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Photopigments and functional carbohydrates from Cyanidiales Delicia Yunita Rahman, D.

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Chapter 1 Introduction

 Chapter 1

Role of microalgae in the global oxygen and carbon cycles

In the beginning there was nothing. At least there was no oxygen. For more than two billion 4 years, the Earth’s atmosphere was devoid of oxygen. The primordial atmosphere contained helium and hydrogen with traces of carbon dioxide, methane and ammonia (Pavlov and Kasting, 2002; Zahnle et al., 2006; Zerkle et al., 2012). The first oxygen-producing microorganisms appeared around 2.7 billion years (Rasmussen et al., 2008), while the first oxygen started to accumulate in the atmosphere some 2.4 to 2.1 billion years ago. It is widely accepted that cyanobacteria were the first oxygenic photosynthetic microorganism (Brocks et al., 2003 a and b). The initially produced oxygen was immediately bound by the vast amounts of free iron in the ocean to form iron oxides that precipitated at the sea floor, resulting in what is called the Banded Iron Formations. Only when all the free iron was bound, oxygen escaped from the ocean water and atmospheric oxygen levels began to rise. This sudden increase in atmospheric oxygen levels is known as the Great Oxidation Event (Lyons et al., 2014) leaving clear fingerprints in rock records. This was also the time of the first mass extinction, as all Life known so far consisted of anaerobic microorganisms for which oxygen is toxic.

Some 1.2 to 1.1 billion years ago, the first eukaryotic microalgae (Rhodophyta) occurred in freshwater lakes, contributing to a further rise of atmospheric oxygen levels (Raven et al., 2012). A significant oxygen increase occurred in the Neoproterozoic era (0.54-1.0 billion years ago), driven by the oxygenation of the deep ocean. This era became a turning point in the development of the modern atmospheric oxygen system (Shield-Zhou and Och, 2011), as many oxygenic photosynthetic organism appeared: the Florideophyceae (phylum Rhodophyta) about 600 million years ago, followed by Ulvophyceae 540 million years ago and the Trebouxiphyceae 450 million years ago (Raven et al., 2012). It was only in the Devonian Period, (416-397 million years ago) that plants evolved, contributing to a further rise in atmospheric oxygen levels. Microalgae, cyanobacteria, and plants are the current day oxygen producers, maintaining atmospheric oxygen levels at 21 percent (Shield-Zhou and Och, 2011).

 Introduction

Microalgae also play a dominant role in the carbon cycle, fixing more than 100 Gigatonnes of carbon dioxide each year. It is estimated that around 40% of the carbon dioxide fixation on our planet occurs in the upper part of the oceans (Falkowski, 1994), whereas land plants account for only 15% of all carbon dioxide fixed each year (Beer et al., 2010). The 5 prochlorophyte Prochlorococcus marinus is one of the most abundant phototrophs in the open ocean (Chisholm et al., 1988; Karl, 2002), contributing between 9 and 82% to the net primary production of the Pacific Ocean (Liu et al., 1997). P. marinus is probably the most abundant photosynthetic organism on our Planet (Partensky et al., 1999). In contrast to oxygen, carbon dioxide has always been present in the Earth’s atmosphere. The current days levels of carbon dioxide of 400 ppm are low compared to levels of 6,000 ppm some 600 to 400 million years ago or the 3,000 ppm present 200 to 150 million years ago (Pearson and Palmer, 2000). From 60 million years ago, carbon dioxide levels have been falling to about 280 ppm in the mid of the 19th century.

As most photosynthesizing organisms evolved during times when the atmospheric carbon dioxide levels were high, Rubisco, which is the primary carbon dioxide fixing protein in all photosynthetic organism and thereby the most abundant protein on our Planet, has a poor affinity for carbon dioxide. The Km (a measure of the affinity) of the cyanobacterial Rubisco for carbon dioxide is about 200 µM. With the current day carbon dioxide levels, it is estimated that Rubisco operates at 30% or less of its maximum capacity (Moroney, 1999). In the addition, Rubisco also uses oxygen as a substrate. With the current day atmospheric levels of oxygen being much higher than carbon dioxide, this further contributes to Rubisco being a very inefficient carbon dioxide fixing enzyme.

Therefore, the challenge for all photosynthetic organisms is to concentrate the carbon dioxide at the site of the Rubisco. However, as membranes are permeable for carbon - dioxide this is not an easy task. Microalgae solve this by concentrating the HCO3 internally, thus limiting the diffusion through the membranes. However, Rubisco requires - carbon dioxide, so the HCO3 first has to be converted to carbon dioxide. This is done by the enzyme carbonic anhydrase, which is tightly packed with the Rubisco protein. In cyanobacteria the Rubisco and carbonic anhydrase are packed in so called carboxysomes, small, protein surrounded vesicle-like structures (Moroney, 1999; Price et al., 1992).  Chapter 1

Photosynthesis

Photosynthesis, the process in which light energy (sunlight) is converted into chemical 6 energy, is found in plants and photosynthetic microorganism including microalgae and cyanobacteria (Zhou and Su, 2014). Photosynthesis has existed and formed the Earth’s environment for more than 3.5 billion years. It is the driving process responsible for producing and maintaining the atmospheric oxygen concentrations, the foundation for all aerobic forms of life (Raven, 1997). Photosynthesis is driven by the capture of photons (light), which when absorbed by the light harvesting complex cause charge separation and electron ejection. The electrons then drive the dissociation of water generating protons/electrons and oxygen as a waste product. The protons and the associated electrons facilitate the reduction of carbon dioxide into organic compounds (Williams and Laurens, 2010; Razzak et al., 2013). The general equation for photosynthesis can be writing as:

6CO2 + 6H2O + photons → C6H12O6 + 6O2

The photosynthesis sequence contains two steps, (i) the light reaction, which only occurs when the cells are illuminated, and (ii) the dark reaction or light-independent reaction, also known as carbon-fixation, that occurs both in the presence and absence of light (Williams and Laurens, 2010; Ho et al., 2011). In eukaryotes, photosynthesis occurs in the chloroplast, a membrane enclosed organelle. The membranes contain stroma, an aqueous fluid that is composed of stacks of flattened disks bound by membranes called thylakoids. The thylakoids are the site of photosynthesis, containing protein complexes including pigments that absorb the light energy (Razzak et al., 2013).

In general, there are three different ways by which plants and algae assimilate CO2: (i) C3,

in which photosynthesis only uses the Calvin cycle to fix CO2 using Rubisco and taking place inside the chloroplast in mesophyll cells. Typical C3 plants are terrestrial plants including many important crops such as rice, wheat, and potato (Matsuoka et al., 2001); (ii)

C4, in which photosynthetic activities are partitioned between mesophyll and bundle sheath cells that are anatomically and structurally different. Maize, sugarcane, and sorghum are

examples of C4 plants; (iii) CAM (Crassulacean Acid Metabolism), in which the stroma are closed during the day and open only at night when the temperature decreases and humidity  Introduction

rises (Ruan et al., 2012; Wang et al., 2012). Microalgae use the C3 way of CO2 assimilation (Xu et al., 2012).

7 Light-dependent reaction photosynthesis

The first step of photosynthesis is the capture of light by photosystem II and converting photons to ATP and NADPH by photosystem I (Fig. 1A). The overall equation for the light- dependent reaction is:

+ + 2H2O + 2NADP + 3ADP + 3P + light → 2NADPH + 2H + 3ATP + O2

Light is absorbed by chlorophyll located in the thylakoid membrane followed by the release of electrons. Light energy is used to synthesize adenosine triphosphate (ATP) and nicotinamide adenine dinucleotide phosphate (NADPH) (Williams and Laurens, 2010; Klinthong et al., 2015). The synthesis of these molecules is done in a cyclic or a non-cyclic pathway. Microalgae and cyanobacteria mainly use the non-cyclic pathway (Razzak et al., 2013).

The electrons flow away from the chlorophyll molecules drawing electrons from water. This happens when the chlorophyll molecules at the core of the photosystem II have accumulated sufficient excitation energy from the antenna pigments. The Z-scheme generates a chemiosmotic potential across the membranes. During photophosphorylation, the ATP synthase enzyme uses the chemiosmotic potential to generate ATP. The NADPH generated in photosystem I is a product of the terminal redox reaction in the Z-scheme (Williams and Laurens, 2010; Razzak et al., 2013).

Light-independent reaction photosynthesis

The NADPH and ATP produced in light reaction are used in the enzymatic light- independent reaction, the part of photosynthesis that enables the incorporation of CO2 into organic material (Fig 1B). The enzyme ribulose bisphosphate carboxylase oxygenase

(Rubisco) plays a key role in capturing of the CO2 from the atmosphere (Williams and  Chapter 1

Lauren, 2010; Razzak et al., 2013). The light-independent reaction also known as Calvin cycle (Fig. 1B), has the overall reaction is:

+ + 8 3CO2 + 9ATP + 6NADPH + 6H → C3H6O3-phosphate + 9ADP + 8P + 6NADP + 3H2O

The first step is the carboxylation of sugar ribose 1:5 bisphosphate (Ru5BP) by Rubisco. Carboxylation of Ru5BP in the Calvin cycle generates two molecules of three-carbon compounds, 3-phosphoglyceric acid (3-PGA) or also called glycerate 3-phosphate (GP) (Klinthong et al., 2015). In the presence of ATP and NADPH, the GP molecules are phosphorylated to 1,3-bisphosphoglycerate and reduced to glyceraldehyde-3-phosphate (G3P). These set of three carbon compounds are the building block for the synthesis of all organic cellular compounds (Williams and Laurens, 2010).

A B

Figure 1. Photosynthesis in the light-dependent reaction (A) and the light-independent reaction (B) (Based on William and Laurens, 2010)

 Introduction

Photopigments

Photosynthesis is performed in different ways in different species, but the process always starts with light absorption by light-harvesting or photosynthetic pigments that capture light 9 and transfer this into chemical energy via photosynthetic electron transport (Barber, 2009; Mulders et al., 2014). Light harvesting pigments are divided into primary and accessory/secondary light harvesting pigments (Lemoine and Schoefs, 2010). Primary light-harvesting pigments are functionally and structurally bound to the reaction centre of the photosystem. Accessory light-harvesting pigments are more loosely attached to the outside of the photosynthetic apparatus, either intertwined with the primary light- harvesting pigments or sticking out of the system as antennae. For an efficient transfer of light energy, these accessory pigments need to be in close proximity to the primary light- harvesting pigments (Telfer et al., 2008; Lemoine and Schoefs, 2010). The constitution of the photosynthetic pigments gives the typical colour to microalgae and is one of phenotypic characteristics used to classified microalgae (Jeffrey et al., 2001).

Chlorophylls

The photosynthetic pigments of microalgae are grouped into three major classes based on their structural differences: chlorophylls, carotenoids, and phycobiliproteins. Chlorophylls consist of 5 aromatic rings, the chlorin part, surrounding a magnesium ion. Four of the five aromatic, heterocyclic rings are pyrroles containing carbon and nitrogen. To one of these pyrroles a diterpene alcohol or phytol is attached, aiding in the association of the chlorophyll molecule to a membrane surrounding (Mulders et al., 2014). There are five different types of chlorophyll: a, b, c, d, and f, with slight variation in the substituent on one of the pyrrole rings (Fig. 2) (Chen et al., 2010).

In chlorophyll b, a formyl group (CHO) replaces the methyl group (CH3) in the ring II of chlorophyll a (Chen et al., 2010), whereas chlorophyll d and f have a formyl group at resp. the C3 and C2 in the ring I (Chen et al., 2010; Willows et al., 2013). Chlorophyll c lacks the hydrocarbon tail (Mulders et al., 2014). These relatively small differences in the chlorophyll structures give significant differences in the absorption spectrum of each of the  Chapter 1

chlorophylls. Chlorophyll a absorbs light in the violet-blue and orange-red spectrum, reflecting light of the yellow-green spectrum, explaining its green appearance. Chlorophyll b absorbs mainly blue light giving it a brilliant green color. Chlorophyll c is yellow green, 10 chlorophyll d absorbs far red light resulting in a brilliant or forest green colour, and chlorophyll f absorbs light of the far red spectrum giving it an emerald green colour (Chen et al., 2010). Chlorophyll a is found in all photosynthetic organisms from microalgae to higher plants, as it is the primary light harvesting pigment. Chlorophyll b is found in the Chlorophyta and Euglenophyta, chlorophyll c is found in Diatoms, Cryptophyta, and Dinophyta, chlorophyll d in red algae and cyanobacteria (Begum et al., 2016). Chlorophyll f has been found in stromatolites, very ancient calcified microbial mats containing cyanobacteria (Holomicronema hongdechloris) found in Shark Bay, Western Australia (Willows et al., 2013).

Figure 2. Chemical structure of chlorophyll a

 Introduction

Chlorophyll biosynthesis

Chlorophyll biosynthesis is divided into four main parts: (1) the formation of 5- aminolevulinic acid (ALA), (2) the formation of protoporphyrin IX (Proto IX) from eight 11 molecules of ALA, (3) the Mg-porphyrin branch to chlorophyll, and (4) the heme- synthesizing branch (Papenbrock and Grimm, 2001). 5-aminolevulinic acid is the first universal tetrapyrrole precursor. Then ALA-dehydratase (porphobilinogen synthase) catalyses the asymmetric condensation of two ALA molecules to form monopyrrole porphobilinogen (Fig. 3 step 1-3), with the release of two molecules of H2O (Caterfranco and Beale, 1983). Four monopyrrole porphobilinogens are polymerized by hydroxymethylbilane synthase (also known as porphobilinogen deaminase and pre- uroporphyrinogen synthase) into the linear tetrapyrrole molecule, hydroxymethylbilane. After an immediate isomerisation, this linear tetrapyrrole molecule gives rise to the ring molecule uroporphyrinogen III. To synthesize the heme and chlorophyll, side chains of the porphyrin ring of uroporphyrinogen III are further decarboxylated and oxidized to form proto IX (Fig. 4, step 1-6) (Beale, 1999; Papenbrock and Grimm, 2001). Then Mg2+ is inserted into the multicomponent complex of protoporphyrin IX monomethyl ester (Fig. 4, step 7-8). The next steps are the formation of the isocyclic ring (Fig. 4, step 9-11), oxidation of the 6-methyl propionate group to a 6-methyl β-ketopropionate (Beale, 1999; Papenbrock and Grimm, 2001), attachment of α-methylene to the γ-meso bridge of the porphyrin, divinyl protochlorophyllide (Castelfranco and Beale, 1983), and reduction to chlorophyllide α by protochlorophyllide oxidoreductase (Fig. 4, step 12-13). The final step of chlorophyll a formation is esterification of the 7-propionic acid with the C20 polyisoprene alcohol phytol (Fig. 4, step 14) (Beale, 1999).

Figure 3. Earlier biosynthesis pathway of chlorophyll, formation of 5-Aminolevulinic acid (ALA). The enzymes involve in individual reaction are: (1) glutamyl-tRNA synthetase; (2) glutamyl-tRNA reductase; (3) glutamate 1-semialdehyde aminotransferase. Based on Beale, 1999; Papenbrock and Grimm, 2001.  Chapter 1

12

Figure 4. Chlorophyll biosynthesis pathway. The enzyme involved are: (1) porphobilinogen synthase; (2) hydroxymethylbilane synthase; (3) uroporphyrinogen III synthase; (4) uroporphyrinogen III decarboxylase; (5) coproporphyrinogen III oxidative decarboxylase; (6) protoporphyrinogen IX oxidase; (7) protoporphyrin IX Mg-chelatase; (8) S-adenosyl-L- methionine:Mg-protoporphyrin IX methyltransferase; (9-11) Mg-protoporphyrin IX monomethyl ester oxidative cyclase; (12) light-dependent NADPH: protochlorophyllide oxidoreductase; (13) divinyl (proto)chlorophyllide 4-vinyl reductase; (14) chlorophyll synthase. Based on Beale, 1999; Papenbrock and Grimm, 2001.  Introduction

Carotenoids

The second major class of photosynthetic pigments is the carotenoids. Carotenoids are long hydrocarbon chains (C40 backbone), made up of eight isoprene units to form a polyene 13 chain with conjugated double bond (Fig. 5). This structure establishes an extended p- electron system that accounts for the carotenoid’s ability to absorb ultraviolet radiation and visible light (Grossman et al., 2004). Carotenoids are divided into carotenes, which are unsaturated hydrocarbons, and xanthophylls, which present one or more functional groups containing oxygen (Cuellar-Bermudez et al., 2015). There are over 400 known carotenoids, but beta-carotene, lutein, and astaxanthin are the primary carotenoids found in microalgae, distributed within the chloroplast along with chlorophyll. Primary carotenoids such as lutein are an integral part of the light-harvesting complex of the photosynthetic systems and function as light absorber in the 400-550 nm region of the visible spectrum, in which chlorophyll does not perform efficiently. The carotenoids quench excess energy and expand the light-absorbing spectrum. In addition, carotenoids protect chlorophyll from photodamage (Mulders et al., 2014; Varela et al., 2015). These secondary carotenoids are made in large amounts and are present in oil droplets forming a protective layer in conditions of stress. These secondary carotenoids give the typical red or pink colour to microalgae, colouring snow, lakes and shallow water reddish.

Figure 5. Overall structure of carotenoids.

The carotenoid biosynthesis is located in the chloroplast while some specific steps take place in the cytoplasm (Valera et al., 2015). The biosynthesis steps are divergent among various organisms (Takaichi, 2011), however all the pathways initiate from a C5 building block, the common precursor of all isoprenoids, the isopentenyl pyrophosphate (IPP) or its isomer, dimethylallyl diphosphate (DMAPP) (Valera et al., 2015). These precursors are produced from either the cytosolic mevalonic acid (MVA) pathway (utilized acetyl-CoA) or the plastid methylerythritol 4-phosphate (MEP) pathway (utilizing pyruvate and G3P)  Chapter 1

(Gong and Bassi, 2016). The chain elongation is catalysed by prenyl transferase that

synthesizes geranyl PP (GPP, C10), farnesyl PP (FPP, C15), or geranylgeranyl PP (GGPP,

C20). These products are the precursor of mono-, di-, and tri- terpenes and carotenoids. The

14 FPP (C15) and the GGPP (C20) are the immediate precursor of C30 and C40 carotenoids (Paniagua-Michel et al., 2012: Varela et al., 2015).

Phycobilisome

Phycobilisomes (PBS) are large supramolecular complexes located on the outer surface of the thylakoids membrane serving as light-harvesting antenna that transfer energy to the photosynthetic reaction centre (Fig. 6) (Glazer, 1994: Chang et al., 2015). The light is captured by phycobiliproteins at wavelengths at which chlorophyll has low absorbance (Eisele et al., 2000). PBSs are composed of two, three or five stacked cylindrical subassemblies of allophycocyanin. Multiple phycocyanin molecules form the first layer of peripheral rods, the second layer is formed by multiple phycoerythrin molecules (Samsonoff and MacColl, 2001: Sun and Wang, 2011). The absorption maximum of allophycocyanin is 650 nm, of phycocyanin 620 nm, and phycoerythrin 540 nm (MacColl, 1998).

Figure 6. Model of a tricylindrical phycobilisome. The phycobilisome is shown attached to photosystem II. A tricylindrical core is shown, with two bottom cylinders attached to the thylakoid membrane. (Based on MacColl, 1998). AP: allophycocyanin, PC: phycocyanin, PE: phycoerythrin.  Introduction

Phycocyanin

Phycocyanin is actually composed of two components, a chromophore and a protein. The chromophore is a tetrapyrrole, known as bilin, which gives the typical blue colour to the 15 phycocyanin (Fig. 7). This chromophore is attached to a conserved cysteine amino acid residue (Cys-84) in the apoprotein by a thioether linkage (Troxler et al., 1981; Zhao et al., 2006). Next to this conserved binding site, there are additional binding sites in the apoprotein near the N-terminus at position 50 and the C-terminus at position 150. Besides these covalent linkages between the apoprotein and the chromophore, there are a number of non-covalent interactions that are essential for proper functioning of phycocyanin. Free biliproteins in solution take a flexible, helical conformation while native biliproteins assume a rigid, extended confirmation (Zhao et al., 2011). The apoprotein of phycocyanin consists of two homologous subunits, α and β that aggregate into a heterodimer, referred to as a monomer as this is the basic building block of the apoprotein. Three of these monomers aggregate into a triangular-shaped “trimer”, which is actually a heterohexamer, which can further aggregate into a heterododecamer/”hexamer” (12 subunits) (Schirmer et al., 1986). These hexamers are the structural unit of the phycocyanin in the phycobilisome complex.

Figure 7. Chemical structure of phycocyanobilin.

Phycocyanin is found in a range of cyanobacteria and several Rhodophyta (red algae) (Edwards et al., 1996). Cyanobacteria are better known as blue-green algae, as the abundance of phycocyanin (blue) and allophycocyanin (green) gives them a blue-green  Chapter 1

appearance. Examples of cyanobacteria producing phycocyanin are Spirulina platensis (Pushparaj et al., 1997; Jimenez et al., 2003), S. fusiformis (Minkova et al., 2003), Anabaena sp. (Moreno et al., 2003), Lyngbya sp. (Patel et al., 2005), and Nostoc sp. (Reis 16 et al., 1998). It has been reported that the freshwater cryptomonad Chroomonas sp. also produces a phycocyanin (Mörshel and Wehrmeyer, 1975). However, with an absorption maximum of 645 nm, the Chroomonas sp. pigment is actually an allophycocyanin. Rhodophyta producing phycocyanin are Porphyridium cruentum (Ma et al., 2003) Galdieria sulphuraria (Gross and Schnarrenberger, 1995), Cyanidium caldarium (Samsonoff and MacColl, 2001) and Cyanidioschyzon merolae (Albertano et al., 2000).

The protein part of phycobiliprotein is synthesized as any protein from DNA via mRNA, which is translated by the ribosomes to a protein. The protein can be post translationally modified (Gantt, 1981; Belford et al., 1983; Grossman et al., 1986). In some cyanobacterial species, the cpc promoter regulates the expression of the phycocyanin apoprotein encoding genes. The genes cpcA and cpcB encode the phycocyanin α and β subunits. Both genes are located in the cpc operon and are translated from the same mRNA transcript (de Lorimier et al., 1984; Bryant et al., 1985). In red algae, the phycobiliprotein encoding genes are located on the plastid genes (Ohta et al., 2003). Once ready, the apoprotein is ligated to the phycobilin via a single thioether bond (Beale, 1993).

Phycobilin biosynthesis

A heme, synthesized from 5-aminolevulinic acid (ALA), is the precursor in the synthesis of phycobilin. The synthesis of heme from ALA is similar to the earlier steps of tetrapyrrole formation in chlorophyll biosynthesis (Fig. 3, step 1-3 and Fig. 4, step 1-6). In the first step, the biliverdin IXa is converted to 3Z-phycocyanobilin, the dominant phycobilin isomer, by 3Z-phycocyanobilin:ferrodoxin oxidoreductase. Then in the last step the phycocyanobilin is coupled to the phycocyanin apoprotein at a specific cysteine residue via a thioether bond by phycocyanobilin lyase (Fig. 8) (Brown et al., 1989: Cornejo and Beale, 1997; Tooley et al., 2001).  Introduction

17

Figure 8. Phycocyanobilin biosynthetic pathway in cyanobacteria and algae (based on Brown et al, 1989; Manirafasha et al., 2016)

Phycocyanin application and large-scale production

There is a growing interest in natural dyes for food, confectionery and beverages as consumers become more and more skeptical about synthetic food colorants. Synthetic dyes are linked to negative health effects and in particular seem to be correlated to children’s behavioural problems (McCann et al., 2007; Arnold et al., 2012). Plants and (micro)algae are promising candidates for finding new natural colorants as they contain a wide range of pigments to capture light. Phycocyanin from Spirulina is used commercially as food and cosmetic colorant in Japan (Yamaguchi, 1997). Very recently, a blue pigment extract of S. platensis containing phycocyanin was approved by the FDA (CFR Code of Regulation Title 21, part 73, section 73.530) for use in confectionary, ice creams, and several other food products. Besides use in food as colorant, phycocyanin is also used as fluorescent dye in immunological assays, as label in cell sorting, gel electrophoresis and gel exclusion chromatography (Glaxer, 1994; Zhang et al., 2002: McQuaid et al., 2011; Singh et al., 2011). Such applications demand high purity phycocyanin, with an absorption ratio  Chapter 1

(A620/A280) of 4 or higher, whereas in food applications ratios of less than 1 are acceptable. Commercially available, food grade phycocyanin has a purity factor of around 0.7 (Cisneros and Rito-Palomares, 2004). Ammonium sulphate precipitation whether 18 combined with a chromatography step or not is commonly used to raise the purity factor (Minkova et al., 2003; Chaiklahan et al., 2011; Sørensen et al., 2013; Kumar et al., 2014; Santiago-Santos et al., 2004).

The first recorded use of Spirulina as a food supplement goes back to the Aztecs. Spanish conquistadores reported that fisherman harvested a biomass from a lake near the capital Tenochtitlan and made a blue-green cake from it. Still today, blue-green algae are found in the lake Texcoco in Mexico (Sánchez et al., 2003). Also local people around lake Chad on the edge of Sahara at the border of Nigeria, Chad, Niger, and Cameroon, for ages harvested blue green biomass from the lake, drying it and selling it on the local food markets. Spirulina was rediscovered in the 1960s as a wonder food or what we nowadays call a super food, due to its high protein content (60 to 70%) in combination with considerable amounts of polyunsaturated fatty acids (1 to 5%) (Vonshak and Richmond, 1988). In Mexico on 1967, for example, the company Sosa-Texcoco Ltd harvested Spirulina from Lake Texcoco, located at 2,200 meters altitude, and turned it into a dried Spirulina powder. Nowadays, harvesting of Spirulina from natural lakes is done in many countries including Mexico, China, Vietnam, Chad, and Myanmar (Shimamatsu, 2004; Habib et al., 2008).

Growing Spirulina in open ponds or tubular bioreactors gives higher yields and better control over the production. Most companies selling Spirulina powder use open ponds, as this is relatively cheap and can be done at large scale. Shallow raceway ponds with paddle wheels to keep the algae in solution and provide air/carbon dioxide can be as large as 5,000 square meters. Large-scale open pond cultivation of Spirulina is done at places with much sunlight such as Hawaii, the Asia Pacific Rim (Lee, 1997), and Spain (Jimenez et al., 2003). Annual production per site is in the order of several hundred tons of dry Spirulina; Earthrise Nutritionals produces over 500 tons of dry Spirulina powder per year at their side in California. Commercial Spirulina production in photobioreactors has not been established, as it is too costly. Open pond cultivation suffers from two major limitations; the first being  Introduction

the problem of infections as hygiene with open ponds is difficult to control and the second being productivity as light and the solubility of carbon dioxide in water are limiting factors.

Many microalgae can grow heterotrophically without light on an organic carbon compound 19 and oxygen (see Bumbak et al., 2011 for an overview). This mode of growth is very common for large-scale cultivation of bacteria, yeast and fungi to produce enzymes, yeast extract, or antibiotics. It is done in closed steel vessel which gives full control over the production and at the same time allows strict control over hygiene conditions. Galdieria sulphuraria is particularly interesting to produce phycocyanin, as it can grow auto, mixo, and heterotrophically (Gross and Schnarrenberger, 1995). G. sulphuraria grows well on various sugars and sugar alcohols, including glucose, galactose and glycerol (Oesterhelt et al., 1999). High cell densities varying from 80 to 110 g L-1 for a fed-batch culture were obtained on as high as 166 g L-1 glucose, yielding 250 to 440 mg L-1 phycocyanin (Schmidt et al., 2005; Graverholt and Eriksen, 2007).

Carbon Storage

The final product from photosynthesis is the carbohydrate glucose which is either converted into energy, processed into other cellular components, or converted into a polymer that serves as energy storage (Chen et al., 2013). Storage carbohydrates drive metabolic processes in periods of darkness when there is no light available to fuel the cells (see Markou et al., 2012). Microalgae are relatively efficient in photo conversion and accumulate high amounts of carbohydrate storage compounds, sometimes to more than 50% of the dry weight (Ho et al., 2011). The major storage carbohydrates in microalgae are starch, glycogen, and cellulose (Nakamura et al., 2005; Chen et al., 2013).

Starch is the primary storage carbohydrate in green algae and higher plants. Starch is a non- soluble, crystalline glucose polymer that is synthesized in the plastids (Ball and Preis, 1982; Ball and Morell, 2003). Starch consists of two different α-polyglucans, amylose and amylopectin. The difference between amylose and amylopectin is in the degree of branching (Vitova et al., 2015). Amylose, the minor component of starch, is a relatively long and linear molecule with α(1→4)-glycosidic bonds and an average of degree of polymerization (DPn) of 900-3000 glucose residues. Whereas amylopectin, the major  Chapter 1

component of starch, is a branched molecule which is composed of short α(1→4) linked chains with branching of α(1→6)-glycosidic bonds occurring every 15-45 glycosyl units.

Amylopectin has a DPn of about 4800-15900, with a molecular mass five times bigger than 20 amylose (Buléon et al., 1998; Tester et al., 2004; Vamadevan and Bertoft, 2015).

Certain microalgae synthesize glycogen as a carbon and energy storage compound. Glycogen is mainly found in animals, fungi, bacteria, and archaea (Ball and Morell, 2003). Glycogen is a water-soluble branched glucose polymer consisting of a linear backbone of α(1→4) linked glucose with side chains attached at α(1→6)-glycosidic bonds. The glycogen structure resembles amylopectin to some extent but it has a higher number of branches (approx. 8-13%), depending on the glycogen source (Matsui et al., 1993; Wang and Wise, 2011; Martinez-Garcia et al., 2016). Each main branch supports two branches, except for the outer unbranched chains. This branching pattern allows for spherical growth of the particle to form a globular structure, the most accepted structural model for glycogen (Manner, 1991; Ball et al., 2011). In this respect, glycogen is significantly different from amylopectin.

In red microalgae, the main energy storage molecule is floridean starch. It is given this name as the first floridean starch was characterized from Floridiophyceae, a class of the Rhodophyta (Barry et al., 1949). Unlike the higher plants and chlorophytes that store starch in their plastids, red macro- and microalgae synthesize and store their starch in the cytosol (Nyvall et al., 1999; Viola et al., 2001). Floridean starch shares metabolic and structural features with both glycogen and plant starch. The synthesis of starch and glycogen requires the same basic elements, a glucose donor (sugar nucleotide), a linear chain receptor for the glucose molecule attachment to form the polymer, a synthesizing enzyme which catalyzes the α(1→4) linked formation, and a branching enzyme which is responsible for the introduction of the α(1→6) branching point. The synthesis of floridean starch in red algae proceeds mainly via uridine-diphosphate-glucose (UDP-glucose) as glycosyl donor; this pathway is analogous to glycogen synthesis in fungi an animal (Viola et al., 2001). However, some species synthesize floridean starch via ADP-glucose (Nagashima et al., 1971; Nyvall et al., 1999).

 Introduction

Initially, it was described that the floridean starch lacks amylose and resembles amylopectin rather than glycogen (Flemming et al., 1956; Meeuse et al., 1960; Manner and Wright, 1962). However, later reports described that amylose was found in the floridean starch extracted from some species (McCracken and Cain, 1981; Shimonaga et al., 2007). The 21 structure and characteristics of floridean starch from different red alga species cannot be generalized, as it shows a wide variation in the structure. In some multicellular red algae, floridean starch is more similar with amylopectin in terms of average chain length, granule formation, and crystallinity. In other, unicellular red algae, the floridean starch is more a semi-amylopectin, and even a glycogen-type polyglucan (Peat et al., 1959; Yu et al., 2002; Shimonaga et al., 2007).

Carbon fixation in red microalgae also produces low molecular weight carbohydrates, mainly floridoside (galactosylglycerol or 2-O-α-D-galactopyranosylglycerol) (Courtois et al., 2008), except for the order Ceramiales that produces mannosylglycerate (Kremer, 1978; Raven et al., 1990; Li et al., 2002). Floridoside is an important soluble or transportable form of carbon between cells in analogy with sucrose in higher plants (Ekman et al., 1991). The other role of floridoside is that of osmoregulator, regulating in the internal osmotic value in response to changes in salinity (Kauss, 1968; Pade et al., 2015). Floridoside is synthesized in the cytosol via UDP-galactose, derived from UDP-glucose and glyceraldehyde-3-phosphate (see Viola et al., 2001).

Cyanidiales

Habitat and challenges

The Cyanidiales are one of the orders within the phylum Rhodophyta. They are unique among the microalgae as they thrive in acidic environments with a pH ranging from 0.5 to 3 and at moderately high-temperature up to 56oC. Members of the Cyanidiales are typically associated with geothermal active regions (Fig. 9 and Table 1) (Seckbach, 1999; Gross et al., 2001) but have also been isolated from the acidic river basin of the Rio Tinto, a red colored acidic river in Spain (Moreira et al., 1994), and peat land in the Czech Republic (Gross et al., 2002). Besides the high acidity, these environments usually have high levels  Chapter 1

of iron, copper, nickel and other heavy metals (Gross et al., 1998; Nordstrom and Alpers, 1999). Sites with geothermal activity pose additional challenges to the organism living there due to toxic gases, elevated temperatures, and low water tension in soils and rocks 22 (Brock, 1978; Yun, 1986). Acidic moderate temperature environments such as peat bogs, acidic mining lakes, and rivers can be deeply coloured by humic acids and hematite, blocking sunlight and reducing photosynthesis (Gyure et al., 1987; Simate and Ndlovu, 2014). The species diversity at such acidic locations is low and at pH values below 2.5 only a handful of microorganisms including bacteria, archaea, and microalgae survive and even thrive (Schwartz and Schwartz, 1965; Brock, 1978).

Figure 9. The distribution of Cyanidiales in the world; G. sulphuraria; G. maxima; G. partita; C. caldarium; G. maxima and G. daedala; C.caldarium and G. partita; C.caldarium and G. sulphuraria; C. caldarium and C. merolae; G. sulphuraria and C. merolae; the ring of fire area.

Living in highly acidic habitats poses considerable challenges, as the organisms have to coop with high proton concentrations. In spite of the low external pH, the cytosol of acidophiles such as Cyanidium caldarium has been shown to be neutral (Beardall and Entwistle, 1984). The consequence of having a neutral cytosolic pH while living in an  Introduction

acidic environment is that a strong proton gradient of 1:1 million or more spans the plasma membrane. Acidophilic microalgae coop with this challenge by having a plasma membrane that is fairly impermeable to protons, requiring only small amounts of energy to pump protons against the passive influx (Gimmler, 1992). However, when the energy supply 23 becomes limiting, such as when it is dark or when there is no oxygen, the cytosol tends to acidify rapidly (Gimmler et al., 1989). The red microalgae C. caldarium and G. sulphuraria, on the contrary, have developed mechanisms to cope with such limitations without acidifying the cytosol as they can survive several months of darkness (unpublished observation, reported in Gross, 2000).

Table 1. Locations from which Cyanidiales species have been isolated

Species Location References Galdieria Galdieria sulphuraria Playon de Ahuachapan, Elsalvador; Mount Lawu, Java, Indonesia; Skalafell, Iceland; Mount Shasta, California; Gross et al., 2001; Yoon et Rio Tinto, Spain; al., 2006 a Azores, Portugal; Mexicali, Mexico; Pisciarelli, Italy; Yangmingshan, Taiwan Yellowstone National Park Toplin et al., 2008 Soos, Czech Republic Gross et al., 2002 Larderello, Tuscany Ciniglia et al., 2004 Galdieria partita The Geysers, Sonoma Co. California; Mount Uzon, Kamchatka, Russia Gross et al., 2001 Kodakarajima Island, Japan Galdieria daedala Kunashir Island, Russia Gross et al., 2001 Galdieria maxima Kunashir Island, Russia Gross et al., 2001 Iceland Ciniglia et al., 2014 Cyanidium Cyanidium caldarium Kamchatka, Russia; Japan; and Gross et al., 2001 Mount Lawu, Java, Indonesia New Zealand Toplin et al., 2008 Larderello, Tuscany Yoon, 2006 a Cyanidioschyzon Cyanidioschyzon merolae Pisciarelli, Italy Gross et al., 2001 Yellowstone National Park; Toplin et al., 2008 Japan

 Chapter 1

Another challenge acidophiles face is the supply of carbon dioxide. The main source of carbon dioxide is the atmosphere, but carbon dioxide can also come from ground water, photo-oxidation of organics, respiratory activity of the sediment, and volcanic gases. The 24 acidic pH strongly affects the availability of dissolved inorganic carbon (DIC) (Azov, 1982). In freshwater at 18oC, the solubility of carbon dioxide is 10-15 μM (Gross, 2000) and is unaffected by the pH. At a low pH all DIC is present in the form of carbon dioxide - and very little in the form of HCO3 . When at a low pH the carbon dioxide is rapidly - consumed by photosynthesis, no dissolved HCO3 is available to replenish the fixed carbon dioxide. This negative effect is counterbalanced by a high diffusion rate of atmospheric carbon dioxide; at neutral pH at 18oC, equilibrium is reached between atmospheric and dissolved carbon dioxide within 3.5 min. At acidic pH (pH 2) and 18oC it only takes 0.42 s (Brinkman et al., 1933). Thus red microalgae living in the water column ‘see’ a much faster replenishment of carbon dioxide than microalgae living at neutral pH. In spite of the favorable solubility of carbon dioxide at low pH, the amount of carbon dioxide present in the water column is very limited due to slow diffusion rates. Growing endolithically on rocks, sand, or soil circumvents this challenge because atmospheric carbon dioxide can diffuse very quickly to the microalgal cells. That is why very likely G. sulphuraria prefers endolithic habitats in volcanic areas (Gross et al., 1998; Yoon et al., 2006a) and peat lands (Gross et al., 2002).

Cyanidiales genera

The red algae or Rhodophyta (Greek, ῥόδον rhodon, "rose" and φυτόν phyton, "plant") are a diverse group of eukaryotic, photosynthetic uni- and multicellular organisms, so called because many members have a reddish colour due to the presence of phycoerythrin (Kursar and Alberte., 1983; Lee, 2008). Some very common Rhodophytes are the typical red seaweeds found on the beaches, such as Gracilaria multipartite (Murano, 1995), Chondrus crispus (aka Irish moss) (Karsten et al., 1998), and Porphyra umbilicalis (Laver) (Figueroa et al., 1995). The phylum Rhodophytes is divided into two subphyla, Cyanidiophytina and Rhodophytina. The latter consists of the six classes: Floridophyceae, Bangiophyceae, Rhodellophyceae, Compsopogonophyceae, Stylonematophyceae, and Phorphyridiophyceae. The former contains only one class, the Cyanidiophyceae, which  Introduction

only has one order named Cyanidiales (Yoon et al., 2006b) that is further divided into the genera Cyanidium, Cyanidioschyzon and Galdieria (Table 2).

25 Table 2. Principal morphology, biochemical and ecophysiological features of three genera from Cyanidiales (Albertano et al., 2000)

Feature C. merolae C. caldarium G. sulphuraria Morphology Shape Oval, club-like Spherical Spherical Size (µm) 1.5 x 3.5 2-5 3-11 Reproduction Binary fission with 2 Endospores with 4 Endospores with 4-32 daughter cells daughter cells daughter cells Cell wall Absent (unclear) Present Present Chloroplast One polymorph with One spherical with One multilobed with double envelope double envelope double envelope (number and shape) Mitochondria One and spherical One and concave One and lencord to net-like (number and shape) vacuoles Absent Absent present Biochemical ct-DNA form Bead-like at central Rod-shaped at central Ring-shaped at peripheral Phycobiliproteins c-phycocyanin c-phycocyanin c-phycocyanin Nuclear DNA content 22.5 x 105 nucleotides 0.8 x 105 nucleotides 35.3 x 105 nucleotides Storage glucans Phytoglycogen, iso- Phytoglycogen, iso- Floridean starch and floridosides and floridosides Linolenic acid Absent Absent Present Ecophysiological

NH4 uptake Yes Yes Yes

NO3 uptake Yes Yes No Salt tolerance % 3 3-4 10 pH optimum 1.5 1.5 2 Temperature optimum 45 45 37 (oC) Mixotrophy no no Yes

 Chapter 1

One of the first reports on a member of the Cyanidiales was by Geitler (1933), who described a unicellular microalgae from a thermal acidic area on what he called the Sunda Islands, the old German name for Indonesia. Later, this isolate was assigned to Cyanidium 26 caldarium (Hirose, 1958). Merolae and co-authors (1981) clarified the classification of isolates similar to C. caldarium and proposed three different species, which co-exist in the same habitats: C. caldarium Geitler, G. sulphuraria (Galdieri) Merola and C. merolae De Luca, Taddei et Varano. Confusion on the classification of various strains in culture collections and isolates from volcanic areas in Europe, Asia and America has persisted as all these isolates were assigned to C. caldarium. Seckbach and coworkers (1991) assigned some of these isolates to G. sulphuraria. Currently, the genera Cyanidium and Cyanidioschyzon only contain one species, while five species have been assigned to the genus Galdieria: G. sulphuraria, G. maxima, G. partita, G. daedala, and G. phlegrea (Sentsova, 1991; Pinto et al., 2007). Detailed morphological, ecophysiological, ultrastructural and phylogenetic analysis suggests that there are actually only two Galdieria species, G. maxima and the species complex G. sulphuraria, which encompasses G. partita, G. daedala, G. phlegrea, and G. sulphuraria (Ciniglia et al., 2014; Hsieh et al., 2015).

Cyanidium caldarium

Cyanidium caldarium is the only species of the genus Cyanidium (Lee, 2008). C. caldarium cells contains one nucleus, one mitochondrion, and one chloroplast (Fig. 10). Highly organized thylakoid membranes surround the chloroplast and nucleus. Phycocyanin is the most abundant phycobiliprotein in this species. The C. caldarium cell has a spherical shape about 2 to 5 µm with a rigid cell wall and multiplies by endospores with 4 daughter cells. In this species, plastid DNAs is located in the centre of the spherical chloroplast. C.

caldarium grows exclusively autotrophically and uses both ammonium (NH4) and nitrate

(NO3). (Kuroiwa, 1998; Albertano et al., 2000).  Introduction

27

C

M

Figure 10. Diagram of Cyanidium caldarium (Lee, 2008). (S) Starch; (W) wall; (N) nucleus; (M) mitochondria; (C) chloroplast.

Cyanidioschyzon merolae

Cyanidioschyzon merolae is a unicellular, strict photosynthetic organism that contains a single nucleus, mitochondrion, and plastid (Fig. 11). It has no cell wall. C. merolae utilizes ammonium as well as nitrate (Albertano et al., 2000). C. merolae have one polymorph chloroplast with double envelope, one spherical mitochondrion, and no vacuoles (Albertano et al., 2000). The differences of C. merolae with other Cyanidiales species are the club shape, the lack of a cell wall, and division by binary fission (Lee, 2008). Cells are approximately 1.5 – 3.5 µm. This species is less tolerant to salts compared to other Cyanidium species. Phycocyanin is the most abundant phycobiliprotein in C. merolae (Albertano et al., 2000). Cunningham et al., (2006) explored C. merolae for carotenoid biosynthesis, as it has the simplest group of chlorophylls and carotenoids found in any eukaryotic photosynthetic organism.

 Chapter 1

28

Figure 11. Left: diagram of Cyanidioschyzon merolae (Lee, 2008). Right: Microscopic view of C. merolae.

Galdieria sulphuraria

Galdieria sulphuraria are round shaped and the largest in size among order Cyanidiales, reaching 11 µm (Fig. 12). They have a cell wall, one multilobed chloroplast with double envelope, one mitochondrion, one vacuole, and they form endospores with 4 - 32 daughter cells (Albertano et al., 2000; Lee, 2008). The plastid DNA is located in a ring configuration in the periphery of an irregularly shaped chloroplast (Kuroiwa, 1998). Phycocyanin is the most abundant phycobiliprotein (Albertano et al., 2000). Members of the genus Galdieria grow autotrophically as well as heterotrophically. G. sulphuraria can utilize 27 different sugars and polyols (Oesterhelt et al., 1999). The ability to grow heterotrophically enables G. sulphuraria to grow on and in soil or rock pockets of volcanic areas with limited water available using organic compounds released by other microorganisms in the endolithic community (Gross, 1999; Ferris et al., 2005). G. sulphuraria is salt tolerant up to 10%; the most salt tolerant strain is G. sulphuraria 002 isolated from Naples (Pinto et al., 2003).

G. sulphuraria accumulates highly branched glycogen, named floridean starch, as its energy storage (Shimonaga et al., 2008; Martinez-Garcia et al., 2016). G. sulphuraria also produces low molecular weight carbohydrates (floridoside and isofloridoside) which contribute to the osmotic acclimation of the cells (Pade et al., 2015). For pigment  Introduction

production, G. sulphuraria has been studied for carotenoid (Graziani et al., 2013), and phycocyanin (Gross and Schnarrenberger, 1995; Sloth et al., 2006; Moon et al., 2014).

29

Figure 12. Galdieria sulphuraria; Arrow: plastoglobules, Scale bar = 1 µm (Pinto et al., 2003); Right: light microscope overview

Galdieria maxima

Galdieria maxima cells have the largest size among the members of Galdieria (Fig. 13). Although genotypically distinct from the G. sulphuraria cluster, they are morphologically very similar (Gross et al., 2001). G. maxima cells are around 7.0 – 14.0 µm with 2, 4, or 8 endospores (Pinto et al., 2003). G. maxima and G. sulphuraria has been studied for highly branched storage polyglucan (Stadnichuck et al., 2007; Martinez-Garcia et al., 2016).

Figure 13. Galdieria maxima, scale bar = 1 µm (Pinto et al., 2003)

 Chapter 1

Other Galdieria species

There are three other species in the genus Galdieria. G. partita and G. daedala that have 30 been isolated from acid springs in Russia (Sentsova, 1991). It is very difficult, if not impossible to distinguish these two isolates from G. sulphuraria or G. maxima. It has been argued that these isolates are morphologically variable ecotypes of one and the same Galdieria species (Ciniglia et al., 1999). Then there is the species Galdieria phlegrea, isolated from Campi Flegrei (Naples, Italy). This species also has the same morphology and reproduction characteristics as the others Galdieria species, but it clearly differs in the nucleotide sequence of the rbcL gene, and thus is considered a separate species (Pinto et al., 2007).

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