 (Pröschold et al., 2005).

The production of complicated eukaryotic proteins with specific post translational modifications is considered advantageous when directed to the nucleus of C. reinhardtii (Jinkerson & Jonikas, 2015). Nuclear transformation also allows production of therapeutic proteins with signal peptides for targeted secretion directly into the growth media (Rasala et al., 2014). Unfortunately however, nuclear transgene expression within Chlamydomonas is limited by its low heterologous protein yields (Lauersen et al., 2015). Low expression levels are often seen in the nuclear region of microalgae due to random integration of transgenes (Scranton et al., 2016). In a study that analysed transgene insertion sites in the C. reinhardtii genome, no predisposition for particular insertion sites was identified (Jinkerson & Jonikas, 2015). The only gene bias that was recognized was that transgenes integrated less into exons than into non-essential genes, since disrupting essential genes would most likely yield non-viable mutants. Therefore the position in the genome where the gene integrates may influence its expression levels (Rasala et al., 2014).

If the transgene position within the nuclear region is not favourable, silencing machinery within C. reinhardtii might supress expression at the translational or post-translational level (López‐Paz et al., 2017). If multiple gene copies are inserted into the genome, then they may be inactivated by DNA methylation or histone modification (Jinkerson & Jonikas, 2015). To circumvent the silencing machinery, the elements within the transgene expression cassette must be modified to represent genes native to Chlamydomonas spp. Utilising endogenous algal promoters, introns and other elements within the gene cassette may help increase levels of expression (Scranton et al., 2016).

The only commercial microalgae plasmid currently available is the pChlamy vector series for C. reinhardtii (Thermo Fisher Scientific, Massachusetts, United States). The pChlamy4 vector contains a unique set of genetic elements that makes its expression toolkit far superior to alternate expression vectors for microalgae. Within the pChalmy4 vector are the endogenous algal Hsp70A- rbcS2 promoter, the endogenous algal intron-1 and the 3’ UTR terminator of rbcS2, the foot-and-

30 Chapter 2 mouth disease-virus (FMDV) 2A peptide sequence and the zeocin she ble resistance gene. The vector also contains both a N-terminal and C-terminal polyhistidine (6× His) tag. This tag remains on the N- or C-terminal when the heterologous protein is translated and allows for easy purification of the recombinant protein from the culture medium by immobilized metal affinity columns (Eichler- Stahlberg et al., 2009). After purification, the Tobacco Etch Virus (TEV) recognition site may be cleaved by a protease to remove the N-terminal 6x His tag from the protein (Sun et al., 2010).

To obtain higher levels of nuclear recombinant gene expression- the transgene may be codon optimised to the Chlamydomonas genome (Ramos‐Martinez et al., 2017). The redundancy of the universal genetic code allows six nucleotide triplets to encode the same amino acid. However, the frequency of degenerate tRNAs can vary in different organisms with some being more prevalent than others. The consequence of this redundancy is that many organisms display a clear codon bias (Nakamura et al. 2000). The genome project of C. reinhardtii allows insight into the endogenous gene structure (Franklin et al., 2002). With an average GC content of 68% in the nuclear coding sequences, a strong codon bias is prevalent in C. reinhardtii (Merchant et al., 2007); with A or U being the most prevalent nucleotide in the third position of the codon (Nakamura et al., 2000). Codon optimisation of transgenes to represent that of the C. reinhardtii genome would allow heterologous genes to be transcribed more efficiently if predominant degenerate tRNAs are used (Gustafsson et al., 2004).

With the optimal expression vector and gene codon optimisation, nuclear transformed C. reinhardtii clones may be obtained at a high frequency; but many researchers, including Schroda and co- workers (Schroda et al., 2000) state how difficult it is to identify those clones which express the gene of interest at desirably high levels. Hence, fluorescent protein expression is often used to simplify screening for high expressing transformants.

Genes coding for fluorescent proteins across the colour spectrum have been successfully expressed when integrated into C. reinhardtii genome. These include: the blue mTagBFP, cyan mCerulean, green CrGFP, yellow Venus, orange tdTomato and red mCherry proteins (Rasala et al., 2013). A comparative analysis revealed that the tdTomato protein fluoresced the highest; but it also had the lowest transformation efficiency. The tdTomato protein is twice the size of the other ﬂuorescent proteins which may complicate the translation and expression of the gene (Rasala et al., 2013). The most widely used fluorescent protein in microalgae research is the Green Fluorescent Protein (GFP) (Ghanbari Motlagh et al., 2016). Its sequence was previously codon-optimized to represent that of the C. reinhardtii nuclear genome (Crgfp) with a GC content of 61.33% which significantly increases its expression (Ghanbari Motlagh et al., 2016, Rasala et al., 2013). This was the first synthetic gene expressed in microalgae (Josef-Engert-Straße, 2015).

The gfp gene was first cloned from jellyfish Aequorea victoria in 1992 and used later that year as a reporter gene within E. coli (Cormier, 1992). Since then GFP has gained widespread use as reliable reporter with the main advantage that it may visualize patterns of gene expression in vivo (Soboleski et al., 2005). The green fluorescence of transgenic cells can be visually detected by Ultraviolet (UV) light and by spectrofluorometers to quantify the expression of GFP (Halfhill et al., 2005). The CrGFP protein is neither toxic to E. coli (Cormier, 1992) nor C. reinhardtii cells (Soboleski et al., 2005) and it may be detected above green microalgae autofluorescence (Ghanbari Motlagh et al., 2016). Once GFP proteins are produced they are quite stable and resistant to internal proteolysis or degradation

31 Chapter 2 brought on by external conditions such as high light, chemical exposure or the high turbidity that microalgae are cultured under (Shaner et al., 2005). Screening for CrGFP-producing C. reinhardtii cells is also more cost effective, time efficient and less labour-intensive than other reporter genes (Rasala et al., 2013). Overall it is considered one of the most reliable, sensitive and reproducible reporter proteins for gene expression (Soboleski et al., 2005).

The aim of this Chapter was to electro-transform the codon optimised Crgfp gene in pChlamy4 into the nuclear genome of the C. reinhardtii CC-125 and CC-400 strains. Molecular cloning of Crgfp into the pChlamy4 vector was achieved by an intermediary step of restriction enzyme subcloning into the pBluescript plasmid. Transgene integration within the transformants was confirmed on the DNA level through PCR amplification of the Crgfp gene, and then the production of the CrGFP protein was examined by fluorescence microscopy within live transformed C. reinhardtii cells.

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2.2 Materials and Methods Most recipes were made according to Protocol Online (www.protocol-online.org) unless stated otherwise. For all kit protocols, sterile reverse osmosis Millipore (Merck, South Africa) double distilled water (ddH2O) was used for elution instead of the elution buffer provided.

Bacterial strains and plasmids The gene of interest was the codon optimised Crgfp gene, contained in plasmid pCrGFP (Appendix A, Figure A1) obtained from the Chlamydomonas Resource Centre, based at the University of Minnesota (https://www.chlamycollection.org). The Crgfp transgene was first subcloned into the pBSK plasmid (Thermo Fisher Scientific, Massachusetts, United States) (Appendix A, Figure A2). The final vector for nuclear transformation was the pChlamy4 plasmid (Thermo Fisher Scientific, Massachusetts, United States) (Appendix A, Figure A3 and Figure A4). To ensure adequate amounts of vector for subsequent reactions, the plasmids were first transformed into the electro-competent DH10B Escherichia coli cells (Ausubel et al., 1992) as indicated below.

Bacterial transformations Electro-competent cells were thawed on ice and 0.5 µg/μL of the plasmid or inactivated ligation mixture (concentration seen in later sections) was aliquoted into the polypropylene tube containing 80 μl of the cells and gently mixed by tapping. After 5 mins the mixture was aliquoted into a chilled 1 mm Pulser® transformation cuvette (Bio-Rad, California, United States). The cells were pulse- electroporated with the settings: 1.8 kV, 25 μF, and 200 Ω with the Genepulser I (Bio-Rad, California, United States) and thereafter immediately suspended in 100 µl fresh Super Optimal Broth (SOC) media (0.5% Yeast Extract, 2% Tryptone, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4, 20 mM Glucose). The SOC media solution was transferred into a sterile McCartney bottle and incubated at 37°C, shaking at 200 rpm for an hour. Various volumes of the transformed cells were plated out (25 µl, 50 µl, 100 µl, 200 µl) onto ampicillin 15% agar plates. The plates were incubated overnight at 37°C, and an individual colony was picked and inoculated into 5 ml Lysogeny Broth (LB) media (10 mg/ml Bacto-tryptone, 5 mg/ml yeast extract, 10 mg/ml NaCl, 2% w/v dextrose) containing 100 μg/ml ampicillin and left to grow at 37°C shaking at 200 rpm for 12 hours. The E. coli culture was aliquoted into 2 ml polypropylene tubes with an equal volume of 50% glycerol and stored at -80°C.

Chlamydomonas strains and growth conditions The C. reinhardtii strains CC-125 and CC-400 (Table 2.1) were obtained from the Chlamydomonas Resource Centre. TAP medium with revised trace elements (Merchant et al., 2006) was used to grow the C. reinhardtii strains. The cultures were grown in Erlenmeyer flasks shaking at 200 rpm, at a temperature of 25 °C with 250 μmol –m.-2.s- 1 ﬂuorescent white light (Harris, 2009).

Table 2.1: The detailed genotypes and fundamental cell properties of two C. reinhardtii strains CC-125 and CC-400 used in this study.

Strains Alternate Genotype Intact cell Flagella- Size (μm) Reference names wall dependent motility CC-125 137c mt+ mt+ nit1 + + 6.96 ± 0.37 (Gallaher nit2 et al., 2015) CC-400 Cw15 mt+ mt+ cw - - 4.27 ± 0.24 (Fan et al., 2017)

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Primer design Primers, for PCR amplification of the Crgfp gene and for screening for transgene insertion into the pChalmy4 plasmid, were designed using Snapgene Software (GSL Biotech, Chicago) with the DNA sequences of the relevant transgene and plasmids (Table 2.2). These primers were subsequently analysed with the IDTDNA primer design tool (https://eu.idtdna.com/calc/analyzer) for hairpin loops, self-dimers, and heterodimers. None of concern were found and the primers were further analysed regarding the following values as parameters:

• Melting temperature (Tm) between 57°C and 70°C. • GC content between 30-80% • Less than 5°C difference in melting temperature between primer pairs

Table 2.2: Primers used in this study for amplification of the Crgfp transgene.

Primer Primer sequence (5’-3’) Length GC Annealing Restriction name (restriction enzyme sequence (bp) content temperature enzyme site underlined) (%) Tm (°C) CrGFP- TAG↓GATCCAAGGGCGAGGAGCTGTTC 32 59.4 69.1 XbaI For ACCGGT CrGFP- TCT↓CTAGATTACTTGTACAGCTCG 41 48.8 67 BamHI Rev TCCATGCCGTGGGTGAT pChlam GGAGAGCAACCCGGGCCCC 19 79 66 None y- For pChlam GCCTCCATTTACACGGAGCGGCTGC 25 64 67 None y-Rev

CrGFPp GACGTGGAGAGCAACCCGGGCCCCGAA 35 65.71 72 None 4-For TTC-CTCG CrGFPp GATCCAACGAGCGCCTCCATTTACACGG 40 60 71.3 None 4-Rev AGCGGCTGCAG * All primers were diluted to 10 mM working stock solutions. ** ↓ Indicates restriction enzyme insertion sites

For PCR amplification of the Crgfp gene, primers CrGFP-For and CrGFP-Rev were longer than recommended since they included a leader sequence (3-6 bp) and restriction enzyme site (6-8 bp) along with the hybridisation sequence (17-22 bp). The hybridisation sequence is the region of the primer which binds to the target gene sequence to be amplified. The recognition site allows for specific restriction enzymes to cut the sequence at this point, in this study the restriction enzyme sites were BamHI in the CrGFP-For primer and XbaI in the CrGFP-Rev primer. These restriction sites were chosen since they are not present within the Crgfp gene, are within only the multiple cloning site (MCS) of the pChlamy4 plasmid and may be utilised with the same buffer in the same conditions for restriction digests with two enzymes. The leader sequence is comprised of base pairs added onto the 5' end of the primer to assist the restriction enzyme binding and digestion.

The pChlamy-For and pChlamy-Rev primers set were designed to anneal outside the MCS region of the pChlamy4 plasmid. These primers were used in a PCR amplification to detect whether the Crgfp transgene was integrated into the pChlamy4 vector by size (the PCR product is 200 bp if no

34 Chapter 2 integration occurs and 917 bp if cloning is successful). The third primer set, CrGFPp4-For and CrGFPp4-Rev was designed to not include a restriction enzyme site and was used in sequencing reactions to validate the insertion of the Crgfp gene within the pChlamy4 vector.

Agarose gel electrophoresis Unless otherwise stated, all DNA samples (plasmid DNA or PCR products) were prepared with the addition of 1x of loading dye (Thermo Fisher Scientific, Massachusetts, United States). Samples were loaded and run using a 1% agarose gel made with Seakem® LE Agarose (Lonza Rockland, USA), 1x Tris, Acetic acid and EDTA (TAE) buffer (40 mM Tris, 20 mM Acetate, 1 mM EDTA, pH of 8.6) with 10 µg/ml Pronasafe (Laboratorios Conda, Spain). Loaded gels were electrophorized at 100 V for 1 hour in 1x TAE buffer (Sigma Aldrich, South Africa). For estimation of the DNA sizes of the DNA extractions, samples were compared to the GeneRuler 1 kb DNA Ladder (Thermo Fisher Scientific, Massachusetts, United States). The Chemidoc MP-R Gel documentation system (Bio-Rad, California, United States) was used to view all agarose gels in this Chapter.

Following gel electrophoresis, the PCR amplicons or plasmids were excised and purified using the

Zymoclean gel DNA recovery kit (Zymo Research, California, United States) with ddH2O for elution. The quantity and quality of all plasmid DNA and PCR products were determined by the NanodropTM 2000 spectrophotometer (Thermo Fisher Scientific, Massachusetts, United States) by obtaining the OD 260/280 and OD 260/230 absorbance ratios.

2.2.1 Subcloning Crgfp into pBluescript High fidelity Taq polymerase was used to amplify the Crgfp gene from the pCrGFP plasmid. A gradient of 60 to 70°C was set for Crgfp primers using the gradient function of the Mastercycler Gradient Universal Block (Eppendorf, Hamburg Germany). Each PCR reaction contained 0.3 μM CrGFP-For and CrGFP-Rev primers, 20 ng template Crgfp plasmid DNA, 1 U Hifi DNA Polymerase (Kapa Biosystems, South Africa), 0.3 mM dNTP Mix (Kapa Biosystems, South Africa), 1x High fidelity

GC Buffer (Kapa Biosystems, South Africa) and ddH2O to a final volume of 50 µl. The PCR cycling conditions were as follows: denaturation at 95°C for 3 min, 35 amplification cycles (98°C for 20s, then either 60°C, 63°C, 65°C, 68°C or 70°C) for 15s and 72°C for 1 min and a final extension step at 72°C for 1 min. The PCR products were viewed on a 1% agarose gel. Once the PCR reactions were considered optimal, the Crgfp gene was amplified for cloning and the PCR products were viewed on a 1% agarose gel.

The Crgfp PCR product was ligated into pBSK linearized with restriction enzyme EcoRV (Thermo Fisher Scientific, Massachusetts, United States). The Fast-link DNA Ligase kit (Epicentre, Wisconsin) was used for ligation. The ligation reaction contained 1X Fast-Link Buffer, 1.5 mM ATP, 30 ng of the pBSK plasmid, 21.3 ng of the Crgfp insert, 1 U/µl Fast link ligase and ddH2O water to a final volume of 15 μl. The molar ratio of Crgfp insert to pBSK vector was calculated to be 3 to 1. The ligation reaction was incubated at 4°C overnight, then inactivated at 70°C for 15 min.

The Crgfp-pBSK ligation reaction was electro-transformed into the E. coli DH10B cells and various volumes of the transformed cells were plated (25 µl, 50 μl, 100 μl, 200 μl) onto LB-IPTG-X-gal-Amp agar (LB media, 0.1 mM IPTG, 80 μg/ml X-gal, 100 μg/ml Amp) plates. The plates were incubated overnight at 37°C whereafter individual white colonies were transferred onto fresh LB-IPTG-X-gal- Amp agar plates. Randomly selected white colonies, with blue colonies as a negative control, were subjected to colony PCR.

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Transformants (white colonies) were tested via colony PCR for the presence of the Crgfp gene using CrGFP-For and CrGFP-Rev primers (Table 2.2). These and all subsequent bacterial colony PCR reactions utilised the KAPA 2G Robust PCR (Kapa Biosystems, South Africa) components. Each 10 µl reaction contained a transformed E. coli colony, 0.5 μM forward and reverse primers, 1 U Kapa 2G DNA Polymerase (Kapa Biosystems, South Africa), 0.3 mM dNTP Mix (Kapa Biosystems, South Africa),

1x KAPA 2G Buffer (Kapa Biosystems, South Africa), ddH2O and 5% DMSO. Previously PCR amplified Crgfp, 15 ng, was used as a positive PCR control with no template also utilised as a negative PCR control. The PCR cycling conditions were as follows: denaturation at 95°C for 3 min, 30 amplification cycles (98°C for 20 sec, 70°C for 15 sec and 72°C for 1 min) and a final extension step at 72°C for 5 min. The PCR products were then viewed on a 1% agarose gel.

The E. coli colonies which gave the desired PCR product underwent small scale plasmid extractions with the Zyppy Plasmid Miniprep kit (Zymo Research, California, United States). These plasmids were subjected to PCR using KAPA 2G with 20 ng of pure plasmid DNA as template, with the remainder of the PCR protocol as described above for colony PCR.

2.2.2 Cloning Crgfp into the pChlamy4 plasmid The Crgfp gene isolated from CrGFP-pBSK was ligated into the final vector, pChlamy4. To first confirm that the plasmid received was pChlamy4, restriction digestion of the plasmid was performed with approximately 400 ng of the pChlamy4 plasmid, 0.5 U/µl BamHI (Thermo Fisher Scientific, Massachusetts, United States) and 1 U/µl HindIII (Thermo Fischer Scientific, Massachusetts, United States) restriction enzymes. All digests were made up to a final volume of 20 µl in accordance to the Thermo Fisher Double Digest Calculator (www.thermofisher.com/za/en/home/brands/thermo- scientific/molecular-biology/thermo-scientific-restriction-modifying-enzymes/restriction-enzymes- thermo-scientific/double-digest-calculator-thermo-scientific.html) with 1x Tango buffer. The digest was incubated at 37°C for 3 hours and then the digested DNA was viewed following electrophoresis on a 1% agarose gel.

Restriction enzyme double digests were performed with 700 ng of the Crgfp-pBSK plasmid and 700 ng of the pChalmy4 plasmid with 0.5 U/µl BamHI and 0.5 U/µl XbaI restriction enzymes, made up to a final volume of 20 µl with 1x Tango buffer (Scientific, 2017). No plasmid, no enzyme and single digests were included as controls for both plasmids. The digests were incubated at 37°C for 3 hours and then the digested DNA was viewed on a 1% agarose gel and purified.

The Crgfp gene was ligated into the pChlamy4 plasmid using Fast-link DNA Ligase, with an insert to plasmid molar ratio of 3:1 with 33 ng pChlamy4 plasmid and 19.69 ng Crgfp insert. Following inactivation, 1µl of the Crgfp-pChlamy4 ligation reaction was transformed into E. coli DH10B cells via electroporation, as described above. The transformation mix was then plated onto LB agar plates containing ampicillin (100 µg/ml). The plates were incubated overnight at 37°C. Individual colonies were transferred onto fresh LB agar plates containing ampicillin (100 µg/ml) and screened using KAPA 2G colony PCR, as described above (Section 2.2.1). A single positive bacterial colony containing the Crgfp-pChalmy4 plasmid was grown for large scale plasmid extraction with the QIAGEN® Plasmid Maxi Kit protocol (Qiagen, South Africa).

To screen the isolated plasmid for the Crgfp gene, KAPA 2G PCR was performed with two different primer sets. The first primer set amplified the Crgfp region using CrGFP-For and CrGFP-Rev primers and the same PCR conditions were repeated with 20 ng of Crgfp plasmid DNA as a template, all the

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PCR products were viewed on 1% agarose gels. The Crgfp-pChlamy4 plasmid DNA was sent to Inqaba biotech for Sanger sequencing, using the sequencing primers CrGFPp4-For and CrGFPp4-Rev. These generated sequences were aligned on MEGA6 software (Tamura K, 2013) using the Muscle Alignment function for DNA.

2.2.3 Nuclear transformation of Chlamydomonas by electroporation The microalgae strains C. reinhardtii CC-125 and CC-400 were subjected to a zeocin kill curve experiment. The algal isolates, with a plating volume of 100 μl, were cultured in triplicate in a 6-well microplate (Corning, New York, United States) on solid 5 ml TAP 15% agar supplemented with the zeocin antibiotic present in the following concentrations: 0, 3, 5 and 10 mg/L.. Four evenly spaced 1 mm apertures in the plastic lid allowed for gaseous exchange whilst reducing contamination. The strains were left to grow for 14 consecutive days at a temperature of 25°C with 150 μmol –m.-2.s- 1 of continuous ﬂuorescent white light. Microalgal growth and mortality response to zeocin was visually scored at each concentration.

The Crgfp-pChlamy4 plasmid (600 ng) was linearized by performing multiple restriction enzyme digest with, 0.5 U/μl ScaI (Thermo Fisher Scientific, Massachusetts, United States) in 1x Tango buffer. Multiple digests were required to generate sufficient linearized plasmid for the transformation. No plasmid, no enzyme and single digests were included as controls. The digests were incubated at 37°C for 9 hours, inactivated at 80°C for 20 minutes and then 2 μl of each digested DNA sample was viewed on a 1% agarose gel. The linearized pChlamy4 restriction digestions were purified using the Isolate II PCR and gel kit (Bioline, South Africa). The digests were then pooled together to final quantities of 1, 2, 4 and 6 µg.

Transformation of the C. reinhardtii CC-125 and CC-400 strains was performed as indicated by the GeneArt™ Chlamydomonas Protein Expression Vector manual (Catalogue No. A24231, Thermo Fisher Scientific, Massachusetts, United States) with specific alterations to the cell concentrations and electroporation parameters, as described below. The C. reinhardtii cells were grown on TAP medium until the suspension reached a density of 1 × 106 cells/mL, counted using an algal haemocytometer (Thermo Fisher Scientific, Massachusetts, United States).

The C. reinhardtii cells were collected by centrifugation, for 10 min at 2,000 × g, and resuspended in 250 μl GeneArt™ MAX Efficiency™ Transformation Reagent (Thermo Fisher Scientific, Massachusetts, United States) to four final concentrations for each strain: 2 × 109 cells/mL, 3 × 107 cells/mL, 1 × 106 and 2 × 105 cells/mL. Different amounts of ScaI-linearized plasmid DNA (1, 2, 4 and 6 µg) as well as the empty pChlamy4 vector and a negative control (ddH2O) were aliquoted into the microalgae cell suspension and left to incubate on ice for 1 min (instead of 5 min) for subsequent transformations. The 250 μl transformation mixture was then transferred into a 4 mm ice cold electroporation cuvette (Bio-Rad, California, United States).

The cells were pulse-electroporated with the Bio-Rad Genepulser I, according to the GeneArt™ Chlamydomonas Protein Expression Vector manual at 500 V, a capacitance initially of 25 µF and then 500 µF, and 800 Ω. These electro-parameters were then changed to 1,500 V, a capacitance of 25 µF and 1,600 Ω. The cells recovered in the dark at 16˚C (as opposed to room temperature) for 15 min then transferred into a sterile conical tube containing 10 mL room temperature TAP-40 mM sucrose solution and left to incubate overnight at 26°C with 50 μmol m-2 s-1 white light.

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The conical tubes were centrifuged, for 5 min at 2,000 × g, the supernatant discarded and C. reinhardtii pellet resuspended in 200 µL TAP medium at room temperature. This entire solution for each transformation was plated and spread onto TAP 15% agar zeocin plates. The plates were left to grow at 26°C with 50 μmol m-2 s- 1 white light for 3 weeks until C. reinhardtii colonies were clearly visible. These colonies were transferred onto fresh TAP 15% agar zeocin plates.

2.2.4 Analysis of CrGFP microalgae transformants To ensure that the C. reinhardtii cells did not naturally fluoresce green, fluorescence of untransformed and transformed microalgae colonies were viewed under the Dual Fluorescent Protein (DFP) Flashlight-1 (NIGHTSEA, Lexington), with UV wavelength and filter sets at an excitation of 360 – 380 nm and emission at 415 nm longpass.

The Chelex-100 Colony PCR technique was utilised for screening the C. reinhardtii colonies (Cao et al., 2009). For both the CC-125 and CC-400 strains, approximately 110 randomly picked putatively transformed colonies, untransformed colonies, and a PCR positive control (the CrGFP-pChlamy4 plasmid) were subjected to PCR to detect the CrGFP transgene. Each colony was suspended in 50 µl 5% chelex-100 and vortexed for 5 – 10s, and allowed to cool on ice for 1 min. The tubes were boiled for 10 min at 100˚C in the Multigene Thermocycler (Labnet, South Africa). The tubes were then centrifuged for 1 min at 1000 x g to collect the liquid and 1 µl of the solution was used in the 10 µl KAPA 2G PCR reaction with CrGFP primers (Section 2.2.1). This colony PCR screen was repeated for four transgenic strains of CC-125 and CC-400 after 6 weeks of sub-culturing in liquid TAP zeocin media to verify whether the nuclear transformation was stable.

Four colonies with the brightest fluorescence of the positive transformed CC-125 and CC-400 microalgae colonies -confirmed by PCR- were selected for upscaling in liquid TAP medium supplemented with zeocin to a density of approximately 1 × 106 cells/mL. Four cultures of each transgenic CC-125 and CC-400 strain, were subjected to a growth curve analysis with the untransformed cultures as a control. To inoculate the 6 well microplate, 500 μl was aliquoted into the well containing 5 ml liquid TAP medium, at a pH of 7; with three biological replicates each. The plates were placed onto a Heidolph titramax shaker (Labotec, South Africa) set shaking at 200 rpm, at a temperature of 25°C with 250 μmol m-2.s-1 of 8 hour fluorescent white light: 16 hour dark phase cycles for 12 days. Microalgae cell density was measured each day, by aliquoting 100 μl of each replicate into a 96-well plate and reading the OD on the BIOTEK power XS multi-well plate reader (ADP Weltevreden Park, South Africa). The Gen 5 1.06 program was used to read the OD of each well at a wavelength of 750 nm.

Cells were visually inspected for CrGFP fluorescence by fluorescence microscopy using an Olympus IX71 microscope (Olympus, Massachusetts), equipped with an Olympus U-RFL-T mercury lamp (460– 495nm excitation ﬁlter and a 510–550nm Emission ﬁlter). The images were captured with a F-View camera and AnalySIS getIT software. Additionally, compound light microscopy- with the Olympus BX40 microscope (Olympus, Massachusetts) was used as a visual inspection of the cells from the transgenic lines compared to the untransformed C. reinhardtii CC-125 and CC-400 cells. For each microscopic inspection, 5 µL of each suspension sample was mounted on a slide and covered with a coverslip.

Fluorescent determinations were made using a Nunclon 96 microplate Flat Bottom Black Polystyrol plate (Thermo Fisher Scientific, Massachusetts, United States) with no cover, the fluorescence

38 Chapter 2 microplate reader (Hidex Sense, Finland) with Hidex Sense program software (V_1.30_05/2007_I500F) on an external PC controlling reader function. Determinations of CrGFP concentrations were made using the filter sets 340 nm, 25 nm bandpass excitation filter and a 535 nm, 10 nm bandpass emission filter without dilutions. Separate plates were used for each C. reinhardtii strain and both compared to a single standard GFP curve.

Four selected transformed colonies, as well as an untransformed colony, for each C. reinhardtii strain CC-125 and CC-400 were cultured with TAP zeocin till a whole cell concentration of 1× 107 mL−1, was reached for each. The quantity of CrGFP was determined by comparing its fluorescence with that of purified E. coli recombinant GFP (rGFP) standard (Catalogue No. 11814524001, Sigma-Aldrich, South Africa) at a known concentration. To minimize protein-protein variation rGFP was diluted in TAP for a series of dilutions to generate the standard curve. Ten concentrations (0-150 ng/ml) were used to generate the standard curve to cover the range of the unknown samples (Table 2.3). To account for baseline variation and natural autofluorescence of C. reinhardtii, fluorescence signal of the transformed cultures was normalised to the untransformed CC-125 and CC-400 microalgae. The fluorescence of the transformed strains was measured for four biological replicates with three technical replicates each.

Table 2.3: The rGFP dilutions utilised to generate the fluorescent standard curve in this study.

Well TAP buffer Volume of 1,000 ng/µl Final rGFP standard number (µl) rGFP (µl) concentration (ng/µl) 1 100 0 0 2 99.7 0.32 3.125 3 99.4 0.625 6.25 4 98.75 1.25 12.5 5 97.5 2.5 25 6 95 5 50 7 92.5 7.5 75 8 90 10 100 9 87.5 12.5 125 10 85 15 150

The numerical data generated was processed with statistical analysis on Microsoft Excel v1808 with all data presented as the mean ± standard error of the mean. The method of least squares was used for the regression analysis to determine the goodness of fit (r2) for the rGFP standard curve. A one- way ANOVA analysis, measured at a p ≤ 0.05 level of statistical significance, was used to detect significant differences between the untransformed and transgenic C. reinhardtii CC-125 and CC-400 cultures.

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2.3 Results

2.3.1 Transgene Crgfp subcloned into pBluescript plasmid

The optimal annealing temperature for PCR amplification of Crgfp was found to be 70°C (Appendix B, Figure B1). Thereafter, high fidelity PCR was performed at this annealing temperature and yielded an expected amplicon size of 727 bp for Crgfp (Figure 2.1).

Figure 2.1: KAPA HiFi PCR amplification of the Crgfp gene from the pCrGFP plasmid visualised on a 1% agarose gel. M = 1 kb GeneRuler DNA ladder (Thermo Fisher). Lanes 2-7: products of identical PCR reactions with an annealing temperature of 70°C. No amplification can be seen in the negative water control (lane 1), indicating no contamination. The desired band is prominent at the expected size of 727 bp.

The restriction digest of pBSK plasmid with EcoRV was performed previously at the CSIR to create a blunt insertion site (Roth, 2018). The amplified Crgfp PCR product was a blunt ended product produced by High fidelity Taq polymerase which was then ligated into the pBSK vector by blunt ended cloning. Following transformation of E. coli DH10B with the inactivated ligation reaction, five white colonies, indicative of an insert interrupting the lacZ activity (Alting-Mees et al., 1995), were selected (Appendix C, Figure C1) from the LB-IPTG-X-gal-Amp agar plates. Colony PCR confirmed that all colonies were positive for the Crgfp gene (Figure 2.2).

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Figure 2.2: A colony PCR amplification of the Crgfp region in Crgfp-pBSK containing E. coli colonies on a 1% agarose gel. M= 1 kb GeneRuler DNA ladder (Thermo Fisher). The desired band is prominent at the expected size of 727 bp seen in the control lane (1) and in all the white Crgfp-pBSK E. coli colonies (1- 5). No amplification can be seen in the no template control lane (6) indicating no contamination.

High fidelity Taq polymerase was utilised for amplifying the Crgfp gene for cloning, but high fidelity Taq polymerase is not necessary for transformant colony screening PCR assays; therefore, the 2G KAPA PCR was used for all subsequent screens. However, amplification of GC rich sequences is difficult due to the high stability and exertion for the primers to bind (Wurch et al., 2000). The 2G KAPA PCR was optimised with the addition of 5% DMSO to the PCR reactions. When DMSO binds to the DNA it allows the cytosine residues to change conformation and the DNA becomes more labile to heat (Tjernberg et al., 2006). As a result, the Tm was lowered, and the GC rich primers could anneal to the template to enhance amplification.

The six colonies, identified by PCR as containing the Crgfp-pBSK plasmid (Figure 2.2), were cultivated in ampicillin LB broth, and the plasmids isolated. The Crgfp-pBSK plasmid with the highest purity (260/280=1.90; 260/230=2.02) and concentration (187.9 ng/µl) was used for subsequent cloning reactions.

2.3.2 Transgene Crgfp cloned into pChlamy4 plasmid A restriction digest confirmed the pChlamy4 plasmid identity. The size and number of the fragments obtained for the respective digest with HindIII and BamHI yielded the corresponding fragments sizes that were to be expected of pChlamy4 (Appendix B, Figure B2). Further restriction digests were set up for both the CrGFP-pBSK plasmid and for the pChlamy4 recipient plasmid. The advantage of subcloning as opposed to cloning the PCR product directly into pChlamy4 is that Crgfp is cut out of the pBSK vector in its entirety with a band seen at the expected size of 717 bp separate from the linearized vector at 2,961 bp (Figure 2.3) indicating complete digestion.

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Figure 2.3: Restriction enzyme double digest of recombinant plasmid Crgfp-pBSK with BamHI and XbaI. The restriction digests were run with no plasmid control (1), no enzyme control (2) and Crgfp-pBSK double digested with BamHI and XbaI (3). The products were run with the 1kb Generuler DNA Ladder (M, Thermo Fisher) which indicates the vector backbone at 2,961 bp with the Crgfp gene, a faint band around 717 bp.

The same restriction enzymes were used to digest the pChalmy4 vector, yielding a prominent linear band at 3,640 bp (Figure 2.4). Both the Crgfp gene and linearized pChlamy4 vector were carefully cut from the gel and purified. The Crgfp transgene was then ligated into the pChlamy4 vector to form CrGFP-pChlamy4 (Figure 2.5).

Figure 2.4: Restriction enzyme double digest of plasmid pChlamy4 with BamHI and XbaI. The restriction digests were run for pChlamy4 with no enzyme control (1), no plasmid control (2), single digest with BamHI (3), single digest with XbaI (4) and double digested with both (5,6). The products were run with the 1kb Generuler DNA Ladder (M, Thermo Fisher) which indicates linearized DNA at approximately 3,500 bp for lane (6), lanes 4 and 5 have small amount of linearized DNA (3) and Lane 3 is virtually uncut (very little linearized DNA at 3,600 bp).

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Figure 2.5: A schematic diagram showing the Crgfp transgene cloned into the pChlamy4 vector with the BamHI and XbaI restriction enzyme sites located within the MCS.

Following transformation of E. coli DH10B colonies with the Crgfp-pChlamy4 ligation, thirteen of the colonies which formed on the ampicillin selection LB agar plates (Appendix C, Figure C2), were subjected to colony KAPA 2G PCR to determine the presence of the Crgfp gene. Seven colonies were positive for the expected 717 bp gene (Appendix B, Fig B3). These DNA bands were faint, so all the colonies were grown in ampicillin containing LB broth, the plasmids extracted, and PCR verification repeated. The isolated plasmids were screened for the codon optimised Crgfp gene, using KAPA 2G with two different primer sets (Table 2.2). The first primer set amplified the Crgfp region of 717 bp using CrGFP-For and CrGFP-Rev primers (Figure 2.6A), and the second PCR utilised the pChlamy4-For and pChlamy4-Rev primers, which were designed for annealing just outside the insertion site on pChalmy4, with an anticipated product size of 917 bp (Figure 2.6B). All the colonies were positive for both primer sets.

Figure 2.6: PCR amplification of the Crgfp region of Crgfp-pChlamy4 plasmids using (A) CrGFP-For and CrGFP- Rev primers and (B) pChlamy4-For and pChlamy4-Rev primers on a 1% agarose gel. M = 1 kb GeneRuler DNA ladder (Thermo Fisher). No amplification is visible for the water control lanes (1) indicating no contamination or primer dimers. The desired DNA band is prominent at the expected size of 717 bp for the positive control lane (2A) containing PCR amplified Crgfp DNA and in all the CrGFP-pChlamy plasmids (lanes 3 - 14).

The Crgfp-pChlamy4 plasmid extract with the highest concentration (169.9 ng/µl) and purity (260/280=1.93; 260/230=2.20) was used in further experimental steps. Plasmid DNA from two isolations was sequenced by Inqaba Biotech (Sanger sequencing method) using the CrGFPp4-For and CrGFPp4-Rev primers (Table 2.2). The results (Figure 2.7) confirmed that the cloned Crgfp gene was correct and that no errors arose during PCR and sub-cloning.

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Figure 2.7: Confirmation of the correct Crgfp-pChlamy4 vector by Sanger sequencing alignment of the sequenced Crgfp-pChlamy4 vector. Separate reactions containing the GFPp4-For and GFPp4-Rev primers for two Crgfp-pChlamy4 vectors were analysed on CLC Genomics software. 2.3.3 Nuclear transformation of Chlamydomonas by electroporation

The CrGFP-pChlamy4 plasmid was linearized by performing a restriction enzyme digest with ScaI. A no plasmid control and a no enzyme control were included with the digests and viewed with the products on an 1% agarose gel (Figure 2.8). The linearized CrGFP-pChlamy4 vector can be visualised in lane 3 at an expected size of 4,367 bp.

Figure 2.8: Restriction enzyme digest of recombinant plasmid Crgfp-pChlamy4 with ScaI. The restriction digest was run with no plasmid control (1), no enzyme control (2) and Crgfp-pChlamy4 digested with ScaI (3). The products were run with the 1kb GeneRuler DNA Ladder (M) with no contamination observed (1), undigested supercoiled vector (2) and the linearized vector at 4,364 bp (3).

The C. reinhardtii isolates CC-125 and CC-400 were tested to ensure they were susceptible to the antibiotic marker, zeocin. Natural resistance would prevent positive selection of the microalgae transformed with the pChlamy4 vector containing the Sh ble gene. A dose-dependent relationship was observed for each C. reinhardtii strain (Figure 2.9). The concentration 5 mg/ml was effective against the CC-125 strain and as little as 3 mg/ml was required for the CC-400 strain. To keep conditions constant, 5 mg/ml zeocin was used for the subsequent selection of all microalgal transformants.

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Figure 2.9: A zeocin kill curve of C. reinhardtii strains (A) CC-125 and (B) CC-400 grown in triplicate on TAP 15% agar at concentrations of 0, 3, 5 and 10 mg/L zeocin antibiotic.

The C. reinhardtii CC-125 and CC-400 cells transformed with linearized CrGFP-pChlamy4 were recovered on TAP zeocin 15% agar plates after electroporation. The transformations with the electroporation conditions stated in the GeneArt™ Chlamydomonas Protein Expression Vector manual failed to produce any colonies after transformation; therefore, electroporation parameters were optimised according to previously successful literature as discoursed in the discussion.

The amount of CrGFP-pChlamy4 linearized plasmid DNA per 250 µL of C. reinhardtii cell suspension was found to influence electroporation (Table 2.4). An increase in DNA volume resulted in arcing and increased time constants. By decreasing the DNA added and decreasing time constants, the 2 μg DNA loading was considered the most ideal for transformation conditions.

Table 2.4: Influence of Crgfp-pChlamy4 linearized plasmid volume on electroporation arcing and number of colonies produced on a plate.

Amount of linearized Arcing Time constant (t) Number of colonies DNA (µg) 0 (ddH2O control) - 4.5 No colonies 1 - 1.4 No colonies 2 - 2.6 200-350 colonies 4 + 7.4 Lawn of colonies 6 + 11.2 Lawn of colonies

The concentration of C. reinhardtii cells also influenced the growth of the recovered transformants when transformed with 2 µg Crgfp-pChlamy4 linearized plasmid DNA. The higher concentration of 2 × 109 cells/mL of cells yielded a lawn of transformants after three weeks of growth (Figure 2.10A), however these colonies were mostly false positives according to colony PCR (data not shown). The cells transformed at low concentrations, 2 × 105 cells/mL, failed to grow after three weeks (data not shown). The concentration of 1 × 106 cells/mL yielded some colonies (Figure 2.10C), but the highest number of transformants was obtained when transformed at a cell density of 3 × 107 cells/mL as single colonies appear on the surface of the plate after three weeks of growth (Figure 2.10B). As expected, the CC-125 and CC-400 transformation without Crgfp-pChlamy4 linearized plasmid DNA did not yield any colonies when grown on TAP zeocin 15% agar plates and the transformation with the empty vector, pChlamy4 did (data not shown).

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Strain (A) 2 × 109 cells/mL (B) 3 × 107 cells/mL (C) 1 × 106 cells/mL

CC-125

CC-400

Figure 2.10: Colonies after electroporation transformation of the CC-125 and CC-400 strains with concentration of (A) 2 × 109 cells/mL, (B) 3 × 107 cells/mL and (C) 1 × 106 cells/mL; plated on TAP 15% agar medium containing 5 µg/mL zeocin.

Once the C. reinhardtii colonies had a size of approximately 1 mm they were picked and subjected to colony PCR. To bypass tedious genomic DNA extractions for numerous samples, the Chelex-100 Chlamydomonas sp. colony PCR was successfully utilised (Cao et al., 2009). For both the CC-125 and CC-400 strains, approximately 110 randomly picked transformants were screened and compared to the untransformed colonies and the PCR positive control, Crgfp-pChlamy4 plasmid DNA (Appendix D, Figure D1 and D2).

The number of transformants selected on the TAP zeocin 15% agar plates and percentage of positive transformants were calculated for each strain (Table 2.5). The transformation efficiency was calculated as the number of colony forming units (cfu) which would be produced by transforming 1 µg of Crgfp-pChlamy4 plasmid into a given volume of 3 x 107 C. reinhardtii cells. The percentage of Crgfp gene positive transformants was calculated only from those screened by PCR. The CC-125 strain had more colonies growing on the plate, but fewer of these were positive for the Crgfp gene when compared to the CC-400 strain which had fewer recovered colonies but a greater percentage of CrGFP transformants.

Table 2.5: Transformation data for the C. reinhardtii strains CC-125 and CC-400. At least 109 independent transformants for each strain were analysed for the Crgfp gene by colony PCR.

Chlamydomonas Number of Relative Microalgae Gene Percentage strains cfu’s per µg transformation screened positive gene positive DNA* efficiency CC-125 350 12.93 109 80 73.4% CC-400 212 7.83 111 97 87.4% *2 µg of linear DNA was used per electroporation

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The positive colonies from the selection plates of each strain were transferred onto fresh TAP medium plates and left to grow for another three weeks to enhance homoplastic state of transformation (Appendix E). Four selected positive colonies of each C. reinhardtii strain were picked from the selection plates and grown under continuous sub-culturing in liquid TAP containing zeocin. These cultures were used for subsequent experiments over a period of six weeks.

A growth study was performed on the transgenic CC-125, CC-400 and untransformed strains within both liquid TAP media with zeocin and liquid TAP media without zeocin. The growth analysis indicated no significant growth curve differences between the CC-125 and CC-400 CrGFP transformed and untransformed strains in liquid TAP medium (Figure 2.11A). The growth pattern remained the same for the transformed strains when cultured with zeocin but, as expected, the untransformed strains did not grow (Figure 2.11B).

0,7 CrGFP CC125 Chlamydomonas 0,6 reinhardtii 0,5 CrGFP CC400 Chlamydomonas 0,4 reinhardtii CC125 Chlamydomonas 0,3 reinhardtii

OD OD nm) (750 0,2 CC400 Chlamydomonas 0,1 reinhardtii 0 0 2 4 6 8 10 12

Time (Days) A

0,7 CrGFP CC125 0,6 Chlamydomonas reinhardtii 0,5 CrGFP CC400 Chlamydomonas 0,4 reinhardtii CC125 0,3 Chlamydomonas

reinhardtii OD OD nm) (750 0,2 CC400 Chlamydomonas reinhardtii 0,1

0 0 2 4 6 8 10 12 Time (Days) B

Figure 2.11: Growth curves of the untransformed and Crgfp transformed C. reinhardtii strains, CC-125 and CC-400 strains in (A) liquid TAP media and (B) liquid TAP media supplemented with zeocin.

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Neither the untransformed CC-125 nor the CC-400 C. reinhardtii strain exhibited natural green fluorescence under the DFP Flashlight (Appendix F, Figures F1 and F2). The fluorescence microscopy indicated the production of CrGFP in the transformed C. reinhardtii CC-125 and CC-400 cell cultures compared to the untransformed strains (Figure 2.12). In the transformants, the plasmid was directed to the nuclear genome with the translated CrGFP protein having no specific protein localisation signal sequence. This resulted in most of the protein being projected to accumulate in the cytosol resulting in the transformed cells having a green fluorescent appearance. The red fluorescence in the untransformed strains was mainly emitted by the chlorophylls of the photosystem II in the chloroplast (Rasala et al., 2013). To detect possible morphological alterations at cellular level, the transformed CC-125 and CC-400 cells were also observed under a compound microscope with no signiﬁcant differences to the untransformed cultures regarding cell morphology, size and shape.

Strain (A) Transformed (B) Transformed with a fluorescent (C) Untransformed signal of CrGFP CC-125

CC-400

Figure 2.12: The C. reinhardtii CC-125 and CC-400 transformed cells seen with (A) light microscopy and fluorescence microscopy of (B) transformed and (C) untransformed cells, with a fluorescent signal in (B) of CrGFP derived from the CrGFP-pChlamy4 plasmid. Scale bar: 50 μm.

The unaltered growth curves of the untransformed and transformed C. reinhardtii strains, led us to assume that neither the Crgfp nor she ble gene or protein is toxic to C. reinhardtii and there is no observed detrimental effect due to the expression of these genes. There was no observable influence on cell morphology due to the expression of the Crgfp transgene.

The ﬂuorescence microplate analysis of four selected transformed colonies for each C. reinhardtii strain CC-125 and CC-400, at a cell density of 1 x 107 detected ﬂuorescence signals. Their Relative Fluorescent Units (RFUs), at the appropriate excitation/emission wavelengths, were normalised with the untransformed strains, with one plate used per strain. The quantity of CrGFP produced for each transformant was then determined by comparing its fluorescence against a rGFP standard curve (Appendix G, Figure G1). The standard curve was created using the rGFP standard and had a high degree of confidence (R2 = 0.996). The purified rGFP (produced from E. coli) has similar properties to CrGFP that it also contains a 6x His-Tag and has a molecular weight of ∼28 kDa; even so, protein- protein variation is inevitable. The CrGFP signal within transformed C. reinhardtii cells could also be masking or enhancing compounds within the cells compared to the purified rGFP standard protein and therefore concentration calculated should only be taken as a relative and not an actual concentration determination.

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12000 12000

10000 10000

8000 8000

6000 6000 4000 4000 2000 2000 2.18 0.66 0.88 3.90 1.47 0.45 1.85 1.31 0 0

Relative Fluorescent Units Fluorescent Relative Relative Fluorescent Units Fluorescent Relative CC-400-A CC-400-B CC-400-C CC-400-D CC-125-A CC-125-B CC-125-C CC-125-D

C. reinhardtii CC-400 Transformants C. reinhardtii CC-125 Tansformants

Figure 2.13: Quantiﬁcation of the relative CrGFP ﬂuorescence of four selected (A) CC-400 and (B) CC-125 transformed whole cell transformants normalised to their untransformed C. reinhardtii counterpart. Emission has been quantified at 535 nm upon excitation at 340 nm. The concentration of CrGFP (ng/ 100 µl well) is indicated per column for each transformant calculated from the same rGFP standard curve. Two separate plates were used for each C. reinhardtii strain, whole cell concentrations were 1× 107 mL−1 and the RFU values represent the mean ± standard deviation (n = 3).

The average CrGFP fluorescence was not the same for each transformed C. reinhardtii clone. A maximum difference of 13-fold was seen with clone CrGFP-CC400-D showing a dramatically higher fluorescence. The minimum fluorescence, clone CrGFP-CC125-B, was still 4-fold greater than the untransformed C. reinhardtii cells. In terms of detection limits as little as 0.45 ng up to 3.9 ng of CrGFP was detected per 100 µl well (Figure 2.13). Strain specific differences were also noted with regard to their protein concentration in relation to fluorescence

To verify whether the nuclear transformation was stable, four selected transgenic strains of CC-125 and CC-400 were evaluated after six weeks of sub-culturing in liquid TAP media with zeocin. Evaluations showed zeocin resistance and green fluorescence using a DFP Flashlight-1 (Appendix F, Figure F3). Stable transformation was further confirmed by colony PCR for CC-125 (Figure 2.14A) and CC-400 (Figure 2.14B).

Figure 2.14: The C. reinhardtii (A) CC-125 and (B) CC-400 colony PCR screen for Crgfp-pChlamy4 transformants on a 1% agarose gel after 6 weeks of liquid TAP culturing with zeocin. M = 1 kb GeneRuler DNA ladder (Thermo Fisher). Crgfp primers CrGFP- For and CrGFP-Rev were utilised and, in each gel,, no amplification is observed in the negative ddH2O control lanes (1) and in the untransformed culture (3). The desired DNA band is present at the expected size of 717 bp for the positive control lanes containing CrGFP- pChlamy plasmid DNA (2) as well as in the transformed cultures (lane 4-7).

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2.4 Discussion The transformation of the Crgfp transgene into the nuclear region of the C. reinhardtii CC-125 and CC-400 strains was successful. The molecular sub-cloning restriction enzyme approach (through the pBluescript plasmid) of the Crgfp gene into the pChlamy4 vector produced no mutations. This could be accounted for by the high fidelity Taq polymerase; known for its minimal probability of mutations in downstream cloning and expression (Stevenson & Brown, 2015).

The Crgfp transgene in pChlamy4 was then successfully electro-transformed into the nuclear region of both C. reinhardtii strains. The electroporation requirements were optimised as initially the transformation parameters outlined in the pChlamy4 GeneArt™ Chlamydomonas Protein Expression Vector manual failed to produce transformants. When considering electroporation as a tool for transformation, there are several factors which affect the overall efficiency. These variables include those associated with the electroporation device settings, such as voltage, temperature, capacitance and resistance of the electric current. Optimisation of the cell concentrations, DNA concentrations, the electroporation buffer, electroporation recovery and very importantly the type of cells used were also considered. The electroporation parameters in this study were specifically altered from the manual to account for the older model of the Bio-Rad™ Gene Pulser™ I device used. With guidance from previously successful studies, the voltage was increased, temperature reduced, and the resistance adjusted as necessary.

In many studies, a higher voltage than the suggested 500 V (Scientific, 2016), had a greater transformation efficiency in Chlamydomonas sp. (Shimogawara et al., 1998). In a study where a NEPA21 electroporator was used, the C. reinhardtii CC-125 strain required a higher voltage input to induce pore formation compared to other wild-type strains (Yamano et al., 2013). A study where the voltage, permeability and viability of C. reinhardtii cells were tested, demonstrated the most optimal voltage to be 2,200 V (Muñoz et al., 2018). This corresponds to our research where transformants were obtained for the higher voltage of 1,500 kV. A higher voltage is also required for smaller cells (Kotnik et al., 2015) and the microalgae cells vary in their diameter depending on their growth phase. Therefore it was important to transform cells during the early log phase of culturing so that cell wall thickness or life stage complications were not a limiting factor (Ghanbari Motlagh et al., 2016). The C. reinhardtii CC-400 strain was observed to have a smaller diameter and the optimal voltage required for this strain should be more carefully examined.

The field strength comprises a range of parameters including, cell size, cuvette gap size and temperature which allows the voltage to be delivered across the electrode gap and permeate the cell membrane (Muñoz et al., 2018). A field strength parameter we are able to change, is temperature and this can be done during and post-electroporation. Within the Thermo Scientific protocol, the cuvette was chilled prior to electroporation and then left to incubate at room temperature. If the cuvette is left at room temperature, the amount of pulses, pulse duration and high voltage exposure could heat up and kill the cells (Chow & Tung, 1999). Therefore a lower temperature was utilised in this study, as it circumvented the consequences of over-heating. A lower temperature also allows the membrane pores to remain open for longer allowing more exogenous DNA to enter the cell (Kotnik et al., 2015). From previous studies, the optimal temperature for recovery of C. reinhardtii after electroporation was considered to be 10-20˚C (Shimogawara et al., 1998) and 16˚C (Wittkopp, 2018). In this study the cuvette was pre-chilled on ice before transformation, and afterwards the cells were recovered at 16˚C. Temperature and voltage are

50 Chapter 2 directly proportional (Chungjatupornchai et al., 2016) and lowering this temperature was another reason the applied voltage was increased in this study.

The Thermo Fisher protocol for algal transformation requires a capacitance of 50 µF, however, the Bio-Rad™ Gene Pulser™ I device only reaches a capacity of 25 µF or -with the addition of an external capacitor- 500 µF. Capacitance is the ability to store electrical energy, measured in units of farads (F) (Bio -Rrad, 1988). The exponential decay waves emitted by the electroporator generate an optimal- peak electrical pulse by allowing a capacitor to store its electrical energy and then completely discharging it. The electroporated cell has a high voltage supply administered which slowly declines over time (Bio-Rad, 1990). The time taken is dependent on the exponential decay wave system which has a time constant (T) pulse which is the rate at which the voltage decays to 1/e (approximately 37%) of the peak value (Figure 2.15).

Figure 2.15: The exponential decay waveform as a measure of T pulse which is the rate of which the voltage decays to 1/e (~37%) of the peak value (BioRad, 1988).

A greater pulse length would increase the pore formation and uptake of exogenous DNA (Kotnik et al., 2015). To calculate the time constant (BioRad, 1988), the values of resistance (R) and capacitance (C) are required such that:

푇 = 푅퐶

The capacitance required in the Thermo Fisher manual was 50 μF, which corresponds to a time constant of 40,000 ms with their suggested voltage (500 V) and resistance (800 Ω). To compensate for the capacitance, the resistance of the Bio-Rad™ Gene Pulser™ I was adjusted to 1,600 Ω to achieve the same time constant.

The resistance may also be affected by the buffer of the cell samples for electroporation. The aim of the buffer is to allow the cells to absorb water and swell right before they pulse allowing the cell membrane to be more permeable (BioRad, 1988) and it also enhances stability and activity over time (Kotnik et al., 2015). However when the ionic strength is too high then the salts could be damaging to some cells and cause an electrical short circuit, known as arcing (Sustarsic et al., 2014). When considering a solution, both the osmolarity and volume have an inversely proportional effect on the resistance. When the volume or ionic strength increases, the resistance decreases, and the potential of arcing is greater (BTX, 2017).

Most buffers used for electroporation are highly conductive, such as phosphate buffered saline or other standard culture media which may contain serum or hypo-osmolar buffers (Bio-Rad, 1988). The buffer used in this study was the GeneArt™ MAX Efficiency™ Transformation Reagent. In this study the cells arced when a greater volume of the Crgfp-pChlamy4 linearized plasmid was utilised. This could have been due to the use of Crgfp-pChlamy4 ScaI restriction enzyme digest reactions

51 Chapter 2 directly, which could have contained an excess number of ions. Therefore, a PCR Clean-up kit was to purify and elute the linearized plasmid DNA utilised with ddH2O for all experimental steps. The clean-up step would reduce the amount of contaminating salts within the final electroporation solution and minimise arcing. The optimal concentration and volume of the Crgfp-pChlamy4 plasmid used for C. reinhardtii transformation in this study was 2 µg in 2 µl. Consideration was taken to not reduce the volumes of DNA to less than 1 µg/µl since the transformation efficiency could easily be evaluated as a function of DNA concentration (Yamano et al., 2013).

The Crgfp-pChlamy4 plasmid was linearized before transformation, as previous studies found it to increase the transformation efficiency up to 2-fold (Ghanbari Motlagh et al., 2016). Linearized DNA reduces the probability of the insert becoming rearranged or being accompanied by a large deletion in the nearby genomic DNA, therefore reducing transcriptional silencing (Allnutt et al., 2000, Coll, 2006, Ghanbari Motlagh et al., 2016)

The Thermo Fisher protocol requires the Crgfp-pChlamy4 DNA to be incubated with the C. reinhardtii cells in transformation reagent buffer for five minutes before transformation. However in previous studies, it was found that DNA binding to the cells was not necessary and incubating for longer than a minute did not increase the transformation efficiency (Dower et al., 1988). In fact, preincubation of DNA and the microalgal cells could be detrimental if nucleases are released by the microalgae cells. Endonucleases were found to be released upon detergent applications in both cell wall containing, CC-125 and mutant strains CC-400 (D’Souza et al., 2018). The nucleases accumulate in exponential phase and appear to be membrane bound since they are released when the cells are treated with cell-wall degrading lytic enzymes (D’Souza et al., 2018). The GeneArt™ MAX Efficiency™ Transformation Reagent contains lytic enzymes to remove the cell wall of C. reinhardtii, this could release endonucleases into the solution and degrade the Crgfp-pChlamy4 DNA. Previous studies found that divalent cations (Ca+ 2, Mg+2, Cu+2 and Zn+2) may prevent DNA degradation (Kotnik et al., 2015). We did not consider the addition of these to the electroporation buffer since the divalent cations would change the ionic strength, resistance, turgidity as well as affect the permeability and survival of cells (Kotnik et al., 2015). Instead, we reduced the incubation time from five minutes as suggested (Scientific, 2016), to one minute, reducing the DNA exposure to the endonucleases.

The concentration of cells was also a factor which could affect the transformation and plating efficiency on solid TAP agar with zeocin after transformation. When the cell concentration was too low, few cells survived. On the other hand, when the cell concentration was too high, a lawn of colonies, of which the majority were untransformed, appeared. The high cell concentrations could have caused the cells to stack and reduce contact with the antibiotic zeocin, reducing its efficiency (Muñoz et al., 2018). The optimal cell concentration for transformation was indeed, mid-exponential phase (grown to a density of 1 x106 cells/mL) as suggested by the Thermo Fisher protocol and transformed at a density of 3 x107 cells/mL. This mid- exponential growth phase is the most optimal for accepting DNA into the nuclear genome (Cerutti et al., 1997). It is evident that transformation efficiencies are not just dependent on the electroporation parameters, but also on the cells themselves; the type of cell and their physiology, which in turn allows for cellular acceptance of foreign DNA during electroporation.

This study compared the untransformed CC-125 and CC-400 strains to those transformed with Crgfp- pChlamy through microscopy and growth analysis within liquid TAP media and on solid TAP media (both with and without zeocin). No physiological or morphological differences were observed

52 Chapter 2 between the transformed and untransformed strains. Previous growth curves of recombinant C. reinhardtii transformed with a very similar expression vector containing a: 5’-UTR, element, Hsp70A-rbcS2 promoter, genes (ble, aphVIII and gfp), and a 3’-UTR element also had the same growth properties as wild type C. reinhardtii (Noor-Mohammadi et al., 2014). We can conclude that the production of CrGFP does not have a harmful effect on cell growth, health or physiology (Rasala et al., 2013). However, differences were noted for the morphology, growth and transformation efficiency between the two C. reinhardtii strains CC-400 and CC-125.

The CC-400 cells lack a cell wall which make them significantly smaller than the CC-125 cells (Taghavi & Robinson, 2016). The CC-400 strain grew at a faster rate, reaching a higher density than CC-125, which has also been seen in previous studies (Fan et al., 2017, Taghavi & Robinson, 2016). Taghavi and Robinson further linked this to the five-fold faster chlorophyll production rate for CC-400 compared to CC-125 (Taghavi & Robinson, 2016). The growth of the CC-400 on solid TAP agar demonstrated that these cells were prone to drying out as a result of the absence of a cell wall (Harris, 2009). Cell walls protect microalgae from external conditions such as changes in temperature, light, pH, harmful chemicals as well as exogenous DNA (Janssen, 2002).

The CC-125 was naturally more resistant to zeocin than the CC-400 strain, and the latter had fewer colonies after transformation. However, the CC-400 strain had a greater percentage of transformants that were Crgfp positive and had higher CrGFP fluorescence than CC-125. This could be attributed to the lack of a cell wall, which means more external DNA could enter the cell to integrate into its nuclear genome. A higher percentage of the CC-400 colonies were positively transformed since their natural resistance to zeocin was lower at the same zeocin concentration as the CC-125 strains, and in so doing providing a more stringent selection criterion for positive clones (Garcia-Echauri & Cardineau, 2015). In a study by Doron and co-workers they discovered that cell wall deficient strains are not necessary for electro-transformation but they have a predominantly greater transformation efficiency (Doron et al., 2016). The cell wall of CC-125 which makes it more tolerable to zeocin, could also make it more resistant to exogenous DNA integration, especially since in this study, the cell wall degrading autolysin incubation time was decreased before electroporation.

The most successful selectable markers for stable nuclear transformation within multiple species of microalgae are antibiotic resistance genes, specifically the zeocin resistance gene (Garcia-Echauri & Cardineau, 2015, Stevens et al., 1996). In this study, CC-125 and CC-400 had a linear dose-dependent relationship with zeocin, the most optimal selection concentration regarded as 5 mg/L. Similar results were seen in a previous study as zeocin had a direct effect on the double stranded DNA breaks within the C. reinhardtii strain CC-400 (Chankova et al., 2007). A dose-dependent relationship was also observed as C. reinhardtii cell survival decreased exponentially from a zeocin concentration of 10- 50 mg/L (Chankova et al., 2007). The concentration recommended by Thermo Fisher Scientific for C. reinhardtii transformation is also 5 mg/L. It would be more beneficial to have the zeocin concentration as low as possible so that up-scaling the microalgae growth would be cost-effective. The lower the antibiotic concentration required to maintain transformation, the lower the costs that would be involved in the maintenance of transgenic cultures (Garcia-Echauri & Cardineau, 2015).

The antibiotic resistance for zeocin is present within the pChlamy4 vector as the Sh ble gene. Higher levels of recombinant protein production have been reported when Sh ble gene was used as a selection marker in C. reinhardtii when compared to other antibiotic selection markers (Calmels et

53 Chapter 2 al., 1991, Garcia-Echauri & Cardineau, 2015, Chankova et al., 2007, Stevens et al., 1996). The FMDV 2A-sh ble fusion protein utilised in this study was also used by Rasala et al. and from their studies they stated that reducing the levels of zeocin had minimal effect on gene expression and greatly improved the transformation efﬁciency (Rasala et al., 2013).This is supported by other studies where the sh ble gene was fused to a FMDV 2A peptide sequence, high expression levels of approximately 0.25% total soluble protein, were obtained (Rasala et al., 2012).

We observed differences in protein production among the transformed cultures which may be attributed to copy number and randomness of transgene integration into the nuclear genome (Slaninová et al., 2008). The CrGFP accumulated to detectable levels within both C. reinhardtii CC- 125 and CC-400 strains, with greater production and fluorescence detected in CC-400. The protein seemed to be efﬁciently processed to yield mature, unfused CrGFP that localized throughout the cytoplasm. In this study, the CrGFP transformed C. reinhardtii strains fluoresced from 4-fold for the CC-125 strain, to 13-fold for the CC-400 strain, when compared to the untransformed counterparts. When compared to the standard rGFP curve it was estimated that the transformed cultures produced CrGFP from 0.45 to 3.9 ng per 100 µl well.

The results in this study are comparable to previous reported data on the expression of codon optimised Crgfp genes integrated into the nucleus of C. reinhardtii. An integration vector with psad, rbcS2 and β2 tubulin promoters and their respective 3΄ and 5΄ UTR’s achieved a 1.1-fold increase in fluorescence compared to the wildtype (Noor-Mohammadi et al., 2014). In addition a integration vector with genetic elements Hsp70A-rbcS2 enhancer/promoter and 3΄UTR rbcS2 introns achieved a 3-fold increase in fluorescence compared to the wildtype (Rasala et al., 2013). Further research by Ghanbari et al. confirmed that the Hsp70A-rbcS2 enhancer/promoter and the introns rbcS2 3΄UTR introns (used within this study) achieved the greatest production as they achieved a 28-fold increase in fluorescence compared to the wildtype (Ghanbari Motlagh et al., 2016).

The high levels of nuclear expression may be linked to the genetic elements contained within the pChlamy4 vector. The rbcS2 promoter fused to the Hsp70A enhancer element increases transgene expression (Schroda et al., 2000). The element serves as a transcriptional activator, conferring light induction and high levels of stable transgene products (Schroda et al., 2000). In several studies the Hsp70A-rbcS2 enhancer/promoter yielded greater protein production than any other promoter and is considered the strongest algal promoter to date (Garcia-Echauri & Cardineau, 2015, Schroda et al., 2000, Fischer & Rochaix, 2001, Perozeni et al., 2018).

In microalgae, gene expression is further enhanced when introns near the 5′ end are included just downstream of the ATG site in the gene construct (Gruber et al., 1996). To test the elements which confer high transgene activity, 25 synthetic algal promoters were developed for C. reinhardtii (Scranton et al., 2016). The first intron of the endogenous C. rehihardtii rbcS2 resulted in the highest recombinant protein yields (Scranton et al., 2016). This intron inserted into the sh ble gene specifically increased expression levels by up to 30-fold (Fischer & Rochaix, 2001). The 3’ UTR used for all transgenes in the construct was also sourced from the rbcS2 gene and is recognised as one of the most efficient endogenous microalgae terminators. It enables transgene stability by regulating the mRNA stability and polyadenylation signals of the transgene (Díaz‐Santos et al., 2013).

Another technique to increase the level of expression within nuclear transformants is to link the transgene to the resistance of a selective agent. When the she ble antibiotic resistant marker gene is

54 Chapter 2 fused to the gene of interest by the FMDV 2A linker peptide, the transformant’s resistance to zeocin is directly coupled to the translation of the desired protein (Collin et al., 2019). The FMDV encodes a short ∼20 amino acid sequence, and when translated, the last two amino acids fail to form a peptide bond and the fused genes from a single mRNA forms two separate proteins (Rasala et al., 2014). Previous research has proven this approach to be a highly effective with a 100-fold increase in the expression of fungal xylanase (Rasala et al., 2012). This system also maintains its high expression levels for long periods, even after the selection agent has been removed (Rasala et al., 2013).

The Crgfp transgene was a convenient and accurate analytical indicator of transformation success and transgene expression. It is a non-invasive, cost effective manner to quantify the amount of CrGFP protein produced within the transformed C. reinhardtii cells in real time. The CrGFP is a sensitive indicator of expression and as we observed with the rGFP standard curve, its fluorescence increased linearly to protein concentration. Similarly, fluorescence increases in direct proportion to the Crgfp mRNA abundance and gene copy number, and is also a reliable measure of underlying differences in gene expression (Soboleski et al., 2005, Halfhill et al., 2005, Rasala et al., 2013, Persson et al., 2002). Depending on its application, fluorescence quantification may surpass RT-qPCR which only measures transcription levels and does not quantify the protein expression (Franklin et al., 2002). As a laboratory-based protein analysis tool, GFP-based protein estimates are also far less time and labour intensive than either enzyme-linked immunosorbent assay (ELISA) (Halfhill et al., 2005) or western blot analysis (Khan et al., 2016).

However, many factors beyond copy number may affect the fluorescence of GFP; rate of maturation, protein mis-folding and protein modification are internal factors that should also be considered (Rasala et al., 2013, Ghanbari Motlagh et al., 2016). It is necessary to note that GFP fluorescence in whole canola plants differed depending on the age of the leaves (Halfhill et al., 2005). Therefore, it was crucial that CrGFP analysis in this study was done on healthy microalgae cultures during the same growth phase at the same cell density. Fluorescence comparison may be limited to the same species, grown under the same conditions (Persson et al., 2002). Due to a variety of laboratory variations, it is not advised to compare actual CrGFP values to other experiments but only as an indication of relative expression and production.

Green CrGFP is the least fluorescent protein, compared to the spectra of fluorescent proteins available for expression within C. reinhardtii (blue mTagBFP, cyan mCerulean, yellow Venus, orange tdTomato and red mCherry) (Rasala et al., 2013). This may be attributed to point mutations within the CrGFP gene and the green spectrum background autofluorescence of green microalgae (Rasala et al., 2013). However, CrGFP was readily available and since the study sought to determine whether microalgal isolates were transformable, a CrGFP accumulation with detectable fluorescence in live- cell microscopy was more than adequate for this study. The CrGFP gene was cloned into the pChlamy4 vector which was stably transformed, with electroporation optimisation, into the nucleus of the C. reinhardtii CC-125 and CC-400 strains. The two strains differed in their response the zeocin selection agent, growth and production of CrGFP.

Study limitations and future considerations Now that we can electro-transform C. reinhardtii to successfully express CrGFP within the nuclear genome utilising our current optimised laboratory conditions, we may consider the future application of microalgae producing recombinant proteins. Fluorescence is an effective mechanism

55 Chapter 2 for screening high expression level transformants and for directly monitoring protein turnover in vivo (Cronin & Hampton, 1999, Ghanbari Motlagh et al., 2016). The fluorescence of the Crgfp gene is maintained when the gene is fused to a recombinant protein (Soboleski et al., 2005) and we may fuse it to a protein of industrial value.

Due to time constraints we optimised the electroporation according to previously successful C. reinhardtii transformation events found in literature, if more time were available all transformation parameters could be tested under the Taguchi principle (Heiat et al., 2012). This would allow us to identify the combination of electroporation components for the best possible outcome. Limited time also had us consider the evaluation of protein expression through fluorescence as adequate. If desired, the analysis may be taken further and the Crgfp transcript may be analysed by RT-qPCR and the CrGFP protein by SDS-PAGE, Western blot and purified by the already fused histidine tag for ELISA protein purification to quantify the total soluble protein produced.

The NEPA21 electroporator should be considered for future electroporation events. In a recent study with C. reinhardtii strains, a higher transformation efficiency and lower cell damage has been observed with NEPA21 compared to the BioRad Gene-Pulser series (Yamano et al., 2013). This electroporator does not require cell walls to be removed, which enables the transformation of mutant-free more robust strains. No cell wall preparation would be required, and time would be saved. Large DNA fragments up to approximately 7,800 bp have been successfully transformed into the C. reinhardtii strains using this method (Yamano et al., 2013). Such large transgene insertions have also allowed for multigene co-expression within the C. reinhardtii nuclear region (Rasala et al., 2014) and may hold promise for the co-expression of an entire biosynthetic pathway or multiple components of a complex protein (Collin et al., 2019).

2.5 Conclusion The Crgfp gene was cloned, and integrated, into the pChlamy4 expression vector. Transformants were obtained in the nuclear genome of both CC-125 and CC-400 strains of C. reinhardtii and the gene expression was further analysed by fluorescence spectrophotometry and microscopy. The successful expression may be attributed to the pChlamy4 molecular toolkit designed to enhance expression within microalgae, the codon optimised Crgfp gene, the optimised electro-parameters and the physiological state of the cells before and after transformation. The integrated Crgfp transgene was not lost from the cultures and transformation was considered stable. Electroporation transformation has proven successful in both cell wall and cell wall deficient C. reinhardtii strains. The CC-400 strain had a faster growth rate in liquid culture, was more sensitive to zeocin (reducing the cost of culturing) and had higher percentage of positive gene transformants. Therefore, the CC- 400 under our optimised electro-parameters has the potential to produce CrGFP tagged recombinant proteins of value to the biotechnology industry. In this chapter a foundation was laid for microalgae biotechnology research at the CSIR in South Africa, with C. reinhardtii as the reference organism with potential of protocol transferal to South African isolates.

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Stevens DR, Purton S, Rochaix JD, 1996. The bacterial phleomycin resistance geneble as a dominant selectable marker in Chlamydomonas. Molecular and General Genetics 251, 23-30. Stevenson J, Brown AJ, 2015. Universal CG cloning of polymerase chain reaction products. Analytical Biochemistry 471, 80-2. Sun J, Hopkins RC, Jenney Jr FE, Mcternan PM, Adams MW, 2010. Heterologous expression and maturation of an NADP-dependent [NiFe]-hydrogenase: a key enzyme in biofuel production. PloS One 5, e10526. Sustarsic M, Plochowietz A, Aigrain L, Yuzenkova Y, Zenkin N, Kapanidis A, 2014. Optimized delivery of fluorescently labeled proteins in live bacteria using electroporation. Histochemistry and Cell Biology 142, 113-24. Taghavi N, Robinson G, 2016. Improving the optimum yield and growth of Chlamydomonas reinhardtii CC125 and CW15 using various carbon sources and growth regimes. African Journal of Biotechnology 15, 1083-100. Tamura K SG, Peterson D, Filipski A, Kumar S, 2013. MEGA6: Molecular evolutionary genetics analysis version 6.0. Molecular Biology and Evolution 30, 2725-9. Tjernberg A, Markova N, Griffiths WJ, Hallén D, 2006. DMSO-related effects in protein characterization. Journal of Biomolecular Screening 11, 131-7. Werner TP, Amrhein N and Freimoser FM, 2007. Inorganic polyphosphate occurs in the cell wall of Chlamydomonas reinhardtii and accumulates during cytokinesis. BMC Plant Biology 7, 51. Wittkopp TM, 2018. Nuclear Transformation of Chlamydomonas reinhardtii by electroporation. The Salk Institute for Biological Studies. https://bio-protocol.org/pdf/bio-protocol2837.pdf. Wurch T, Lestienne F, Pauwels P, 2000. Optimisation of overlap extension PCR-based mutagenesis of a GC-rich DNA template: Application to the human α2C-adrenoceptor cDNA. Biotechnology Letters 22, 1603-9. Yamano T, Iguchi H, Fukuzawa H, 2013. Rapid transformation of Chlamydomonas reinhardtii without cell-wall removal. Journal of Bioscience and Bioengineering 115, 691-4.

Chapter 3 Identification and growth of South African microalgae

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Abstract South Africa is a biodiversity hotspot with many endemic species being associated within localised biomes. South Africa has higher temperatures and accessible solar energy than many European and North American countries whom are currently cultivating microalgae. There is a possibility that novel indigenous South African algae may contain strains that are more capable of rapid growth and solar- energy conversion than the biotechnology microalgal model, Chlamydomonas reinhardtii. The aim of this study was to select South African microalgal isolates based on their growth potential. Forty unicellular microalgae with many or large chloroplasts and a thin cell wall were selected from the microalgal culture collection of South Africa (MiCCSA), and these underwent growth analysis. The nine best growing isolates were then directly compared to C. reinhardtii CC-400 and CC-125 under phototrophic and mixotrophic growth conditions. These nine isolates were then taxonomically identified by 18S rDNA Sanger sequencing as Tetradesmus obliquus, Tetradesmus dimorphus, Chlorella sorokiniana, Chlorella sp., Chlorella vulgaris, and Raphidocelis subcapitata species. Contamination during culturing was reduced, and the microalgae C. reinhardtii CC-125, T. obliquus, T. dimorphus, Chlorella sp., C. sorokiniana and C. vulgaris were successfully cryopreserved at -80˚C for long term storage.

Keywords: South African microalgae, MiCCSA, growth rates, biomass, Sanger sequencing, light, AF6 media, TAP media, antibiotics, cryopreservation

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3.1 Introduction Production of heterologous proteins from microalgae may be dependent on the microalgal biomass accumulation rate if the impact of the metabolic burden is low. Optimizing resource inputs and maintaining high productivity are key components to control the quantity and cost of microalgae production (Jia et al., 2015). Microalgae which can produce high cell densities on minimal media under simple culture conditions may minimize the space required to grow, may be easily upscaled and would therefore have great economic value in the heterologous protein expression industry (Wan et al., 2011). The accurate determination of microalgal growth in culture is critical to many downstream processes and should be performed for new strains prior to any genetic engineering experiments.

An important factor in the analysis of microalgae biomass is to relate the growth in terms of measurable parameters, such as concentration and optical density (OD) of the cultures. These biological variables of absorbance can be easily and continuously monitored through spectrophotometry (Govindarajan et al., 2010, Jia et al., 2015). In closed systems such as with batch cultures, microalgal growth may be summarised into five different phases, namely: (1) the lag phase, (2) the exponential/ log phase, (3) the declining growth phase, (4) the stationary phase and (5) the death phase (Figure 3.1).

Figure 3.1: The five phases associated with the growth of a closed system microalgal culture. Adapted from (Levine, 2018).

The lag phase is the physiological adaptation phase of the cells to the current environment, so that the microalgal metabolism would be able to support growth. This includes an increase of the levels of enzymes and metabolites required for cell division and carbon fixation (Morris & Glover, 1974). Once the microalgal cells have adapted to utilise the available nutrients and light for cellular division, the cells enter the exponential phase and start to multiply rapidly (Andersen, 2005). This phase may depend on the microalgal growth rate capacity, inoculum size and the medium and culturing conditions (Khona et al., 2016). Microalgal growth capacity is the maximum biomass the microalgae may reach under the specified culture conditions (Khona et al., 2016). Whereas the inoculum size affects the exponential phase, since the growth rate is constant and the more culture there is to start with, the more biomass will accumulate within a shorter period (Andersen, 2005). The decline phase occurs after the culture experiences a limiting factor (nutrient, light) or a toxic factor build up; then the cellular division of the microalgae slows down (Heijerick et al., 2002). The stationary phase is when the microalgal culture has reached the maximum carrying capacity under the current cultivation conditions. The specific death rate and growth rate are balanced and the cell density plateaus (Velichkova et al., 2013). Once the water quality degrades, nutrients diminish, and the

Chapter 3 culture conditions are unable to sustain the microalgae, then the culture will eventually collapse. This is known as the death phase and in laboratory environments it is important to realise that culture collapse may also occur due to overheating, a pH imbalance or a biotic contamination (Han et al., 2012).- crashes can al

For biotechnology purposes, cultures should be grown until their maximum density in the culture, before the declining growth stage, then removed if working in batch culture systems, in order to prevent toxic metabolite build up or an undesirable change in the microalga’s biochemical composition (Barsanti & Gualtieri, 2014). Many factors can influence the specific growth rate which may lead to greater biomass production, the most important culturing conditions include light exposure and the culture medium (Blair et al., 2014).

It is the knowledge of the individual cell cycle, and how it would react to a variety of nutrients, which may influence what medium it would be most optimal on. Media which support the growth of freshwater microalgae include AF6 (Watanabe, 2000), Tris-acetate-phosphate (TAP) (Merchant et al., 2006), BG-11 (Rippka et al., 1979), Allen (Allen, 1968), Bristole/ Bold’s Basal Medium (Bold, 1949), Chu’s medium (Stein et al., 1973), Waris medium (Waris, 1953) and Fitzgerald medium (Liu &

Bangert, 2015). Microalgal biomass has a typical biochemical ratio of C106:N16:P1 (Fields et al., 2014), and while each media is different, it is essential that these components, amongst other micronutrients and metals, are obtained for algal growth. The most ideal media would be that which allows the highest growth rate for the microalgal isolates while containing the fewest constituents to reduce the cost of large-scale production.

Most media are designed for specific microalgal strains and would not work for a wide range of microalgae or are too expensive (Liu & Bangert, 2015). The AF6 medium was readily used at the CSIR; it is a general freshwater medium that supports the growth of a wide diversity of photoautotrophic algae (Birungi & Chirwa, 2014). In this study, the microalgae examined were isolated across different regions of South Africa, the microalgae had not been previously characterised and a media which can support the growth of a large diversity of species was necessary (Gupta et al., 2015).

Microalgae are generally photoautotrophic which means they utilise light energy and water to fix

CO2 as its carbon source (Suzuki & Johnson, 2001). However, some microalgae may utilise organic compounds, such as glucose, acetate, methanol or glycerol, as carbon sources instead (Morales- Sánchez et al., 2015, Khan et al., 2016). Although this cultivation would be simple to scale up with greater biomass production, it is also expensive (Wan et al., 2011), and prone to contamination (Gao et al., 2019). With the abundance of solar energy in South Africa, photoautotrophic microalgae would provide the most benefit.

Alternatively, mixotrophic microalgal growth may combine the advantages of both phototrophic and heterotrophic strategies by using solar energy as well as an organic carbon source (Wan et al., 2011). Mixotrophic conditions are predicted to shorten the growth cycle of microalgae and increase their final biomass yield (Wan et al., 2011). One of the most well-known and inexpensive mixotrophic media is TAP media. Its carbon source is acetate -which is easily diffused into the microalgal cell cytosol- and does not require large transporters such as the transport of glucose or sucrose into the cell (Wan et al., 2011). Acetate was also found to be the best carbon source for mixotrophic growth of many microalgae species (Gao et al., 2019, Taghavi & Robinson, 2016).

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Besides the carbon source, there are many other differences between AF6 and TAP medium: they have different sources of nitrogen with predominantly stable nitrate in AF6 and less stable ammonium in TAP medium, their buffers differ with 2-(N-morpholino) ethanesulfonic acid (MES) in AF6 and Tris in TAP, as well as their quantity of macronutrients and microelements. AF6 media has a lower concentration of each main nutrient component than TAP (Figure 3.2) which is considered more expensive. The TAP media also contains up to two-fold more trace metals but AF6 contains vitamins which TAP lacks.

7 7

3 2.7 AF6 Medium TAP medium 2

Concentration Final in Medium (mM) 1 0.83 0.45 0.29 0.012 0.10 0.07 0 Nitrogen Magnesium/ Sulphur Potassium/ Calcium/ Chlorine Phosphorus Nutrient Component

Figure 3.2: The comparison of nitrogen, magnesium, sulphur, phosphorous, chlorine and calcium nutrient concentrations levels utilised in AF6 and TAP medium.

Optimal nutrient concentrations and growth conditions for microalgal biomass production are often species-dependent (Gao et al., 2019). The Chlorophyta are the most diverse group of algae with over 8,000 species worldwide (Gerber et al., 2006). Globally, South Africa boasts as one of the most bio- diverse regions, with tremendous freshwater species richness; yet only 34 known South African freshwater species have been taxonomically identified (Gerber et al., 2006). Currently there is a Microalgal Culture Collection of South Africa (MiCCSA) in association with the CSIR, Evolutionary Studies Institute at the Wits University and South African National Phycology Culture Collection (SANPCC) consisting of approximately 500 microalgae isolated from freshwater sources across South Africa. Although some of these microalgae have been described by the Institutes mentioned above, the majority are unidentified with uncharacterised growth capacity and biochemical profiles.

The classification of algae has developed over the years from microscopic techniques, to a physiological approach and now comprises of many analytical methods. Originally green algae were identified based on their morphology, but their ultrastructural characteristics are difficult to differentiate (Champenois et al., 2015). Many algal isolates, such as those from the Chlorella genus, can be characterised and identified according to their physiology (Krauss and Shihira, 2010). Some

Chapter 3 species have a functional profile with a distinct response to varying growth conditions. However, it is difficult to duplicate the exact conditions within a laboratory and inevitably there would be slight differentiations in humidity, temperature, light and medium composition which could alter the algal response and lead to incorrect identification (Krauss and Shihira, 2010).

A more accurate and sensitive method of comparison to identify microalgae is genetic sequence analysis which can directly compare sequences at the base pair level (Lin et al., 2012). Furthermore, if sequences do not match any existing species, they may be recorded and used as a future barcode reference. The sequence data used for identification and barcoding must include a DNA region, which has been genetically conserved. Conserved sequences often used are the small and large subunit ribosomal RNA (rRNA) genes as well as the internal transcribed spacer (ITS) of ribosomal DNA (Champenois et al., 2015). The most extensively used molecular marker for algae identification has been proven to be the 18s rRNA gene (Zhang et al., 2014, Ali et al., 2001). The 18S ribosomal RNA is a component of the 40S small eukaryotic ribosomal subunit, one of the basic components of all eukaryotic cells which makes it an important marker in phylogenetic studies and in environmental biodiversity screening (Ali et al., 2001).

It is difficult to simulate all the growth variables of the ecological niche from which a species has evolved. Therefore, those species which can increase their biomass exponentially under easily reproducible growth conditions would be considered favourable for biotechnology applications. In this Chapter, the aim was to identify South African microalgae with the greatest growth rates for future heterologous protein production potential. Forty microalgal isolates obtained from the MiCCSA database were screened considering their morphology and growth performance. Isolates with the highest growth rates were identified using genetic sequencing and their growth further characterised under mixotrophic conditions with the inclusion of antibiotics. Finally, selected isolates were cryopreserved, and all microalgal characteristics documented with their potential for biotechnological applications taken into consideration.

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3.2 Materials and Methods

Microalgae culture growth and maintenance For the duration of the study, microalgal cultures were routinely sub-cultured in a laminar flow cabinet under sterile conditions. Inoculum (1%) from a stationary phase culture was transferred by sterilized micropipettes to a prelabelled Erlenmeyer flask containing 200 mL fresh, sterile AF6 (Watanabe, 2000)/ TAP (Merchant et al., 2007) media. The cultures were grown under optimal conditions to the late log-phase, shaking in the Titramax shaker (Heidolph, Germany) at 135 rpm, at a temperature of 25°C with 250 μmol m-2.s- 1 of continuous ﬂuorescent white light. After the cultures reached stationary phase, they were then maintained under sub-optimal conditions and shaken to prevent photo-oxidative stress (Andersen, 2005) using a Titramax shaker set to 110 rpm, at a temperature of 23°C with 100 μmol m-2.s- 1 of continuous ﬂuorescent white light. Light intensity was measured using a calibrated Lux Meter Android Application (2018) on a Samsung S6 handheld device. The lux was then converted to photosynthetic photon flux density (μmol m-2.s- 1) using the conversion factor of 0.0135, supplied for cool white fluorescent lamps (Thimijan & Heins, 1983).

3.2.1 South African microalgal isolates The MiCCSA culture collection held at the CSIR, contains microalgal isolates collected across South Africa (Figure 3.3), cultured on AF6 agar slants with a matching microscopic picture record for each isolate. From this record, 40 algal isolates were selected for growth curve analysis. These green microalgae were chosen since the cells were unicellular, non-colonial, with large or many chloroplasts and a thin cell wall. Compound light microscopy, utilising the Olympus BX40 microscope (Olympus, Massachusetts, United States), confirmed the microalgal isolates were those previously recorded, and the algal isolates were identified according to their morphological characteristics. The classifications were performed according to the freshwater algae guides: The Freshwater Algae (Prescott, 1954); Freshwater Algae: Identification and Use as Bioindicators (Bellinger and Sigee, 2015) and, A Guide for the Identification of Microscopic Algae in South African Freshwaters (Gerber et al., 2006).

Figure 3.3: The respective locations of over 500 Indigenous freshwater microalgae isolated across South Africa as part of the MiCCSA database. The provinces are abbreviated as, NC= Northern Cape, WC= Western Cape, EC= Eastern Cape, NW = North West, FSA = Free State region A, FSB= Free State region B, GPA= Gauteng, KZN= KwaZulu Natal, MP= Mpumalanga and LPB= Limpopo. Technology Innovation Agency project number available upon request.

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3.2.2 Microalgae growth analysis Growth curve analysis of selected microalgal cultures were established from each microalgal isolate slant to ensure that subsequent growth analysis inoculations would be with a uniform amount. A loop of each culture was inoculated into 10 mL of AF6 media contained in a McCartney bottle and left to grow for 10 consecutive days on a Titramax shaker at 135 rpm, at a temperature of 25°C with 250 μmol m-2.s- 1 of continuous ﬂuorescent white light.

After uniform monoseptic cultures were established, the cultures were centrifuged for 10 min at

5,000 × g then the cultures were diluted accordingly for an inoculation of approximately 0.1 at OD750 to ensure equivalent growth comparisons among the different cultures. The wells of each clear six well Costar® microplate (Corning, New York, United States) were filled with 5 ml liquid AF6 medium, whereafter 500 μl starter culture was inoculated into each well (Figure 3.4), to obtain three biological replicates for each isolate. The plates were placed onto a Titramax shaker set as shaking at 200 rpm, at a temperature of 25°C and 250 μmol m.-2s- 1 of continuous fluorescent white light. The experiment was repeated three more times with the algal isolates under: 1) a lower intensity of 100 μmol m-2.s-1 continuous fluorescent white light, 2) a light intensity of 250 μmol m-2.s-1 of continuous fluorescent white light but with a stock culture inoculum increase of 20%; and 3) 20% algal inoculum under 250 μmol m-2.s-1 of 8 hour light: 16 hour dark phase cycles.

Figure 3.4: Schematic diagram of the six well microplate containing three biological replicates of two algal strains used for microplate growth study experiments. A total of 21 plates were used to assess the growth of 40 selected South African microalgae and two C. reinhardtii strains.

The optimal growth conditions were then repeated for the nine South African microalgal isolates which grew the best, as well as the C. reinhardtii CC-125 and CC-400 strains (acclimatised to AF6). These selected South African microalgal isolates were then acclimatised from AF6 to TAP medium with revised trace elements (Merchant et al., 2006) and grown under the same culture conditions. To minimise contamination and prevent intra-well mixing, the plates were covered by NUNC plastic cover slips with punctured holes to allow for gas transfer. Evaporation loss was prevented by having the initial media volume marked on the side and lost water was replaced with ddH2O as needed. The microplates were distributed randomly throughout the incubator to prevent environment bias skewing the results.

All the cultures were monitored through photographic observations and the OD750 was measured once a day for 12 days. A volume of 100 μl of each replicate was aliquoted into a 96-well microplate (Thermo Fisher Scientific, South Africa) and the BIOTEK power XS multi-well plate reader (ADP Weltevreden Park, South Africa) using the Gen 5 1.06 program to read the OD of each well at a wavelength of 750 nm (corresponding to the microalgae biomass detection).

The culture density was also measured by counting the cells on a 4x4 grid algal hemocytometer (Thermo Fisher Scientific, Massachusetts, United States) under the Olympus BX40 microscope. Each

Chapter 3 microalgal culture was diluted, and the cells counted at OD750 when the absorbance vales were (0.2, 0.4, 0.6, 0.8, 1, 1.2, 1.4) to create a calibration curve. Microsoft Excel v1808 was used to calculate the number of cells based on dilution factor and physical parameters of the hemocytometer.

The specific growth rates (µmax) for each microalgal isolate was calculated from the exponential slope of the sigmoid plot based on the raw data (Hall et al., 2013).

푙푛 ( 푁 −푁 ) Specific growth rate: µ = 1 0 (푡1−푡0)

푁1 = Cell density at a point towards the end of the isolate’s exponential phase 푁0 = Cell density at the start of the isolate’s exponential phase 푡1 = Day 푁1 was taken 푡0 = Day 푁0 was taken

3.2.3 Biomass comparison of South African microalgal cultures To estimate biomass production, the dry weight (mg/mL) of the nine South African microalgal isolates as well as the C. reinhardtii CC-125 and CC-400 strains was also measured. The microalgae were cultured in 100 ml of either TAP or AF6 medium in Erlenmeyer flasks on a Titramax shaker set to 135 rpm, at a temperature of 25°C with 250 μmol m-2.s- 1 of continuous ﬂuorescent white light. All Erlenmeyer flasks in this study were covered with a permeable filter paper to prevent contamination whilst allowing for gaseous exchange. After 14 days, 15 ml of the culture was aliquoted into a pre- weighed conical tube and centrifuged for 10 min at 5,000 × g. The supernatant was discarded, and the cell pellet dried overnight to a constant weight in an EcoTherm Economy incubation oven (Labotec, South Africa) set to 40˚C. The weight of the conical tube with the dried algae culture was recorded and the biomass was calculated (Blair et al., 2014).

1000 Microalgal biomass (mg): 푤 = (푤 − 푤 ) 푥 1 2 푉

푤1 = weight of empty conical tube (mg) 푤2 = weight of dried microalgae and the tube (mg) 푉 = culture volume measures (mL)

Microalgae growth data was collected and analysed by variance analysis with a one-way ANOVA performed using Microsoft excel software v1808. All samples had a minimum of three biological replicates.

3.2.4 Classification of Algae by 18S rRNA DNA Sequencing Genomic DNA was extracted from nine selected South African microalgal isolates. The Invisorb Spin Tissue Kit (DSTR1032100100S, Stratec, Germany) was used per the manual’s instruction with the modification of bead beating during the lysing step to break down the glycoprotein cell walls. The microalgae suspensions were centrifuged at 10,000 x g for 10 min through a spin filter column and after the final elution step, the tubes were left open on the bench for one hour to allow all ethanol to evaporate.

Chapter 3

The 18S rDNA region of the selected microalgal isolates, were amplified for subsequent Sanger sequencing. The 18s rRNA primer pair sequences used in this study (Table 3.1) (Duong et al., 2015) were synthesised by Inqaba Biotec, South Africa. The PCR reactions contained 0.5 μM forward and reverse primers, 20 ng genomic DNA, 1x Ampliqon Taq DNA Polymerase (Ampliqon, Denmark) and ddH2O to a final volume of 50 µl. The PCR cycling conditions were as follows: denaturation at 94°C for 5 min, 35 amplification cycles (94°C for 30 s, 55°C for 30 s and 72°C for 1 min) and a final extension step at 72°C for 10 min.

Table 3.1: The 18S rRNA primers used in this study for the classification of microalgae.

Primer name Primer sequence (5’-3’) Length GC content Annealing (bp) (%) temperature (°C) M18S-rRNA- For GCGGTAATTCCAGCTCCAATAGC 23 52.2 60.5 M18S-rRNA-Rev GACCATACTCCCCCCGGAACC 21 67.7 64.9

PCR products were visualised following electrophoresis through 1.5% agarose sodium borate gels. Gels were prepared with Seakem® LE Agarose (Lonza Rockland, USA) and 1x Sodium Borate buffer (with 0.2 µg/ml EtBr). The gels were electrophorized at 150 V for 15 min, in 1x Sodium Borate buffer (Sigma Aldrich, Missouri, United States). For DNA band size determinations, the PCR product extraction samples were run alongside a Fast DNA ladder (New England Biolabs, Massachusetts, United States). DNA on the agarose gels were viewed using the Sigma UV-transilluminator (Sigma Aldrich, Missouri, United States).

The BigDye sequencing reaction was set up as follows: 400 ng of the PCR product, 1x Sequence Buffer (Thermo Fisher Scientific, Massachusetts, United States), 10 μM M18S-rRNA forward primer with ddH2O to a final reaction volume of 10 µl. The PCR cycling conditions were as follows: denaturation at 94°C for 5 min, 25 amplification cycles (94°C for 30 s, 55°C for 10 s and 60°C for 4 min) and a final extension step at 72°C for 10 min. The PCR products were purified by Sephadex G50 columns (Etchevers, 2007) and eluted with ddH2O into a 0.2 mL Eppendorf tube and sent to the ABIsystems DNA Sanger Sequencing Facility at the University of Pretoria, South Africa.

The DNA sequencing data was analysed and edited using BioEdit v7.0.5, sequence alignment editor software (http://mbio.ncsu.edu/BioEdit/bioedit.html) and compared using BLASTn searches (Karlin & Altschul, 1990) on the National Centre for Biotechnology Information (NCBI) (https://blast.ncbi.nlm.nih.gov). Sequence alignments were made online with MAFFT version 7 (https://mafft.cbrc.jp/alignment/server) and further edited with BioEdit v7.0.5.

The evolutionary history of each microalgal isolate was inferred by the maximum likelihood method (Tamura & Nei, 1993) using MEGA6 software (Tamura K, 2013) to calculate the pairwise genetic distances for all the species based on their 18S rDNA sequence. For comparative analysis, sequences with the same genera as isolates from this study were downloaded from GenBank, with their accession numbers shown in Table 3.2. These sequences were all freshwater green microalgae (Chlorophyceae), therefore the marine green algae, Ulva prolifera (Ulvophyceae) was used as an outlier to root the phylogram.

Chapter 3

Table 3.2: List of additional microalgae GenBank accession numbers used in this study to construct the maximum likelihood phylogram for 18S rDNA comparison.

Taxon 18S rRNA accession number C. reinhardtii FR865562.1 Tetradesmus dimorphus MK041923.1 Scenedesmus bijugus KX495068.1 Ulva prolifera HQ850569

3.2.5 Controlling contamination in microalgae cultures After prolonged culture incubation (over a period of 6-18 months) fungal and bacterial contamination was observed on the TAP medium cultures. A pilot study was undertaken to find the source of the contamination, control it and obtain cultures free from obvious contamination once more.

To test for airborne contamination within the laboratory, solid TAP 15% agar, was made in petri- plates and exposed to the laboratory room at intervals of 1 minute, 5 minutes, 15 minutes and 1 hour. These dishes were then incubated at 22 °C with 100 μmol m-2.s-1 of continuous ﬂuorescent white light and contamination visually observed after 7 days.

To eliminate bacterial and fungal contamination an antibiotic-antimycotic solution was created. The solution contained carbenicillin (500 mg/L), kanamycin (200 mg/L) and carbendazim (40 μg/mL) used previously for microalgal cultures (Guillard, 2005, Mahan et al., 2005). The mixture was filter sterilized with a 0.22 Millex micron filter (Merck, New Jersey, United States) and spread over solid TAP 15% agar plates. Microalgae grown on TAP media without the addition of any antibiotics was used a control. The 300 µL of contaminated C. reinhardtii CC-125 and CC-400 liquid cultures were spread across these plates and incubated at 22 °C with 100 μmol m.-2.s-1 of continuous ﬂuorescent white light. Single microalgal colonies were streaked onto TAP antibiotic-antimycotic plates and then streaked onto normal TAP media. From this a single cell was used to inoculate the liquid TAP, culture media for stock cultures.

3.2.6 Microalgae growth analysis with zeocin The natural resistance of algal isolates (NWA 23.4, FSA 22.3, LPB 14.1, GPA 9.3, GPA 9.1, WC 4.1, NC 72) to the antibiotic, zeocin was tested. The antibiotic kill-curve also allowed the optimal zeocin concentration to be determined for future selection in transformation studies. The algal isolates, 100 μl, were cultured in triplicate in the 6-well 5 ml microplates on solid TAP 15% agar supplemented with the zeocin antibiotic present in the following concentrations: 0, 3, 5 and 10 mg/L. Four evenly spaced 1 mm apertures in the plastic lid allowed for gaseous exchange whilst reducing contamination. The algal isolates were left to cultivate for 14 consecutive days at a temperature of 25 °C with 150 μmol m-2.s- 1 of continuous ﬂuorescent white light. Visual scoring was used to analyse the microalgal isolate response to zeocin at each concentration.

Chapter 3

3.2.7 Microalgae cryopreservation The GeneArt® Cryopreservation Kit for Algae (Thermo Fischer Scientific, Massachusetts, United States) was used to cryopreserve the microalgal isolates used in this study: FSA 22.3, GPA 18.1, NC 72, LPB 14.1, GPA 9.3, WC 4.1, GPA 9.1, GPA 19.1, NWA 23.4, as well as the C. reinhardtii untransformed and CrGFP transformed CC-125 and CC-400 strains (Chapter 2); with a few modifications.

The cells were inoculated into 20 ml TAP medium supplemented with 500 µL of Cryopreservation Reagent A and grown with shaking in the Titramax shaker at 135 rpm, at a temperature of 25°C with 250 μmol m-2.s- 1 of continuous ﬂuorescent white light. When the cells reached a density of 1 x 106 cells/mL, they were harvested by centrifugation at 2,500 x g for 5 minutes and the supernatant was removed. The cells were resuspended in Cryopreservation Reagent B to a final concentration of 2.5 × 107 cells/mL and left to incubate for 30 minutes at room temperature. The cell suspension was aliquoted (240 µL) into 2 ml cryovials and incubated for 30 minutes further. These vials were then placed into a Mr. Frosty® Nalgene freezing container (Thermo Fisher Scientific, Massachusetts, United States) at –80°C for four hours. The microalgae containing cryovials were then removed from the Mr. Frosty® freezing container and placed directly into the -80˚C for permanent storage.

After three weeks of storage, the cryopreserved cells were thawed and rejuvenated as per manufacturer’s instructions with some modifications. The vials were removed and placed on ice for an hour, then the cryovials were placed in a 35°C water bath and swirled for 2 mins for slow thawing. From the cryovial, 230 µL of the thawed cells were aliquoted into sterile MCartney tubes containing 20 mL of room temperature TAP medium. The tubes were incubated on the rotary shaker set to 110 rpm at 25°C with low light, 50 μmol m-2.s- 1, until sufficient microalgal biomass was visible for visual comparison.

Chapter 3

3.3 Results

3.3.1 Compound light microscopy From the MiCCSA database of over 500 isolates, the microalgae cells were distinguished by their size, shape, colour, and presence of chloroplast organelles to select 40 isolates from the database of 500 (Appendix H, Table H1). Microalgae were selected with a thin cell wall, were predominantly green, small and unicellular. Microalgal isolates which were excluded were filamentous, had a thick gelatinous layer surrounding the cells, aggregated in clumps and contained large vacuoles. The physiological morphology of the identified isolates as well as the limitations in this study of utilising microscopy for microalgae identification is discussed further in Appendix H.

3.3.2 Microalgae growth analysis

In our study, we measured the OD750 of the culture as an in vivo proxy of the cell number, dry weight biomass was also determined at the end of a culture period. When the culture is not limited by light or nutrients, then the OD measurements should give a good representation of exponential growth (Andersen, 2005, Chioccioli et al., 2014). The growth rate and final dry biomass may indicate the productivity potential of the batch culture system for that particular isolate (Rocha et al., 2017, Wood et al., 2005).

Initial experiments utilised 20 isolates with their OD measured over a period of 4-12 days on AF6 media, before the microalgae became photo-inhibited and the experiments were disregarded as unsuccessful (Appendix I, Figures I1, I2 and I3). Thereafter, four trials were performed to optimise the growth parameters of the selected South African microalgal isolates. The first trial included a continuous supply of ﬂuorescent white light at 250 μmol m-2.s- 1. Exponential growth was observed over the first two days, which then plateaued from day 3 till day 5 for most isolates (Appendix I,

Figure I1). The microalgae which grew the best was isolate GPA 18.1, with a maximum OD750 of 0.229 on day 5. The growth analysis experiment was then repeated with a lower intensity of 100 μmol m- 2.s- 1 continuous ﬂuorescent white light. The lower light intensity did not improve the growth curve and by day three most isolates were photo-inhibited once more (Appendix I, Figure I2). The microalga which obtained the maximum OD750 of 0.284 on day 4 was isolate GPA 9.3. The third experiment included the original light intensity of 250 μmol m-2.s- 1 of continuous ﬂuorescent white light but with twice the amount of inoculum with a stock culture at 20% instead of 10%. This increase was motivated by the idea that by having twice as many cells to start off, with the cultures would be able to self-shade and be under less light stress. Over a period of 12 days however, it was clear that most isolates were still photo-inhibited (Appendix I, Figure I3). The isolate GPA 18.1 representing a complete sigmoid growth curve with a maximum OD750 of 0.356 on day 12.

The final growth analysis experiment utilised 20% algal inoculum under 250 μmol m-2.s- 1 of 8 hour light: 16 hour dark phase cycles. These cultures grew well, with a growth curve obtained for most isolates (Appendix I, Figure I4). The experiment was repeated with the 20 additional isolates, (Appendix I, Figure I5), concluding which South African microalgal isolates had high growth rates and high biomass potential within the 12-day period. These included isolates from the provinces, Free State (FSA 22.3, FSB 18.1), Gauteng (GPA 9.1, GPA 19.1, GPA 18.1), Northern Cape (NC 72), Limpopo (LPB 14.1) and the Western Cape (WC 4.1). The visual inspection of the 40 microalgal stock cultures grown in AF6 medium in Erlenmeyer flasks validates this choice as the first nine microalgae which had the best growth were the most green and healthy (Figure 3.5).

Chapter 3

Figure 3.5: Stock cultures of the selected 40 South African microalgae isolates grown on AF6 media after four weeks under 250 μmol m-2.s- 1 of continuous light. The first nine microalgae were the isolates identified as those which grew the best; these cultures were far denser and healthier than the rest of the microalgal cultures.

These final nine selected microalgal isolates were re-cultured under the same conditions on AF6 medium with C. reinhardtii strains CC-125 and CC-400 included in the growth study (Figure 3.6). Most of the strains showed a similar growth curve progression with approximately 2/3 days of lag phase and 5/8 days of exponential phase, followed by a declining phase or stationary phase for the remainder of the period. The CC-125 strain had poor growth, only surpassing the growth of isolate NWA 23.4 and had a longer lag, and shorter exponential phase compared to the rest of the isolates. The CC-400 experienced a faster specific growth rate than CC-125 but most South African microalgal isolates outcompeted both the C. reinhardtii control strains on AF6 medium.

1,2 Chlorella sp. FSA 22.3

1 Chlorella sorokiniana GPA 18.1 Chlorella vulgaris NC 72 0,8 Tetradesmus obliquus LPB 14.1 Chlorella sorokiniana GPA 0,6 9.3

Chlorella sp. WC 4.1 OD OD nm) (750 0,4 Tetradesmus obliquus GPA 9.1 Tetradesmus dimorphus 0,2 GPA 19.1 Raphidocelis subcapitata NWA 23.4 Chlamydomonas reinhardtii 0 CC125 0 2 4 6 8 10 12 Chlamydomonas reinhardtii Time (Days) CC400

Figure 3.6: Growth curve analysis of 11 microalgal isolates with 8-hour light (250 μmol m-2.s-1)/ 16-hour dark phase cycles in AF6 medium. The cells were cultured in AF6 media and the OD750 nm was measured every day for 12 days. The microalgae which grew the best were the isolates NC 72, WC 4.1, WC 1.1 and FSA 22.3. The values represent the mean ± standard deviation (n = 3).

Chapter 3

The 11 microalgae were subsequently cultivated under the same culture conditions utilising TAP medium in place of AF6. On the TAP medium, most of the microalgal isolates grew much faster with a greater and longer exponential phase (Figure 3.7). Here CC-125 grew far better than on the AF6 medium, but it still did not surpass the growth of most South African isolates. A shorter lag phase with longer exponential phase was seen for most isolates. Both GPA 9.3 and GPA 18.1 having not reached stationary phase during this growth period. The visual comparison of microalgal stock cultures grown on TAP and AF6 media also indicate greener, more dense cultures for TAP media (Appendix I, Figure I6).

1,2 Chlorella sp. FSA 22.3

Chlorella sorokiniana GPA 18.1 1 Chlorella vulgaris NC 72

Tetradesmus obliquus LPB 0,8 14.1 Chlorella sorokiniana GPA 9.3 0,6 Chlorella sp. WC 4.1

Tetradesmus obliquus GPA

OD OD nm) (750 9.1 0,4 Tetradesmus dimorphus GPA 19.1 Raphidocelis subcapitata 0,2 NWA 23.4 Chlamydomonas reinhardtii CC125 Chlamydomonas reinhardtii 0 CC400 0 2 4 6 8 10 12 Time (Days)

Figure 3.7: Growth curve analysis of 11 microalgal isolates with 8-hour light (250 μmol m-2.s-1)/ 16 hour dark phase cycles in TAP medium. The cells were cultured in TAP media and the OD750 nm was measured every day for 12 days. The microalgae which grew the best were the isolates NC 72, WC 4.1, WC 1.1 and FSA 22.3. The values represent the mean ± standard deviation (n = 3).

The growth curves of microalgae on AF6 medium (Figure 3.6) appeared more scattered than those of TAP medium (Figure 3.7). A correlation curve between the cell counts and OD dilution measurements for each microalgal isolate (Appendix I, Figure I7), indicated a good linear regression trend. From this graph, the expected cell number for each isolate was estimated for the maximum cell density each culture reached (Table 3.3). The specific growth rate and maximum OD750 was also calculated for each isolate (Table 3.3).

−1 On AF6 media the fastest growth rate was for WC 4.1 with 0.26 day , with a maximum OD750nm of 0.74 on day 12, and cell density estimated from the calibration curve to be 7.5 x 106 cells/mL. On TAP media, the isolate with the highest growth rate was GPA 18.1 with 0.36 day−1, with a maximum 6 OD750 of 1.02 on day 12, estimated by the calibration curve to be 13 x 10 cells/mL. Here the maximum biomass concentration represents the carrying capacity of that microalgal strain under the current culture conditions.

Chapter 3

Table 3.3: The growth rate, maximum OD cell density and estimated cell concentration of each microalgal isolate grown on either AF6 or TAP media.

AF6 medium TAP medium Isolate Species Growth rate (day−1) Maximum Estimated cell Growth rate (day−1) Maximum Estimated cell biomass concentration biomass concentration concentration (OD (cells/mL x 106) concentration (OD (cells/mL x 106) 750 nm) 750 nm) GPA 9.3 C. sorokiniana 0.24 0.62 ± 0.01 6.7 0.35 1.01 ± 0.03 13.1 GPA 18.1 C. sorokiniana 0.23 0.55 ± 0.01 6.2 0.36 1.02 ± 0.02 13.2 NC 72 C. vulgaris 0.25 0.71 ± 0.06 7.5 0.32 0.99 ± 0.003 12.3 WC 4.1 Chlorella sp. 0.26 0.74 ± 0.05 7.5 0.34 0.98 ± 0.01 10.7 LPB 14.1 T. obliquus 0.22 0.65 ± 0.02 2.6 0.24 0.69 ± 0.01 2.9 FSA 22.3 Chlorella sp. 0.21 0.67 ± 0.05 8.9 0.31 0.92 ± 0.03 11.3 GPA 9.1 T. obliquus 0.21 0.61 ± 0.05 3.2 0.23 0.65 ± 0.01 3.4 CC400 C. reinhardtii 0.18 0.46 ± 0,01 3.4 0.25 0.64 ± 0.03 3.9 GPA 19.1 T. dimorphus 0.15 0.62 ± 0.04 2.1 0.21 0.65 ± 0.04 2.0 CC125 C. reinhardtii 0.13 0.28 ± 0.04 3.2 0.21 0.49 ± 0.03 3.4 NWA 23.4 R. subcapitata 0.09 0.39 ±0.01 0.8 0.19 0.43 ± 0.04 1.2

An important parameter in algal culture is the direct biomass production of a microalgal strain. Since it is difficult to measure a small volume of cells grown in microwell plates, the dry weight biomass was measured on a larger scale. Each isolate was cultured in an Erlenmeyer flask of either TAP or AF6 media and the dry- weight biomass after 14 days of culturing was calculated per isolate. Most microalgae had a much greater biomass on TAP than AF6 media (Figure 3.8). On the AF6 medium, the highest biomass of 0.95 mg/mL was achieved for the isolate LPB 14.1 and on the TAP medium the highest biomass of 1.298 mg/mL was achieved for the isolate GPA 9.3.

Chapter 3

1400 1200 1000 800 600

400 Biomass(mg/L) AF6 200 TAP 0

Microalgal Isolates

Figure 3.8: The biomass production for each microalgal isolate on AF6 and TAP media after a period of 14 days. The values represent the mean ± standard deviation (n = 3).

At the end of the cultivation period, there was a clear difference in cell density and biomass within each medium for the different isolates. The results revealed that the South African isolates showed good growth potential in both TAP and AF6 media. There was also a discrepancy between microalgae photobleaching when grown within 6 well microplates (Appendix I, Figure I8) compared to Erlenmeyer flasks (Appendix I, Figure I9). This further highlighted that the microalgae responded to light and nutrient changes differently, resulting in isolate specific growth trends and maximum biomass concentration.

3.3.3 Classification of algae by 18S rRNA DNA sequencing The nine South African microalgal isolates with high growth potential (FSA 22.3, GPA 9.3, GPA 9.1, GPA 19.1, FSB 18.1, GPA 18.1, NC 72, WC 4.1 and LPB 14.1) were identified based on Sanger sequencing of the 18SrRNA gene and subsequent comparison with sequences of microalgal isolates in Genbank. Various genomic DNA extraction methods were tested (Appendix J) but the Invisorb Spin Tissue Kit extracted the highest amount of DNA with the best quality.

After microalgal genome DNA was extracted, partial 18S rRNA gene sequences (412–488 bp) were PCR amplified (Appendix J, Figure J1), sequenced and BLASTn aligned to similar sequences on the GenBank Database. Table 3.4 below describes the various identity values along with their statistical measure of identification Expect (E) values. Most of the microalgae had a high sequence similarity >98%, and E value of 0.0 indicating a significant match with the 18S rRNA sequence; this was supported by the microscopic examination, allowing for the species name to be adopted (Yarza et al., 2014). However, isolates GPA 9.1 and GPA 9.3 did not meet this threshold with only a sequence similarity of 87% (E value 1e-150) and 89% (E value 1e-87) respectively. The max score is the score of the sequence that aligns best to the query sequence while the total score is the sum of scores of all aligned sequences (Lipman et al., 1984). These scores are different between the isolates because each query sequence was aligned to a separate database of sequences. In most cases the max score and total score are equal, which occurs when only a single alignment is present.

Chapter 3

Table 3.4: Identification results from the BLASTn (NCBI) analysis for each South African microalgal isolate, based on the 18S rDNA sequence compared with sequences on GenBank. Isolates were assigned species names if sequence similarity with Genbank sequences was higher than 97% and E values greater than 1e-5.

Isolate Description as stated in the BLASTn top Sequence Max score Total score Query E value Identity Genbank number match length cover % accession number GPA 9.3 Chlorella sorokiniana strain LU5 18S 412 544 544 100% 1e-150 89% JQ360515.1 ribosomal RNA gene, partial sequence

GPA 9.1 Tetradesmus obliquus isolate NC-M7 18S 435 334 489 98% 1e-87 87% KY637056.1 ribosomal RNA gene, partial sequence

GPA 18.1 Chlorella sorokiniana 18S rRNA gene, 488 881 881 100% 0.0 100% X73993.1 strain SAG GPA 19.1 Tetradesmus dimorphus gene for 18S 485 870 870 100% 0.0 99% LC192134.1 ribosomal RNA, partial sequence, strain: NIES-93 LPB 14.1 Scenedesmus bijugus 18S ribosomal RNA 483 872 872 100% 0.0 100% MF069190.1 gene, partial sequence

NC 72 Chlorella vulgaris isolate 18s rRNA small 488 881 881 100% 0.0 100% MF686452.1 subunit ribosomal RNA gene, partial sequence

WC 4.1 Auxenochlorella pyrenoidosa strain PT1 485 875 875 100% 0.0 100% KX752082.1 18S ribosomal RNA gene, partial sequence FSA 22.3 Auxenochlorella pyrenoidosa strain PT1 485 870 870 100% 0.0 100% KX752082.1 18S ribosomal RNA gene, partial sequence NWA 23.4 Selenastrum capricornutum small subunit 487 879 879 100% 0.0 100% AF169628.1 ribosomal RNA gene, complete sequence

Chapter 3

Much uncertainty exists around the nomenclature of microalgae. Therefore the most recent taxonomic description for the species indicated on the global algal database, AlgaeBase [www.algaebase.org (Guiry, 2017)] was adopted in this study as their current taxonomic names.

The algal isolate from Gauteng: GPA 9.1 was identified (without certainty based on the E value) as Tetradesmus obliquus (Turpin); GPA 19.1 was identified as Tetradesmus dimorphus (Turpin) strain NIES-93; GPA 9.3 (without certainty based on the E value) and GPA 18.1, were both characterised as the Chlorella sorokiniana strain LU5 and strain SAG. From Limpopo, the algal isolate LPB 14.1 was identified as Scenedesmus bijugus (Turpin); a mislabelled name referring to Scenedesmus bijugatus (Kützing) which is currently regarded as a synonym of Tetradesmus obliquus (Turpin) (Guiry, 2017). From the Northern Cape, the isolate NC 72 was identified as Chlorella vulgaris. From the North West province, the isolate NWA 23.4 was identified as Selenastrum capricornutum which was renamed to Pseudokirchneriella subcapitata (Korshikov) and then later Raphidocelis subcapitata but either of the latter names are accepted in the algal community (Guiry, 2017, Yamagishi et al., 2017). The algal isolate, WC 4.1 from the Western Cape and FSA 22.3 from the Free State were both identified as Auxenochlorella pyrenoidosa strain PT1 18S. There is much confusion over Auxenchlorella spp.; as it was initially described by Chick in 1903 (Guiry, 2017) and in 1965 it was regarded a separate genus from Chlorella spp. due to its thiamine growth requirement (Rodó & Molinari-Novoa, 2017). This taxon is often mentioned but does not have distinguishable morphological characteristics; when the UTEX Culture Collection of Algae (University of Texas) was re-examined, many of the A. pyrendoisa strains belonged to different species of Chlorella, namely: C. vulgaris, C. sorokiniana and C. fusca (Champenois et al., 2015). This suggests A. pyrendoisa refers to strains of uncertain taxonomic status and further investigations are required. In this study, it was postulated that A. pyrenoidosa is not a bona fide species but in fact misidentified C. sorokiniana (Personal communication, Spicer A, 2019). For this study, the isolates were simply labelled as Chlorella sp. to avoid any misidentifications.

The evolutionary histories of these isolates were inferred by using the maximum likelihood method. The tree with the highest log likelihood was adopted with the percentage of trees in which the associated taxa clustered together shown next to the branches (Figure 3.9). The analysis involved 14 18S rDNA nucleotide sequences, including: the nine South African isolates, four related microalgae and one microalga outlier. Relatedness was inferred and with the combined support of our morphological and molecular data, the taxonomic identification to the species level may be considered accurate. There is strong support for the grouping of the Chlorella and Auxenochlorella, the Scenedesmus and Tetradesmus species with the Raphidocelis genus with inferred relatedness between these two clades. The two isolates with higher E values than 0 (GPA 9.1 and GPA 9.3) were in the same clade as their assumed genera but with poor support for LPB 14.1 as T. obliquus when compared to the reference sequence. South African species have not been well characterised and are poorly represented on Genbank, therefore slight DNA variations are possible between different strains of the same genera. However, insufficient information was available to form a reputable conclusion as to species identity.

Chapter 3

Figure 3.9: Maximum likelihood molecular phylogenetic analysis of 18s rRNA green microalgal isolates. Included are sequences of other green microalgae as well as the brown algae outlier, U. prolifera from the GenBank database (indicated with *). The numbers at the nodes indicate bootstrap support as analysed on Mega 6 software. 3.3.4 Controlling contamination in microalgae cultures For the duration of the study it was necessary to maintain healthy, physiologically, morphologically, and genetically representative populations of each microalgal strain. Therefore, the metabolically active microalgal cultures were routinely sub-cultured in a laminar flow under sterile conditions. However, after prolonged culturing on the TAP medium these microalgal cultures became contaminated. Healthy stock cultures would turn from bright green (Figure 3.10A) to a cloudy white- yellow (Figure 3.10B), indicating that the algae were under severe stress. Contamination was prevalent on agar plates (Figure 3.10C) and visualised within our cultures under the Olympus BX40 microscope (Figure 3.10D-F). Contamination must however be distinguished from natural microalgal growth characteristics, such as palmelloids (Appendix I, Figure I11).

Chapter 3

Figure 3.10: Contamination of TAP medium microalgal stock cultures after prolonged sub-culture of 6-18 months. (A) Healthy cultures of CC-125 and CC-400. (B) Unhealthy and contaminated cultures of CC-125 and CC-400. (C) Contamination observed on TAP 15% agar plates when exposed to laboratory air for 5 mins (D) Contamination of CC-125 viewed under the Olympus BX 40 microscope; Scale bar 20 µM. (E) Contamination of CC-125 viewed under the Olympus BX 40 microscope; scale bar 50 µM. (F) Contamination of CC-400 viewed under the Olympus BX 40 microscope; scale bar 50 µM.

As a representative of the working conditions, airborne contamination was tested by exposing the TAP agar plates to the air for certain time intervals on the bench. Numerous round, yellow bacterial colonies as well as white and grey fungi were present after just 5 minutes of exposure (Figure 3.10C) with increasing colonies as exposure time increased and no growth for media unexposed to the air (data not shown). This eliminates any media preparation issues and infers that the environment was heavily contaminated. The fact that similar contamination was seen within the Erlenmeyer flasks (covered with permeable filter paper) handled under sterile conditions in the laminar flow hood but cultured within the laboratory, suggests that either air-borne contaminants smaller than the 0.44 um filter paper were present; or contaminants were introduced during inoculation of media with slow growing bacteria.

To eliminate bacterial and fungal contamination, the microalgae were streaked onto TAP agar plates containing an antibiotic-antimycotic solution. A slight inhibition effect of the microalgal growth on the cocktail of antibiotics compared to the control plates was noted, but under the microscopic inspection, the microalgal cells appeared to not have any physiological or morphological changes. The cells were transferred for single cell isolation and single cells inoculated into liquid TAP medium. This was an appropriate temporary method to eradicate contamination before subsequent experiments were continued. Unfortunately, contamination reoccurred after a period of incubation and this antibiotic-antimycotic procedure had to be repeated every few weeks.

Chapter 3

3.3.5 Microalgae growth analysis with zeocin The microalgal isolates were tested to see if they were susceptible to the antibiotic zeocin for potential future transformation reactions. Natural resistance would prevent positive selection of the transformed microalgae present with the pChlamy4 vector containing the Sh ble gene. A dose- dependent relationship was observed for most of the South African microalgal isolates (Figure 3.11).

Figure 3.11: A zeocin kill curve of the South Africa microalgal isolates GPA 18.1, LPB 14.1, FSA 22.3, WC 4.1, GPA 9.3, GPA 19.1, NWA 23.4 and NC 72 grown in triplicate on TAP 15% agar at concentrations of 0, 3, 5 and 10 mg/L Zeocin antibiotic.

Two South African microalgal isolates FSA 22.3, WC 4.1 had the most prevalent algal growth at 0 mg/L and the most stunted growth observed at 10 mg/L. This indicated that the microalgae had a linear dose-dependent relationship with the zeocin antibiotic. The isolates, GPA 18.1 and LPB 14.1 were the most resistant to zeocin with colonies observed at 10 mg/L exhibiting a natural high level of resistance to zeocin with only slight reduction in growth as the zeocin concentration increased. The isolates, GPA 9.3 and 19.1, had less growth at 0 mg/L with hardly any at 3, 5 and 10 mg/L, whilst cultures NWA 23.4 and NC 72 had very stunted growth on the TAP agar plates at all concentrations tested, including 0 mg/L.

Chapter 3

3.3.6 Microalgae cryopreservation For long term microalgae storage after all necessary work was completed, the South African microalgal isolates as well as the C. reinhardtii CC-125 and CC-400 strains (transformed and untransformed) were cryopreserved. To check for viability, the cryopreserved microalgal cells were subsequently thawed and grown in TAP medium for two weeks to observe growth (Figure 3.12). The microalgal isolates FSA 22.3, GPA 18.1, NC 72, GPA 9.3, WC 4.1, CC-125 and CrGFP-CC-125 were rejuvenated successfully; high levels of post-thaw viability were obtained for these cultures with no noticeable inhibition of subsequent growth-capability. The microalgal isolates LPB 14.1 and GPA 9.1 grew, but the cultures were discoloured. The NWA 23.4 culture grew but then died shortly after inoculation, and CC-400 and CrGFP transformed CC-400 cultures did not seem to grow at all. Cryopreservation success may therefore be highly variable among species. The transgenic CC-125 strain did not produce a CrGFP signal nor was the CrGFP gene detected with colony PCR (data not shown).

FSA 22.3 GPA 18.1 NC 72 GPA 9.3 WC 4.1 CC-125 CC-125- LPB 14.1 GPA 9.1 NWA 23.4 CC-400 CrGFP

Figure 3.12: Microalgae rejuvenation growth in liquid TAP media after cryopreservation.

Chapter 3

3.4 Discussion Microalgae were selected from the MiCCSA culture collection based on criteria for future transformation and heterologous protein production: unicellular, with large or many chloroplasts and a thin cell wall. Unicellular microalgae are easier to transform and a single cell type culture would be more uniform in their recombinant protein accumulation (Specht et al., 2010). Green isolates with a greater chloroplast to cell ratio would allow for rapid conversion of solar energy to biomass accumulation (Wood et al., 2005, Pacheco et al., 2018). Furthermore, considering future electroporation transformation, microalgal isolates with no or a thinner cell wall are considered easier to transform (Jinkerson & Jonikas, 2015) as those with thick cell walls would prevent the vector DNA from entering the cell (Muñoz et al., 2018).

In this study, the South African microalgae species, C. sorokiniana, C. vulgaris, Chlorella sp. and T. obliquus had greater growth than the C. reinhardtii control CC-400. The species T. dimorphus had a lower growth rate than CC-400 but still surpassed the growth of C. reinhardtii CC-125 strain, whilst R. subcapitata had the lowest growth rate. Microalgal growth depends on a variety of factors, such as the type of alga (its cellular composition), the growth medium and physiological conditions under which it was cultured (Blair et al., 2014). At the start of the growth study, it was discovered that light was an important factor for phototrophic algae. Microalgal isolates grown in the first three growth experiments with continuous light became photobleached and ceased to grow after a few days. Even continuous low light failed to be successful for most microalgal isolates. Green microalgae are photosynthetic, so one would expect a higher light intensity to allow for greater growth rate and biomass (Singh & Singh, 2015). Most green microalgae were previously found to grow at an optimum light intensity of, 40-300 μmol m-2.s- 1 (Andersen, 2005). However, if exposed to a high or continuous lower light intensities for prolonged periods, then the growth of Chlorophyta species may cease (Janssen, 2002).

Too much light can lead to the production of reactive oxygen species (ROS) within the algal cells, which would lead to photosystem damage (Janssen, 2002); if continued it may lead to complete cell death (Gorelova et al., 2019). Green microalgal growth was found to be inhibited when light saturation reaches approximately 80-260 μmol m-2.s- 1 (Latala, 1991); however this is species-specific as isolates respond to light intensity or photoperiod differently. The specific growth rate of Tetradesmus sp. increases with light intensity with an upper limit of 420 μmol m-2.s- 1 (Singh & Singh, 2015); R. subcapitata up to 460 μmol m-2.s- 1 (Gutierrez‐Wing et al., 2012); C. vulgaris up to 100 μmol m-2.s- 1 (Khoeyi et al., 2012) but C. sorokiniana, could effectively grow under high light intensities up to 1,500 μmol m-2.s- 1 (Kliphuis et al., 2010). In this study under continuous light many of these isolates became photobleached with as little at 100 μmol m-2.s- 1 continuous light. Only C. sorokiniana GPA 18.1 and Chlorella sp. FSA 22.3 isolates did not exhibit as drastic photo-inhibition when cultivated under continuous light. The transformed and untransformed C. reinhardtii cultures in Chapter Two did not experience photobleaching when they underwent their growth curve analysis. This may be explained by the TAP medium, which has been shown to have a direct effect on the photosystem II of Chlamydomonas species, causing it to be less susceptible to photoinhibition (Roach et al., 2013).

In this study microalgal growth was successful only when a diurnal light cycle was introduced. Diurnal light cycles were found to have a higher energy conversion than continuous light, within microalgae (Carreres et al., 2018). Photosynthesis is governed by two reactions: the photochemical

Chapter 3 phase which is light dependent and produces compounds (ATP, NADPH) which are then used in the biochemical light independent phase (Janssen, 2002). Photosynthetic microalgae synchronise their metabolic activities to the diurnal light availability to maximize carbon fixation during the day (Suzuki & Johnson, 2001). The dark period allows microalgae to recover from any photo-inhibitory damage (Janssen, 2002) as well as to schedule light sensitive processes, such as DNA synthesis and cell division (Carreres et al., 2018). However, microalgae grown under continuous light within the Erlenmeyer flasks did not experience photoinhibition due to light saturation (further explained in Appendix I, Figure I10). In this study, when the OD was sampled sporadically, the growth curves appeared more scattered (Figure 3.6) than when they were sampled at the same time during each photoperiod (Figure 3.7). Cytokinesis occurs periodically in the light-dark cycle and sampling at the same time reduces scatter and results in more accurate growth curves (Wood et al., 2005).

Other light options in relevance to microalgal growth is pulsed light and the type of light wavelength. Pulsed light includes dark–light cycles at several frequencies (Vejrazka et al., 2012), the effect of which is species specific; in previous studies growth of T. obliquus cells were enhanced (Gris et al., 2014) but growth of C. reinhardtii cells were not (Vejrazka et al., 2012). The type of light wavelength (clear, blue, red and green light wavelengths) also effects microalgae growth (Jia et al., 2015). A study on these wavelengths for C. vulgaris (Blair et al., 2014) and Nannochloropsis spp. (Vadiveloo et al., 2017) observed a greater biomass productivity on blue light than white light. Light may also change the biochemical composition of microalgae: high light intensity changes the composition from protein and carbohydrates to greater lipid accumulation (Gorelova et al., 2019). The light intensity which provides the most optimum growth for recombinant protein production should be determined. In a recent study, the greatest GFP recombinant protein yield was obtained when C. reinhardtii was cultured under very low light, (35 μmol m-2.s- 1) with a diel cycle (Pacheco et al., 2018).

The photosynthetic eﬃciency of microalgae deﬁnes the upper limit on potential productivity of a cultivation system (Rocha et al., 2017). Light however is not an independent abiotic factor on microalgal growth but is intrinsically related to temperature (Fields et al., 2014) and nutrient availability (Latala, 1991). In this study greater microalgal growth was observed when cultured on TAP, containing acetate, compared to phototrophic AF6 media. These results are comparable to previous studies where greater growth was observed under acetate mixotrophic conditions for Chlamydomonas sp. (Deng et al., 2011, Rocha et al., 2017), C. vulgaris (Deng et al., 2011, Li et al., 2012, Sharma et al., 2016, Zhang, 2016, Rocha et al., 2017), C. pyrendoisa (Zhang, 2016), C. sorokiniana (Rocha et al., 2017) and R. subcapitata (Wang et al., 2017). The Tetradesmus sp. had a similar growth rate on TAP and AF6 medium and is considered capable of mixotrophic growth with no increase compared to photoautotrophic growth (Rocha et al., 2017). Acetate does not always promote growth, at higher concentrations it may even be toxic to microalgal cells (Sharma et al., 2016).

The type of nitrogen required for growth may be species specific; both Chlamydomonas sp. and Chlorella sp. grow better when ammonium is supplied (Deng et al., 2011, Wan et al., 2011, Zhang, 2016). The C. reinhardtii CC-125 strain contains the nit1 and nit2 mutations in its genome which renders it unable to produce nitrate reductase; without ammonium the cells would stop growing (Deng et al., 2011, Pröschold et al., 2005). AF6 had a small concentration of ammonium to be utilised by CC-125 but these mutations are not present in the C. reinhardtii CC-400 strain, which grew better

Chapter 3 on the AF6 medium. The Tetradesmus sp. and R. subcapitata isolates had similar growth in AF6 compared to TAP. This can be attributed to their significant preference for nitrate over ammonium balancing out the additional supply of nutrients provided in TAP (Mandal et al., 2018, Velichkova et al., 2013).

Higher nitrogen concentrations are favourable for increasing microalgal biomass (Sharma et al., 2016, Velichkova et al., 2013), a trend currently seen in 56 eukaryotic photoautotrophs (Fields et al., 2014). Nitrogen deficiency was visually observed in our experiments as pale green cultures (Figure 3.5), since a lack of nitrogen reduces the amount of chlorophyll synthesis (Andersen, 2005). On the other end of the spectrum, an excessive nitrogen concentration could lead to deleterious effects and poor biomass yields (Sharma et al., 2016). In TAP medium, cells might struggle to keep ammonium out to maintain an adequate intracellular pH level. Surplus ammonium concentrations could be turned to the respiration toxin ammonia, which could cause chloroplasts to bleach and cultures to crash (Griffiths et al., 2011).

Although phosphorous is considered one of the main macronutrients; it is not required in the medium (Zhang, 2016). Microalgae have a typical N:P ratio of 16:1 (Gour et al., 2018) and the green microalgae in the class Chlorophyceace, have an internal source which they utilise first (Zhang, 2016). Micronutrients and trace metals also influence growth with significantly less of these in AF6 than in TAP medium. Citric acid however, was only present in AF6 media, this acid solubilizes salts and increases their availability to microalgal cells, allowing for greater growth rates (Gour et al., 2018). AF6 media is also often used for the long term storage of algal culture collections since it contains vitamins (Richmond & Hu, 2013).

The dry weight biomass obtained at the end of the culture period for species, Tetradesmus sp. and R. subcapitata was far greater than expected based on the cell density measured by OD750. Often OD may give an overestimation of the growth data (Zhang, 2016); therefore calibration curves were created to try and standardize the OD in relation to cell size and number for each isolate. In a previous study with Tetradesmus sp., greater biomass was obtained in contrast to the number of cells, and the biomass was far greater than that of Chlorella sp. (Rocha et al., 2017). Optical density as an indication of growth could be underestimated due to cell aggregation and morphology (Tam & Wong, 1989). Colonial clustering is commonly observed in both Tetradesmus spp. (Zhang, 2016) and Raphidocelis spp. (Krienitz et al., 2001). Under microscopic observation (Appendix H, Table H1) the South African Tetradesmus sp. had a cell size of approximately (10 μm) and R. subcapitata (7-10 μm), both far larger than that of Chlorella sp. (3-6 μm) or C. reinhardtii species (4-6 μm). Similarly, the CC- 400 grew faster with a higher final cell density, but the CC-125 strain had a greater final biomass yield, in this instance the cell-wall may account for the difference (Fan et al., 2017). In addition to the variation in cell size, the biochemical composition (protein, amino acids, carbohydrate, lipid, fatty acids, chlorophyll and carotenoids) for each microalgae is different (Sharma et al., 2016) and could also influence the final biomass weight.

Not all microalgae in this study had the same nutrient requirements, indicating the existence of functional diversity. For example, different sources of nitrogen would be available in an aquatic ecosystem; often in nutrient poor water ammonium is predominant whilst nitrate is the principal nitrogen source in eutrophic waterways (Glibert et al., 2016). In nature, nutrient availability may affect the biodiversity of species and the fitness of an organism may be due to the environment and their genetics (Andersen, 2005).

Chapter 3

Microalgae in this study were isolated from freshwaters in South Africa and identified by genetic sequencing. The 18S rRNA universal primers used in this study have been previously used to categorise algae of Chlorophyceae and Trebouxiophyceae with species Desmodesmus sp., Scenedesmus dimorphus, and Scenedesmus communis (Duong et al., 2015). Due to South Africa’s location, we expected the isolates to include novel species; however, the combined molecular and morphological analysis identified the fast-growing South African isolates as well-known, high- biomass-yielding microalgae. The microalgae identified belonged to either the class Chlorophyceae as C. sorokiniana, C. vulgaris, Chlorella sp. and R. subcapitata, or to the class Trebouxiophyceae as T. obliquus or T. dimporphus. There is a slight possibility that sequences with lower percentage identities, GPA 9.1 and GPA 9.3 could possibly be new species related to T. obliquus and C. sorokiniana but sequencing of more genes and multilocus sequence analysis would be required to reveal this.

Some isolates were duplicates of the same genera; both GPA 9.1 and LPB 14.1 were considered T. obliquus, while WC 4.1 and FSA 22.3 were taken for A. pyerndoisa. We would expect same species from the same region if there is a high abundance of those species in that area, but the species were isolated from separate locations across the country: from Gauteng, Limpopo, the Western Cape and the Free State. The isolates might also not be identical to previous isolates described in GenBank because of the geographical isolation between freshwater sources in South Africa and those frequently found in GenBank (North America, Europe and Asia). Different strains of the same species may have different biochemical profiles, as we saw with C. reinhardtii CC-125 and CC-400. However, in this study, the best growing microalgal isolates were screened for, therefore it makes sense that they represent some of the most predominant microalgae species in industry today.

The microalgae, Chlorella sp., Tetradesmus sp. and Raphidocelis sp., are well known for their rapid asexual growth and are currently used in commercial applications such as biofuel production (Cazzaniga et al., 2014, Guo et al., 2013), wastewater treatment (Dzhambazov et al., 2002, Breuer et al., 2015, de-Bashan et al., 2008), animal and human feed (Guo et al., 2013, Cazzaniga et al., 2014), aquaculture (Ahmad et al., 2018), ecological bioassays (Yamagishi et al., 2017) as well as cosmetic and pharmaceutical industries (Suzuki et al., 2018). Of these genera, nuclear genome sequencing has been recently completed for C. vulgaris (Guarnieri et al., 2018), T. obliquus (Carreres et al., 2017) and R. subcapitata (Suzuki et al., 2018). In this study, the Chlorella sp. were resistant to high light conditions; the species is also known to be incredibly robust with a range of cultivation conditions (Xu & Hu, 2013) which make them desirable for commercial biotechnology applications (Cazzaniga et al., 2014).

The development of a Chlorella sp. heterologous protein expression system has been delayed due to the lack of a stable transformation technique or sophisticated genetic toolbox (Liu & Chen, 2014), but research is now expanding into developing a molecular toolkit for C. vulgaris (Kim et al., 2018). In none of the previous genetic transformations were the transgenes driven by endogenous algal promoters (Dawson et al., 1997, Chow & Tung, 1999, Hawkins & Nakamura, 1999, Niu et al., 2011, San Cha et al., 2012). As discussed in Chapter Two, endogenous algal regulatory elements might greatly increase transgene expression and transforming these isolates with the pChlamy4 vector containing the Hsp70A-rbcS2 enhancer/promoter and the desired gene could greatly progress microalgal biotechnology research. In this study, the South African microalgae were subjected to a

Chapter 3 zeocin kill curve to determine whether zeocin selection by the pChlamy4 plasmid would be feasible for future transformation events.

Many algae have natural resistance to most antibiotics commonly used for genetic transformation; an undesirable trait when considering the selection of positive gene transformants (Gomma et al., 2015). In our study, the response of the South African isolates to zeocin was species dependent. Most microalgal growth was inhibited at 5 mg/L zeocin, indicating potential transformation feasibility with the pChlamy4 vector. The concentration of 5 mg/L zeocin was also considered the most optimal concentration on the genetically transformed Tetraselmis chuii green microalgae (Úbeda-Mínguez et al., 2015). The isolates, C. sorokiniana GPA 18.1 and T. obliquus LPB 14.1 appeared to have natural resistance to zeocin, however, previous studies observed that Chlorella sp. (Wang, 2007) and Scenedesmus almeriensis (Dautor, 2014) were sensitive to zeocin. This might be due to a biochemical difference between the South African microalgae and those species characterised before, or the unstable zeocin in these plates degraded due to excessive light exposure (Díaz‐Santos et al., 2013). Antibiotics are not just required for transformation selection, but may be necessary to maintain stable, contaminant free microalgal cultures.

Often cultured microalgae may contain bacterial and fungal contamination (Harris, 2009) which could easily outgrow the desired microalgal species (Andersen, 2005). If contamination is too high, the conditions may become toxic, and the microalgal culture would collapse (Andersen, 2005). In general, fungal contamination may be worse than bacterial contamination since it may overgrow the microalgae and is harder to eradicate (Mahan et al., 2005). In this study contamination was predominately observed in TAP media containing the organic carbon source (Wan et al., 2011). Contamination is also often seen in vitamin-free cultures as a symbiosis between vitamin producing bacteria and algae (Warren et al., 2002). However, most liquid TAP medium microalgal cultures became stressed and died which suggests that the bacteria present was not symbiotic but pathogenic, toxic or simply outcompeting for nutrients.

In this study, we adapted a method to reduce the bacterial and fungal contamination for C. reinhardtii cultures grown on TAP media. Previous research was combined to produce an antibiotic-antimycotic cocktail mixture (Harris, 2009, Sreedharan et al., 2018, Preisig & Andersen, 2005). The antibiotics kanamycin and carbenicillin kill bacteria; the former by inhibiting protein synthesis, whilst the latter by inhibiting peptidoglycan cross-links in cell wall synthesis (Sambrook et al., 2001). The carbendazim within this cocktail is considered an inexpensive, broad spectrum fungicide which interferes with cell division and transport by binding directly to the fungal microtubules (Mahan et al., 2005).

After a period following antibiotic exposure, the cultures would once again go cloudy and exhibit signs of contamination. We discovered that the predominant source of bacterial and fungal contamination was the air within the laboratory. In June 2017 the phycology laboratory was downscaled and relocated to a new workroom. During this study the air vents in the new laboratory space were often under construction. This left the laboratory vulnerable to contamination from the workers and the unsterile outside air. In future, extra caution should be taken to ensure sterile conditions are maintained. The higher tolerance that microalgae have to antibiotics compared to bacteria (Yu et al., 2017), allows the viability of antibiotic supplementation in media. However, careful consideration should be taken since antibiotics could apply a selective pressure on the microalgae and bacterial antibiotic resistance could accumulate (Andersen, 2005).

Chapter 3

Antibiotics could inhibit undesired pathogens as well as act as a transformation selectable marker without drastically affecting microalgal growth. However, the use of antibiotics and fungicides significantly increase operating costs, and should only be used when necessary (Sreedharan et al., 2018, Chen & Melis, 2013). Alternative methods should be considered for the long-term maintenance of microalgal cultures.

For this study the microalgal cultures were maintained for approximately 18 months by sub- culturing. Agar plates and serial transfers requires extensive labour, consumable costs and time, with a greater risk of mislabelling, contamination and strain deterioration through genetic drift (Andersen, 2005). In this study the GeneArt® Cryopreservation Kit for Algae was used to preserve the nine indigenous, two C. reinhardtii (CC-125 and CC-400) and two CrGFP transformed C. reinhardtii (CC-125 and CC-400) microalgal cultures. Cryopreservation freezes samples at very low temperatures, to make cells metabolically inactive for long term storage (Hipkin et al., 2014). This metabolically inactive state preserves the morphological, physiological, biochemical and genetic state of microalgal stock cultures (Day et al., 2017).

As with most of the experiments within this study, cryopreservation and the recovery of the microalgae was species-specific (Stock et al., 2018). The cryopreservation kit we utilised was optimised for Chlamydomonas sp. and Chlorella sp. and as expected microalgal isolates which grew well after storage at -80˚C were these, as well as the Tetradesmus sp. Their cells appeared visually healthy but cellular morphology and growth rate differences should be analysed between the stock and rejuvenated cells to ensure no structural or functional features were lost (Stock et al., 2018). Species-specific cell features could influence their ability to survive ultra-low temperatures such as their cell-wall polysaccharide composition, degree of cell wall silicification and cell size (Stock et al., 2018).

The CC-400 strain did not rejuvenate after freezing; without the protection of a cell wall, organelles within the cytoplasm most likely suffered from permanent damage or cell lysis. The other unrejuvenated isolate, R. subcapitata is sensitive to toxicants (Rojíčková & Maršálek, 1999) and is most likely also intolerable to freeze-thawing. Cryopreservation of microalgae is still very experimental and the underlying physiology of cell injury during freezing and thawing is not fully understood (Stock et al., 2018). In the GeneArt® Cryopreservation manual, thawed cultures were immediately cultured in light. The microalgae which did not grow well most likely underwent photo- oxidative stress (Hipkin et al., 2014). A suggested method to improve viability and reduce light- induced stress would include an incubation period in the dark for 24 hours before and after cryopreservation (Day et al., 2017, Hipkin et al., 2014). Our transgenic CrGFP CC-125 culture was rejuvenated but failed to produce a CrGFP signal with no Crgfp gene detected on the DNA level. There is very little information regarding the cryopreservation of transgenic microalgae, the first successful cryopreservation of a transgenic microalgae was for a diatom (Hipkin et al., 2014) with almost no subsequent studies.

According to Hipkin, the only practical long-term preservation of microalgae is cryopreservation. In their studies, microalgae post-thaw obtained the highest cell viability levels of 75% (Hipkin et al., 2014). The viability of microalgae cells after thawing has been recorded to be stable for at least 20 years (Day et al., 1997). Although not successful for all species in this study, cryopreservation has become a standard approach for large, global microalgal culture collections (Day et al., 2017) and should be researched further for the long-term storage of the MiCCSA database.

Chapter 3

Study limitations and future considerations Microalgal culturing is still a small niche and further developments will depend on continued research. Many of our results followed the trend established by previous studies using similar nutrient and cultivation conditions. However, the OD and biomass values obtained in this study were slightly lower than those previously reported and could be attributed to difference in the laboratory conditions and equipment available for microalgal cultivation. Another explanation could be that despite observed taxonomic agreement between the morphological and molecular results, the geological location could impact the microalgae’s physiological characteristics (Hadi et al., 2016).

To eliminate OD-cell number calibration curves, future growth experiments could utilise flow cytometry. This method is just as sensitive as OD to measure cell concentration with the advantage that it takes cellular characteristics, such as cell size (which varied among the isolates in this study)- into consideration (Chioccioli et al., 2014).

The most ideal medium would be that which supplies the highest growth rate for the microalgal isolates while containing the least constituents, to reduce the cost of large-scale production. This study validates that it is possible to scale down the chemical costs for microalgal cultivation on AF6 rather than TAP media. Photoautotrophic growth is inexpensive with less contamination, whilst mixotrophic cultivation is a desirable option for greater biomass yields for larger scale cultures, as well as increased recombinant protein production (Chai et al., 2018). A future prospect could be a two-stage cultivation where photoautotrophic growth is followed by mixotrophic growth at higher light intensities for greater biomass and recombinant protein production with reduced risk of contamination (Yen & Chang, 2013). Another option that does not seem to be previously tested but might be of value, is supplementing our AF6 medium with acetate.

Future studies should compartmentalise the media into the individual nutrient components and test these as well as the physical conditions for the most optimal inexpensive cultivation (Eriksen, 2008). Otherwise optimal growth conditions can also be determined and developed for each algal isolate of interest. Furthermore, media has been found to alter the biochemical composition of microalgae and culture conditions should be considered for the greatest production of recombinant proteins in C. reinhardtii (Harrison et al., 1990, Juneja et al., 2013, George et al., 2014).

In this study during the screening and identification of microalgae, it became clear that there are gaps in microalgal taxonomy. No Chlorophyta or other microalgal 18S rRNA sequences were found on the NCBI refSeq database, instead, the comparison sequences used were taken from GenBank. The GenBank database is an archive of sequences submitted from the public that are not curated and may be redundant (Benson et al., 2018). Sequences from this study were not submitted to GenBank, since many had an E value of 0 and submitting would only increase the redundancy of sequences currently available. However, the taxonomic identification of these organisms will be submitted to the MiCCSA database to contribute to the list of unique South African isolates.

Although the 18S region is fundamental for global identification of microalgae (Bradley et al., 2016), in future it would be best to consider utilising more than one genomic region (such as the ITS, cox1 and rbcl genes) for accurate taxonomic placement (Champenois et al., 2015). The local isolates identified in this study only represent a tiny fraction of the diversity available, thus ongoing screening and identification should continue. For future large-scale identification of the entire 500 MiCCSA database, a more rapid and economical method should be considered. Matrix Assisted Laser

Chapter 3

Desorption/Ionization Time-of-Flight Mass Spectrometry would be able to differentiate microalgal mixtures at the species as well as strain levels; with a higher taxonomic resolution than molecular techniques (Barbano et al., 2015).

3.5 Conclusion This study aimed at screening and identifying South African microalgae to determine their feasibility for future biotechnological research. Nine isolates from the MiCCSA database which had high growth potential, were selected. Physiological and metabolic characterization of the isolates were performed to provide relevant cultivation information. The microalgae were studied with their growth properties in inexpensive phototrophic media, under mixotrophic media and with the antibiotic selectable marker zeocin. Very little information is available regarding the growth of any green microalgae on AF6 media, so the results in this study may prove valuable to other interested microalgae researchers or industries. In order of the greatest growth potential, microalgae which grew best were identified by Sanger sequencing as C. sorokiniana, Chlorella sp., C. vulgaris, T. obliquus, T. dimorphus and R. subcapitata species.

In this study, the South African Chlorella spp. isolates had a greater growth rate in both AF6 and TAP medium than Chlamydomonas spp. controls. The C. sorokiniana species are considered superior for commercial cultivation as they are more resistant to high light and shear stress (Taunt et al., 2018) and is therefore a good candidate for heterologous protein production. Culture conditions in the South African laboratory were refined by eliminating commonly reoccurring contamination using an antibiotic-antimycotic mixture. Furthermore, the validity of cryopreservation was tested for the long-term storage viability of South African microalgal isolates. Culture collections should not only contain the indigenous isolates as a record of biodiversity, but it would be valuable to have physiological data regarding these isolates. Therefore, all information regarding the isolate origin, taxonomic identity, morphological features and growth characteristics, would be able to add to the knowledge to the MiCCSA database both for ecological conservation as well as potential use in the biotechnology industry.

Chapter 3

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Chapter 4 General conclusion: Future potential of microalgae biotechnology in South Africa

Chapter 4

4.1 Introduction Microalgal biotechnology is developing at an extraordinary rate with this study using literature from around the globe, including the United Kingdom, North America, the Netherlands, Czech Republic, Spain, Brazil, Australia, South Africa and China. Microalgal biotechnology is attracting more and more investment from organisations as a means for producing high-quality sustainable biofuels, human and animal nutrition, high value products, bioplastics, biochemicals, fertilisers, pharmaceuticals and cosmetics. Despite all the advancement; the commercialisation of microalgae technologies is still poor in developing countries, where it may offer the most value. With its low- cost, easily scalable, low maintenance technologies, microalgae could impact the socio-economy as a sustainable resource and help conserve the environment.

4.1.1 Conclusion to this study In Chapter Two, we stably transformed the Crgfp transgene into the nuclear genome of Chlamydomonas reinhardtii CC-125 and CC-400 strains, validating the feasibility of future microalgal biotechnology research in South Africa. In Chapter Three, South African microalgae, in order of the greatest growth potential, were identified as Chlorella sorokiniana, Chlorella sp., Chlorella vulgaris, Tetradesmus obliquus, Tetradesmus dimorphus and Raphidocelis subcapitata species. All the indigenous microalgae except for R. subcapitata, had greater specific growth rates and biomass concentration than the C. reinhardtii controls on both phototrophic minimal AF6 media and nutrient rich, mixotrophic TAP media. Although these isolates were identified with a very high match similarity to microalgal isolate sequences in the Genbank, it must be considered that the geographical difference could influence the biochemical profiles of the isolates as slightly different to those isolated elsewhere in the world. Microalgae culturing techniques were refined to eliminate contamination for long term maintenance of stock cultures and the isolates C. reinhardtii, C. sorokiniana, Chlorella sp., C. vulgaris, T. obliquus, T. dimorphus, were cryopreserved for future microalgal research. The Chlorella spp. had the greatest specific growth rate and could tolerate high light, temperature and slow freezing conditions associated with commercial culturing; justifying it as a microalgae option for many future biotechnology applications.

Subsequent conclusions and future direction of this study According to Rocha and co-workers, genetic transformation for protein expression is possible for most Chlorophyta microalgae (Rocha et al., 2017). Therefore, the optimised electroporation procedures presented in Chapter Two were utilised to transform Crgfp-pChlamy4 into all three Chlorella spp. isolated in Chapter Three. However, none of these transformations were successful (no data shown). The most likely reason is that Chlorella spp. have rigid cell walls that inhibit the exogenous DNA from entering the cell and incorporating within the nucleus. The cell wall structure and composition is different for many microalgae species (Yamano et al., 2013). Unlike the cellulose hydroxyproline-rich glycoproteins cell walls of Chlamydomonas spp. (Burczyk et al., 1999), Chlorella spp. cell walls contain glucosamine, rhamnose and arabinose polysaccharides (Muñoz et al., 2018).

The cell wall is not a new limitation; since the 1990’s the electro-transformation studies of microalgae observed that transformation efficiency decreases with increasing cell wall thickness (Kotnik et al., 2015). Some algal cell walls are highly resistant to the reorientation required for pore formation; but if the voltage is too high it would lead to heat damage and cell death (Kotnik et al., 2015). Therefore, the voltage required for transformation is species specific.

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Transformation between species may also be influenced by the life cycle, reproduction strategy and ultrastructure of the microalgae (Chungjatupornchai et al., 2016). In previous Chlorella spp. transformations, enzymes such as chitinases and lysozymes, specific for the degradation of N- acetylglucosamine containing polymers in the cell wall, were used (Gerken et al., 2013). By removing the cell wall, electro-transformation of Chlorella spp. is more likely (Kumar et al., 2018). This highlights that there is a need for the development of robust and stable genetic transformation methods for most microalgal strains (Díaz‐Santos et al., 2013). Future research could harness positive Chlorella spp. transformation results from literature to troubleshoot transformation for the South African microalgal isolates.

Since microalgal biotechnology has many benefits over current expression systems, production of heterologous proteins of industrial value should initially be attempted in the C. reinhardtii CC-400 strain. Currently the CSIR utilises yeast, bacteria and tobacco protein expression systems; specializing in the production of antibodies, reagent proteins and animal vaccines. For example, the CSIR currently produces the RabiVir™ monoclonal antibody in tobacco for the post-exposure prophylaxis application against rabies (CSIR, 2018). Recombinant proteins may be fused to CrGFP and the electro-transformation protocol established in Chapter Two could be used for transformation and production of heterologous proteins. The production of the recombinant protein could also be optimised by adjusting the culture conditions, analysing the life cycle and process design to maximize productivity, efficiency, and cost ratios of microalgal expression system. Future research could then be extended to include expressing these proteins within the chloroplast genome of the C. reinhardtii CC-400 strain.

In this study the low transgene expression in the nuclear genome of C. reinhardtii was overcome by codon optimisation and the use of endogenous algal elements. A prospect for higher transgene production could be to utilise synthetic biology to create completely novel genetic arrangements (Taunt et al., 2018). Novel synthetic promoters and UTRs could be designed which combine multiple positive elements and exclude all negative regulatory elements. Such a synthetic chloroplast genome has already been developed for use in C. reinhardtii (O’Neill et al., 2012). We could further aid the insertion of exogenous DNA into highly transcribed regions of the genome by Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 technology (Jinkerson & Jonikas, 2015). As proof of concept, the CRISPR/Cas9 system has been transiently and stably transformed into the marine diatom Phaeodactylum tricornutum (Sharma AK, 2018). CRISPR has also been used to create a knock out mutant of the zeaxanthin epoxidase gene in C. reinhardtii (Baek et al., 2018).

Ethical standards of genetically modified microalgae The C. reinhardtii cells transformed in this study harbour a foreign DNA element, the Crgfp gene, within the nuclear genome, and are therefore considered genetically modified organisms (GMO) (Ayele, 2007). South Africa regulates genetic engineering according to the GMO Act (No 15 of 1997), a legal framework to ensure that the development of modern biotechnology takes place under safe conditions (Mayet, 2000). The C. reinhardtii microalgae has been classified as a GRAS organism by the United States Food and Drug Administration (Murbach et al., 2018), but to minimise any potential risk- the Safety Guidelines from the Biosafety in Microbiological and Biomedical Laboratories, 5th ed., were adhered to in this study (Chosewood, 2007).

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For future consideration, a scientifically-based risk assessment should be conducted before any algal derived protein may be released for commercialisation (Mayet, 2000). Once considered safe the public must be notified before release of any microalgal GMO products (Ayele, 2007). In this study any environmental risks associated with GMOs were eliminated since the microalgae were cultured and contained within the laboratory and potential scale up will remain in a closed system. The transgenic algal species will not be able to contaminate those in nature and no interbreeding should occur. Therefore, the risk of outcrossing (the threat of transgene drift into the environment) is predicted as nonviable and biodiversity will not be affected (Jaffe, 2004). If we wish to upscale the transgenic microalgae, it would be best to maintain the microalgae a closed transparent vessel (Mayet, 2000).

Large scale cultivation To comply with the South African GMO act of 1997, recombinant protein expressing microalgae would be best grown within a photobioreactor (Figure 4.1A). Photobioreactors may reduce foreign contamination experienced within culture flasks in this study or prevalent within open pond systems (Gupta et al., 2015). The C. reinhardtii or South African microalgae’s’ ideal growth parameters, such as light intensity, nutrient levels, and temperature, may be controlled to establish the most optimal economical algae productivity (Gao et al., 2017, Demirbas & Demirbas, 2011). Photobioreactors are thought to require extensive infrastructure due to artificial lighting and maintenance, unavailable for developing countries (Rosenberg et al., 2008). However, an efficient, scalable, easily manageable and affordable alternative closed cultivation system utilises sterile-disposable polythene hanging bags (Figure 4.1B). When incorporated with a simple medium such as AF6, the cost of this system is approximately $0.002 per litre (Taunt et al., 2018, Levine, 2018). These bags may be enriched with

CO2 and instead of artificial lights, photovoltaic devices may capture sunlight -abundant in South Africa- and emit the solar energy through LED lights. In this manner the exact light of optimal intensity and wavelength can be achieved for the best biomass and recombinant protein production (Loera‐Quezada et al., 2016).

A B Figure 4.1: Closed systems for cultivating microalgae (A) traditional photobioreactor systems for algae culturing. (Philippidis, 2015) and (B) microalgae growth in 40 L hanging bags (Dowd, ND).

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4.1.2 Future applications in heterologous protein production Potential applications of transgenic microalgae are not just limited to production of recombinant molecules, but may be used as whole cells for viral, bacterial and insect control. Microalgae’s GRAS status makes them suitable for the large-scale production of oral vaccines as an inexpensive, accessible, solution to many third world diseases (Specht et al., 2010). Transgenic microalgae may help with the struggle against malaria with transgenic C. reinhardtii successfully developed as a mosquito larvicide (Kang et al., 2017).

C. reinhardtii has been researched as a system to deliver oral vaccines to fish, animals and even humans. These vaccines are easy to transport and administer to rural locations as they do not require specialised staff or constant refrigeration (Chebolu S, 2009, Kwon KC, 2019). Successful vaccines utilising transgenic microalgae have been designed and tested on rabbits and trout- both eliciting a proper immune response when delivered orally (Harakuni T, 2005, Sayre et al., 2017). To comply with regulations, bio-encapsulated transgenic microalgae may be cultivated within a closed system and then the biomass freeze-dried to ensure they are inanimate before they are released into the environment (Charoonnart et al., 2018). This is of special interest to South Africa’s aquaculture industry since the South African Marine Aquaculture Centre industry was discontinued in 2015 (Wang et al., 2018). At the 31st Congress of The Phycological Society of Southern Africa it was conferred that aquaculture in South Africa is failing due to disease (SANPCC, 2018). This is an entire industry in South Africa where genetically modified microalgal vaccines could act as an environmentally sustainable solution.

4.1.3 Microalgae as a sustainable resource The importance of this project proceeds beyond just a platform for future microalgae biotechnology research. The use of wild type indigenous algal strains would address many international key issues regarding the sustainable use of resources and climate change. “Our planet is running out of resources”; the effect climate change will have on our economy, environment and global society is discussed in the 2018 report, the Green New Deal. World leaders state that we need a sustainable approach for water usage, fuel and food production that reduces the effects of climate change (Carlock, 2018).

Microalgae have approximately a nine-fold greater photosynthetic efficiency than terrestrial land plants (Duong et al., 2015, Norsker et al., 2011, Kirst & Melis, 2018) which allows for a faster growth rate and greater biomass production using less resources (Miao et al., 2004). Furthermore, the production of microalgae has the potential to transform current liabilities into valuable assets. Microalgae can capture carbon from the atmosphere (Sydney et al., 2019), wastewater can be bio- remediated (Osundeko et al., 2019), microalgae growth in open ponds or photobioreactors can utilise non-arable land (Zhuang et al., 2018) and the microalgal biomass generated has huge commercial potential as biofuel, bioplastics or animal feed (Khan et al., 2018).

In this study the growth of South African microalgal isolates were screened for growth on AF6 media to include algae with nitrate reductase function. This allows a more diverse set of future applications not restricted to ammonium supplemented media. A future prospect could be to use wastewater as the medium. Microalgae in this study, Chlorella sp. (Bohutskyi et al., 2016), Chlamydomonas sp. (Kong et al., 2010), Tetradesmus sp. (Bohutskyi et al., 2016) and Raphidocelis sp. (Kalka, 2012) have been successfully cultivated in wastewater. Approximately 20 mg/L nitrogen and 4 mg/L phosphorus

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Chapter 4 is contained within domestic wastewater, very similar to AF6 medium, an ideal amount for microalgal growth (Rahman et al., 2015). The microalgae could potentially remove nitrogen, phosphorus, and heavy metals upon cultivation, leaving cleaner, bio-remediated water (Blair et al.,

2014). If CO2 emissions from an industrial source were directly used to supplement microalgal cultures, it may provide an excellent, inexpensive medium for microalgae with the added environmental benefit of wastewater bioremediation and lessening fossil fuel emissions in the process.

In South Africa’s National Research Developmental Strategy, emphasis is placed on how the improvement in science and technology should allow for economic improvement, increase in quality of life and sustainability for our nation (Wang et al., 2018). South Africa aims to focus on applying bio-innovations to generate sustainable economic, social and environmental development. Microalgae has a potentially important application in the emerging bio-economy strategy of South Africa where biological materials are used to create products. Microalgae spans across the bio- economy spectrum from biorefinery, to agriculture, to aquaculture, environmental sciences with wastewater treatment and even health through the production of easily accessible, easily administered vaccines and therapeutic proteins (Ferreira et al., 2018).

Microalgae holds large economic value, with their worth only predicted to increase. In January 2018, a report from Meticulous Research™ forecasts that by 2022 the worldwide algae products market will reach $3,368.42 million with a compounded annually growth rate of 6.7% during the period of 2017-2022 (Meticulous, 2018). There is currently an inverse relationship between the volume of microalgal products produced and the cost of production; the higher the volume, the lower the economic value (Figure 4.3). For example, microalgae biofuels, falls into the high volume, low cost margin and therapeutic proteins falls into the small volume, high value spectrum (Norsker et al., 2011). Therefore, although fuels are the largest commodity on this planet (Lewis, 2007)- we first need to focus on establishing microalgal biotechnology.

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Figure 4.2: The input for microalgae biomass may produce a range of products which have an inverse relationship between the volume and cost of products produced. Adapted from (Levine, 2018).

South Africa has the potential to innovate its own biotechnology applications to create value and public benefit. Microalgae products may transcend to society with scientific research institutes acting as a technology transfer unit- an innovation chasm from research to industry. Therefore, the goal for South Africa should be to establish microalgal biotechnology focusing on high value recombinant protein products, such as HIV neutralising antibodies, which would generate the most money per litre and would provide the greatest socio-economic impact. Once microalgae have proven their success it would be easier to gain support from research institutions, government and the public to expand microalgae research into delivering sustainable waste water treatment, biofuels, aquaculture and agricultural applications.

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4.2 Final Conclusion South Africa has great potential in the field of microalgae biotechnology. In this study we established an electroporation transformation protocol to produce a recombinant CrGFP protein system within the C. reinhardtii CC-125 and CC-400 microalgae strains. South African microalgae with the most promising growth potential were characterised on AF6 and TAP medium and genetically identified as C. sorokiniana, Chlorella sp., C. vulgaris, T. obliquus, T. dimorphus and R. subcapitata. This project set the foundation for the expression of industrially desired recombinant proteins in C. reinhardtii; as well as considering the potential nuclear transformation in select indigenous species such as Chlorella sp. There is a great need for future microalgae research to optimise microalgae protein expression systems and to develop products of value. It is essential that we start investing in microalgal biotechnology research and so that we may deliver tangible sustainable products as solutions to society’s largest challenges of resource depletion, food, fuel and water security as well as climate change. The long-term reward will be invaluable to our society and the global community at large.

"The greatest threat to our planet is the belief that someone else will save it" Robert Swan

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4.3 References Ayele S, 2007. The legitimation of GMO governance in Africa. Science and Public Policy 34, 239-49. Baek K, Yu J, Jeong J, Sim SJ, Bae S, Jin E, 2018. Photoautotrophic production of macular pigment in a Chlamydomonas reinhardtii strain generated by using DNA‐free CRISPR‐Cas9 RNP‐mediated mutagenesis. Biotechnology and Bioengineering 115, 719-28. Blair MF, Kokabian B, Gude VG, 2014. Light and growth medium effect on Chlorella vulgaris biomass production. Journal of Environmental Chemical Engineering 2, 665-74. Bohutskyi P, Kligerman DC, Byers N, Nasr LK, Cua C, Chow S, Su C, Tang Y, Betenbaugh MJ, Bouwer EJ, 2016. Effects of inoculum size, light intensity, and dose of anaerobic digestion centrate on growth and productivity of Chlorella and Scenedesmus microalgae and their poly-culture in primary and secondary wastewater. Algal Research 19, 278-90. Burczyk J, Śmietana B, Termińska-Pabis K, Zych M, Kowalowski P, 1999. Comparison of nitrogen content amino acid composition and glucosamine content of cell walls of various chlorococcalean algae. Phytochemistry 51, 491-7. Carlock G ME, Mcelwee S, 2018. A Green New Deal. A Progressive Vision for Environmental Sustainability and Economic Stability Policy Report by Data for Progress. www.dataforprogress.org/green-new-deal. Charoonnart P, Purton S, Saksmerprome V, 2018. Applications of microalgal biotechnology for disease control in aquaculture. Biology 7, 24. Chebolu S, Daniell H, 2009. Chloroplast-derived vaccine antigens and biopharmaceuticals: expression, folding, assembly and functionality. Plant-Produced Microbial Vaccines 332, 33-54. Chosewood LC, 2007. Biosafety in microbiological and biomedical laboratories. National Institutes of Health, Diane Publishing 4, . Chungjatupornchai W, Kitraksa P, Fa-Aroonsawat S, 2016. Stable nuclear transformation of the oleaginous microalga Neochloris oleoabundans by electroporation. Journal of Applied Phycology 28, 191-9. CSIR, 2018. Technology Available for Licensing. South African Council for Scientific and Indutrial Research. www.csir.co.za/technology-available-for-licencing. Demirbas A, Demirbas MF, 2011. Importance of algae oil as a source of biodiesel. Energy Conversion and Management 52, 163-170. Díaz‐Santos E, Vega M, Vila M, Vigara J, León R, 2013. Efficiency of different heterologous promoters in the unicellular microalga Chlamydomonas reinhardtii. Biotechnology Progress 29, 319-28. Dowd T, ND. Growth of Haematococcus pluvialis in 40L hanging bags. Phyconet: Supreme Biotechnologies Ltd. www.phyconet.org.uk/. Duong VT, Ahmed F, Thomas-Hall SR, Quigley S, Nowak E, Schenk PM, 2015. High protein-and high lipid-producing microalgae from Northern Australia as potential feedstock for animal feed and biodiesel. Frontiers in Bioengineering and Biotechnology 3, 53. Ferreira A, Marques P, Ribeiro B, Assemany P, de Mendonça HV, Barata A, Oliveira AC, Reis A, Pinheiro HM, Gouveia L, 2018. Combining biotechnology with circular bioeconomy: from poultry, swine, cattle, brewery, dairy and urban wastewaters to biohydrogen. Environmental Research 164, 32-8. Gao X, Kong B, Vigil RD, 2017. Comprehensive computational model for combining fluid hydrodynamics, light transport and biomass growth in a Taylor vortex algal photobioreactor: Lagrangian approach. Bioresource Technology 224, 523-30. Gerken HG, Donohoe B, Knoshaug EP, 2013. Enzymatic cell wall degradation of Chlorella vulgaris and other microalgae for biofuels production. Planta 237, 239-53. Gupta PL, Lee SM, Choi HJ, 2015. A mini review: photobioreactors for large scale algal cultivation. World Journal of Microbiology and Biotechnology 31, 1409-17. Harakuni T, Sugawa H, Komesu A, Tadano M, Arakawa T, 2005. Heteropentameric cholera toxin B subunit chimeric molecules genetically fused to a vaccine antigen induce systemic and

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mucosal immune responses: a potential new strategy to target recombinant vaccine antigens to mucosal immune systems. Infect Immunology 73, 5654–65. Jaffe G, 2004. Regulating transgenic crops: a comparative analysis of different regulatory processes. Transgenic Research 13, 5-19. Jinkerson RE, Jonikas MC, 2015. Molecular techniques to interrogate and edit the Chlamydomonas nuclear genome. The Plant Journal 82, 393-412. Kalka J, 2012. Landfill leachate toxicity removal in combined treatment with municipal wastewater. The Scientific World Journal. www.hindawi.com/journals/tswj/2012/202897/abs/. Kang S, Odom OW, Thangamani S, Herrin DL, 2017. Toward mosquito control with a green alga: expression of Cry toxins of Bacillus thuringiensis subsp. israelensis (Bti) in the chloroplast of Chlamydomonas. Journal of Applied Phycology 29, 1377-89. Khan MI, Shin JH, Kim JD, 2018. The promising future of microalgae: current status, challenges, and optimization of a sustainable and renewable industry for biofuels, feed, and other products. Microbial Cell Factories 17, 36. Kirst H, Melis A, 2018. Improving photosynthetic solar energy conversion efficiency: the truncated light-harvesting antenna (TLA) concept. Microalgal Hydrogen Production: Achievements and Perspectives, 335-53. Kong QX, Li L, Martinez B, Chen P, Ruan R, 2010. Culture of microalgae Chlamydomonas reinhardtii in wastewater for biomass feedstock production. Applied Biochemistry and Biotechnology 160, 9. Kotnik T, Frey W, Sack M, Meglič SH, Peterka M, Miklavčič D, 2015. Electroporation-based applications in biotechnology. Trends in Biotechnology 33, 480-8. Kumar M, Jeon J, Choi J, Kim S-R, 2018. Rapid and efficient genetic transformation of the green microalga Chlorella vulgaris. Journal of Applied Phycology 30, 1735-45. Kwon LA, Fox D, Jegathese S, 2019. An evaluation of microalgae as a recombinant protein oral delivery platform for fish using green fluorescent protein (GFP). Fish & Shellfish Immunology 87, 414-20. Levine SMI, 2018. Introduction to Algae. Coursera. www.coursera.org/learn/algae. Lewis NS, 2007. Powering the planet. Materials Research Society Bulletin 32, 808-20. Loera‐Quezada MM, Leyva‐González MA, Velázquez‐Juárez G, Sanchez‐Calderón L, Do Nascimento M, López‐Arredondo D, Herrera‐Estrella L, 2016. A novel genetic engineering platform for the effective management of biological contaminants for the production of microalgae. Plant Biotechnology Journal 14, 2066-76. Mayet M, 2000. Analysis of South Africa’s GMO Act of 1997. Biowatch South Africa. https://acbio.org.za/wp-content/uploads/2015/02/SA_GMO_Leg_short.pdf. Meticulous, 2018. Algae Products Market– Global Opportunity Analysis And Industry Forecast (2017- 2022). Meticulous Research®. www.meticulousresearch.com/product/algae-products-market- forecast-2022/. Muñoz CF, de Jaeger L, Sturme MH, Lip KY, Olijslager JW, Springer J, Wolbert EJ, Martens DE, Eggink G, Weusthuis RA, Wijffels RH, 2018. Improved DNA/protein delivery in microalgae–A simple and reliable method for the prediction of optimal electroporation settings. Algal Research 33, 448-55. Murbach TS, Glávits R, Endres JR, Hirka G, Vértesi A, Béres E, Szakonyiné IP, 2018. A toxicological evaluation of Chlamydomonas reinhardtii, a green algae. International Journal of Toxicology 37, 53-62. Norsker NH, Barbosa MJ, Vermuë MH, Wijffels RH, 2011. Microalgal production—a close look at the economics. Biotechnology Advances 29, 24-7. Osundeko O, Ansolia P, Gupta SK, Bag P, Bajhaiya AK, 2019. Promises and challenges of growing microalgae in wastewater. Water Conservation, Recycling and Reuse: Issues and Challenges. Springer, 29-53.

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Rahman A, Putman RJ, Inan K, Sal FA, Sathish A, Smith T, Nielsen C, Sims RC, Miller CD, 2015. Polyhydroxybutyrate production using a wastewater microalgae based media. Algal Research 8, 95-8. Rocha RP, Machado M, Vaz MG, Vinson CC, Leite M, Richard R, Mendes LB, Araujo WL, Caldana C, Martins MA, Williams TC, 2017. Exploring the metabolic and physiological diversity of native microalgal strains (Chlorophyta) isolated from tropical freshwater reservoirs. Algal Research 28, 139-50. Rosenberg JN, Oyler GA, Wilkinson L, Betenbaugh MJ, 2008. A green light for engineered algae: redirecting metabolism to fuel a biotechnology revolution. Current Opinion in Biotechnology 19, 430-6. SANPCC. Aquaculture and bioremediation. Proceedings of the 31st Congress of The Phycological Society of Southern Africa, 2018: Phycological Society of Southern Africa. Sayre RT, Wagner RE, Siripornadulsil S, Farias C, 2017. Transgenic algae for delivering antigens to an animal. U.S. Patent No. 7,410,637. Sharma Ak NM, Sparstad T, Bones Am and Winge P, 2018. Transgene-free genome editing in marine algae by bacterial conjugation – comparison with biolistic CRISPR/Cas9 transformation. Scientific Reports 8, 14401. Specht E, Miyake-Stoner S, Mayfield S, 2010. Micro-algae come of age as a platform for recombinant protein production. Biotechnology Letters 32, 1373-83. Sydney EB, Sydney ACN, De Carvalho JC, Soccol CR, 2019. Potential carbon fixation of industrially important microalgae. Biofuels from Algae. Elsevier 2, 67-88. Taunt HN, Stoffels L, Purton S, 2018. Green biologics: The algal chloroplast as a platform for making biopharmaceuticals. Bioengineered 9, 48-54. Wang Z, Zhang D, Ma Z, 2018. South Africa Report on Science, Technology and Innovation. BRICS Innovative Competitiveness Report 2017. Springer, 197-217. Yamano T, Iguchi H, Fukuzawa H, 2013. Rapid transformation of Chlamydomonas reinhardtii without cell-wall removal. Journal of Bioscience and Bioengineering 115, 691-4. Zhuang LL, Yu D, Zhang J, Liu FF, Wu YH, Zhang TY, Dao GH, Hu HY, 2018. The characteristics and influencing factors of the attached microalgae cultivation: a review. Renewable and Sustainable Energy Reviews 94, 1110-9.

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Appendices Supplementary materials of Chapters 2-3

Appendices

Appendix A Cloning vectors

Plasmid and vectors used in this study to facilitate the cloning of the codon optimised Crgfp gene into the pChlamy4 vector. The Crgfp gene, from the pCrGFP plasmid (Figure A1), was PCR amplified and ligated into the intermediate pBluescript II SK(+) (pBSK) plasmid (Figure A2) for blue-white selection of positive transformants in Escherichia coli. The Crgfp gene was then restriction-digest cloned into the pChlamy4 vector (Figure A3), to form the final Crgfp-pChlamy4 plasmid (Figure A3 and A4) which was electro-transformed into the nuclear region of the Chlamydomonas reinhardtii CC-125 and CC-400 strains.

Figure A1: A schematic representation of the pCrGFP Figure A2: A schematic representation of the pBSK plasmid plasmid containing the C. reinhardtii optimised Crgfp. showcasing the MCS and vector features (Addgene, 2017). The pCrGFP plasmid contains the codon optimised CrGFP and a 3'-UTR of the rbcS2 gene from C. reinhardtii ligated into the MCS of the vector pUC19 (Entelechon GmbH, Regensburg, Germany)

Figure A3: Schematic diagram of the pChlamy4 plasmid generated by Snapgene illustrating the enzyme recognition sites within its MCS and the primers (Snapgene, 2017).

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Figure A4: A schematic representation of the pChlam4 vector features (Thermo Fisher Scientific, Massachusetts, United States).

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Appendix B PCR and restriction digest agarose gels

The agarose gels below provide evidence for the successful cloning process of the Crgfp transgene into the pChlamy4 plasmid. The gradient PCR results of the Crgfp gene show the optimum annealing temperature of 70˚C (Figure B1) for subsequent PCRs. The pChlamy4 vector was validated through restriction digestion (Figure B2). Incomplete digestion was seen for the single digests in lanes 2 and 3. In lane 4, the double restriction digest of pChlamy4 with HindIII and BamHI depicts DNA bands at the expected sizes of 2,553 bp and 1,087 bp. Colony PCR of the Crgfp gene region specified faint bands of the of the positive Crgfp-pChlamy4 plasmid clones (Figure B3).

Figure B1: KAPA HiFi Gradient PCR gene amplification Figure B2: Restriction enzyme double digest analysis of Crgfp from the pCrGFP plasmid. A temperature gradient pChlamy4 vector with HindIII and BamHI. M = 1 kb gel using CrGFP-For and CrGFP-Rev primers a range 60- GeneRuler DNA ladder (Thermo Fisher). The restriction 70°C annealing temperatures on a 1% agarose gel. M = 1 digests were run for pChlamy4 with no plasmid control kb GeneRuler DNA ladder (Thermo Fisher). The annealing (1), single digest with BamHI (2), single digest with HindIII temperatures were 70°C (1). 68°C (2), 65°C (3), 63°C (4), (3), double digested with both HindIII and BamHI (4). 60°C (5). No amplification can be viewed in the ddH20 Lanes 2 and 3 indicate linearized vector at 3,640 bp with control lane (6) run at 68°C indicating no contamination. A some undigested vector, and lane 4 shows the expected DNA band can be seen for each gradient at the expected bands of 2,553 and 1,087 bp. Crgfp size of 727 bp and a faint band at 6,000 bp can be seen in lane 5 and 4,500 bp for lanes 2,3 and 4, indicative of visible template DNA. Lane 1 had no nonspecific amplification.

Figure B3: Colony PCR gene amplification of the Crgfp gene in the Crgfp-pChlamy4 plasmids with primers CrGFP-For and CrGFP-Rev, on a 1% agarose gel. M = 1 kb GeneRuler DNA ladder (Thermo Fisher). No amplification is noted in the ddH20 control lane (1) indicating no contamination. The desired amplified Crgfp PCR product was visible at the expected size of 717 bp for the positive control (2) and recombinant colonies (3,4,5,7,8,11 and 12). No amplification in lanes 6, 9, 10, 13 and 14 indicate the absence of the Crgfp gene.

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Appendix C Selection plates of the Crgfp cloned into E. coli

The recombinant Crgfp-pBSK E. coli DH10B cells were selected by blue-white selection on IPTG-X-gal plates, and ampicillin resistance (Figure C1). The recombinant Crgfp-pChlamy4 E. coli DH10B cells were selected by using ampicillin resistance alone (Figure C2).

For blue-white screening, the pBSK phagemid vector was used for cloning the Crgfp purified PCR product. The plasmid not only contains an ampicillin resistance gene but also the lacZ gene, which encodes a fragment of the α-peptide- 146 amino acids of the enzyme β-galactosidase. This normally results in a blue colony when grown on IPTG and X-gal. The bacterial colonies containing the Crgfp- pBSK plasmid were grown on selective medium containing ampicillin, the colourless chromogenic compound X-gal (5-bromo-4-chloro-3indolyl-β-d-galactoside) and an inducing agent IPTG (Isopropyl β-D-1-thiogalactopyranosid). When X-gal is used as a substrate, the functional β-galactosidase catalyses the hydrolysis of X-gal, converting the colourless substrate into a blue coloured product and blue colonies result. Within the lacZ gene coding sequence is a MCS which can be used for inserting DNA fragments during cloning. When a foreign DNA sequence is inserted into the MCS, the coding sequence of the α-peptide is disrupted. This would produce non-functional β-galactosidase which would be unable to catalyse the hydrolysis of X-gal and the resulting colonies are white. Therefore, negative (blue) and positive (white) recombinant transformants were easily distinguished. When combined with ampicillin selection, the white colonies indicate successful cloning of the Crgfp gene into the pBSK plasmid (Alting-Mees et al., 1995).

Figure C1: Recombinant E. coli DH10B containing Crgfp- Figure C2: Recombinant E. coli DH10B containing CrGFP- pBSK, after colony PCR. LB agar plate showing the result of a pChlamy4 after colony PCR. LB agar plate of transferred blue-white screen using X-gal, IPTG and ampicillin with recombinant CrGFP-pChlamy4 cells on solid LB agar recombinant Crgfp-pBSK cells (white) and non-recombinant containing ampicillin. cells (blue).

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Appendix D Chlamydomonas CC-125 and CC-400 colony PCR screen for Crgfp transformants

The transformed C. reinhardtii colonies selected on the TAP agar zeocin plates were screened using colony PCR to identify positive transformants. The C. reinhardtii CC-125 strain, colony PCR analysis (Figure D1) indicates that out of 109 colonies, 80 were positive indicative of a transformation efficiency of 73.4%.

The C. reinhardtii strain,CC-400 colony PCR screen (Figure D2) provides evidence that out of the 111 colonies evaluated, 97 were positive which is indicative a transformation efficiency of 87.4%.

Figure D1: Colony PCR of Crgfp gene within transformed C. reinhardtii CC-125 cells on a 1% agarose gel. M= 1 kb GeneRuler DNA ladder (M, Thermo Fisher). In each agarose gel, the desired DNA band is visible at the expected size of 717 bp seen in positive control lanes containing Crgfp genomic DNA (1) No amplification can be seen in the no template control lanes (2) indicating no contamination. Positive transformants are visible in the remaining lanes by the expected 717 bp band, and negative transformants by the lack of one.

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Figure D2: Colony PCR of Crgfp within transformed C. reinhardtii CC-400 cells on a 1% agarose gel. M= 1 kb GeneRuler DNA ladder (M, Thermo Fisher). In each gel, the desired PCR product is visible at the expected size of 717 bp including the control lanes containing Crgfp genomic DNA (1) No amplification is visible in the no template control lanes (2) indicative of no contamination. Positive transformants are visible in the remaining lanes by the expected 717 bp band, and negative transformants by the lack of one.

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Appendix E Microalgae streaked on TAP zeocin plates for selection

Throughout this study microalgae were cultured on TAP plates with and without zeocin. For selection, screening and maintenance transformed C. reinhardtii colonies were streaked onto TAP 15% agar plates containing zeocin. Plating provided an easy method to maintain cultures after transformation with growth observed after 2 weeks and maintained for approximately 5 weeks, before drying out (Figure E1).

During growth and maintenance of these streaked plates, microalgae culturing observations were made, with careful consideration when placing agar plates in the laboratory. With a shared laboratory environment- plates were often stacked or moved from their position in the light. Interestingly the C. reinhardtii CC-125 screen plate, (Figure E2), was stacked at the bottom where not enough light was able to penetrate to the culture. The microalgae lost its green colour, illustrating a reduction in chlorophyll. The solid agar plates contained acetate and therefore the C. reinhardtii CC- 125 strain was able to continue growth despite the lack of light (Taghavi & Robinson, 2016).

Similar observations were noted for the C. reinhardtii CC-400 plate, (Figure E3), which started to lose chlorophyll towards the borders of the culture. It is also important to note that it is critical invert plates when being cultured. Incorrect storage could cause condensation related streaking across a plate resulting in individual colonies being mixed (Figure 3E).

Figure E1: After transformation and Figure E2: Chlorophyll deficiency Figure E3: Cell colony mixing due to colony PCR, positive Crgfp colonies seen in C. reinhardtii strain CC-125 condensation in C. reinhardtii strain of C. reinhardtii CC-125 and CC-400 cultured under dark conditions, CC-400, plated on TAP agar medium were transferred onto TAP agar plated on TAP agar medium containing 5 µg/mL zeocin. medium containing 5 µg/mL zeocin. containing 5 µg/mL zeocin.

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Appendix F

Fluorescence of Chlamydomonas untransformed and Crgfp transformed cultures

To ensure the untransformed C. reinhardtii strains did not autofluoresce, the cultures were evaluated under UV light with the DFP Flashlight-1 (NIGHTSEA, Lexington). Only the red autofluorescence of the chlorophylls can be seen for the untransformed C. reinhardtii CC-125 (Figure F1) and C. reinhardtii CC-400 strains (Figure F2). Larger cultures were used to evaluate the microalgae growth and stability of nuclear integration after 6 weeks (Figure F3).

Figure F1: The untransformed C. reinhardtii CC-125 strain Figure F2: The untransformed C. reinhardtii CC-400 strain under normal (left) and UV (right) light. under normal (left) and UV (right) light.

Figure F3: C. reinhardtii CC-400 strain untransformed (fluorescing red) and transformed (fluorescing green) cells viewed by a DFP UV Flashlight-1.

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Appendix G Standard curve of rGFP

A standard curve was created using commercial recombinant GFP which has a known protein concentration (Figure G1). The standard GFP was used to quantify the CrGFP protein produced within the CrGFP transformed C. reinhardtii CC-125 and CC-400 cultures.

350000

300000

250000

200000 RFU Linear (RFU) 150000

100000 R² = 0,9955

50000 y = 2097,3x + 2443,8 Relative Relative Fluorescent Units (485 nm/535 nm)

0 0 20 40 60 80 100 120 140 160 GFP (ng/µl)

Figure G1: The standard concentration curve of purified E. coli recombinant GFP (Sigma-Aldrich, South Africa), measured on a HIDEX microplate reader. Dilutions were made using TAP medium up to a final volume of 100 ul/ well. The fluorescence was determined using a 485 nm, 10 nm excitation filter and a 535 nm, 20 nm emission filter. Goodness of fit linear line of regression, R2= 0,996.

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Appendix H Microscopic identification of the 40 microalgae isolates

From the MiCCSA database 40 microalgal isolates were chosen based on compound microscopy of desired structural and cellular characteristics (Table H1). These isolates were all unicellular, with large or many chloroplasts and a thin cell wall. They also had a reduced gelatinous sheath and limited cellular aggregation. The potential genus of the 40 microalgal isolates were estimated based on their morphology. The morphological descriptions of the nine microalgae identified in this study are further discussed as well as the limitations to microscopic identification. Compound microscopy also identified pallmeloid formation in C. reinhardtii CC- 125 cultures under stress (Figure I1).

Table H1: The 40 South African microalgal isolates from the CSIR culture collection screened in a growth curve in this study.

Province Microscopic image Shape of cell Cell wall or Chloroplasts Average Congruency Potential genus code and (magnification: 100x with 20 μm filamentous sheath approximate isolate scale line) size of number diameter (μm)

FSA Round Thin smooth cell One large 03-06 Congregation of two Chlorella 2.2 wall green, or more cells parietal, pyrenoid present

FSA Round Very thin smooth Many small 21-26 Singular Volvox 8.1 cell wall

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FSA Round Apparent cell wall One large cup 6-10 Singular Chlamydomonas 17.1 shaped; pyrenoid present

FSA Ovoid Thick and smooth Many small 10-12 Congregation in pairs Oocytis 21.1 cell wall

FSA Round Firm cell wall One large 03-06 Singular Chlorella 22.3

FSA Round Smooth cell wall Large parietal 03-06 Singular Chlorella 33.1

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FSA Round Thin cell wall, thick Many 10-20 Singular Volvox 42.1 mucilage envelope chloroplasts

FSA Round Apparent cell wall One large cup 05-10 Singular Chlamydomonas 44.4 shaped; pyrenoid present

FSB Ovoid Thin, transparent Many small 04-06 Singular Oocytis 18.1 with mucilage with envelope pyrenoids present

FSB Round Thin cell wall, thin One large 06-20 Singular Pandorina 23.1 undefinitive mucilage envelope shape

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FSB Round Smooth, apparent One large 04-06 Singular Chlorella 24.1 cell wall with mucilage envelope

FSB Round-ovoid Thin cell wall thicker One or few 10-12 Two or more cells Oocystis 27.5 “lemon at polar ends large, congregate (could be shaped” pyrenoid dividing) present in some

FSB Sigmoid Thin transparent cell Two large 17-20 Singular Selenustrum 40.1 wall

FSB Round-ovoid Thin cell wall thicker One or few 05-07 Two colonies Oocystis 41.2 “lemon at polar ends large congregate shaped”

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GPA Ovoid Smooth firm cell One large 02-07 Singular Chlorella 1.2 wall

GPA Sigmoid Thin cell wall Many small; 07-10 Singular Scenedesmus 9.1 clearly visible pyrenoids

GPA Round- ovoid Transparent One or few 03-07 Cells congregate Coelastrum 9.3 undefinitive gelatinous layer large

GPA Round Thin, apparent cell One large, in 04-07 Singular but some Chlorella 14.1 wall with mucilage some cells congregate envelope singlepyrenoid clearly visible

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GPA Round Thin apparent cell One large 03-05 Singular Chlorella 18.1 wall with mucilage envelope

GPA Ovoid Thin apparent cell One or few 06-08 Singular Oocytis 19.1 wall large

GPA Round-ovoid Thin cell wall, One or few 07-12 Gelatinous colonies Cosmarium 24.2 Undefinitive surrounding large gelatinous matrix

GPA Sigmoid Thin, delicate cell Many medium 03-11 Singular Chlorogonium 25.1 wall

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LPA Ovoid Thin “wrinkled’ cell Large cup- 09-14 Singular Pandorina 4.2 wall with broad shaped zone of clear chloroplast mucilage

LPA Sickle shape Thin cell wall Single 10-13 Singular Closterium 7.1 chloroplast with many scattered pyrenoids

LPA Ovoid Transparent Large cup- 04-15 Singular with two or Pandorina 7.3 mucilage envelope shaped more cells congregate chloroplast (could be dividing)

LPA Undefinitive Gelatinous matrix Many 10-17 Congregated within Coelastrum 15.2 gelatinous layer

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LPB Ovoid Transparent One large 10-14 Singular or Scenedesmus 14.1 gelatinous layer parietal; congregate in colonies clearly visible of 2-4 pyrenoids

LPB 52.3 Cylindrical Transparent One large 10-14 Congregate Scenedesmus gelatinous layer parietal; in colonies of 2-8 clearly visible pyrenoids

LPB Elongated Transparent Single, large 07-10 Singular Scenedesmus 58.3 cylindrical gelatinous layer and plate like

MP Round-ovoid Apparent cell wall, One large 05-15 Congregate Coelastrum 53.1 transparent gelatinous layer

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NC Round Thin cell wall One large; 04-06 Singular Chlorella 2.3 pyrenoid present

NC Round Thin cell wall One large, 03-07 Singular Chlorella 7.1 cupshaped

NC Round Thin cell wall One large; 03-10 Singular Chlorella 37.5 one or more pyrenoids present

NC Round Smooth cell wall One large cup 03-06 Singular Chlorella 37.7 shaped

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NC Round Thin gelatinous layer Large 08-10 Singular Chlorella 72

NWA C-shaped Mucilage envelope Parietal 07-10 Singular Kirchneriella 23.4

NWA Round-oval Transparent One large; 05-08 Singular Oocystis 27.1 undefinitive gelatinous layer one pyrenoid present

WCA Round Thin cell wall One large 04-07 Singular Oocystis 4.1

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WCC Round Thin cell wall Many 05-07 Singular Chlorella 1.1

WCC Round Thin cell wall One large; red 04-05 Singular Duniella 6.1 eyespot visible

WCC Cylindrical Apparent cell wall Single large, 06-11 Singular Scenedesmus 21.2 plate like

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Morphological descriptions of microalgae identified in this study

Chlorella sp. The Chlorella spp. are coccoid single spherical, flattened or spindle-shaped cells ranging from 1-6 μm, they lack flagella which renders them immotile. Usually only one thin chromatophore is present and may be structured as a spherical cup shape green in colour and their pyrenoid may be visible or not (Krauss and Shihira, 2010).

Raphidocelis subcapitata The unicellular coccoid green alga, R. subcapitata has cells which are symmetrically crescent-shaped, and the outer convex walls may differentiate by size and curvature. Their length varies between 8 and 14 µm, with a width range of 2 and 3 µm. Normally these cells are singular but may form close clusters from 4 to 32 cells free of gelatinous envelopes (Prescott, 1954).

Tetradesmus sp. Tetradesmus spp. exist as nonmotile unicells; and are often found in coenobia of four or eight cells within a trilaminar sheath composed of algaenans (Lürling, 1999).

Microscopic limitation and future considerations In this study the stage of growth of was not accounted for during the microscopy investigation. The growth phase could indicate whether they are adapting, dividing, in stationary phase or dying which could affect their morphology. Cell cycle could influence the congregation, size and colour of the microalgae. For example, the average size of C. reinhardtii cells are about 10 µm, but significant variation in cell size may be observed depending on its growth phase (Harris, 2009). In this study microalgae in different cell cycles were detected, for example, the isolate LPA 15.2, appears to be a mother cell busy dividing which renders it larger than usual. To accurately account for morphological differences associated with growth phase, the microalgae should perhaps be counted in the future so that they may be microscopically viewed under the same cellular density and therefore growth phase.

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Appendix I Growth analysis of South African microalgal isolates

The 40 selected South African microalgae isolates underwent a growth study analysis. The OD750 was measured under different culturing conditions (Figures I1-5). After growth optimisation, the physiological growth comparison for the nine best growing microalgae isolates grown on either AF6 or TAP medium were evaluated (Figure I6). A calibration curve was created for the nine South African microalgae isolates which grew the best, along with C. reinhardtii controls, CC-400 and CC- 125, expressing the cell concentration of each isolate at an absorbance of 750 nm (Figure I7).

The discrepancy between microalgae photobleaching grown within 6 well microplates (Figure I8) compared to Erlenmeyer flasks (Figure I9) is discussed as a result of light saturation (Figure I10). To further culturing techniques, compound microscopy identified pallmeloid formation in C. reinhardtii CC-125 cultures under stress (Figure I11) compared to natural culture characteristics.

0,25 FSA 2.2 FSA 21.1 FSA 22.3 0,2 FSA 33.1 FSA 42.1 FSB 18.1 0,15 FSB 23.1 FSB 27.5 FSB 40.1 FSB 41.2 0,1

OD OD nm) (750 GPA 18.1 GPA 14.1 GPA 9.3 0,05 GPA 1.2 GPA 9.1 GPA 25.1 0 GPA 24.2 Day1 1 Day2 2 Day3 3 Day4 4 Day5 5 GPA 19.1 LPA 7.1 Time (Days) NC 11.6

Figure I1: Growth curve analysis of 20 South African green microalgae cultured with 250 μmol m-2.s- 1 continuous light, with 10% inoculum. The cells were cultured in AF6 media and the OD750 was measured every day for 5 days before growth appeared stagnant. The microalgal isolates which grew the best were isolates GPA 18.1, GPA 9.3 and FSA 33.1. The values represent the mean ± standard deviation (n = 3).

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0,35 FSA 2.2 GPA 1.2 0,3 FSB 41.2 FSA 33.1 FSA 42.1 0,25 FSB 18.1 FSB 23.1 0,2 FSB 27.5 FSB 40.1 0,15 GPA 9.3

OD OD (750nm) GPA 18.1 0,1 GPA 14.1 FSA 22.3 FSA 21.1 0,05 GPA 9.1 GPA 25.1 0 GPA 24.2 Day1 1 Day2 2 Day3 3 Day4 4 GPA 19.1 LPA 7.1 Time (Days) NC 11.6

Figure I2: Growth curve analysis of 20 South African green microalgae cultured with 100 μmol m-2.s-1 continuous light, with 10% inoculum. The cells were cultured in AF6 media and the OD750 was measured every day for 4 days before growth appeared stagnant. The microalgal isolates which grew the best were isolates GPA 9.3, GPA 18.1 and FSA 22.3. The values represent the mean ± standard deviation (n = 3).

0,45 FSA 2.2 FSA 21.1 0,4 FSA 22.3 0,35 FSA 33.1 FSA 42.1 0,3 FSB 18.1 FSB 23.1 0,25 FSB 27.5 0,2 FSB 40.1

FSB 41.2 OD OD nm) (750 0,15 GPA 18.1 0,1 GPA 14.1 GPA 9.3 0,05 GPA 1.2 GPA 9.1 0 GPA 25.1 Day1 1 Day2 2 Day3 3 Day4 4 Day5 5 Day6 6 Day7 7 Day8 8 Day9 9 Day10 Day11 Day12 GPA 24.2 10 11 12 GPA 19.1 Time (Days) LPA 7.1 NC 11.6

Figure I3: Growth curve analysis of 20 South African green microalgae cultured with 250 μmol m-2.s-1 continuous light, with 20% inoculum. The cells were cultured in AF6 media and the OD750 was measured every day for 12 days before growth appeared stagnant. The microalgal isolates which grew the best were isolates GPA 18.1, FSA 22.3 and GPA 9.3. The values represent the mean ± standard deviation (n = 3).

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0,8 FSA 2.2 FSA 21.1 0,7 FSA 22.3 FSA 33.1 0,6 FSA 42.1 FSB 18.1 0,5 FSB 23.1 FSB 27.5 0,4 FSB 40.1 FSB 41.2

OD OD (750nm) 0,3 GPA 18.1 0,2 GPA 14.1 GPA 9.3 0,1 GPA 1.2 GPA 9.1 0 GPA 25.1 Day1 1 Day2 2 Day3 3 Day4 4 Day5 5 Day6 6 Day7 7 Day8 8 Day9 9 10Day 11Day Day12 GPA 24.2 10 11 12 GPA 19.1 Time (Days) LPA 7.1 NC 11.6

Figure I4: Growth curve analysis of 20 South African green microalgae cultures with 8-hour light (250 μmol m-2.s-1)/ 16 hour dark phase cycles, with 20% inoculum. The cells were cultured in AF6 media and the OD750 was measured every day for 12 days. The microalgae isolates which grew the best were: FSA 22.3, GPA 9.3, GPA 9.1, GPA 19.1, FSB 18.1 and GPA 18.1. The values represent the mean ± standard deviation (n = 3).

0,9 WCC 1.1 WC 4.1 0,8 WCC 6.1 0,7 WCC 21.2 FSA 2.2 0,6 GPA 14.1 NC 2.3 0,5 NC 5.1 0,4 NC 37.5

NC 37.7 OD OD nm) (750 0,3 NC 72 NWA 23.4 0,2 LPA 4.2 0,1 LPA 7.3 LPA 15.2 0 LPB 12.1 Day1 1 Day2 2 Day3 3 Day4 4 Day5 5 Day6 6 Day7 7 Day8 8 Day9 9 Day10 Day11 Day12 LPB 14.1 10 11 12 LPB 60.1 Time (Days) LPB 58.3 MP 53.1

Figure I5: Growth curve analysis of 20 South African green microalgae cultures with 8-hour light (250 μmol m-2.s-1)/ 16 hour dark phase cycles, with 20% inoculum. The cells were cultured in AF6 media and the OD750 was measured every day for 12 days. The microalgal isolates which grew the best were the isolates NC 72, WC 4.1, WC 1.1 and LPB 14.1. The values represent the mean ± standard deviation (n = 3).

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Figure I6: Stock cultures of the nine selected South African microalgae isolates and C. reinhardtii CC-125 and CC-400 strains grown on AF6 and TAP media after four weeks under 250 μmol m-2.s-1 continuous light. These nine microalgae were the isolates identified as those which grew the best in our growth studies; which correlates to the visual inspection- the cultures are far denser and healthier.

25 Tetradesmus dimorphus GPA 19.1 Tetradesmus obliquus LPB 14.1/ 20 GPA 9.1 Raphidocelis subcapitata NWA 23.4 15 Chlamydomonas

reinhardtii CC125

cells/mL 6

10 Chlamydomonas reinhardtii CC400

Chlorella vulgaris NC 72

Cell Cell Numberx10 5

Chlorella sp. FSA 22.3 0 0 0,1 0,2 0,3 0,4 0,5 0,6 0,7 0,8 0,9 1 1,1 1,2 1,3 1,4 1,5 Chlorella sorokiniana GPA 18.1 -5 Chlorella sp. WC Optical Density (750 nm) 4.1

Figure I7: The calibration curve for the South African microalgal isolates and the C. reinhardtii controls (CC-400 and CC- 125) expressing the cell concentration of each isolate at an absorbance of OD750. The linear regression value for each isolate was as followed: Isolate R. subcapitata NWA 23.4 R²= 0,8913; T. obliquus LPB 14.1 R²= 0,8953; T. obliquus GPA 9.1 R²= 0,9335; T. dimorphus GPA 19.1 R²= 0,9322; C. reinhardtii CC125 R²= 0,9394; C. reinhardtii CC400 R²= 0,9241; C. vulgaris NC 72 R²= 0,9843; Chlorella. sp. FSA 22.3 R²= 0,9786; C. sorokiniana GPA 9.3 R²= 0,9671; C. sorokiniana GPA 18.1 R²= 0,9715; Chlorella. sp. WC 4.1 R²= 0,9746.

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Growth experiments in microplates utilised light which easily penetrated the entire plate (Figure I8). However, the microalgal stock cultures were grown in 250 ml Erlenmeyer flasks under continuous 250 μmol –m.-2.s- 1 light exposure (Figure I9), and no photoinhibition occurred.

Figure I8: The set-up of the six well microplates under Figure I9: An experimental setup of batch cultures of for which most growth curve experiments were performed. determining the biomass. Stock cultures were also maintained in these larger, Erlenmeyer flasks.

The successful growth of the microalgae within the Erlenmeyer flasks can be explained by higher cell biomass densities, which affected the light path and created a photic and dark zone within the flasks (Figure I10). The photic zone is the outer layer of the flask most exposed to high light conditions. The algae in this zone have enough light exposure for photosynthesis. Just beyond the photic zone, is the dark zone, where the algae do not photosynthesize (Janssen, 2002). These flasks were continuously mixed, which allowed the microalgae cells to pass through both zones, undergoing both light and dark cycles. Therefore, photoinhibition does not occur under continuous light at larger volumes in the Erlenmeyer flasks.

Figure I10: The light path in stock cultures experience limited photoinhibition due to the higher cell densities, resulting in two zones, the photic zone (light saturated) and the dark zone (no photosynthesis) (Stapelberg, 2018).

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Distinguishing between contamination and culturing characteristics The physiology of green microalgae cells change when they undergo stress caused by biotic factors, such as contamination or the biotoxins these contaminants produce (Khona et al., 2016). The C. reinhardtii stress response has been well studied with three phases on the physiological level. The first and most instant response is the loss of the flagella (Hessen et al., 1995). Succeeding this, the cells form palmelloids where they revert from “free-living” to a “colonial” state (Lurling & Beekman, 2006) until finally cell death transpires (Pérez-Pérez et al., 2010). In our study, the cultures in the presence of contaminants were indeed under stress with the confirmation of the cells aggregating and forming palmelloids (Figure I11). Palmelloids are varying number of cells clustered within a common membrane. For protection they excrete exopolysaccharides, undergo abnormal cell division and thicken their individual cell walls (Khona et al., 2016). The C. reinhardtii strain, CC-400, had a higher mortality under contamination, which could be contributed to its lack of a cell wall (Taghavi & Robinson, 2016). It is imperative that the cultures are kept healthy and free of contaminants, salt, temperature or light stress to avoid cellular morphological changes which could hinder biomass accumulation or the transformation process.

Figure I11: Pallmeloid formation in the C. reinhardtii CC-125 culture due to contamination-related stress. Scale bar = 50 µm.

It is also important to differentiate contamination and stress with the physiology of cells when they are transferred onto solid media. In liquid media, C. reinhardtii is incredibly motile but upon culturing on solid agar media C. reinhardtii cells become deflagellated, immobilised and appear “soupy” (Khona et al., 2016). A gelatinous sheath forms around the colonies which does not indicate contamination but is considered a natural occurrence in Chlorophytes (Khona et al., 2016).

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Appendix J Genomic DNA extraction and PCR of 18S rDNA gene

To identify the South African microalgae, Sanger sequencing of the 18S rDNA region required genomic DNA. DNA was extracted by varies methods from the South African microalgal isolates. These include the CTAB protocol (Fawley & Fawley, 2004), the CTAB protocol modified with and without lysis glass bashing beads, Proteinease K and a column shredder. The following DNA extraction kits were also tested: ZR Plant/ Seed DNA MiniPrep (Zymo Research, California, United States), Quick gDNA MiniPrep (Zymo Research, California, United States), Invisorb Spin Tissue (Lasec, South Africa), DNeasy Blood & Tissue (Qiagen, Germany), Plant DNA minikit (Qiagen, Germany) and the DNeasy PowerPlant Pro (Qiagen, Germany). The kit which provided the greatest quantity and quality of genomic microalgal DNA was that of Invisorb Spin Tissue kit (DSTR1032100100S, Stratec, Germany). The PCR results of this kit further indicated the best PCR product DNA concentration of 10 ng. The PCR products derived from this optimised reaction were utilised for Sanger sequencing (Figure J1).

B C

Figure J1: The 18S rDNA PCR amplification from the genomic DNA of nine South African algal isolates run on 1.5% agarose sodium borate gels. M= Fast DNA ladder (New England Biolabs). The DNA samples in (A) and (B) were amplified with a 3-set dilution series (10 µg, 1 µg, 0.1 µg) of template DNA for each. (A) GPA 9.1 (lanes 1-3), GPA 9.3 (lanes 4-6), GPA 18.1 (lanes 7-9) and GPA 19.1 (lanes 10-12). (B) GPA 9.3 (lanes 1-3). (C) LPB 14.1 (lane 1), NWA 23.4 (lane 2), FSA 22.3 (lane 3) and NC 72 (lane 4). The anticipated amplicon size is approximately 450 bp. The last lane of each gel (13/4/7) contained the ddH2O controls (no amplification). The 10 ng concentration resulted in the most distinct DNA band for each isolate and was used for downstream Sanger sequencing applications.

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