Purification and Identification of Selenium-Containing C-Phycocyanin from Spirulina

Purification and Identification of Selenium-containing C-phycocyanin from Spirulina:

Implications for Bioaccumulation and Ecotoxicity

A Thesis Submitted to the Committee on Graduate Studies in Partial Fulfillment of the Requirements of the Degree of Master of Science in the Faculty of Arts and Science

TRENT UNIVERSITY

Peterborough, Ontario, Canada

Environmental and Life Sciences M.Sc. Graduate Program

January 2015

Abstract

Purification and Identification of Selenium-containing C-phycocyanin from Spirulina: Implications for Bioaccumulation and Ecotoxicity

Jana Farell

Selenium is an essential trace nutrient to many organisms, yet in high concentrations it is toxic. Organic selenium is more bioavailable to aquatic biota than inorganic selenium, but is usually found in much lower concentrations. Algae are known to biotransform inorganic selenium into several organo-selenium compounds, but it is unknown whether any of these bioaccumulate in the food chain. In this study, selenium was incorporated into the methionine residues of an algal photosynthetic protein, c- phycocyanin from Spirulina spp. The extent of selenium incorporation was quantified by inductively coupled plasma-mass spectrometry (ICP-MS), and the protein was identified using electrospray mass spectrometry (ES-MS).

C-phycocyanin was isolated and purified from Spirulina with a final recovery of

20-30 % of the total c-phycocyanin present. Selenomethionine replaced 92.8% ± 1.22 of the methionine residues in c-phycocyanin when grown in 2.5 ppm sodium selenite. ES-

MS was used to obtain protein spectra, and pure c-phycocyanin was identified. Data of full scans provided estimated masses of both protein subunits—α-chain measured at

18,036 Da; β-chain measured at 19,250 Da—close to the theoretical masses. Protein fragmentation by collision-induced dissociation and electron capture dissociation provided approximately 52 % amino acid sequence match with c-phycocyanin from

Spirulina platensis. This study demonstrates the incorporation of selenium into an algal protein, and the identification of c-phycocyanin using electrospray ionization-mass spectrometry.

ii Keywords: organo-selenium, selenomethionine, c-phycocyanin, electrospray mass spectrometry

iii Acknowledgements

I would like to thank everyone who has contributed to my thesis. For their guidance and support, I would first like to thank my supervisor, Dirk Wallschläger, and my supervisory committee, Steven Rafferty and Neil Emery. A big thank you to

Jacqueline London for help in the lab, particularly with temperamental nebulizers.

Thanks to Naomi Stock for instrument training, troubleshooting, and protein mass spectrometry suggestions. A thank you to Bastian Georg and Michael Doran for instrument training and assistance. I would like to thank Katherine Kellersberger, without whose help obtaining protein mass spectra at Bruker Daltonic, this thesis would be woefully short. A special thanks to Sarah D’Amario for having patience with the trials of method development, and for having enthusiasm in the face of bleak results. Thanks to

Kelly LeBlanc for algal culturing help, and other tips throughout my thesis. And thanks to Stephanie Jackman for CE tutorials, even though I never got around to using the instrument.

iv Table of Contents

Page number

Abstract ii

Acknowledgements iv

List of Figures and Tables vii

I. Introduction 1 1.1 Selenium as a trace element in natural waters 1.2 Selenium ecotoxicity and biogeochemistry 1.3 Selenium incorporation into proteins 1.4 Protein biodegradation 1.5 C-phycocyanin in Spirulina 1.6 Recombinant proteins 1.7 Protein mass spectrometry 1.8 Experimental objectives

II. Methods 15 2.1 Recombinant c-phycocyanin expression 2.2 Algal growth conditions 2.3 C-phycocyanin purification from Spirulina 2.4 Inductively coupled plasma-mass spectrometry 2.5 Electrospray mass spectrometry

III. Results and Discussion 22 3.1 Recombinant c-phycocyanin 3.2 C-phycocyanin yield and recovery from Spirulina 3.3 Selenium incorporation into c-phycocyanin 3.4 Evidence of c-phycocyanin 3.4.1 SDS-PAGE 3.4.2 Electrospray mass spectrometry 3.4.3 Selenium-containing c-phycocyanin

VI. Conclusions and outlook 47

References 49

vi List of Figures Page number Figure 1: Selenium biotransformation 3 Figure 2: C-phycocyanin crystal structure 9 Figure 3: Phycocyanobilin structure 10 Figure 4: Spirulina growth curve 17 Figure 5: SDS-PAGE of recombinant protein expression 23 Figure 6: SDS-PAGE of purified algal c-phycocyanin 32 Figure 7: ES-MS full scan of denatured c-phycocyanin 36 Figure 8: ES-MS fine structure 37 Figure 9: ES-MS full scan of non-denatured c-phycocyanin 40 Figure 10: ECD of non-denatured c-phycocyanin 41 Figure 11: CID of both side chains showing phycocyanobilin 42 Figure 12: ES-MS of a selenium-containing protein 45 Figure 13: ES-MS full scan of non-denatured Se-CPC and CPC 46

List of Tables

Table 1: Algal growth and protein concentration 25 Table 2: Protein mass balance 28 Table 3: Selenium incorporation into c-phycocyanin 29 Table 4: Approximate c-phycocyanin mass 33 Table 5: Fragmentation sequencing 39

vii 1

I. Introduction

1.1 Selenium as a trace element in natural waters

Over the past several decades, selenium has garnered increasing attention from the environmental community due to toxic effects to aquatic biota even at low concentrations. Selenium in most ecosystems is found in low concentrations. It is naturally occurring in the environment in shale deposits and other sedimentary rock formations (Presser et al., 1994), as well as in some sulfidic ores. Natural leaching by weathering is usually a slow process and so not a large factor in selenium toxicity.

Conversely, anthropogenic activity can cause large selenium influxes to the environment.

Some sources of anthropogenic selenium include irrigation, agricultural, and industrial wastewaters (Martin et al., 2011; Petrov et al., 2011). Additionally, commercial and pharmaceutical plants can be a source of selenium-contaminated wastewaters (Young et al., 2010).

Dissolved selenium is primarily found in natural waters as selenate (SeVI) or selenite (SeIV), and to a lesser extent, selenide (Se-II) (Maher, et al., 2010). Selenium speciation in aqueous environments is predominantly determined by the reduction potential in the water (Cutter, 1982; Dungan and Frankenberger, 1999). Aside from reduction potential, chemical speciation is generally determined by microbially mediated transformations of mostly inorganic selenium into both inorganic and organic selenium species. This influences trophic transfer and bioaccumulation (Maher, et al., 2010).

Since selenium ecotoxicity is determined by a number of factors, such as the chemical and biological species present, selenium contamination is assessed on a site- specific basis (Maher et al., 2010). Some selenium-contaminated sites have been reported 2 to have significantly higher than background concentrations; background concentrations for most sites are generally defined as < 1 µg/L. For example, in the Elk River Valley, total selenium concentrations in surface waters were reported to be over 20 µg/L (Martin et al., 2011). Such sites have been the focus of bioremediation efforts (Jones et al., 2009).

1.2 Selenium ecotoxicity and biogeochemistry

Selenium shares many chemical properties with sulfur, which, due to sulfur’s biological prevalence, makes selenium an ecotoxicological concern. Selenium’s similar chemistry with sulfur means that it can compete with sulfur in biological uptake.

Selenium is an essential trace element for many living organisms, yet it has a narrow nutritional margin. Indeed, its biological essentiality is in particular proteins, specifically as selenocysteine, the selenium analogue of sulfur containing cysteine. In most aquatic environments, selenium concentrations are low enough that they do not pose a threat to aquatic biota (Young et al., 2010).

Aquatic organisms are known to take up selenium analogues of sulfur molecules; some aquatic plants can convert selenate and selenite into organic selenium compounds.

In most aquatic environments, concentrations of these inorganic selenium compounds are too low to pose a toxicity threat to higher organisms. However, since organic selenium is much more bioavailable, it likely poses a larger bioaccumulation risk, even though it is found in much lower concentrations than its inorganic counterparts (Besser et al., 1993).

Selenium bioconcentration occurs most dramatically in the first trophic transfer: from water to algae, selenium concentrations can increase by up to six orders of magnitude (Young et al., 2010). Algal selenium uptake is an active process, and, due to selenium’s chemical similarities to sulfur, selenium can be transported intracellularly in 3 place of sulfur through sulfate channels. Some algae are able to resist selenium toxicity even at high concentrations by producing volatile selenium compounds or by reduction to the relatively less toxic elemental selenium (Janz et al., 2010). Conversion to selenides happens rapidly after uptake of inorganic selenium, which is then converted to selenocysteine or selenomethionine (see Figure 1).

Figure 1. Biotransformation of inorganic selenium into selenocysteine and selenomethionine.

Algae in particular have been shown to significantly influence the chemical speciation. Organisms lower on the food chain, such as algae and macroinvertebrates, have high selenium tolerances compared to vertebrates. Aquatic organisms, such as fish, and their predators, such as birds, are particularly susceptible to selenium toxicity.

Bioaccumulation in higher organisms likely happens through the substitution of sulfur amino acids with selenium-containing amino acids. Selenium bioconcentration can occur 4 when selenium replaces sulfur non-specifically in enzymes and proteins. Selenium toxicity presents in oviparous, or egg-laying, vertebrates as teratogenesis and mutagenesis. In vertebrates, excess selenium may interfere with the mechanisms that regulate oxidative stress and causes increased rates of oxidative cell damage. This is somewhat paradoxical since selenocysteine is essential in enzymes, such as glutathione peroxidase, that maintain normal oxidative stress levels. Exposure to high levels of selenium can also suppress immune response (Janz et al., 2010). In embryos, selenomethionine accumulates in the yolk; metabolism of this selenomethionine can produce more reactive selenium species, such as methylselenol, that contribute to oxidative stress by forming the superoxide radical, which, in turn, causes developmental deformities (Palace et al., 2004).

While most selenium in ambient waters is inorganic (Chapman et al., 2010), the organic selenium fraction poses a higher risk for bioaccumulation and toxicity, particularly in higher organisms (Hamilton et al., 1990; Besser et al., 1993). Several studies (Hamilton et al., 1990; Ingersoll et al., 1990; Besser et al., 1993) have shown that organic selenium is more bioavailable than inorganic selenium. Selenium’s toxic threshold and bioaccumulation ability are dependent upon both the biological and the chemical species present. Many of the inorganic forms of selenium present in aqueous environments have been identified whereas organo-selenium compounds implicated in bioaccumulation are often left unidentified—characterized as just organic selenium— leaving a gap in the understanding of which organo-selenium compounds are responsible for biomagnification in aqueous environments. 5

The rate of selenium uptake and biotransformation in algae varies not only with the biological and chemical species, but also on the sulfur concentrations present (Li et al., 2003; Young et al., 2010). Many freshwater algal species preferentially take up selenate over selenite, but some, such as Spirulina, prefer selenite. Sulfur has much more biological use than selenium and competes for uptake. Indeed, even in low concentrations, sulfur is somewhat preferentially incorporated over selenium (Li et al.,

2003).

We do know that algae produce several types of organic selenium compounds ranging in size from small, such as organo-selenides, as small as 0.1 kDa, to large, such as selenium-containing proteins, several hundred thousand Daltons in size (Behne and

Kyriakopolous, 2001). Some of the small selenium compounds identified, such as the methylated selenium compounds dimethylselenide and dimethylselenide, are volatile species that may play an important role in biogeochemical cycling (Besser et al., 1989;

Wallschläger and Feldmann, 2010), but are not likely candidates for aquatic bioaccumulation since they volatilize out of water. Selenocysteine, the “21st” amino acid, is unstable in water and dimerizes to the more stable selenocystine (Peachey et al., 2009).

However, selenomethionine is soluble and stable in water, and has been shown to exist in natural waters (LeBlanc et al., 2014).

1.3 Selenium incorporation into proteins

Selenium incorporates into three discrete types of proteins: selenium-binding proteins, selenoproteins, and selenium-containing proteins. Selenium-binding proteins have been studied extensively (Bansal et al., 1989; Yang and Sytkowski, 1998; Raucci et al., 2011), and attempts have been made to characterize the nature of the selenium 6 binding. These types of proteins will not be discussed further since selenium is not incorporated into amino acids, and the true nature of the selenium binding in these proteins is unknown and not yet implicated in bioaccumulation. Selenoproteins are proteins that incorporate selenium into one or more cysteine residues. Since selenocysteine is essential for protein function and this type of selenium incorporation is genetically encoded, selenoproteins are also not a biotoxicity concern (Behne and

Kyriakopoulos, 2001). The third type of protein is the most likely candidate for bioaccumulation in the food chain: selenium-containing proteins. In selenium-containing proteins, selenium incorporation is inessential and highly variable. Selenium can indiscriminately replace the sulfur atom in methionine to form selenomethionine. Since incorporation is unregulated, the extent of selenomethionine replacement of methionine is dependent upon the amount of available selenium relative to sulfur.

For three quarters of a century, researchers have implicated the replacement of methionine with selenomethionine in selenium toxicity in plants. Aquatic animals do not have the ability to synthesize selenomethionine and so selenomethionine from other sources, such as algae, may be largely responsible for selenium toxicity in these animals.

Ingested selenomethionine that remains unmetabolized is incorporated into proteins very efficiently (Schrauzer, 2000).

1.4 Protein biodegradation

When algae die, the cells rupture, releasing their contents. This release includes water-soluble proteins entering the aquatic environment. Proteins are a highly labile form of organic carbon; they make up approximately 10 % of total dissolved organic carbon 7

(Knicker and Hatcher, 1997), and are readily degraded by microbial metabolism (Taylor,

1995). Dissolved cellular enzymes that are released when the cells break open do not account for substantial protein degradation; most degradation is biotic, though some can be chemical or photochemical (Hollibaugh and Azam, 1983; Wiegner and Steitzinger,

2001). Algal protein exudate is an important source of labile organic nitrogen, which may account for its rapid microbial decomposition (Ghosh and Leff, 2013). Microbial activity in natural waters often results in protein breakdown over the course of a few hours to a couple of days depending on several factors such as temperature, microbial community makeup, and substrate concentration. Once proteins are degraded into smaller polypeptide chains, these smaller molecules are even more susceptible to microbial breakdown, and can be used up rapidly as a carbon source for microorganisms (minutes to hours) (Keil and Kirchman, 1994).

It is expected that selenium-containing proteins, like their sulfur-containing counterparts, are prone to biodegradation. So, water-soluble selenium-containing protein degradation products are likely an important source of bioavailable organic selenium in the water column in the form of small selenium-containing polypeptides and free selenomethionine. As a result of the breakdown of selenium-containing proteins, selenium bioavailability at the base of the food chain would increase drastically, leading to biomagnification in higher trophic levels. So, even with a predominantly inorganic selenium source, which is not particularly bioavailable to higher organisms, its incorporation into selenium-containing proteins may have an important effect on aquatic ecosystems.

1.5 C-phycocyanin in Spirulina

Spirulina spp., a blue-green alga that is well known for its nutritional and medicinal properties, is common and grows in many regions around the world, with a range from tropical regions to the North Sea (Ciferri, 1983). It is a main source of food for aquatic life in ponds where it is abundant. Spirulina has been shown to take up selenium and incorporate it into its proteins (Sarada et al., 1999; Pronina et al., 2002; Li et al., 2003; Patil et al., 2008), which makes it an ideal organism to use for studying environmentally relevant organo-selenium compounds.

Several studies (Novoselov et al., 2002; Pronina et al., 2002; Li et al., 2003; Chen et al., 2006a; Chen et al., 2006b; Huang et al., 2007) have determined how much selenium Spirulina can tolerate in growth media as well as how much and where it is incorporated intracellularly; most is incorporated into proteins. Generally, while both inorganic and organic forms of selenium do have stimulating effects on algal growth

(Besser et al., 1993), over 20 ppm selenium in growth media can be toxic. In cultures with very high selenium concentrations, algal cells turn red because of the reduction of selenate or selenite to elemental selenium, and cell growth is stunted (Li et al., 2003).

C-phycocyanin (Figure 2), a protein known to non-specifically incorporate selenium into some, if not all, of its ten methionine residues, is one type of protein in

Spirulina’s phycobilisomes. Phycobilisomes are a class of light-harvesting protein complexes found in red and blue-green algae that assume similar roles to that of the chlorophyll b-protein complexes present in higher plants (Ciferri and Tiboni, 1985).

These pigment-protein complexes are responsible for approximately half of the light harvested in blue-green algae. Chlorophyll a, along with several other photosynthetic 9 pigments present in blue-green algae, can only harness energy from the blue end of the spectrum; phycobilisomes compliment this by harvesting light in the region that chlorophyll a does not cover, specifically green and yellow-orange light (Piñero-Estrada et al., 2001; Habib et al., 2008; Moraes et al., 2011).

Figure 2. Crystal structure of c-phycocyanin, composed of a trimer of heterodimers (αβ)3 with attached chromophores; isolated from Themosynechococcus vulcans. The side chains show similar structure and have complimentary tertiary structure or folding (Brejc et al., 1995), though they differ in primary structure (amino acid sequence) and the number of attached chromophores. Image from Research Collaboratory for Structural Bioinformatics (RCSB) Protein Database.

C-phycocyanin is comprised of an α-chain and a β-chain, linked through hydrogen bonding. These interactions are normal protein-protein interactions that comprise quaternary protein structure. The affinity of these side chains for one another causes them to form the heterodimer configuration of the protein (αβ). Further associations of these heterodimers frequently occurs such that (αβ)3 and (αβ)6 species 10 form the active light harvesting proteins of the phycobilisomes (Lenong et al., 2001).

Linker proteins are also involved in these structures, but are found in much lower concentrations and do not actively harvest light (Brejc et al., 1995; Stec et al., 1999).

These complexes contain proteins covalently bonded to chromophores; specifically, phycocyanobilin (Figure 3) binds to c-phycocyanin. Phycocyanobilin is a tetrapyrrole that binds to a cysteine residue through a cysteinyl thioether linkage. Each chain is covalently bonded to at least one chromophore: the α-chain contains one phycocyanobilin, the β-chain contains two, resulting in the quaternary structure of the holo-protein (here, this refers to the intact protein, or apo-protein, with its chromophore attached). Each is covalently bonded to a specific cysteine residue (Romay et al., 2003).

Phycocyanobilin has a blue colour and is stable while attached to the apo-protein, but degrades and loses colour quickly when free in solution (Coyler et al., 2005).

Figure 3. Structure of phycocyanobilin showing peptide linkage (image from Coyler et al., 2005).

In Spirulina, the proteins that actively harvest light in the phycobilisomes are allophycocyanin and c-phycocyanin in a 1:10 ratio, and together account for about 50 % of the water-soluble proteins (Seo et al., 2013). C-phycocyanin is found in large concentrations in Spirulina (Minkova et al., 2003; Silveira et al., 2007). However, 11 relative to other protein sources, proteins isolated from Spirulina are somewhat deficient in sulfur amino acids (Ciferri, 1983; Ciferri and Tiboni, 1985).

C-phycocyanin is stable at low temperatures and up to 30 °C for at least seven days and over a pH range of 5.0-7.5 (Sarada et al., 1999). At concentrations above 0.1

µM, the holo-protein aggregates into multimers (Eriksen, 2008). Due to its highly hydrophilic nature, the presence of high concentrations of organic solvents can also cause aggregation and even precipitation out of solution (Simó et al., 2005). Blue-green algae cell walls can resist rupture, so mechanical shearing, such as sonication, is often necessary to break open cells (Moraes et al., 2011). Once lysed, the water-soluble phycobiliproteins are released and can be collected in the supernatant. The amino acid sequence of c-phycocyanin is similar among different species of blue-green and red algae

(Binder et al., 1972; Santiago-Santos et al., 2004; Eriksen, 2008).

1.6 Recombinant proteins

Recombinant protein expression in Escherichia coli is a common method for the production of large quantities of protein from other sources. Previous studies (Bryant et al., 1985; Cai et al., 2001; Guan et al., 2007; Wang et al., 2007) have successfully expressed phycobiliproteins as recombinant proteins in E. coli. One advantage of recombinant protein expression is the large quantities of easily purified protein. Another advantage is that it is possible to completely substitute selenomethionine for methionine using simplified media supplemented with selenomethionine (Guerrero et al., 2001). The c-phycocyanin β-subunit contains six methionine residues, two more than the α-subunit

(UniProt, 2013c-d), and so more selenium can be incorporated as selenomethionine.

Furthermore, the β-chain has been expressed and purified in E. coli BL21, and unlike the 12

α-chain, which self-aggregates, the β-chain can be expressed as a soluble monomer

(Wang et al., 2007).

A vector plasmid, essentially a string of DNA, is inserted into E. coli via electroporation. An electric charge is applied to create an electric field across the cells.

This field disrupts the cell membranes enough that microscopic pores form for a very brief time (~5 ms), which allows the plasmid, containing the genetic information of the protein that will be synthesized, to flow into the cells. When these modified cells reproduce, they will replicate the plasmid DNA along with their chromosomal DNA

(Neumann et al., 1982). This material is then plated with an antibacterial substance. The cells that have incorporated the plasmid are resistant to the antibacterial substance, and they start to grow colonies; all the cells that did not receive plasmid will die off. These colonies can be harvested and grown into the log growth phase. Hence, the resulting liquid cell culture will be homogenous E. coli that can be modified to produce the protein encoded in the plasmid.

1.7 Protein mass spectrometry

Over the past fifty years, protein characterization and identification has been facilitated by mass spectrometry. Electrospray ionization mass spectrometry employs a soft ionization method that allows for analysis of large (hundreds of kDa) intact molecules. Unlike many other forms of mass spectrometry that use a harsher ionization method, electrospray protein spectra show a single intact molecule (protein or peptide) at

n+ multiple charge states. Each charge state denotes a different number of protons, MHn ; the more protons a protein can accept, the lower it will appear on the mass to charge scale, making the detection of some very large proteins possible at a low m/z range (de 13

Hoffman and Stroobant, 2007). Many proteins are able to accept multiple charges, so a distinct pattern is present in the spectrum. Adjacent peaks are representative of consecutive charge states, and the total molecular mass can be calculated from the charge and mass shown in the spectrum.

Protein mass spectrometry can be used as a powerful protein-sequencing tool; by fragmenting different charge states, amino acid sequences can be assigned to the resultant fragments. Tandem mass spectrometry (MS/MS) is essential in this work. Collision- induced fragmentation (CID) MS/MS is most commonly used for protein fragmentation.

The first mass analyzer is used to select the ions for fragmentation, which are then collided with neutral gas molecules in the collision chamber before being analyzed by the second mass analyzer (de Hoffman and Stroobant, 2007). Alternatively, before mass spectrometry, the protein can be digested with an enzyme and applied to a liquid chromatographic column. The peptide fragments can be matched to an amino acid sequence (Zubarev et al., 2000), often achieving close to complete sequence match.

1.8 Experimental objectives

In aquatic environments with elevated selenium levels, high rates of selenium incorporation into algal proteins may be an important source of bioavailable selenomethionine. Since proteins are an excellent source of labile organic carbon in freshwater systems, the degradation of selenium-containing proteins may contribute to selenium bioaccumulation and ecotoxicity by providing a highly bioavailable organic selenium source to the bottom of the food chain.

The aim of this project was to determine whether Spirulina could be used as a source of selenium-containing c-phycocyanin, and whether complete selenium 14 substitution could be achieved by incorporation into methionine residues. This was investigated through method development for selenium incorporation as well as characterization and identification by electrospray mass spectrometry.

The development of a method for acquiring selenium-containing proteins would allow for gathering a better understanding of the importance of organo-selenium compounds in aquatic environments. Purified selenium-containing c-phycocyanin could be spiked into natural waters to explore the degradation rates of the selenium-substituted proteins, and selenium-containing degradation products that may be implicated in selenium bioaccumulation in aquatic environments could be identified. Ultimately, the goals of this work are to help provide a better understanding of selenium bioavailability at the first trophic level.

II. Methods

2.1 Recombinant c-phycocyanin expression

A plasmid vector conferring kanamycin antibacterial resistance and containing the amino acid sequence for c-phycocyanin β-chain from Spirulina platensis optimized for expression in E. coli was obtained from DNA 2.0 (Menlo Park, CA). The vector contained a high-expression pJExpress401 backbone with a T5 promoter. Once obtained, the vector was extracted from the filter paper with a slightly basic buffer, spun down, and stored at -20 °C. The concentration was measured by nanodrop (~17 ng/µL) and high purity (260 nm/280 nm close to 1.8). Transformation into E. coli BL21 competent cells was successful; frozen cell stocks were made and stored at -80 °C until use.

To optimize protein expression, transformed cells were streaked onto a kanamycin plate and single colonies were used to start overnight cultures; aliquots of the overnight cultures, grown at 37 °C with continuous shaking, were used to inoculate 50 mL cultures. For protein expression, cells were grown to OD600 = 0.4-0.8 in Luria broth

(LB) containing 1:1000 kanamycin. Protein expression was induced with 1 mM isopropyl

β-D-1-thiogalactopyranoside (IPTG) for 4 hours at 37 °C. A noninduced control was run in parallel. Cells were harvested by centrifugation at 3500 g for 15 min, then flash frozen and stored at -80 °C until use. Harvested cell cultures were thawed and protein extracted using 4:1 B-PER reagent to cell pellet volume. After 10 min incubation at room temperature extracts were spun down at 17000 g for 15 min; supernatants were collected and pellets discarded. Aliquots from these extracts were put in loading buffer (4:1, respectively) and stock extracts were stored at -20 °C. Samples in loading buffer were run 16 on a 14 % sodium dodecyl sulfate polyacrylamide gel (SDS PAGE) at room temperature.

Gels were stained with Coomassie brilliant blue G250 and destained in water overnight.

2.2 Algal growth conditions

Spirulina spp. was obtained from the Canadian Phycological Culture Center at the

University of Waterloo (strain CPCC 695 Spirulina sp.). New cultures were inoculated using sterile technique in a laminar flow hood. All materials for the culturing process, including glassware, media, and pipette tips, were autoclaved at 121 °C and 20 psi for 20 to 30 minutes. Foam stoppers were used to cap the flasks to inhibit bacterial contamination. Growth experiments were conducted to achieve optimal growing conditions and maximum selenium incorporation into proteins. Algae were grown in two types of media, Bold’s basal media and BG-11 media to determine which yielded more biomass. Slow-growing stock cultures were maintained in 100 % Bold’s basal media and

100 % BG-11 media, experimental cultures were prepared with 5 mL stock in 100 mL corresponding 10 % media. Small-scale experimental cultures were grown in triplicate.

Cultures were grown at 28 °C on a 12:12 hour light:dark schedule under 30 µE/m2s. For aeration, flasks were gently shaken manually once per day. For Se-rich cultures, sodium selenite was spiked into solution upon inoculation. Concentrations ranged from 1 ppm to

10 ppm selenium. Once the optimal Se concentration was determined, a large-scale culture (10 L) was grown in a 14 L microfermentor (New Brunswick Scientific): 10 %

BG-11 with 2.5 ppm Se, using the inoculation and growing conditions outlined above.

For small-scale cultures, three 100 mL cultures were pooled and harvested as one batch; the large-scale culture was harvested as one batch. According to the literature, highest biomass comes at three weeks after culture inoculation (see Figure 4, Mezzomo et 17 al., 2010), which matched observations of the Spirulina strain obtained from CPCC. The algal strain grown here grew in clusters and not homogenously in solution, so biomass measurements would have been inaccurate even if a separate flask were harvested each day since the initial algal biomass used to inoculate each culture would have differed.

Figure 4. Growth curve for Spirulina. Highest cell counts are at three weeks (figure from Mezzomo et al., 2010). The different curves show differences in growth media.

2.3 C-phycocyanin extraction and purification from Spirulina

Algal cells grew for three weeks before harvesting by filtration using cellulose

Whatman filters. The resulting algal pellet was transferred to a centrifuge tube, washed once with approximately 20 mL deionized water, and re-pelleted by centrifugation

(Sorvall) at 17200 g for 20 minutes. The washed pellet was re-suspended in 15 mL

Millipore water and then either flash frozen in liquid nitrogen or frozen overnight at -80

°C. 18

On the day of cell lysis, cells were thawed. Once at room temperature, algal cells were sonicated (Fisher sonic dismembrator) at 60 % power for ten cycles. Each cycle consisted of alternating twenty second pulses with twenty second intervals of resting on ice. Sonication was carried out at 4 °C. Lysed cells were spun down for 10 minutes at

17000 g (Eppendorf bench-top centrifuge). The 20 mL of supernatant was collected and the crude proteins were precipitated with 50 % w/v ammonium sulfate (5.82 g). This solution was centrifuged at 17200 g for 40 minutes. The resulting pellet was re-dissolved in 15 mL 5 mM potassium phosphate buffer (pH 7.0) and spun at 17000 g for 40 minutes.

The supernatant was collected and used in subsequent steps.

Using ultrafiltration, the blue-green solution was concentrated down to approximately 5 mL, then diluted to 50 mL in 5 mM potassium phosphate three times to rinse out any residual ammonium sulfate. For this, a 50 mL ultrafiltration cell (Amicon stirred cell) was equipped with a 10,000 molecular weight cutoff membrane and pressurized to 75 psi. The blue-green protein solution was collected and applied to a

DEAE-sepharose column (GE Life Sciences). The phycobiliproteins were eluted using a potassium phosphate gradient from 5 mM to 300 mM (pH 7.0) with a flow rate of 1 mL/minute. The chlorophyll a eluted in the first fraction without binding to the column.

Allophycocyanin and c-phycocyanin eluted at 125-150 mM potassium phosphate, confirmed visually by blue color and by UV/Vis spectroscopy. Pooled fractions, 50 mL in total, were concentrated using an ultrafiltration membrane with a 500 molecular weight cutoff; 5 mM (pH 7.0) potassium phosphate buffer was used for ultrafiltration and as the final buffer solution. The final concentrated volume was 10 mL. 19

The separation of allophycocyanin from c-phycocyanin was accomplished through batch sorption of allophycocyanin. Approximately 10 % (v/v) hydroxyapatite

(Bio-Rad) was added to the protein solution and mixed at 4 °C for 20 minutes. Pure c- phycocyanin was obtained by centrifuging this solution at 2,500 g for 10 minutes and collecting the supernatant. Bradford assays were done according to standard procedure

(Bradford, 1976) to quantify protein.

Gel electrophoresis confirmed that the purified protein was in the right molecular mass range (see section 3.4.1). Protein sample (0.1 mg/mL) was denatured with in loading buffer (4:1) and heated for 5 minutes at 70 °C. On the gel, 7 µL of protein ladder

(Mark12TM Unstained Standard, Life Technologies) and 20 µL of purified protein were loaded into adjacent lanes on a 14 % acrylamide gel. Electrophoresis was run at 120 V until the dye front reached the bottom of the gel. Coomassie Brilliant Blue G250 stained the proteins and protein ladder.

To determine dry mass, algal cultures were harvested by vacuum filtration and dried in a muffle furnace at 85 °C for 4 hours. Protein concentration was determined by

Bradford assay.

2.4 Inductively coupled plasma-mass spectrometry

Quantitative analysis of selenium by inductively coupled plasma-mass spectrometry (ICP-MS) is made difficult by isobaric interference of selenium’s most abundant isotope, 80Se, by the polyatomic 40Ar40Ar+ species, and interference at the second most abundant isotope, 78Se by 38Ar40Ar+. To overcome this obstacle, a collision cell or a dynamic reaction cell is employed to break up such interferences. In this 20 research, a collision cell was used. Hydrogen and helium gases are introduced to the skimmer cone and bombard polyatomic atoms from the plasma to break up interferences.

Selenium detection limits decrease since the concentrations of polyatomic argon species are reduced several-fold by this method (Marchante-Gayón et al., 2001).

Other elements are known to have an effect on the strength of the selenium signal.

Organic carbon has been shown to enhance selenium signals in ICP-MS, even at low concentrations (Larsen and Stürup, 1994). Salts have been documented to suppress analyte signals (de Hoffman and Stroobant, 2007), so their addition at relative concentrations to the samples should account for their effect on the samples.

To determine the extent of selenium incorporation into c-phycocyanin, purified

Se-containing protein was diluted to ~3 mg/L. The internal standard contained 5 ppb Mn,

Rh, and In. Organic carbon, in the form of cytochrome c, and salts in the form of potassium phosphate were spiked into the calibration standards at similar concentrations to the sample solutions to account for their effects on the selenium signal. Additionally, selenium spikes were added to matrix-matched samples to test the accuracy of this approach; recoveries from this method were within 2 % of the actual value. The calibration spanned from 2-45 ppb Se. All solutions were acidified to 2 % HNO3. Total selenium concentrations were analyzed by ICP-MS using a collision cell (Thermo

Scientific XSERIES 2). A hydrogen-helium gas mixture was used as the collision gas.

Selenium masses at m/z 76, 77, 78, and 82 were monitored. 78Se was used to calculate the total selenium concentration.

2.5 Electrospray mass spectrometry

Electrospray mass spectrometry (ES-MS) was used to confirm the identity of the purified protein. ES-MS was conducted at the Water Quality Centre (Trent University in

Peterborough, ON) and at Bruker Daltonic (Billerica, MA). Since salts interfere with protein mass spectrometry by creating salt adducts that decrease the intensity of major peaks, prior to electrospray mass spectrometry the purified protein was buffer-exchanged into 10 mM ammonium acetate by ultrafiltration. 500 µL of purified protein was concentrated to 100 µL in 10 kDa microfuge ultrafiltration concentrators at 17000 g

(Eppendorf bench-top centrifuge). This solution was diluted to 500 µL in 10 mM ammonium acetate buffer and spun down at 17000 g five times before mixing either with

0.1 % formic acid in acetonitrile (1:1) to denature the protein, or with 10 % methanol for analysis of the non-denatured protein.

At Trent University, samples were run denatured on AB Sciex QTRAP 5500 and

API 3000 systems. The five desalting washes were still insufficient for distinct protein signals on the API 3000, so the protein samples were further desalted by the same procedure mentioned above, for a total of twenty ammonium acetate washes. Samples were confirmed to be salt free upon inspection of the resultant mass spectrum by comparison to a blank. Analysis of samples was accomplished by direct infusion at 10

µl/min in positive mode encompassing the m/z range 200-1200 and 200-3000, respectively. At Bruker, samples were run on a SolariX XR equipped with a 12 T magnet.

Infusion rate was 2 µL/min. Scans encompassed the m/z range 200 to 4500. Data analysis of these scans was done using DataAnalysis and BioTools software.

III. Results and Discussion

3.1 Recombinant c-phycocyanin

Protein expression levels appeared to be very low, so a variety of different expression conditions were tried. Varying expression temperature (37 °C, 28 °C, 20 °C),

IPTG concentration (from 0.1 mM to 1 mM), media type (Luria Broth, Terrific Broth), and expression time (4 hours, 6 hours, overnight) in different combinations yielded no improvement: the expression conditions detailed in the methods were found to produce the best expression. To eliminate researcher error, possible issues with the cell lines, and reagents, a control plasmid (purple prancer protein) in the same vector backbone was transformed into cells, cultured, expressed, harvested, and run on a gel in parallel with the study vector (see Figure 5).

Expression of the control protein was high; it was the most prominent band on the gel at its known molecular weight (~26.4 kDa). The expression of c-phycocyanin β-chain under all culture conditions was low. The band at approximately 20 kDa was present in lanes 3, 4, and 6 and appeared to be the only band not in the non-induced cell cultures.

When protein expression was induced, the bacteria put the majority of their energy into making the protein encoded in the plasmid, so that protein should be the most abundant.

Since it appeared that CPC-β was being made, but only in small quantities relative to other proteins, it is possible that the protein was being immediately broken down inside the cell. One reason for this phenomenon is that the stability of this protein may be conferred by the cofactor (phycocyanobilin), which E. coli do not make on their own and was not present when c-phycocyanin β-chain is being made in the cell. Cofactors can help proteins hold their active conformations (Mirsky and Pauling, 1936) and when they 23 are not present, the protein can unfold, in which state they are more labile and subject to enzymatic attack.

protein ladder 1 2 3 4 5 6

200 116.3 97.4 66.3

55.4

36.5

31 purple prancer control

CPC-β 21.5

14.4

Figure 5. SDS-PAGE of control plasmid and CPC-β. Lanes 1-3 show cell lysates of freshly transformed competent cells. Lanes 4-6 shwo cell lysates of cells from glycerol stock. Lanes 1-4 show expression at 37 °C for four hours; lanes 5 & 6 show expression at 28 °C overnight. The purple prancer protein was run in lane 1; the other five lanes show CPC-β under different conditions. Lane 2: non-induced CPC-β; lane 3 & 4: CPC-β expressed with 1 mM IPTG; lane 5: non-induced CPC-β; lane 6: CPC-β expressed with 1 mM IPTG.

3.2 C-phycocyanin yield and recovery from Spirulina

The average protein content in Spirulina is 65 % (Babadzhanov et al., 2004), but can range from 50 to over 70 % depending on growing conditions (Ciferri, 1983; Ciferri and Tiboni, 1985). The average protein concentration of the cell lysate measured here averaged 38.41 % of the dry mass. This value is comparable to reported protein extraction (43.75 %) by a similar method (Moraes et al. 2011). The protein percentages reported here are based on the cell lysate and are thus not within the reported literature values of 50-70 % total protein in the algae, suggesting that either a significant fraction of the total algal proteins is not water soluble, or that proteins were not released quantitatively during cell lysis, or a combination of these factors. About half of the proteins are not water-soluble (Minkova et al., 2003), so these are not reflected in the crude lysate approximation of total cellular protein reported. Total protein was not measured, and since the total protein content in the algae cannot be determined from the existing data, c-phycocyanin approximate yield is calculated from algal dry mass. Since c-phycocyanin content is reported to be narrowly variable, 14-16 % of the dry mass

(Moraes et al. 2011), percent yield was calculated from the average, 15 %.

Measured concentrations of purified c-phycocyanin averaged 3.79 % of the dry mass (see Table 1 for selected values) out of the possible 15 % (Minkova et al., 2003;

Huang et al., 2007). The percent recovery (total recovered c-phycocyanin out of theoretically possible c-phycocyanin) ranged from 20-30 % (see Table 1), which is 2-3 times higher than recovery reported by Seo et al. (2013), whose purification was conducted by similar methods and only recovered 10 % of the total c-phycocyanin. By 25 comparison, the recovery in this study is considerably better, even though at least 75 % of c-phycocyanin was lost during purification and or extraction.

Depending on the total protein content of the Spirulina cultured and used in these experiments, at least 75 % and possibly > 90 % of the total c-phycocyanin was lost during cell lysis and/or protein purification. To determine where the losses occur, purified c-phycocyanin can be run through the purification process a second time, and quantified after each step. This will also indirectly provide information on the extraction efficiency of the cell lysis; if 75-90 % of the protein is lost during purification, cell lysis was effective. If not, cell lysis was incomplete. Since one of the limitations to successful electrospray mass spectrometry on the API 3000 was low concentration (see section

3.4.2), it was important to elucidate the main source of protein loss.

Table 1: Growth experiments with different media types and different Se concentrations. Percentages are by dry mass. Unless specified, data represent 3-100 mL small-scale cultures. Media [Se] Dry mass % total protein C-phycocyanin C-phycocyanin C-phycocyanin ppm (mg) in cell lysate (µg) (%) % yield BBM 0 2.0064 26.28 ± 1.31 72.91 ± 3.65 3.63 ± 0.18 24.20 ± 1.21 BBM 1 9.5730 n/a 349.34 ± 17.46 3.65 ± 0.18 24.33 ± 1.22 BG-11 0 9.9560 38.40 ± 1.92 419.14 ± 20.96 4.21 ± 0.21 28.11 ± 1.41 BG-11 1 9.1109 40.07 ± 2.04 380.86 ± 19.04 3.88 ± 0.19 25.87 ± 1.29 BG-11 2.5 8.9331 40.40 ± 2.02 331.20 ± 16.56 3.71 ± 0.19 24.73 ± 1.24 BG-11* 2.5 352.5344 39.87 ± 1.99 12,838.70 ± 3.64 ± 0.18 24.27 ± 1.21 616.94 BG-11 10 5.5146 33.45 ± 1.77 180.44 ± 9.02 3.27 ± 0.16 21.80 ± 1.09 *This is the large-scale (10 L) algal culture.

Several factors could account for lower than expected protein in the crude lysate: cell lysis may have been incomplete, or lipid-soluble and water-soluble proteins were 26 retained in the algal pellet. Regardless, measurement of total protein in the cell lysate would not reflect the lipid-bound portion, which comprises approximately half of the proteins in Spirulina (Minkova et al., 2003). To test for the presence of water-soluble proteins in the sonicated pellet, a salt wash (0.1 M NaCl) was used in an attempt to extract more protein from the algal pellet which resulted in an additional ~200 µg, or about 3 %, more protein. This was deemed insignificant.

Sonication can have negative impacts on protein recovery; during sonication, native protein structure can be destroyed from mechanical shearing and excessive localized heat from cavitation, which can form protein aggregates (Stathopulos et al.,

2004). These aggregates can interfere with successful protein purification. While gel electrophoresis indicates that aggregation was not an issue (see Figure 6 in the next section), major protein losses in the purification steps indicate that native structure may have been partially denatured. However, denaturation would be mild, in the form of loss of some quaternary protein folding, since electrospray mass spectrometry of the non- denatured protein indicates that the subunits were still hydrogen-bonded together after purification.

The multiple ultrafiltration steps to desalt the protein account for another major source of protein loss during purification. Ultrafiltration, while a faster alternative to dialysis, can introduce problems. Ultrafiltration membrane sizes are reported as molecular weight cutoffs; the pores are designed with the assumption that all proteins

(and other macromolecules) above the specified mass are folded similarly and thus have the same size to molecular weight ratio. In the case of non-native proteins (proteins that are not in the same configuration as in vivo), partially denatured proteins (proteins with 27 loss of some subunit complexation and or folding), or proteins with atypical folding, this assumption is problematic since they may be smaller than the membrane pores and flow through. Indeed, the filtrate from ultrafiltration was often observed to contain a slight blue colour, indicative of the presence of phycobiliproteins, though not enough to quantify using Bradford assay. The use of a membrane with a lower molecular weight cutoff membrane (500 Da) to limit losses was abandoned since filtering was recorded at a rate of 5 mL/day, though the filtrate was colourless.

Protein concentration was quantified by Bradford assay to identify which steps were most responsible for losses (Table 2). Data are not shown for the ultrafiltration step in between the ammonium sulfate precipitation and the ion exchange column since the large volume of the multiple washes was too dilute to measure by Bradford assay. The first purification step (ultrafiltration plus the ion exchange column) lead to the greatest protein loss. The reasons are two-fold. One: c-phycocyanin and allophycocyanin account only for 50 % of the water-soluble proteins in Spirulina (Ciferri and Tiboni, 1985; Patil et al., 2008) in a 10:1 ratio, so there should be a reduction in total protein concentration since other proteins are being intentionally removed. Two: the ultrafiltration step may account for some of the other loss. However, the loss from this ultrafiltration step is considerably larger than in the second ultrafiltration step. To apply to the ion exchange column, the protein solution must be in 5 mM potassium phosphate buffer, a lower ionic strength buffer than is necessary for the hydroxyapatite column. So the first buffer exchange was more exhaustive, hence more protein loss. Also, it is likely that some small fraction of the phycobiliproteins elute from the column outside the collected fractions

(potassium phosphate concentrations 125-150 mM).

Table 2: Mass balance for total protein recovery. Purification step Protein (µg) % loss from previous step 1. Crude lysate 6231 ± 311 n/a 2. Ammonium sulfate 4187 ± 209 32.80 ± 1.64 precipitation re-suspension 3. Post ion exchange column 403 ± 20 90.37 ± 4.51 4. Post second ultrafiltration 358 ± 18 11.17 ± 0.55 5. Post hydroxyapatite column 305 ± 15 14.80 ± 0.73 1. Protein concentration in the supernatant after sonication and pelleting 2. After washing ammonium sulfate pellet in 5 mM potassium phosphate buffer 3. After buffer exchanges into 5 mM potassium phosphate buffer and ion exchange column 4. After the second buffer exchange into ~20 mM potassium phosphate buffer 5. Purified c-phycocyanin

The values reported in Table 2 are from the optimized purification protocol. Some issues that accounted for additional losses in the earlier versions of the purification protocol were related to the hydroxyapatite column and the ultrafiltration steps. Initially, buffer exchanges were done using 15 mL spin columns (Millipore), and protein loss in these steps accounted for 50 % loss each time. The use of a pressurized ultrafiltration cell

(as outlined in section 2.3) reduced the losses in the ultrafiltration steps to approximately

10 %. In the hydroxyapatite column step, most (> 90%) of the c-phycocyanin was lost initially; it bound to the column and did not elute. Using a higher ionic strength potassium phosphate buffer, 20 mM, increased the recovery of c-phycocyanin in this step to approximately 95 % (see Table 2).

3.3 Selenium incorporation into c-phycocyanin

Total selenium was measured in purified protein to determine the extent of selenium incorporation into c-phycocyanin. Correlation coefficients for calibration curves were >0.99995. Internal standards were consistent across standards and samples, and 29 matrix-matched spike recovery averaged within 2 % of the standards, so no corrections were necessary. Sodium selenite concentrations in growth media were 1 ppm, 2.5 ppm, 5 ppm, and 10 ppm (Table 3). Data are not provided for the 10 ppm selenite culture due to low algal growth (see Table 2 in previous section). One small-scale culture was grown with 10 ppm sodium selenate.

Table 3: Selenium incorporation into c-phycocyanin from different concentrations and chemical species. [Se] in growth media Se species Se atoms per protein (ppm) 1 IV 8.93 ± 0.98 2.5 IV 9.28 ± 1.02 5 IV 9.37 ± 1.03 10 VI 2.77 ± 0.30

To calculate the average number of selenium atoms incorporated into c- phycocyanin, the concentration measured by ICP-MS was multiplied by the number of times the sample was diluted before analysis, then dividing by the atomic mass of Se.

This was divided by the molarity of the protein solution before dilution in 2 % HNO3.

Here is an example (from 2.5 ppm Se in growth media):

Concentration in ICP-MS sample: 37.64 ppb = 37.64 ng/mL Concentration in undiluted sample: 37.64 ng/mL x 30 = 1129 ng/mL = 1.129 x 10-3 g/L Concentration (molarity) of selenium in undiluted sample: 1.129 x 10-3 g/L x (1mol / 78.96 g) = 1.4301 x 10-5 M Se C-phycocyanin molarity: 57.72 µg/ mL = 0.05772 g/L x (1mol / 37455) = 1.5410 x 10-6 M c-phycocyanin Se atoms per c-phycocyanin: 1.4301 x 10-5 M Se / 1.5410 x 10-6 M c-phycocyanin = 9.28 Se atoms / c-phycocyanin 30

Spirulina grown with 2.5 ppm sodium selenite in BG-11 media yielded the best combination of algal mass and selenium incorporation into c-phycocyanin (see Tables 2 and 3). Error associated with the Bradford assay concentration measurements averaged ±

5 %, error from total selenium measurements averaged ± 2 % from the ICP-MS. C- phycocyanin contains a total of ten methionine residues; when grown with 2.5 ppm sodium selenite, on average, 9.28 ± 1.22 of these are selenomethionine. Spirulina was grown to successfully produce selenium-substituted c-phycocyanin. Selenium incorporation into c-phycocyanin has a large impact on the mass spectrum; this will be discussed further in the following section.

Selenium concentrations in contaminated environments are often at one to two orders of magnitude lower than the 2.5 ppm used in this study to achieve complete substitution of methionine by selenomethionine (Martin et al., 2011). At 1 ppm, Spirulina was found to be a third less effective at incorporating selenium into c-phycocyanin; however, it is unclear whether or not the relationship between selenium concentration and selenium incorporation as selenomethionine is linear. It is possible that a low threshold selenium concentration is necessary for some substitution in proteins, but much higher concentrations are required for complete substitution of selenomethionine for methionine.

If this is the case, then selenium-containing c-phycocyanin, as well as other selenium- containing algal proteins, may play an important role in selenium biogeochemical cycling.

3.4 Evidence of c-phycocyanin

3.4.1 SDS -PAGE

Gel electrophoresis of a concentrated sample of purified protein (0.1 mg/mL) established its approximate mass and purity (Figure 6). Standard protein loading concentrations are around 2 mg/mL for each protein, twenty times more concentrated than the sample shown here, hence the difference in intensity between the ladder and the purified sample.

SDS-PAGE analysis of the purified protein shows the presence of two distinct bands. The masses corresponding to the two bands on the protein ladder to the two bands indicate a pure c-phycocyanin with denatured quaternary structure: the α- and β-chains are visible. SDS-PAGE can provide only approximate molecular masses of proteins (Rath et al., 2008); indeed, both of the bands ran higher on the gel than expected (see Table 4).

The expected α-chain mass is 16-18 kDa and the β-chain has a mass around 18-20 kDa, depending on the source algal species (UniProt, 2013a-d). These masses correspond to the monomer (αβ) rather than any aggregated species such as the trimer (αβ)3 or hexamer

(αβ)6.

Figure 6. SDS-PAGE of purified c-phycocyanin from Spirulina. Gel shows Coomassie Blue dyed proteins and protein ladder (Mark12TM Unstained Standard, Life Technologies). The approximate masses of the α-chain and the β-chain shown here are measured as 18 kDa and 21 kDa, respectively.

Table 4: Approximate mass of c-phycocyanin α- and β-chains as measured by SDS-PAGE and ES-MS (discussed in following section) compared to their respective theoretical masses (UniProt database 2013a-d) from two algal sources. Protein Approximat Calculated* ES- Approximate† Theoretical Theoretical chain e SDS- MS ES-MS mass from mass from PAGE experimental experimental Spirulina Spirulina masses (Da) masses (Da) masses (Da) platensis (Da) jenneri (Da) CPC-α ~18,000 18,036 ± 0.60 18,041 ± 23.45 18,188 18,202 CPC-β ~21,000 19,250 ± 4.45 19,253 ± 25.03 19,267 19,174 *Traditionally calculated masses †Averagine calculated masses

3.4.2 Electrospray mass spectrometry

One of the obstacles of protein electrospray mass spectrometry (ES-MS) is the ionization of large (tens of kDa) molecules. Since many proteins are polar and immiscible in organic solvents, and many buffers and detergents that are compatible with proteins are entirely incompatible with ES-MS, finding an appropriate buffer can be difficult. Salt buffers form adducts which interfere with protein signal and can cause signal suppression

(King et al., 2000).

Attempts were made to use ES-MS to confirm the identity of the protein in the purified sample using an API 3000 system as well as a QTRAP 5500 system. The 5500

QTRAP was abandoned early since the m/z range only goes up to 1200, which is at the low end of the range in which c-phycocyanin is expected to appear based on possible charge states (Herrero et al., 2005; Simó et al., 2005). Charge states identified in the literature range from +9 to +14, with masses in the 900-2200 m/z range. Results from the

API 3000 system were inconclusive due to low mass spectrometer resolution and sensitivity, low protein concentrations, and the presence of salts, which form salt adducts with the protein even at low salt concentrations. 34

Salt adducts are problematic in ES-MS since this instrument reveals several lower intensity peaks, showing each different protein-salt adduct rather than one clearly visible intense peak. In the presence of more than one type of salt and multiple salts per protein, the theoretical single high-intensity peak becomes a smear, indecipherable from background (Jackson et al., 2007). The spectra from the API 3000 suffered from this effect; indeed, no prominent protein peaks were discernable (spectra not shown). After extensive ultrafiltration (20 washes) eliminated as much salt as possible, mixing the protein solution (in 10 mM ammonium acetate) with denaturing buffer (1:1) caused the protein to precipitate out of solution. Since c-phycocyanin is highly water-soluble, the precipitation is likely due to the high organic content of the denaturing solution (0.1 % formic acid in acetonitrile). In addition, the API 3000 system was not equipped with an ion trap. This made it impossible to enhance the signal by pre-concentrating protein ions before they reached the detector.

Spectral data from the SolariX was more informative. Bradford assay measured the protein concentration at 2 pM; dilution in denaturing buffer decreased the final concentration to 1 pM. The following spectra are of c-phycocyanin without selenium incorporated. From direct injection with 1 pM denatured c-phycocyanin, two sets of protein peaks were immediately apparent (Figure 7). Resolution for these scans was around 250,000 mass resolving units, high enough to show fine structure. Fine structure of individual peaks shows Gaussian distribution of isotope peaks characteristic of proteins and peptides (Figure 8).

An obstacle arises during data analysis when approximating protein mass. An approximate estimate for protein mass obtained from the mass spectrum software 35

(DataAnalysis) is obtained by the averagine model. The averagine model provides a profile for a theoretical (or “average”) protein with the formula

C4.9384H7.7583N1.3577O1.4773S0.0417. Using a charge state ruler, the user can select relevant protein peaks, which are then assigned charge states. This is multiplied by multiples of

~1.0022 Da to obtain an approximate molecular mass by using the mass and charge information in the spectrum (Senko et al., 1995; Horn et al., 2000). However, averagine includes only carbon, nitrogen, oxygen, and sulfur at specified ratios and cannot account for variations in protein composition. To overcome this issue, the molecular mass can be calculated the traditional way, using charge states and mass to charge ratios from adjacent peaks (see Table 4) in the following manner:

A = [M + nH+]n+ B = [M + (n + 1)H+](n+1)+ Where A and B are adjacent peaks (A has charge state B+1), M is the mass of the protein, H+ is a proton, and n is the charge state of A.

Protein and peptide signals present a characteristic isotopic distribution, which helps distinguish protein signals from other compounds. The fine structure shows the isotopic variation of the most common elements found in proteins; the isotope distribution of a protein shows all of the relative abundances of its constituent isotopes.

Since carbon is the most abundant element in proteins, the mass difference between the highest peaks in each set should be the mass difference between the carbon isotopes.

Other isotopic signatures can appear in the spectrum as well. This will be important later, when predicting the mass spectrum of the selenium-substituted protein.

x 108! ! Intensity

1000! 1500! 2000! m/z!

Figure 7. Denatured c-phycocyanin electrospray full-scan spectrum. Estimated charges states range from +9 to +20. Peaks at m/z 963.46031 (+20) and 1014.17159 (+19) belong to the β-chain. Scans from SolariX XR at Bruker Daltonic. 36

x 108! ! Intensity

1500.0! 1502.5! 1505.0! 1507.5! m/z! Figure 8. Fine structure of protein peaks at m/z 1499 to 1512 showing normal protein isotopic distribution. Scan from SolariX XR at Bruker Daltonic using resolution of ~250,000 mass resolving units (using the FWHM definition).

Starting at m/z ~900 and scanning to m/z ~2200, a set of peaks exhibiting this typical protein distribution can be seen; this mass range is similar to c-phycocyanin α- and β-chains reported in the literature (Herrero et al., 2005 and Simó et al., 2005).

Approximate masses for the two chains were obtained using the charge state ruler in

DataAnalysis: 18,041 ± 23.45 Da for the α-chain and 19,253 ± 25.03 Da for the β-chain

(Table 4). The mass of one phycocyanobilin (588 Da) was added to the theoretical mass of CPC-α; 1176 Da was added to CPC-β to account for the presence of two chromophores. These approximate values correspond to values in the ES-MS literature 38 for c-phycocyanin extracted from Spirulina platensis: 18,186-18,188 Da for the α-chain,

19,220 Da for the β-chain (Herrero et al., 2005 and Simó et al., 2005).

Peaks were selected for fragmentation by electron-capture dissociation (ECD) and collision-induced dissociation (CID) using continuous accumulation of selected ions

(CASI). Combining data from ECD and CID can provide complimentary data for complete amino acid sequencing, since each provides different types of fragments. ECD and CID were effective methods for fragmenting both chains of the denatured c- phycocyanin. Typically for fragmentation, the highest peaks from the full scan are chosen. However, occasionally these peaks do not fragment as well as some of the less prominent peaks, as was the case here. The set of peaks that contained the peak at m/z

1388.42241 was determined to be the α-chain; the β-chain contained the peak at m/z

1376.04060. The α-chain was more susceptible to fragmentation by both ECD and CID than the β-chain. For the β-chain, CID of the peak at m/z 1376.04060 only fragmented into a few peaks; ECD of m/z 1289.25326 was a slightly more effective fragmentation method for the α-chain (spectra not shown). Amino acid sequencing results from fragmentation are shown in Table 5. The amino acid sequence matches (matched to sequences from UniProt database) account for a 60 % match with the α-chain and a 45 % match with the β-chain.

Amino acid sequencing was attempted with the data from the denatured protein scans to determine the parent algal species, S. platensis or jenneri. The algal strain obtained from CPCC had been conditionally identified as Spirulina jenneri, though this identification remains unconfirmed. The amino acid sequences for c-phycocyanin from S. platensis and S. jenneri are quite similar. From Spirulina platensis amino acid sequence 39 for c-phycocyanin, there was a 52 % match to the denatured protein. Spirulina jenneri amino acid sequence only matched 30 %. Neither the masses nor the primary structure were an exact match to either Spirulina species, so it cannot be confirmed that the original algal culture was either species. The data here show more similarities to S. platensis than S. jenneri, the species identified but unconfirmed by CPCC.

Table 5. Amino acid sequencing from fragmentation of denatured protein. α- or β-chain Fragmentation Fragmented Resultant Possible amino acid type peak (m/z) peak(s) sequence(s) α-chain CID 1388 1049 119-127 1325 32-44, 128-137 1380 51-63 1650 11-25, 107-120 α-chain ECD 1289 1032 2-9 1388 1639 133-148 β-chain CID 1284 1983 43-61, 84-103 1376 1143 1-10 2048 15-33 β-chain ECD 1376 1639 17-27

α-chain; 162 amino acids 10 20 30 40 50 MKTPLTEAVS IADSQGRFLS STEIQVAFGR FRQAKAGLEA AKALTSKADS 60 70 80 90 100 LISGAAQAVY NKFPYTTQMQ GPNYAADQRG KDKCARDIGY YLRMVTYCLI 110 120 130 140 150 AGGTGPMDEY LIAGIDEINR TFELSPSWYI EALKYIKANH GLSGDAAVEA 160 NSYLDYAINA LS

β-chain; 172 amino acids 10 20 30 40 50 MFDAFTKVVS QADTRGEMLS TAQIDALSQM VAESNKRLDA VNRITSNAST 60 70 80 90 100 IVSNAARSLF AEQPQLIAPG GNAYTSRRMA ACLRDMEIIL RYVTYAVFAG 110 120 130 140 150 DASVLEDRCL NGLRETYLAL GTPGSSVAVG VGKMKEAALA IVNDPAGITP 160 170 GDCSALASEI ASYFDRACAA VS 40

Non-denatured c-phycocyanin spectra were more obscured by salt adducts than the denatured protein; in the higher m/z range (>2000), distinct peaks were not discernable. However, desalting the protein in several ammonium acetate washes resulted in more distinguishable peaks, but there was evidence of salts remaining (Figure 9). Due to the number of peaks from sodium and potassium still present, it was not possible to accomplish amino acid sequencing from the non-denatured scans. Additionally, neither

ECD nor CID were very effective fragmentation techniques for the non-denatured protein, though fragmentation from both methods did produce a few product ions. ECD of peak m/z 2870 (Figure 10), resulted in a peak at m/z 3733, which was identified in

BioTools software as corresponding to one of four fragments from the β-chain: amino acid residues 26-60, 27-61, 66-98, or 99-135.

x 108! ! Intensity

2000! 2400! 2800! 3200! m/z! Figure 9. Mass scan of non-denatured c-phycocyanin after four desalting washes. Scans from SolariX XR at Bruker Daltonic.

!!!!!!+8! !!!!!!+7! 2256.85240! 2579.11588!

x 108!

!!!!!!+6! 3211.27233!

!!!!!!+7! 2752.6651! !!!!!!+6! ! 3008.80023! +4! 3733.93625! Intensity

2000! 2500! 3500! 3000! m/z! Figure 10. Non-denatured c-phycocyanin. ECD of m/z 2870. Peak at m/z 3733 is a product peak. Grey arrow shows diminished dissociated peak. Scans from SolariX XR at Bruker Daltonic.

Phycocyanobilin, C33H40N4O6, has a mass of 588.69 g/mol (Cole et al., 1967); there is no corresponding peak in the full-scan mass spectrum of the denatured protein, which indicates that this spectrum (Figure 7) is of the holoprotein. Previous studies

(Buehler et al., 1976; Sarada et al., 1999; Simó et al., 2005) show that under mild denaturing and electrospray ionization conditions, the chromophore remains covalently bonded to the intact protein. However, in CID fragmentation of m/z 1389.145 and

1375.269 (Figure 11), the base peak was found at m/z 587.286, which is indicative of phycocyanobilin loss from the intact protein. 42

+1! 587.28670!

x 108! phycocyanobilin!

+11! 1550.12492!

+14! +3! 1378.05260! 1774.21788! !

+6! 1000.84931! Intensity

+1! 751.39891! +5! 1049.53504! +3! 2104.03387!

500! 1000! 1500! 2000! a. m/z!

+1! 587.28670!

x 108! phycocyanobilin!

+14! 1376.04060! !

+12! 1325.76677! Intensity

+2! 1672.28690!

+1! 2048.94394!

b. 750! 1500! 2250! m/z!

Figure 11. CID of α- and β-chains, both with base peak (phycocyanobilin) at m/z 587.286. Grey arrows indicate fragmented peaks. a.) α-chain CID of m/z 1389.145. b.) β-chain CID of m/z 1375.269. Scans from SolariX XR at Bruker Daltonic. 43

Complete amino acid sequencing was not accomplished with the collected data.

Reported here is top down sequencing, but bottom up sequencing is often done to achieve more sequence coverage. Literature often reports well over 75 % amino acid sequence coverage by bottom up sequencing. This techniques includes an enzymatic digest (usually trypsin) followed by liquid chromatography before electrospray-mass spectrometry is attempted (Zubarev et al., 2000; Encinar et al., 2003). This technique allows for more thorough exploration of each digested fragment, and would likely provide more than 52

% amino acid sequence match.

3.4.3 Selenium-containing c-phycocyanin

Simulation of the isotopic signature of methionine versus selenomethionine on a high-resolution mass spectrometer shows the complexity of selenium-containing protein spectra, which emphasizes the generality of this complexity. The selenomethionine spectrum contains twenty-seven peaks; the methionine spectrum contains ten. In a protein that contains multiple selenium atoms, the spectrum becomes even more complex. It is, however, possible to determine the number of selenomethionine residues that are present in a protein, and even mixtures of different degrees of selenomethionine substitution in the same protein. Each protein with a different degree of selenium substitution would have a different mass and the total protein mass would increase by 48 Da per selenium substitution. Each charge state would be offset by 48 divided by the number of charges; the mass to charge shift would be greater than 1 Da per substitution for any protein with fewer than +48 charge states. 44

Indeed, identification of differing numbers of selenomethionine residues in a single protein have been elucidated (Encinar et al., 2003), and is applicable to the current issue (Figure 12). To facilitate identification, enzymatic digestion followed by liquid chromatography can help to study the protein in more detail, but even with unit resolution, this should not be necessary. Figure 12 shows multiple LC fractions depicting peptides containing different numbers of selenium atoms replacing sulfur. Each peptide signal is offset by m/z 47. Full scans and fragmentation (not shown) of each peak was done individually, but from the resulting mass spectra, we can see that none of peaks of interest would overlap at the same mass to charge ratios. This work was done on instruments with resolution of ~ 20,000 mass resolving units, so ultra high-resolution mass spectrometry is not necessary to distinguish these types of molecules.

As shown in Figure 12, the isotopic pattern of the selenium-containing protein differs from that of the fine structure of proteins without selenium (see Figure 8). This is due to the isotopic makeup of selenium and its distribution in the mass spectrum: 78Se contributes ~ 24 %; 80Se contributes ~ 50 %; plus four other minor isotopes that make up a shared ~ 26 %. So, the fine structure peaks become broader to take into account more isotopic variations. 45

Figure 12. Tryptic digestion of a selenium-containing protein, with different degrees of selenium substitution (figure from Encinar et al., 2003).

The selenium-containing c-phycocyanin electrospray spectrum is much more complex than that shown in Figure 9. Figure 13 shows the non-denatured c-phycocyanin with the non-denatured selenium-containing c-phycocyanin overlain. As expected, there are more peaks in the selenium-containing protein spectrum than in the spectrum without 46 selenium. However, the presence of salt adducts obscures the true nature of the differences in these spectra.

Se-CPC! CPC! ! Intensity[%]

1000! 2000! 3000! m/z! Figure 13. Full scans of non-denatured Se-CPC and CPC full scans. Scans from SolariX XR at Bruker Daltonic.

It is expected that a salt-free selenium-containing c-phycocyanin mass spectrum would resemble the mass spectrum from c-phycocyanin without selenium, though at a higher mass (+ 48 for each substituted selenium) and with more peaks, particularly in the fine structure.

IV. Conclusions and outlook

Experimental findings

This study has shown that Spirulina can be used as a prototype organism for selenium incorporation into c-phycocyanin, likely as selenomethionine. C-phycocyanin was successfully purified and then identified by electrospray mass spectrometry, and selenium was incorporated into most of the methionine residues, as determined by ICP-

MS. The change in selenium speciation from selenite to selenomethionine is of ecotoxicological importance since selenomethionine can bioaccumulate in higher organisms. Preliminary results suggest that selenium-substituted c-phycocyanin can be identified separately from the protein without selenium incorporation, which has implications for identifying selenium-containing proteins and polypeptides in complex mixtures.

Future preliminary degradation experiments

Selenium-containing c-phycocyanin will be spiked into water from the Otonabee

River and degradation over the course of several hours will be monitored by UV-Vis spectroscopy at 280 nm. The decomposed sample will be filtered through a 10 kDa and a

500 MWCO filter and total selenium will be measured in each filtrate and the retained each retained fraction. In the filtrate from the 500 MWCO, selenomethionine will be tested for, as well as selenate and selenite. Since proteins are a labile source of carbon and nitrogen, only small amounts of intact c-phycocyanin are likely to remain after degradation.

Importance of future research

Identification of organo-selenium compounds that may participate in bioaccumulation is important for understanding selenium biogeochemical cycling. To accomplish this, selenium-substituted c-phycocyanin can be degraded in natural waters.

Selenium-containing breakdown products can be quantified by coupling capillary electrophoresis (CE) to ICP-MS, and identified using CE-ES-MS with a high-resolution electrospray instrument. With high-resolution mass spectrometry, identification of selenium-containing compounds in a complex mixture should be possible due to selenium’s isotopic fingerprint. Then, it should be investigated whether or not the identified organo-selenium compounds are formed in natural environments, and if they are formed, it should be determined whether or not they are sufficiently stable to persist long enough to be taken up by other organisms.

It has been suggested (Maher et al., 2010) that algae play an important role in the biotransformation of inorganic selenium into bioavailable organo-selenium; the proposed research would provide insight into this theory. The proposed work would conclusively demonstrate whether or not algal selenium biotransformation is implicated in bioaccumulation at the bottom of the food chain, leading to biomagnification in trophic transfer.

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