Cultivation, extraction and analytics of high-valuable

pigments

Margarida Caetano Afonso

Thesis to obtain the Master of Science Degree in

Biological Engineering

Supervisors: Prof. Dr. José António Leonardo dos Santos

Prof. Dr. Christoph Griesbeck

Examination Committee

Chairperson: Prof. Dr. Jorge Humberto Gomes Leitão

Supervisor: Prof. Dr. José António Leonardo dos Santos

Members of the Committee: Prof. Dr. Pedro Carlos De Barros Fernandes

October 2018

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The work presented in this thesis was performed at Algal Biotechnology Department of

Management Center Innsbruck (Innsbruck, Austria), during the period March-July 2018, under the

supervision of Prof. Dr. Christoph Griesbeck. The thesis was co-supervised at Instituto Superior Técnico

by Prof. Dr. José António Leonardo dos Santos.

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Acknowledgments I would like to express my sincere gratitude to Professor Christoph Griesbeck for the opportunity

of doing my master thesis at the MCI and for the warm welcoming. I would also like to thank professors

Peter Leitner and Daniel Nothdurfter, for all the guidance and support they gave during this work and

for always being available to help me and to discuss the results and the next steps of my work.

To Professor José Santos, my supervisor at IST for the guidance, help and support along the entire

thesis.

To all the members of the Algal Biotechnology Department at the MCI who welcomed me and helped

whenever I needed.

To Vivi and Kevin, for all the moments of fun, happiness and laughter in and out of the laboratory. I wish

you all the luck in the world and I hope we share a lot more unforgettable moments together.

To Tiago my best friend, for his irreplaceable friendship, even though this last step took us through

distant paths, our friendship remains the same.

To all the friends I made in the last five years, IST gave me a lot of headaches but it also gave me a lot

of joy, and by the most part I owe it to you.

Last but not least, I would like to thank my parents, my sister and my grandparents, for their endless

support, love and for the continuous encouragement through all my life and in particular through the

development of this thesis.

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Abstract High valuable pigments have gained attention in the science field, due to their many applications

as strong antioxidants with health-enhancing properties. A rise of consumer’s awareness for the adverse

effects of synthetic products and the correlation between natural products with health lead to a consumer

preference for natural compounds over synthetic ones. Microalgae appear as the solution for a natural

source, as they are known to synthetize a wide range of high valuable molecules and in stress conditions

accumulate them.

This project focuses on the cultivation of Chromochloris sp., a strain known to enhance the

canthaxanthin accumulation, the development of a fractionation procedure and an HPLC technique that

allows the identification and quantification of high valuable pigments.

Six different cultivation conditions were tested regarding algal growth and carotenoid content.

Two-stage cultivation achieved the highest growth rate (0.15 day-1) and the highest cell density (19.4

g/L) suggesting that a photoautotrophic growth followed by a mixotrophic stage enhances algal growth.

High salinity cultivation obtained the highest canthaxanthin content (21.74 mg/g), while N and P

deficiency cultivation obtained the highest astaxanthin content (27.88 mg/g) suggesting that these

conditions induce canthaxanthin and astaxanthin accumulation, respectively.

Six approaches were tested in the fractionation procedure. Solvent acetonitrile and mixture ethyl

acetate: Acetonitrile (2:1) showed high extraction rates and carotenoid content indicating their potential

as fractionation agents. However, the fractionation of astaxanthin from canthaxanthin was not

accomplished.

An HPLC method was developed and allows the identification and quantification of

canthaxanthin free form and one astaxanthin-di-ester.

Keywords: microalgae, Chromochloris sp., carotenoid, astaxanthin, canthaxanthin, cultivation,

extraction, fractionation.

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Resumo Os pigmentos têm ganho atenção na comunidade científica devido às suas variadas aplicações

como antioxidantes. A recente preocupação do consumidor com os efeitos adversos de produtos

sintéticos fazem os consumidores preferir produtos naturais em detrimento dos sintéticos. As

microalgas aparecem como uma fonte natural, pois são organismos capazes de sintetizar uma grande

variedade de pigmentos e em condições de stress de os acumular.

Este projecto foca-se no cultivo da estirpe Chromochloris sp., conhecida por realçar a

acumulação de cantaxantina, no desenvolvimento de um processo de fracionamento e de um método

de HPLC para identificar e quantificar carotenoides.

Seis condições de cultivo foram testadas em termos de crescimento celular e teor de

carotenoides. O cultivo em duas fases obteve a maior taxa de crescimento (0.215 dia-1) e a maior

densidade celular (19.4 g/L) sugerindo que um crescimento fotoautotrófico seguido de uma fase

mixotrófica aumenta o crescimento celular.

O cultivo com elevada salinidade obteve o maior teor em cantaxantina (21.74 mg/g) enquanto

o cultivo com deficiência em N e PO4- obteve o maior teor em astaxantina (27.88 mg/g) sugerindo que

estas condições induzem a acumulação de cantaxantina e astaxantina, respectivamente.

Seis métodos foram testados no processo de fracionamento. O solvente acetonitrilo e a mistura

etil acetato:acetonitrilo (2:1) mostraram um alto teor de carotenoides o que sugere o seu potencial como

agentes fraccionantes. A separação dos pigmentos astaxantina e cantaxantina não foi conseguida.

O método de RP-HPLC foi desenvolvido e permite a identificação e quantificação de dois

pigmentos: cantaxantina (livre) e astaxantina (diester).

Palavras-chave: Microalgas; Chromochloris sp.; Astaxantina; Cantaxantina; Cultivo; Extracção;

Fraccionamento

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Table of Contents

1. INTRODUCTION .............................................................................................................................. 1

1.1 Background ........................................................................................................................... 1

1.2 Literature Overview ............................................................................................................... 1

1.2.1 Microalgae ....................................................................................................................................... 1

1.2.2 Carotenoids ..................................................................................................................................... 2

1.2.3 Astaxanthin ...................................................................................................................................... 6

1.2.4 Canthaxanthin.................................................................................................................................. 8

1.2.5 Microalgae Cultivation Parameters .................................................................................................. 9

1.2.5.1 Carbon Source Manipulation ................................................................................................. 10

1.2.5.2 Nitrogen Deficiency ................................................................................................................ 11

1.2.5.3 Phosphorus Deficiency .......................................................................................................... 11

1.2.5.4 Salt stress .............................................................................................................................. 12

1.2.5.5 High light Intensity ................................................................................................................. 12

1.2.5.6 Temperature and pH .............................................................................................................. 13

1.2.5.7 Two stage cultivation ............................................................................................................. 13

1.2.5.8 Co-culture Strategy ................................................................................................................ 14

1.2.6 Extraction ....................................................................................................................................... 14

1.2.7 Fractionation .................................................................................................................................. 15

1.2.8 Analytics ........................................................................................................................................ 16

1.3 Aim of the thesis .................................................................................................................. 18

2. MATERIALS AND METHODS ...................................................................................................... 19

2.1 Materials .............................................................................................................................. 19

2.1.1 Algal strains ................................................................................................................................... 19

2.1.2 Chemicals ...................................................................................................................................... 19

2.1.3 Solvents ......................................................................................................................................... 19

2.1.4 Glass Beads .................................................................................................................................. 20

2.1.5 Media Composition ........................................................................................................................ 20

2.1.6 Lipopolysaccharide (LPS) assay kit ............................................................................................... 21

2.1.7 Software ........................................................................................................................................ 21

2.2 Methods ............................................................................................................................... 21

2.2.1 General Overview of the process ................................................................................................... 21

2.2.2 Biomass Production ....................................................................................................................... 22

2.2.2.1 Pre-Culture ............................................................................................................................ 22

2.2.2.2 Biomass Production (Scale Up process)................................................................................ 22

2.2.2.3 First Cultivation Setup ............................................................................................................ 22

2.2.2.4 Second Cultivation Setup....................................................................................................... 23

2.2.2.5 Summary ............................................................................................................................... 24

2.2.2.6 Analysis ................................................................................................................................. 24

2.2.2.7 Proof of purity ........................................................................................................................ 25

2.2.3 Harvesting and freeze drying ......................................................................................................... 25

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2.2.3.1 Procedure .............................................................................................................................. 25

2.2.3.2 Harvested Biomass Labeling Code ........................................................................................ 26

2.2.4 Lipid Content Analysis ................................................................................................................... 26

2.2.5 LPS Assay ..................................................................................................................................... 26

2.2.5.1 Procedure .............................................................................................................................. 26

2.2.5.2 LPS Assay Code .................................................................................................................... 28

2.2.5.3 List of LPS assay samples ..................................................................................................... 28

2.2.6 Extraction ....................................................................................................................................... 29

2.2.6.1 Standard Procedure ............................................................................................................... 29

2.2.6.2 Experiment with different glass beads size ............................................................................ 29

2.2.6.3 Extraction Results Analysis.................................................................................................... 29

2.2.7 Fractionation .................................................................................................................................. 29

2.2.7.1 Procedure .............................................................................................................................. 29

2.2.7.2 Fractionation’s sample code .................................................................................................. 30

2.2.8 Quantitative and Qualitative Analysis of high valuable pigments ................................................... 31

2.2.8.1 Procedure .............................................................................................................................. 31

2.2.8.2 HPLC Results Analysis .......................................................................................................... 32

3. RESULTS AND DISCUSSION ...................................................................................................... 33

3.1 Cultivation ............................................................................................................................ 33

3.1.1 Study of the influence of bacterial co-culture in cell growth of Chromochloris sp. and carotenoid

content 33

3.1.1.1 Proof of purity ........................................................................................................................ 34

3.1.1.2 Growth Curve and Rate Analysis ........................................................................................... 35

3.1.2 Study of Stress Conditions: N and PO4- Deficiency and High Salinity in cell growth ...................... 36

3.1.3 Comparison mixotrophic with phototrophic conditions in cell growth ............................................. 37

3.1.4 Study of the impact of photoperiod regime in cell growth .............................................................. 38

3.1.5 Cultivation Summary and Discussion ............................................................................................ 39

3.2 Lipid Content in Chromochloris sp. (1st Cultivation Set) ...................................................... 42

3.3 LPS assay: Study of Toxicity ............................................................................................... 45

3.3.1 Impact of Extraction procedure in Sample Toxicity ........................................................................ 45

3.3.2 Impact of Filtration procedure in Sample Toxicity .......................................................................... 47

3.4 Extraction ............................................................................................................................. 48

3.4.1 Impact of glass beads size ............................................................................................................ 48

3.5 Fractionation ........................................................................................................................ 50

3.5.1 Comparison between different methods ........................................................................................ 51

3.6 Analytics .............................................................................................................................. 54

3.6.1 Method Development ..................................................................................................................... 54

3.6.2 Influence of cultivation conditions in astaxanthin and canthaxanthin content ................................ 57

3.6.3 Comparison between different glass beads size in carotenoid content ......................................... 61

3.6.4 Comparison between different fractionation methods .................................................................... 61

4. CONCLUSIONS AND FUTURE WORK ........................................................................................ 64

5. REFERENCES ............................................................................................................................... 68

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6. ANNEX ........................................................................................................................................... 74

6.1 Bold Basal Media Composition ........................................................................................... 74

6.2 LPS Assay Procedure ......................................................................................................... 75

6.3 Extraction SOP .................................................................................................................... 77

6.4 Chromatogram of Haematococcus pluvialis and Hordeum vulgare samples ................... 78

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List of Figures

Figure 1. 1 Schematic Diagram with the various current applications of microalgae.[7] ........................ 2

Figure 1. 2 Distribution of photosynthetically important or commercially applied pigments, across the

most common microalgal groups. [15] .................................................................................................... 3

Figure 1. 3 Metabolic pathway for the synthesis of carotenoids in most green algae and higher plants.

The astaxanthin synthesis is however limited to the microalgae. [1] ...................................................... 4

Figure 1. 4 Factors that influence the degradation of carotenoid by oxidation and mechanisms

associated to each one. [20].................................................................................................................... 5

Figure 1. 5 Chemical structure of astaxanthin. [24] ................................................................................. 6

Figure 1. 6 Chemical Structure of Astaxanthin stereoisomers. [25] ........................................................ 7

Figure 1. 7 Chemical Structure of free (A), monoester (B) and diester (C) astaxanthin. [30] ................. 7

Figure 1. 8 Canthaxanthin molecular structure. [38] ............................................................................... 8

Figure 2. 1 General Overview of the process of upstream and downstream of Chromochloris sp. carried

out in this thesis in order to analyze high valuable pigments. ............................................................... 21

Figure 2. 2 Picture illustrating the first set up of the cultivation (March to April) of Chromochloris sp. with

the respective identification of each photobioreactor. ........................................................................... 23

Figure 2. 3 Picture illustrating the second set up of the cultivation (May to June) of Chromochloris sp.

with the respective identification of each photobioreactor. .................................................................... 24

Figure 2. 4 Layout of the 96 well plate used in both LPS assays. The wells in blue were filled with sterile

water, the ones in yellow were used to samples, the green line was used to test the standards and the

orange line was filled with the blank. ..................................................................................................... 27

Figure 3. 1 Pictures illustrating the evolution of the first set of cultivation with four PBR’s. A: day 7 of

cultivation for PBR 1, 2 and 3 and day 0 for PBR 4. B: day 19 of cultivation for PBR 1, 2 and 3 and day

11 for PBR 4. C: Harvested biomass at day 30 of cultivation for PBR 1, 2 and 3................................. 34

Figure 3. 2 Microscope pictures of the first cultivation setup, PBR 2 and 4. In PBR 2 Chromochloris sp.

appears in co-culture with bacteria at cultivation day 6 (A) and 21 (B). In both pictures, the black arrow

indicates the bacteria contamination. In PBR 4 showing the purity ensured with no apparent

contamination visible at day 0 of cultivation (C) and at day 13 (D). (Magnification: 1000x) ................. 35

Figure 3. 3 Graphic representation of growth curves for PBR 1 bacterial co-culture (), PBR 2 bacterial

co-culture (), PBR 3 two-stage cultivation () and PBR 4 control () based on OD measurements

taken during the 30 days of cultivation. ................................................................................................. 36

Figure 3. 4 Graphic representation of growth curves for PBR 5 high salinity stress (), PBR 6 N and P

deficiency (), PBR 7 mixotrophic cultivation () and PBR 8 control () based on OD measurements

taken during the 30/45 days of cultivation. ............................................................................................ 36

Figure 3. 5 Graphic representation of growth curves for PBR 3 two stage cultivation (), PBR 7

mixotrophic cultivation () and PBR 8 photoautotrophic cultivation (). The arrow targets the time point

when glucose was added to PBR 3 turning the phototrophic cultivation in mixotrophic cultivation. ..... 37

Figure 3. 6 Microscope pictures of PBR 3: (A) at day 6 of cultivation (before adding glucose), (B) at day

21 of cultivation (after the addition of the C source). (Magnification: 1000x) ........................................ 38

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Figure 3. 7 Graphic representation of growth curves of PBR 4 – control, continuous light () and 8 –

control, 16:8 (L:D) () based on OD measurements taken during the 30/45 days of cultivation. ........ 39

Figure 3. 8 Total lipid content in percentage of Dry Weight obtained for biomass harvested of

Chromochloris sp. in the first cultivation setup. The data obtained is the result of a mean between the

results of two samples and the deviation is represented be the error bars. .......................................... 43

Figure 3. 9 Endotoxicity level assay performed with biomass (B) and extract (E) samples of PBR 1, 2, 3

and 4. The sample concentration described is 0.2 mg/mL and all the samples were analyzed without

filtration (U). The results are expressed as endotoxin units per mililiter, (EU/mL). ............................... 46

Figure 3. 10 Endotoxin level asay of biomass (B), extract (E) and liquid (L) samples of PBR 1, 3.3, 4, 5

and 6. The samples are also classified as filtered (F) or unfiltered (U). The sample concentration chosen

was 0.2 mg/mL and the results are expressed as endotoxin units per mililiter, (EU/mL). .................... 47

Figure 3. 11 Effect of different glass bead sizes in cell disruption. The average extraction yield is reported

as a percentage of Dry Weight and it is the result of the mean between two independent samples.... 49

Figure 3. 12 Microscope pictures of samples from the extraction procedure with different glass beads

size after the first centrifugation cycle (Magnification: 1000x) A: 0.25 – 0.50 mm; B: 0.75 – 1.0 mm; C:

4.0 ± 0.3 mm; D: 0.25 – 0.50 mm + 4.0 ± 0.3 mm ................................................................................. 50

Figure 3. 13 Relative Polarity Scale of solvents, with the interval of solubility of astaxanthin in orange

and canthaxanthin in blue with the region where the solubility of the two carotenoids intersect. ......... 51

Figure 3. 14 Extraction yields of fractionation methods tested. All approaches are based in one replicate

with exception to method A, the standard method, where it was performed two replicates. ................ 52

Figure 3. 15 Chromatograms of canthaxanthin (150 µg/mL) (A) and astaxanthin (500 µg/mL) (B)

standards and respective retention times. The black arrow shows the peaks chosen to quantify in the

samples. ................................................................................................................................................ 55

Figure 3. 16 Calibration curve of canthaxanthin (left) and astaxanthin (right) using the general method

for quantification of pigments in samples. The equations obtained were Area peak= 0.33416

[canthaxanthin] – 0.05557, R² = 0.99998 for canthaxanthin and Area peak = 1.12697[astaxanthin] +

2.00414, R² = 0.999874 for astaxanthin. ............................................................................................... 56

Figure 3. 17 Chromatogram obtained after analysis of N and P deficiency cultivation of Chromochloris

sp. (PBR 5) and retention times identified by the software ChemStation. The analysis was performed by

injecting 5 µL of sample in a C18 5 μm (250x4 mm) column. The flow was kept at 1 mL/min during

washing, injection and elution and the temperate at 30ºC. ................................................................... 57

Figure 3. 18 Chromatogram obtained after analysis of Chromochloris sp. in control conditions (PBR 4)

with retention times identified by software ChemStation. The analysis was performed by injecting 5 µL

of sample in a C18 (5 μm 250x4 mm) column. The flow was kept at 1 mL/min during washing, injection

and elution and the temperate at 30ºC.Canthaxanthin and astaxanthin diester are marked by the black

arrow and unidentified significant peaks are marked by the blue arrow. .............................................. 57

Figure 3. 19 Chromatograms obtained after analysis of Chromochloris sp. in two-stage cultivation at A)

early stationary phase (PBR 3.1) and B) end of stationary phase (PBR 3.2) with retention times identified

by software ChemStation. The analysis was performed by injecting 5 µL of sample in a C18 (5 μm 250x4

mm) column. The flow was kept at 1 mL/min during washing, injection and elution and the temperate at

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30ºC. Canthaxanthin and astaxanthin diester are marked by black arrows and unidentified significant

peaks are marked by blue arrows. ........................................................................................................ 58

Figure 3. 20 Carotenoid content in different cultivation conditions. An average of the carotenoid content

is reported as mg of pigment per g of biomass. .................................................................................... 58

Figure 3. 21 Carotenoid content with different extraction procedures. An average of the carotenoid

content is reported as mg of pigment per g of biomass. ....................................................................... 61

Figure 3. 22 Carotenoid content in the five fractionation procedures studied. ...................................... 62

Figure 6. 1 Chromatograms obtained after the analysis of samples of Hordeum vulgare and

Haematococcus pluvialis by RP-HPLC. The analysis was performed by injecting 5 µL of sample in a

C18 column while the flow was kept at 1 mL/min during washing, injection and elution and the temperate

at 30ºC. The chromatograms were obtained by Innsbruck Botanical Institute. .................................... 78

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List of Tables Table 1. 1 Type of culture and respective energy and carbon source. [43] .......................................... 10

Table 2. 1 Chemicals used and companies from where they were purchased. .................................... 19

Table 2. 2 Solvents used and companies from where they were purchased. ....................................... 20

Table 2. 3 Glass beads used and company from where they were purchased. ................................... 20

Table 2. 4 Growth Medias used in the cultivation of microalgae (Chromochloris sp.) and its composition,

the abbreviation SL stands for stock solution. ....................................................................................... 20

Table 2. 5 Description of each photobioreactor of the cultivation and conditions of this process. ........ 24

The code used to label the harvested biomass for further procedures is as follows in table 2.6:Table 2.

6 Identification of the biomass from each PBR cultivated and harvested. ............................................ 26

Table 2. 7 Identification of the LPS assay samples. ............................................................................. 28

Table 2. 8 Methods used for fractionation and sequence of the solvents/solvent mixtures used in each

case. ...................................................................................................................................................... 30

Table 2. 9 Identification of the extracts obtain in each fractionation method. ....................................... 30

Table 2. 10 Columns used and methods tried in each case. ................................................................ 31

Table 3. 1 Exponential growth rates (µ) and duplication times (tD) of the first cultivation set. .............. 36

Table 3. 2 Exponential growth rates (µ) and duplication times (tD) of the second cultivation set. ........ 37

Table 3. 3 Photobioreactors cultivated, type of cultivation followed, strain and media tested, time of

glucose addition, type of stress induced and pH at the time, as well as, end measurements (OD, DW

and pH). ................................................................................................................................................. 39

Table 3. 4 Fatty acid profile of PBR 1 – 4 expressed as major fatty acids, quantity of saturated FA and

unsaturated fatty acids present in the samples. The data is expressed as a mean of two independent

measurements in percentage of DW. .................................................................................................... 44

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Abbreviations ACS American Chemical Society

ADSI Austrian Drug Screening Institute

BBM Bold Basal Medium

C Carbon

DCM Dichloromethane

DMSO Dimethyl Sulfoxide

DW Dry Weight

EDTA Ethylenediaminetetraacetic acid disodium salt solution

EU Endotoxin Unit

FA Fatty Acid

FHOO FH Wels Upper Austria

HPLC High Performance Liquid Chromatography

L: D Light: Dark

LOD Limit of Detection

LOQ Limit of Quantification

LPS Lipopolysaccharide

MAE Microwave-Assisted Extraction

MCI Management Center Innsbruck

MTBE t-butyl methyl ether

N Nitrogen

OD Optical Density

P Phosphorus

PBR Photobioreactor

PLE Pressurized Liquid Extraction

PSY enzyme phytoene synthase

RP-HPLC Reverse Phase High-Performance Liquid Chromatography

SC-CO2 Supercritical CO2

SCF Supercritical fluids

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SFE Supercritical Extraction

SOP Standard Operating Procedure

tD Doubling time

TLC Thin Layer Chromatography

UAE Ultrasound-Assisted Extraction

UHPLC Ultra High-Performance Liquid Chromatograph

USP United States Pharmacopeia

µ Exponential Growth Rate

1. Introduction

1.1 Background

High-valuable pigments commercialization is a well-established market with chemical

synthetized carotenoids as the principal product. However, throughout the years the preference of the

consumer for natural and environment friendly products and the potential applications in pharmaceutics

and cosmetics have given rise to a search for a sustainable process to produce these high valuable

pigments from a natural source. In this context, microalgae emerge as the obvious choice. [1,2]

Microalgae are excellent hosts for the mass production of carotenoids since these unicellular

microorganisms have high carotenoid content, fast growth and many other advantages. [2] Furthermore,

even though in the last few decades the research on carotenoids from microalgae has advanced it is

still required to further investigate on carotenoid production, the optimal cultivation conditions for each

strain, the optimal purification process, etc. [1–3]

The next section gives an overview of the process of obtaining high valuable pigments in specific

astaxanthin and canthaxanthin, from the source (microalgae) to the cultivation parameters as well as

the downstream process with focus on the analytics. The second chapter introduces the materials and

methods used to develop the practical work done while the third chapter describes the results obtained

and shows the discussion of these results. The fourth section shows the conclusions of this project and

the future prospects and recommendations for further work.

1.2 Literature Overview

1.2.1 Microalgae

Microalgae are unicellular organisms that can survive in a wide range of different habitats such

as fresh and salt water, marine and soil environments. [4,5] They are generally eukaryotic organisms,

even though cyanobacteria that are also considered microalgae are prokaryotic. [5]

Their size ranges from 5 μm to more than 100 μm [6] and regarding their morphology,

microalgae have different shapes and forms, not only among species but also among the different life

stages of one species. Many produce several different morphologies, for example, flagellate, coccoid,

and cyst stages. [4,7] Although, they can be referred as plant-like organisms, microalgae lack roots,

vacular systems, stems and leaves and have simple reproduction organs. [4,7,8]

Microalgae are autotrophic in nature and photosynthetic organisms [7], capable of converting

solar energy into biomass. They have an oxygen-evolving photosynthesis where they use inorganic

compounds (such as CO2) and light energy to synthetize bio-compounds with high nutritional value and

therapeutic functions such as lipids, proteins, carbohydrates, pigments and polymers. [2,5] Their

metabolism reacts to changes in the external environment with changes in the intracellular environment.

Therefore, the manipulation of the culture conditions stimulates the biosynthesis of specific compounds.

[5] These compounds have been investigated through the years to, not only understand their nature but

also to look for substances with applications in different fields of study. [4]

2

Regarding algae biodiversity, it has been estimated that about 200,000-800,000 species exist

of which about 50,000 species are described and from those, only about 100 have been produced at

laboratory scale. This means that there is still an enormous potential for new microalgae and products

that are still undeveloped. [1]

In recent years, the use of microalgae in biotechnology has been increasing with implications in

food, cosmetic and pharmaceutical industries. [9] The main applications of microalgae are showed in

Figure 1.1. Currently, many microalgae are acquiring increasing biotechnological interest because they

can produce several nutraceutical compounds. These molecules can be defined as nutrients from food

products that not only supplement the diet but also facilitate the prevention or treatment of a disease

and/or disorder. [10]

Figure 1. 1 Schematic Diagram with the various current applications of microalgae.[7]

1.2.2 Carotenoids

Carotenoids are in general richly colored molecules consisting of a class of more than 600

naturally occurring organic pigments synthesized by plants, algae and bacteria. They play different

physiological roles and because of that offer huge nutraceutical values. [10,11]

The chemical structure of the more than 600 different carotenoids is derived from a 40 - carbon

polyene chain, which can be considered as the backbone of the molecule. [12] This gives to the

molecules distinctive molecular structures, chemical properties and light-absorption features that are

essential for photosynthesis and, in general, for life in the presence of oxygen. [11,12]

Hydrocarbon carotenoids are denoted as carotenes, but oxygenated derivatives are known

specifically as xanthophylls - with oxygen being present as -OH groups, as oxi-groups or as a

combination of both. [13]

Xanthophylls are hydrophobic molecules; therefore, they are usually associated with

membranes and/or non- covalently bound to specific proteins. The primary carotenoids are generally

located in the thylakoid membrane, while secondary carotenoids are found in lipid vesicles either in the

plastid stroma or in the cytosol. [1,13,14]

A distinction is usually made between primary and secondary xanthophylls: primary ones are

structural and functional components of the cellular photosynthetic apparatus, so they are essential for

survival, while secondary ones encompass those produced by microalgae to large levels, but only after

3

exposure to specific environmental stimuli (via carotenogenesis). [1] Even though many algae are known

for producing these xanthophylls, at present, only alga H. pluvialis is able to synthesize and accumulate

it in large quantities so it can be commercially developed in a large scale. [14]

Green Algae are not only capable of synthesizing all xanthophylls synthesized by higher plants

but also additional ones such as loroxanthin, astaxanthin and canthaxanthin. [13,14] Furthermore, from

an evolutionary perspective, different microalgae synthetize different pigments and most microalgae

groups only contain a subset of pigments (Figure 1.2). [15]

While astaxanthin is only found in the Chlorophyta phylum, with a selective number of

Chlorophyta being able to overproduce this pigment, canthaxanthin is found not only in Chlorophyta but

also in some Cyanophyta. From Chlorophyta phylum are part H. pluvialis, Chlorella zofingiensis the

main producers of astaxanthin as well as, Dactylococcus dissociatus MT1, the most promising source

of canthaxanthin in the present. [15] Furthermore, estimation numbers show that just in the Chlorophyta

phylum there are 3000 species yet to be described, enhancing the fact that there still potential sources

of carotenoids undescribed. [16] The elucidation of the different pigments synthesis by each phylum is

also important in the clarification of the metabolic pathways of the production of carotenoids. [15]

Figure 1. 2 Distribution of photosynthetically important or commercially applied pigments, across the most common microalgal groups. [15]

4

Even though in the past years the understanding of the biochemistry of the carotenogenesis has

made some progress it is still not fully elucidated how the carotenoids are synthesized. [14] Also, the

biosynthesis of carotenoids differs from species to species; however, almost all photosynthetic

microalgae or plant species share the common primary metabolic pathway. The biosynthesis of

carotenoids takes place in the chloroplast, with some specific steps located in the cytoplasm. [2]

The enzyme phytoene synthase (PSY) is among the key enzymes for carotenoid biosynthesis

in photosynthetic organisms since it carries out a rate-limiting step. The expression of PSY and other

synthase genes can be up regulated by environmental stresses, which can increase or decrease the

synthesis of carotenoids (Figure 1. 3). [2,14]

Figure 1. 3 Metabolic pathway for the synthesis of carotenoids in most green algae and higher plants. The astaxanthin synthesis is however limited to the microalgae. [1]

Regarding the purpose of carotenoids, they perform several functions in microalgae: they are

involved in light harvesting, but also contribute to stabilize the structure and aid in the function of

photosynthetic complexes. The intrinsic antioxidant activity of carotenoids constitutes the basis for their

protective action against oxidative stress; however, not all biological activities claimed for carotenoids

relate to their ability to inactivate free radicals and reactive oxygen species. [11,13]

Moreover, even though, carotenoids have widespread applications as food colorants, cosmetics

and feed additives, only recently the benefits of carotenoids for human health were better understood.

5

[17] Similar to the protective roles carotenoids played in microalgae and plants, these pigments also

provide a protective role for humans. The anti-oxidant property in general mediates the harmful effects

of free radicals, hence can potentially protect humans from compromised immune response, premature

aging, certain cancers, cardiovascular diseases, and/or arthritis. [1,17]

In the current days, the chemical synthesis of carotenoids is a well-established market. [1,13]

However, the use of these products in direct human consumption is limited due to potential safety

concerns. While natural carotenoids are usually a complex mixture of various isomers and are usually

found mixed with other bioactive compounds, synthetic carotenoids are dominantly all-trans compounds.

Due to the competitive inhibition among carotenoids for human absorption, the intake of certain synthetic

carotenoid isomers is considered not as safe as the intake of the natural occurring mixtures. [18,19]

Thus, the applications of synthetic carotenoids are limited to animal feed, colorants, preservants, etc.

The natural carotenoids have the advantage of lower toxicity and higher customer preference for

medicine or as supplements. However, due to current production technology limitations, there is still a

low expression of the natural carotenoids in the global market. [3]

A drawback in carotenoid production is their stability. This issue needs to be overcome before

carotenoids can be successfully introduced in the food market. [20] In their natural environment

carotenoids are well protected as they are incorporated in lipoproteins or membranes but in isolation or

in transformation to solution, they are more prone to isomerisation and oxidative degradation. [21] The

most common causes and chemical mechanisms associated with the degradation of carotenoids are

shown in Figure 1.4.

The favourable conditions for oxidation can cause the cleavage of the molecule thereby affecting

the stability, bioavailability and physiological properties of carotenoids. [21,22]

Figure 1. 4 Factors that influence the degradation of carotenoid by oxidation and mechanisms associated to each one. [20]

6

Exposure to light or high temperature are the two main factors that cause carotenoid

degradation by radical mediated oxidation. Moreover, factors as oxygen availability, oxidative enzymes,

lower pH, catalytic metals, presence of unsaturated lipids and pro-oxidants in the biomass accelerates

the degradation of these molecules. [21,23]

1.2.3 Astaxanthin

Astaxanthin also known as 3, 3’-dihydroxy-β, β-carotene-4, 4’-dione is a bright red keto -

carotenoid with high commercial value that is used in functional foods, dietary supplements, beverages,

cosmetics and the nutraceutical industry because of its super-high anti-oxidative activities.

[1,3,18,19,24] The molecular formula of astaxanthin is C40H52O4 and its molar mass is 596.4 g/mol. [24]

Regarding astaxanthin’s structure, the molecule is a hydrocarbon that contains two terminal ring

systems joined by a chain of conjugated double bonds also known as a polyene system (Figure 1.5).

The presence of the hydroxyl- and keto-groups on each ring, explains some unique features, such as

the ability to be esterified, a higher anti-oxidant activity and a more polar configuration than other

carotenoids. [25]

Furthermore, astaxanthin is considered one of the best natural carotenoids on protecting cells,

lipids and membrane lipoproteins against oxidative damage. [24,25] In fact, several studies have

reported astaxanthin’s potential in the prevention and treatment of various diseases, such as diabetes,

cancer, ocular diseases, cardiovascular diseases, skin diseases, male infertility, neurological diseases,

muscular soreness, and hypertension. [24–27]

Figure 1. 5 Chemical structure of astaxanthin. [24]

Astaxanthin can be found in nature, in aquatic environments. The molecule is biosynthesized

by microalgae or phytoplankton, the microalgae are then consumed by zooplankton or crustaceans that

accumulate astaxanthin and in turn are ingested by salmonids. [28] There are diverse natural sources

of astaxanthin, such as microalgae Haematococcus pluvialis, Chlorococcum, Chlorella zofingiensis, red

yeast, Phaffia rhodozyma and bacteria, Paracoccus carotinifaciens. [24]

In nature, astaxanthin can acquire different stereoisomers that differ in the configuration of the

two-hydroxyl groups in the molecule. There are three stereoisomers for astaxanthin: two enantiomers

(3R, 3’R and 3S, 3’S) and a meso form (3R, 3’S) (Figure 1.6). Of all 3 isomers, the meso form is the

most abundant in nature, with different organisms producing different stereoisomeric ratios of

astaxanthin. [25]

7

Since free astaxanthin is particularly sensitive to oxidation, in nature, the molecule can be found

either conjugated to proteins, as in salmon muscle or lobster exoskeleton, or esterified with one or two

fatty acids, which stabilize the molecule. [29] Besides stabilization, it was also proved that esterified

astaxanthin increases biological activities since it can be easily absorbed into the metabolism, when

compared to its free form. [27]

Figure 1. 6 Chemical Structure of Astaxanthin stereoisomers. [25]

The synthetic version of astaxanthin is produced as the unesterified xanthophyll and as a 1:2:1

mixture of the three stereoisomers: 3S-3’S, 3R-3’S and 3R, 3’R. While in nature, more specifically, in H.

pluvialis, astaxanthin can be found in a ratio of 70% monoesters, 25% diesters and 5% free-form with

the most predominantly form being the stereoisomer 3S, 3’S esterified (monoester) (Figure 1.7).

The esters (in Figure 1.7 shown as R groups) present in monoester and diester forms of

astaxanthin are usually fatty acids that differ from species to species. The most predominant form of

monoesters in H. pluvialis are C18:4, C18:3, C18:2, C18:1, C16:0 and C18:0 whereas the main

components of diesters are C18:1/C18:3, C18:1/C18:2, C16:0/C18:2, C18:1/C18:1, C16:0/C16:0 and

C18:0/C18:1. This ultimately leads to an increase of the molecular weight of astaxanthin and prejudices

its quantitative and qualitative analysis. [30]

Figure 1. 7 Chemical Structure of free (A), monoester (B) and diester (C) astaxanthin. [30]

8

As of now, the major market for astaxanthin is as a pigmentation source in aquaculture, primarily

in salmon and trout with an annual worldwide market estimated at US$270 million (2016). [31,32]

However, 95 % of this market consumes synthetic astaxanthin. Compared to this formula the natural

astaxanthin has a much stronger anti-oxidative activity for human health applications, with almost 20

times anti-oxidative power than the chemical synthetized version. [19,24] This allied to the consumer

demand for natural products makes the synthetic pigments much less desirable and provides an

opportunity for the production of natural astaxanthin. [24,28]

With the astaxanthin market in a great demand for food, feed, nutraceutical and pharmaceutical

applications all the efforts are promoted to improve astaxanthin production from biological sources

instead of synthetic ones. In the current bio-based industry, the green microalga Haematococcus

pluvialis has been regarded as ideal producer of commercial natural astaxanthin. However, there has

been some hurdles in the production of astaxanthin by H. pluvialis such as a slow growth and

susceptibility to microbial contamination in the scale up of the cultivation process. [24,26]

In recent years, the green microalga Chlorella zofingiensis has been used as a promising

alternative producer of natural astaxanthin mainly due to its high growth rate and ability to accumulate

astaxanthin under various stress conditions. Furthermore, C. zofingiensis has the additional advantages

of having a higher efficiency cell wall disruption and thus a higher recovery of high-value intracellular

metabolites than H. pluvialis. However, as of now the biomass yield is still higher in H. pluvialis so there

is a need to improve the astaxanthin accumulation in C. zofingiensis that would allow the industrial

production of natural astaxanthin by this microorganism. [33]

1.2.4 Canthaxanthin

Canthaxanthin is a diketo-carotenoid that was first isolated in edible mushrooms. [34] It has also

been found in green algae, bacteria, crustaceans, and various fish. [35] It has a chemical formula of

C40H52O2 and a molecular weight of 564.86 g/mol. Since canthaxanthin absorbs light in the visible region

of (λ=470 nm), the molecule appears as a red-orange lipophilic pigment. [36]

Because of its nonpolar polyene chain, canthaxanthin is insoluble in polar solvents, such as

water and ethanol, and more soluble in non-polar solvents such as acetone, ethyl acetate, toluene,

cyclohexane, 1,2-dichloroethane, and chloroform. The solubility of canthaxanthin improves with the

increasing of the temperature. [15,16] Like astaxanthin, canthaxanthin is also known for its anti-oxidant

properties. This is due to the keto group present in both molecules, in conjugation with the polyene

backbone that is responsible for stabilizing the carbon-centred radicals more effectively than the polyene

backbone alone (Figure 1.8). [36,38]

Figure 1. 8 Canthaxanthin molecular structure. [38]

9

Like astaxanthin, canthaxanthin is used as a coloring agent in poultry feeds and fish feeds. [39]

However, this carotenoid unlike others is still not used as a nutraceutical albeit its functions as a

molecule with potential health benefits have been described, including free radical scavenging,

antioxidant and gene regulatory properties. [15,17] Furthermore, the molecular targets (e.g.,

transcription factors) by which canthaxanthin may mediate the potential health benefits are largely

unknown and need further investigation. Also, the fact that the toxicology safety of this carotenoid has

just been proven, after a past with toxicity concerns regarding the accumulation of canthaxanthin crystals

in the macula lutea membranes of the retina contributes to the preference of astaxanthin over

canthaxanthin. [38,40]

Canthaxanthin worldwide market is estimated at US$75 million as of 2016 and as well as the

other carotenoids, is expected to continue to grow in the next few years, as more and more benefits and

potential applications are discovered in these molecules. [31] However, again like most of carotenoids

these days, the primary and major source of commercial canthaxanthin is chemical. In the case of

canthaxanthin, the use of a chemical source instead of a natural one is due to lower concentrations and

lower yields in the natural environment. However, the consumer demand for natural products over

synthetic products, makes the need for a natural source of canthaxanthin urgent. [41]

The potential natural sources of canthaxanthin are: the green alga Haematococcus pluvialis,

the microalgae Chlorella zofingiensis and Dactylococcus dissociatus MT1, with the last being the most

promising one. [18,19] Even though that for now the commercialization a natural synthetized

canthaxanthin is still unfeasible, the discovery of metabolic pathways conjugated with genetic

engineering and the optimization of cultivation conditions will ultimately lead to a commercially available

source of natural canthaxanthin in the future. [41]

1.2.5 Microalgae Cultivation Parameters

The growing demand for natural sources of carotenoids targets microalgae as the primary

candidate. The need for obtaining the maximum yield of biomass and carotenoid accumulation leads to

an optimization of the microalgae growth as the source of the carotenoid. The strategies to accomplish

this aim are generally to find the optimal cultivation conditions, as well as manipulate the pigment

metabolic pathways and their regulation. [1,15,43]

Focusing on optimizing the cultivation parameters, the use of photobioreactors is the most

common way to cultivate microalgae not only because it allows a fast growth but also because it

minimizes possible contaminations. However, it is necessary to maintain a strict control of cultivation

conditions such as: nutrients, pH, temperature, aeration rate, CO2 concentration, light regime, inoculum

stage and size. [43,44]

There are four major types of cultivation conditions for microalgae: photoautotrophic,

heterotrophic, mixotrophic and photoheterotrophic cultivation (Table 1.1). Essentially, they differ in the

energy and carbon source. The most common type used to cultivate microalgae is the photoautotrophic

that uses light as energy source and inorganic carbon as carbon source. [43,45]

10

Table 1. 1 Type of culture and respective energy and carbon source. [43]

The manipulation of the cultivation conditions allows the optimization of the biomass production

and the carotenoid synthesis. Deviating the culture conditions from the optimal introduces stress in the

algae, which makes them produce or overproduce some type of metabolites to contradict the stress.

Among the stress factors that can be applied there are the nutrimental and the physical. [43] The

nutrimental factors are considered as a manipulation of culture media composition (carbon source,

nitrogen, phosphorus and iron deficiency), while physical are described as a manipulation in the

operation conditions and external factors that affect the microalgae growth (high light intensities,

temperature, pH, salinity, etc.). [43,46,47]

The limitation of essential nutrients such as nitrogen in the culture medium increases the content

of metabolites of interest like lipids and pigments, being this strategy a scalable technology in mass

cultures, providing the same effect as the use of external factors such as light intensity. [43,48]

1.2.5.1 Carbon Source Manipulation

Carbon (C) is the most prominent element of microalgae that accounts for approximately 50%

of the microalgal biomass. Carbon dioxide is the primary inorganic carbon source for photosynthesis of

C. zofingiensis. In photoautotrophic growth conditions, CO2 concentration has a crucial impact on the

growth of C. zofingiensis. Low CO2 is unable to provide enough carbon to support efficient

photosynthesis while high CO2 causes a pH decrease in the culture medium, which may in turn inhibit

the algal growth.[43] Besides the importance of CO2, C. zofingiensis is also capable of using organic

carbon sources such as glucose and acetate being in this case under photoheterotrophic conditions.

[49]

One study investigated the effect of sugars in the growth of C. zofingiensis. The results indicated

that glucose, fructose, mannose and sucrose were efficiently consumed by the alga for rapid growth

whereas lactose and galactose were poorly assimilated and could not support robust algal growth. Also,

in this study, glucose increased the astaxanthin content on the algae while lactose led to the lowest

content of this carotenoid in the cells. [50]

Under heterotrophic conditions and with glucose as the sole carbon source, the cell density and

astaxanthin content of C. zofingiensis is associated with the initial glucose concentration. Increasing the

glucose concentration leads to higher final cell densities until the limit of 30 g/L, where there is no

significant increase in the cell density and can also lead to a decrease of the growth rate. This can be

explained by the concept of substrate inhibition, with the glucose inhibiting the algal growth. [49,51]

The astaxanthin accumulation in C. zofingiensis is also related to the initial glucose

concentration and enhanced astaxanthin content is achieved with high glucose concentration (30 g/L),

11

possibly because high glucose, when assimilated for glycolysis, provides more carbon precursors

entering the astaxanthin biosynthetic pathway. It is worth noting, however, that a further increase in

sugar concentration fails to enhance astaxanthin level, indicating the presence of other limiting factors

in addition to carbon precursors. [33,49,51]

1.2.5.2 Nitrogen Deficiency

Nitrogen (N) is a very important element for algal growth, as a main constituent of protein and

genetic material and is one of the most abundant elements in microalgae. The most common ways of

supplementing N to the media is in the form of nitrate, urea, and ammonia. [43]

In microalgae culture, N stress is carried out either by N depletion or N limitation. Under N

depletion, microalgae grow in a medium lacking an N source, while under N limitation there is a constant

but insufficient supply of N. If nitrogen supply is limited in proportion to other elements, photosynthesis

may continue but the resultant metabolites will include a smaller proportion of nitrogen-rich components,

like carotenoids and more energy-rich components such as lipids and carbohydrates. [43,52]

One study reported the influence of N deficiency in the alga Haematococcus pluvialis under the

conditions of high and low light intensities in astaxanthin accumulation. The results suggested that N

deficiency has a greater effect than light intensity on astaxanthin synthesis. This behaviour proposes

the use of N limitation as an economic strategy for carotenoids production rather than application of high

light intensities. [53]

In heterotrophic cultures of C. zofingiensis, nitrogen availability also plays a critical role in

astaxanthin accumulation. In this kind of cultures, it is necessary to look at the carbon/nitrogen (C/N)

ratio, that controls the switch between protein and lipid synthesis and that is usually employed to show

the combined effect of carbon and nitrogen on astaxanthin synthesis. In fact, it was reported that higher

C/N ratios trigger the accumulation of astaxanthin. This may be explained by the excess carbon that

enters the carotenoid biosynthetic pathway. [43,49]

Other studies with autotrophic cultures, reported that nitrogen is an important factor influencing

intracellular accumulation of secondary carotenoids, especially astaxanthin, and that nitrogen

limitation/starvation generally causes the enhanced synthesis of astaxanthin in C. zofingiensis.

However, it is worth noting that nutrient limitation/deficiency can inhibit algal growth and may even lead

to a complete cessation of growth. [49,54]

1.2.5.3 Phosphorus Deficiency

Phosphorous (P) is an important rate limiting nutrient in many ecosystems. It is an element

required for the normal growth of microalgae and is also associated with the rate of phytoplankton

growth. This nutrient plays a major role in ATP production for energy metabolism, phospholipids

synthesis and the production of other cell components. Phosphorus is supplemented to algae in the

phosphate form, usually as H2PO4− or HPO4

2−. [43,49]

It has been reported, by one study that phosphorus limitation causes a decrease in the

microalgae growth but has no effect in the carotenoid accumulation in Chlorella vulgaris. [55] Also, in C.

12

zofingiensis this nutrient plays a much less prominent role on cell growth and astaxanthin content than

nitrogen, which is evident by the fact that phosphate at the range of 0.04–0.33 g·L−1 causes no significant

change in specific growth rate, final cell density, or astaxanthin content. [49]

In another study, where a comparison between different cultivation conditions was made it was

observed that in heterotrophic cultures the final biomass production under phosphate deficiency was

lower, whereas under mixotrophic and autotrophic conditions maximal phosphate assimilation was

observed. These findings suggested that illumination and autotrophic growth might significantly increase

the metabolic requirement for phosphorous in C. pyrenoidosa. [43,56]

1.2.5.4 Salt stress

The effect of salt stress on biomass and pigment production has been studied mostly in fresh

water microalgae species and it suggests that it can replace the light intensity stress in the accumulation

of carotenoids. [43]

C. zofingiensis is a freshwater alga that has a moderate tolerance to salt: up to 0.1 M NaCl with

just a slight impact on algal growth; increasing the salt concentration to 0.2 M and 0.4 M, however,

reduces significantly the final cell density by 30% and 50%, respectively. [57] Nevertheless, the addition

of 0.2 M salt results in the highest astaxanthin yield, up to 60% higher than that of the salt-free cultures.

[49,57] Similarly, it has been reported almost no difference in cell density with a salt concentration range

of 0–0.08 M NaCl. However, the highest astaxanthin yield was achieved with 0.08M NaCl and further

increase of salt concentration caused severe decrease of the yield. [58] The difference between the two

studies may be explained by the different culture conditions.

Other studies reported that low-light intensity and salt stress leads to a higher accumulation of

canthaxanthin, suggesting that in salt stress conditions, light doesn’t play a role in the accumulation of

canthaxanthin but is essential when it comes to astaxanthin accumulation. [59]

Salinity also plays an important role in the astaxanthin accumulation in H. pluvialis. Studies

reported that high NaCl concentrations caused an increase on the carotenoid accumulation and a

decrease in the algae growth. [60]

1.2.5.5 High light Intensity

The light intensity is one of the most important factors affecting photosynthesis. The amount of

light that microalgae receive influences the way that these organisms conduct their metabolic activities.

Furthermore, numerous studies analysing the light effect on microalgae production of pigments,

unsaturated fatty acids, carbohydrates and proteins content have been carried out. [43]

Not only in the production of metabolites, but light also plays a role in microalgae growth. There

is a range of light intensity for optimal algal growth, bellow which the light is insufficient that leads to a

slow growth and above which photo inhibition occurs and causes a growth attenuation or even cell

death. This range of light intensity varies according to cell density and the optical light paths that are

employed. [47,61]

13

In the case of carotenoid production, it has been documented that high light intensity promotes

the biosynthesis and accumulation of astaxanthin by Haematococcus pluvialis. High light levels

accelerate the growth process of the algae, increasing the rate of nutrient depletion and providing more

energy for astaxanthin biosynthesis. [62] In the case of microalgae C. zofingiensis, that can produce this

pigment in the dark (at lower rates), light is not a triggering factor for astaxanthin production but

enhances the accumulation of astaxanthin. [33,51]

In most studies, high light is not employed alone but accompanied by other stress factor that

induces carotenogenesis for astaxanthin accumulation. [47,51,60,62] However, the stress conditions,

cause a severe reduction in the algae growth and therefore a reduction in the final astaxanthin yield. To

solve this problem, microalgae are usually cultured with nitrogen and optimal light intensity for optimal

biomass production and after reaching the stationary phase, the stresses are induced in a two stage

cultivation strategy. [49]

1.2.5.6 Temperature and pH

One study investigated the effect of temperature in algal growth and astaxanthin accumulation

in C. zofingiensis. The results showed that the optimal temperature for cell growth is in the range of 20–

30°C and for astaxanthin accumulation between 25 and 30°C. [22,23] Regarding the pH, C. zofingiensis

can tolerate a relative wide range of pH (5.5–8.5) and pH 5.5 gives rise to the highest astaxanthin

content. [49]

Regarding algae Haematococcus pluvialis, one study investigated the optimal temperature for

cell growth and astaxanthin accumulation. The results showed that H. pluvialis cultures are sensitive to

changes in temperature and that the optimal temperature for biomass to grow is 20°C, as for astaxanthin

accumulation a temperature of 27°C was considered the optimal. [46] In terms of pH, one study

investigated the optimal value for cell growth and astaxanthin accumulation. The results showed that

pH 7.0 was best in terms of astaxanthin production and content. [63]

1.2.5.7 Two stage cultivation

Two-stage culture strategy has been used for enhancing accumulation of high-valuable

compounds in microalgae cells. In a large scale production of a microalgae metabolite, high cell density

is desirable to reduce the cost of the downstream process. The two-stage strategy technique is

composed by two distinct phases: the first one where occurs the production of green biomass under

optimal growth conditions and the second one exposing the algae to adverse environmental conditions

to induce the accumulation of high valuable pigments. [43]

One study developed a two-stage strategy with microalga Chlorella vulgaris and a first stage of

autotrophic growth followed by a mixotrophic stage. The results indicated a 64% increase of biomass in

the two stage cultivation when compared to a simple autotrophic cultivation. [64]

One other study compared the biomass growth and astaxanthin production in a two-stage

cultivation vs. one stage cultivation with microalga Haematococcus pluivialis. Results showed that a two

stage cultivation obtained a faster algal growth and a higher astaxanthin content. [65]

14

1.2.5.8 Co-culture Strategy

In recent years, investigators have started to question whether an axenic culture is the best way

to cultivate microalgae. This type of cultures are predominantly used in bio-industry, to meet tight safety

regulations. However, maintaining axenic cultures has proved to be expensive and labor intensive, given

the recurrent problem of contamination by bacteria, viruses, fungi, yeast, etc. and in their natural

environment microalgae thrive alongside other organisms. [66]

Microalgae co-culture is a field with potential but is yet to be further investigated. Small scale

trials have been carried out to investigate potential interactions between microalgae and other

organisms. Bacteria have been the focus of the investigation since many bacterial species are

endogenous in non-axenic microalgal cultures. [66]

One study reported an increment in pigment and lipid content, as well as, a higher algal growth

using a co-culture strategy with microalgae Chlorella sp. and microalgae-growth-promoting bacterium

Azospirillum brasilense. [67]

1.2.6 Extraction

One of the main obstacles to fully taking advantage of natural carotenoids potential and channel

it into the market refers to the ability to successfully and efficiently extract the pigment from the algal

cake. The extraction phase can be divided into three main processes: cell disruption; dehydration; and

recovery of the desired metabolite. [68]

A major practical problem in using microalgae as the source of the carotenoid is the need for

cell wall disruption in order to have access to the intracellular compound. Unfortunately, the cell wall is

rigid and extremely resistant to both chemical and physical disruptions. [1,69] The disruption can be

accomplished through a variety of ways, e.g., milling, ultrasound, microwave, freezing, thawing or

chemical attack. [1] Once the wall is disrupted, organic solvents are usually needed to extract the

pigment from the interior. [68]

On the other hand, carotenoids exhibit a variety of polarities, solubilities and chemical stabilities.

Therefore, a suitable solvent system must be selected based on the target carotenoids, which could

selectively and efficiently extract carotenoids with greater purity. [70] In a conventional extraction

process, since most carotenoids possess a high degree of hydrophobicity, their effective extractions

require the use of non-polar solvents, for example n-hexane, dichloromethane, dimethyl ether, diethyl

ether, etc. [70,71] However, solvents like acetone, octane, or mixtures of several organic solvents have

also been studied for the selective extraction of carotenoids. [70] Recently, with a need to maintain the

process as environment friendly as possible the use of several green solvents such as ethanol, limonene

and biphasic mixtures of water and organic solvents has been investigated for recovery of carotenoids.

However, extraction efficiency, selectivity and high solvent consumption still remain a limiting factor in

the conventional solvent extraction process. [68]

Currently, different astaxanthin extraction methods are used in algae, including organic

solvents, pre-treatment of encysted cells (cryogenic grinding or acid/base treatment), enzyme lysis,

15

mechanical disruption and spray drying. [72] In addition, since astaxanthin is a xanthophyll pigment,

around 95% of astaxanthin accumulated in H. pluvialis cells is esterified. Thus, the extraction of

astaxanthin can require a hydrolysis step in order to free the astaxanthin molecule. [24,71]

The current literature identifies that Supercritical Extraction (SFE) is the most extensively

studied non-conventional extraction technique for the recovery of carotenoids from algae and

microalgae. [70] Supercritical fluids (SCF) are now widely used for extraction purposes since they are

more efficient than the traditional extractions. Supercritical CO2 (SC-CO2) is by far the most common

supercritical fluid used in extraction of natural compounds and food processing. Moreover, the addition

of small amounts of other solvents (called co-solvent) increases the solvent power of the SCF. One

previous study tested the usage of the SC-CO2 in extracting astaxanthin from H. pluvialis cells. Results

showed that a supercritical fluid extraction obtains bigger yields that a traditional one and that adding

ethanol as a co-solvent increases even more the extraction yield. [73]

Other alternatives to the conventional extraction technique are the Pressurized Liquid Extraction

(PLE), the Microwave-Assisted Extraction (MAE) or the Ultrasound-Assisted Extraction (UAE). In

general, MAE and UAE are suitable for rapid extraction, whilst PLE reduces solvent consumption.

However, temperatures associated with these techniques can cause degradation of thermolabile

carotenoids and therefore they are not usually chosen to pigment extraction. [70]

Currently, the extraction and purification methods for high-quality lipids, carotenoids like

astaxanthin, have been studied at laboratory and small-scale level. [1] However, the recovery of

intracellular metabolites at large scale is still challenging since not every cell disruption, extraction or

purification methods are scalable. In the case of solvent extraction methods, the solvent has to be

accepted by regulatory agencies for animal or human consumption. Nowadays, the efforts should focus

in the reduction of product loss and equipment and energy costs associated with the extraction and

purification steps. In addition, large-scale downstream processing must be further developed in order to

achieve economically viable and environmentally friendly processes. [68,70]

1.2.7 Fractionation

Since the discovery of pigments from biological sources, the need for a purification process

became a topic of interest for research. [70] However, in the case of microalgae and the carotenoids

synthetized there is still extensive research needed. The variety of microalgae that synthetize pigments

and the parameters that influence the quantity of carotenoids produced play a big role on the process

of purification chosen. Additionally, the similarity between some carotenoids, their light and temperature

sensitivity are drawbacks in the downstream processing. [74]

Even though the process of fractionation has been successfully applied in proteins and lipids,

there is no documentation of fractionation processes to purification of carotenoids from a microalgae

source. One application previously study is the fractionation and identification of chemical constituents

from the leaves crude extracts of M. piperita. In this case, the process of fractionation was described as

a scheme of successive extractions with different solvents that were called fractions. The results showed

that different fractions collected different compounds resulting in a successful fractionation. [75]

16

Another study that research the fractionation of leaves extracts of Nyctanthes arbor tristis used

an approach that was based in a chromatographic step with a mobile phase with two organic solvents

and performed successive elutions increasing the percentage of one solvent over the other. As a result,

different fractions pooled different compounds with biologic potential. [76]

1.2.8 Analytics

The interest for naturally synthetized carotenoids like astaxanthin and canthaxanthin sparked

also interest in developing a method to identify and quantify these molecules. While talking about the

approaches that can be taken, a variety of methods has been employed to detect a wide range of

carotenoids from thin layer chromatography (TLC), to high-pressure liquid chromatography (HPLC) and

a combination of HPLC with mass spectrometry including MALDI-TOF. HPLC in combination with UV-

vis absorption detection is especially known to be the most common method for the separation and

determination of natural carotenoids.

HPLC methods described in the literature differ in resolution, sensitivity, and rapidity based on

the type of column (stationary phase) and solvent gradient (mobile phase) used. [77] Regarding the

HPLC method employed, both normal phase and reverse phase HPLC can be performed. However,

reverse phase HPLC is preferred since normal phase HPLC fails in separating non-polar carotenoids.

In reverse phase HPLC, there is an increase in the interaction between the analyte and the non-polar

stationary phase, which leads to an enhanced resolution of carotenoids.

As it was said before, it is well known that the separation of carotenoids is strongly influenced

by the properties of the stationary phase. The most commonly used columns in reverse phase HPLC

for carotenoid identification are: C8, C18 and C30. [78]

While C8 columns might be a good approach to identify carotenoids in a sample with unknown

carotenoids, the low sensitivity of the column is a disadvantage. Furthermore, C18 columns are usually

preferred to carotenoid separation, with an isocratic or gradient mode. However, C18 columns also have

a drawback: the inability to resolve geometrical isomers and positional isomers. This happens due to

insufficient thickness of the stationary phase that does not allow the penetration of carotenoid molecules,

which resolves in weak solute-bonded phase interactions. [78]

To maximize chromatographic resolution and selectivity, it was developed a method using a

RP-HPLC with a C30 column, which allows the interpretation of the carotenoids and its isomers. Another

ability of using a C30 column is the possibility to resolve cis/trans-carotenoids. Nevertheless, the

efficiency of the C30 column in resolving geometrical isomers is counterweigh with a requirement to do

longer runs (60 minutes or more) for a complete separation of carotenoids. It was also concluded that

the more hydrophobic C30 phase is the more efficient the separation of carotenoids is. [79]

Another approach to improve the identification and quantification of carotenoids was the usage

of the Ultra High-Performance Liquid Chromatography (UHPLC). This technique offers several

advantages over HPLC such as higher peak capacities, smaller peak widths, gain in sensitivity and

higher chromatographic resolution. The shorter analysis times also considerably save mobile phase

17

solvents. The carotenoid separation is performed by UHPLC with a C18 column since C30 columns are

not commercially available for this approach yet. [78,80]

The mobile phase is another parameter to have into consideration. However, it is impossible to

obtain a solvent system that can be applied to all samples. [81] For instance, various mixtures of solvents

have been used with reversed-phase chromatography, including water, methanol, acetonitrile, 2-

propanol, acetone, ethyl acetate, t-butyl methyl ether (MTBE), dichloromethane and chloroform. [78]

One study that focused on investigating a variety of isomers of astaxanthin from extracts of

Haematococcus pluvialis successfully used as mobile phase: solvent A (dichloromethane/ methanol/

acetonitrile/ water, 5.0:85.0:5.5:4.5, v/v) and solvent B (dichloromethane/ methanol/ acetonitrile/ water,

22.0:28.0:45.5:4.5, v/v) with a C18 column. The results showed that it was possible to elute separately

free astaxanthin and astaxanthin esters. Furthermore, a comparison of the spectrum obtained with

published data on the known carotenoids allowed almost total identification of the carotenoids and its

isomers. [82] Up until now, this study and specifically this HPLC method has been reproduced by other

investigators when the goal is to identify and quantify astaxanthin and its esters.

On the other hand, one previous study stated that a drawback in most of the methods using a

reverse phase chromatography is the relatively low retentivity towards the more polar derivatives

(xanthophylls). This implies a change in the mobile phase with substantial amounts of water to be

included in the eluent to ensure enough solute retention. Consequently, this can have a deteriorating

effect on peak shape and even cause partial solute precipitation on the column. [83]

In truth, the choice of the mobile phase has to take into consideration the stationary phase and

the carotenoid to identify and quantify. Besides the choice of the mobile phase, there are other

parameters to have into consideration such as: temperature, injection volume, solvent flow or the elution

mode. While talking about the elution mode it is possible to choose between an isocratic, a gradient or

a step elution. A gradient elution was previously described as a good system to cover the whole range

of carotenoid (polar and non-polar). In a previous study, an isocratic elution was used to study the

identification of lutein and zeaxanthin and the results described the method as a quick method to

screening lutein and zeaxanthin isomers in different food products. [79]

The temperature and the injection volume are two parameters needed to have into

consideration, especially when analysing carotenoids. As is known carotenoids are thermo-degradable,

so high temperatures will likely degrade the pigments and interfere with their analysis. [84] Regarding

the injection volumes, these should be kept to a minimum to prolong column life and reduce band

broadening while the injection solvent should be compatible with the HPLC mobile phase. [69]

The existence of different columns with different diameters, lengths, particle sizes, and different

degrees of effectiveness in the resolution of key pigments, different mobile phases and different modes

of elution makes a comparison of the different existing HPLC methods a subject of discussion.

Furthermore, the existence of different goals while analysing compounds, or the stability of the

carotenoids adds up to the discussion. [77] Moreover, the specificity of the biological matrix defines the

method of extraction, which in turn determines the success of the chromatographic step. [35]

18

1.3 Aim of the thesis

The ultimate goal of MCI’s algal project is to establish an overall economic process for the

production of valuable products from phototrophic microorganisms (algae, microalgae). This project

denominated (co)-Operation SKD is a partnership between the Management Center Innsbruck (MCI),

the FH Wels Upper Austria (FH OÖ) and the Austrian Drug Screening Institute (ADSI). More precisely,

it focus on the development of individual process steps or in competences that will favour the knowledge

of the entire process to allow the offer of purposeful services for potential company partners in the range

of pharmaceutical, cosmetics and food industry.

This thesis is part of this ongoing project and has the overall aim of study the cultivation,

extraction and analytics of high-valuable pigments, in specific astaxanthin and canthaxanthin

synthetized by Chromochloris sp., MCI-31 strain. This strain has demonstrated in the past evidence of

canthaxanthin accumulation in stress conditions and is part of a unique algal collection ASIB 505 from

the Botanical Institute of the University of Innsbruck (Austria) that comprises 1500 aero-terrestrial algae

cultures of the alpine regions of Europe.

The tasks defined for this work were as follows:

Investigation of different growth conditions and stress conditions on biomass production and on

synthesis of high valuable pigments with a focus on astaxanthin and canthaxanthin synthesis;

Optimization of the extraction process, in order to optimize the quantity of carotenoids extracted

by testing the cell disruption efficiency with different glass beads sizes;

Investigation of the lipid content in different cultivation conditions and how it can affect the

extraction procedure;

Investigation of the impact of the extraction procedure in the endotoxicity of the samples and

study the implementation of a filtration procedure to reduce the endotoxin level of the samples

for future applications;

Development of a fractionation process based on organic solvents, their polarity and its impact

in different high valuable pigments that is able to separate different carotenoids in different

fractions, in particular, to separate astaxanthin from canthaxanthin and vice versa;

Development of an analytic method using a reverse phase HPLC technique able to identify and

quantify the synthetized carotenoids.

19

2. Materials and Methods

2.1 Materials

2.1.1 Algal strains

The algal strain used in this work was originally from the ASIB 505 collection from the Botanical

Institute of the University of Innsbruck. The strain used in cultivation is V46 according to the labelling of

ASIB that was re-labelled as MCI-31, when it started to be cultivated in the Management Center

Innsbruck. According to the ASIB collection V46 is classified as Chromochloris sp., after molecular

genotyping tests.

2.1.2 Chemicals

The chemicals used in the practical work are described in Table 2.1. All the chemicals used are

ACS (American Chemical Society) grade with the exception of astaxanthin and canthaxanthin standard

that are USP (United States Pharmacopeia) grade.

Table 2. 1 Chemicals used and companies from where they were purchased.

Chemical Company

Glucose Carl Roth GmbH

Sodium Chloride Carl Roth GmbH

Ferrous(II) Sulfate Heptahydrate Carl Roth GmbH

Sodium Nitrate Carl Roth GmbH

Magnesium Sulfate Heptahydrate Carl Roth GmbH

Potassium Phosphate Dibasic Carl Roth GmbH

Potassium Dihydrogen Phosphate Carl Roth GmbH

Calcium Chloride Dihydrate Carl Roth GmbH

Zinc Sulfate Heptahydrate Carl Roth GmbH

Manganese(II) Chloride Tetrahydrate Carl Roth GmbH

Molybdenum(VI) Oxide Carl Roth GmbH

Copper(II) Sulfate Pentahydrate Carl Roth GmbH

Cobalt(II) Nitrate Hexahydrate Carl Roth GmbH

Boric Acid Carl Roth GmbH

Ethylenediaminetetraacetic Acid Disodium Salt Solution (EDTA)

Carl Roth GmbH

Potassium Hydroxide Carl Roth GmbH

Astaxanthin Standard United States Pharmacopeia

Canthaxanthin Standard Dr. Ehrenstorfer GmbH

2.1.3 Solvents

The solvents used in the extraction, fractionation and analytics were HPLC gradient grade.

Table 2.2 shows the list of solvents used in the procedures and the company from where they were

purchased. Regarding water, the extraction procedure was performed using Type III Water (distilled

water), while in the fractionation and analytics it was used Type I Water (HPLC gradient). In the case of

the LPS assay, the water used was sterile and it was provided in the LPS kit.

20

Table 2. 2 Solvents used and companies from where they were purchased.

Solvents Used Brand

n-Hexane Carl Roth GmbH

Methanol Carl Roth GmbH

2-Propanol Carl Roth GmbH

Ethyl Acetate Carl Roth GmbH

Ethanol Carl Roth GmbH

Dichloromethane (DCM) Carl Roth GmbH

Chloroform Carl Roth GmbH

Acetone Sigma Aldrich

Acetonitrile Carl Roth GmbH

Dimethyl Sulfoxide (DMSO) Sigma Aldrich

Acetic Acid Carl Roth GmbH

Sulphuric Acid Carl Roth GmbH

2.1.4 Glass Beads

The glass beads were used in the extraction and fractionation procedure. Table 2.3 clarifies the

glass beads used and their size as well as the company from where they were purchased.

Table 2. 3 Glass beads used and company from where they were purchased.

Glass Beads Company

Glass Beads (0.25 – 0.50 mm) Carl Roth GmbH

Glass Beads (0.75 – 1.0 mm) Carl Roth GmbH

Glass Beads (4.0 ± 0.3 mm) Carl Roth GmbH

2.1.5 Media Composition

The composition of each media used in this thesis practical work is presented in table 2.4. The

media was added to the photobioreactors and sterilized inside them with steam for 20 minutes at 121°C.

The media was made with deionized water (Type III). The media composition used was based on a

procedure originally from Culture Collection of Cryophilic Algae (CCCryo) (Annex 6.1).

Table 2. 4 Growth Medias used in the cultivation of microalgae (Chromochloris sp.) and its composition, the abbreviation

SL stands for stock solution.

Media Composition

3N-Bold’s Basal Media

(3N-BBM)

SL1 8.82 ×10-3 M NaNO3

SL2 3.04×10-4 M MgSO4.7H2O

SL3 4.28×10-4 M NaCl

SL4 4.31×10-4 M K2HPO4

SL5 1.29×10-3 M KH2PO4

SL6 1.70×10-4 M CaCl2.2H2O

SL7

3.07×10-5 M ZnSO4.7H2O

7.28×10-6 M MnCl2.4H20

4.93×10-6 M MoO3

6.29×10-6 M CuSO4.5H2O

1.68×10-6 M Co(NO3)2.6H2O

SL8 1.85×10-4 M H3BO3

SL9 5.53×10-4 M KOH

1.71×10-4 M EDTA.Na2

21

SL10 1.79×10-5 M FeSO4

.7H2O

1 mL of H2SO4 to acidify

Bold’s Basal Medium (N and

PO4- free)

3N – BBM media without SL1, SL4 and SL5

2.1.6 Lipopolysaccharide (LPS) assay kit

Pierce LAL™ Chromogenic Endotoxin Quantitation Kit from Thermo Scientific™

2.1.7 Software

The software used in this thesis project was:

MikroWin for microplate reading while performing the LPS assays;

ChemStation for LC 3D systems for analyzing chromatograms obtained with the HPLC;

MS-Office Excel for mathematical calculations, tables and graph plotting;

NIS-Elements BR 3.2 for obtaining microscope images.

2.2 Methods

2.2.1 General Overview of the process

Algae were cultivated in photobioreactors (PBR). This type of cultivation system has

advantages, such as: good heat and mass transfer, no moving parts, ease of operation, and low

operating and maintenance costs. [85] In this case, the starting point was a liquid culture in a conical

flask (600 mL), from which there was a scale up to the photobioreactors of 1 L. The cultivation was kept

running until the maximum yield of biomass is obtained and then the algae are harvested. In harvesting,

centrifugation is used to separate the biomass from the wasted media followed by two washing steps.

After harvesting, the biomass is freeze dried and then stored at -20 °C until it is used for extraction or

fractionation processes. After the downstream process, the extract is analyzed by HPLC to identify and

try to quantify the amount of carotenoids synthesized. The overall process can be observed in figure

2.1.

Figure 2. 1 General Overview of the process of upstream and downstream of Chromochloris sp. carried out in this thesis in order to analyze high valuable pigments.

Pre-culture:

Erlenmeyer flask

(600 mL)

Photobioreactor:

Bubble column

(1000 mL)

Harvesting:

centrifugation method

(10 min 3000 rpm)

Freeze Drying:

(27 hours -65 °C)

Storage at -20 °C Extraction Qualitative and

Qualitative Analysis:

HPLC (carotenoids) Fractionation

22

2.2.2 Biomass Production

2.2.2.1 Pre-Culture

The method used for biomass production was the up-scaling of a liquid pre-culture. It is essential

that the cell density is not too low to ensure that cells continue growing after lag phase, otherwise the

cell growth starts to decrease. In order to ensure the correct up-scaling process a sample was taken

from the pre-culture to measure the OD at 750 nm. Then, considering the OD measurement of the pre-

culture and the dilution factor of the photobioreactor it was estimated the amount of starting culture

needed to be added to the PBR so that the starting OD would be approximately 0.50. Proportionality

between biomass concentration and optical density was assumed and Equation 2.1 was used to the

calculation.

𝐶𝐼 × 𝑉𝐼 = 𝐶𝐹 × 𝑉𝐹

where CI and CV represent the biomass concentration in the starting culture and in the PBR,

respectively, while VI and VF represent the volume of culture of the pre culture and the final volume of

the PBR, respectively.

2.2.2.2 Biomass Production (Scale Up process)

Eight photobioreactors were used to cultivate the microalgae divided in two experimental setups.

Each PBR had a setup volume of 1000 mL and an actual working volume of 900 mL. The reactors were

filled with the media and then autoclaved at 121 °C for 20 minutes. The algae cultivation was added to

the PBR with the aid of a sterile syringe (20 mL) and the PBR’s were placed in a clima controlled room,

with a temperature of 21 °C. In both setups, the culture was stirred by air with a flow of 400 L/h (EHEIM

400 230 V~50Hz) and the air was previously filtered through a 0.2 μm filter.

The PBR’s were illuminated with 100-200 μmol.m-2.s-1 of light by Sanlight M30. In setup I,

microalgae were exposed to 24 h of light and in setup II to cycles of 16:8 (Light: Dark). The intensity of

light that the PBR’s were receiving was controlled by a light sensor (Apogee MQ-200).

2.2.2.3 First Cultivation Setup

The first cultivation setup comprises four PBR’s (PBR1 - 4) as it can be seen in figure 2.2. PBR

1 and 2 were cultivated with a starting culture of Chromochloris sp. (MCI-31) in co-culture with

uncharacterized bacteria while PBR 3 and 4 were cultivated with a starting culture of Chromochloris sp.

in unialgae state. The first assay took place between the months of March and April of 2018, and all

PBR’s were cultivated for 30 days. Regarding, the cultivation conditions of the four PBR’s, the type of

cultivation was phototrophic and they were illuminated with 100-200 μmol.m-2.s-1 in a continuous light

regime.

In the case of PBR 3, 5% (m/v) of glucose solution (90 mL) was supplemented to the PBR with

a sterile syringe (50 mL) when the stationary phase was attained (day 16 of cultivation).

(2.1)

23

Figure 2. 2 Picture illustrating the first set up of the cultivation (March to April) of Chromochloris sp. with the respective identification of each photobioreactor.

2.2.2.4 Second Cultivation Setup

The second cultivation setup also functioned with four photobioreactors: PBR 5, 6, 7 and 8 as it

can be seen in figure 2.3. In this case, all PBR’s had the same start culture (Chromochloris sp. unialgae).

This setup functioned between May and June of 2018 also for 30 days with exception to PBR 8 that was

kept running for 45 days. PBR 5, 6 and 7 had a mixotrophic type of cultivation since glucose (5% (m/v))

was added to the media before autoclaving, while PBR 8 had a phototrophic type of cultivation. In this

setup, the photobioreactors were illuminated with 100-200 μmol.m-2.s-1 in light cycles (16:8 (Light:

Dark)).

For PBR 5, after attaining stationary phase the algae were submitted to salt stress (NaCl 0.3 M)

by adding 100 mL of NaCl (3 M).

In the case of PBR 6, after reaching stationary phase the biomass was harvested and a new

media without NaNO3 (N deficiency) and without K2HPO4 and KH2PO4 (P deficiency) was added to the

photobioreactor.

PBR 1 PBR 2 PBR 3 PBR 4

24

Figure 2. 3 Picture illustrating the second set up of the cultivation (May to June) of Chromochloris sp. with the respective identification of each photobioreactor.

2.2.2.5 Summary

Table 2.6 summarizes the experiments performed with the photobioreactors and the cultivation

conditions used in each case.

Table 2. 5 Description of each photobioreactor of the cultivation and conditions of this process.

PBR PBR 1 PBR 2 PBR 3 PBR 4 PBR 5 PBR 6 PBR 7 PBR 8

Assay 1 1 1 1 2 2 2 2

Type of strain

MCI-31 co-culture with

bacteria

MCI-31 co-culture with

bacteria

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

Type of Cultivation

Phototrophic Phototrophic Phototrophic Phototrophic Mixotrophic Mixotrophic Mixotrophic Phototrophic

Media 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM

Glucose added

- - Day 16 - Day 0 Day 0 Day 0 -

Type of stress

- - - - Salt Stress

(0.3 M NaCl) N and PO4

-

depletion - -

Illumination cycle

24 hours 24 hours 24 hours 24 hours 16:8 hours 16:8 hours 16:8 hours 16:8 hours

2.2.2.6 Analysis

To ensure the viability of the cells and to obtain the cell growth profiles, the optical density (OD)

was measured at 750 nm using a spectrophotometer (JENWAY 7315 Spectrophotometer). Samples

with an OD>1 were diluted prior to measurement with sterile media to ensure the linearity between the

OD and the biomass concentration. The results of corrected OD measurements were used to plot the

growth curve and the values of OD in the exponential phase to determine the growth rate (μ) according

to the equation 2.2. The doubling time (tD) was also calculated according to equation 2.3.

PBR 5 PBR 6 PBR 7 PBR 8

25

𝜇 =ln(

𝑂𝐷𝑡2𝑂𝐷𝑡1

)

𝑡2 − 𝑡1

𝑡𝐷 =ln(2)

𝜇

Dry weight (DW) measurements were made at day 0, day of stress-induced and at the last day

of cultivation using a scale (Sartorius MA35). The pH was also measured following the same timeline

as the DW measurements.

2.2.2.7 Proof of purity

In both assays, the algae cultivations were controlled by microscope pictures (Nikon ECLIPSE

50i). In the case of MCI-31 co-culture it was analyzed the ratio of algae vs. bacteria, and if the number

of bacteria was higher than the algae number (indicating an overtake of the culture by bacteria), the

assay would stop, if not the strain was declared as MCI-31 in co-culture with bacteria and the assay

would continue.

On the other hand, in the case of MCI-31 unialgal, if no fungal or bacterial contamination was

detected microscopically, the strain was declared unialgal and the assay would continue.

2.2.3 Harvesting and freeze drying

2.2.3.1 Procedure

Algae from 7 photobioreactors was harvested completely (900 mL) after approximately 30 days

of cultivation, 1 of the PBR’s (PBR 8) was harvested after 45 days of cultivation. In the case of two

photobioreactors (PBR 3 and PBR 7) harvesting also occurred when they attained stationary phase

(early harvesting).

In the case of PBR 3 after the 30 days 800 mL of biomass was harvested but the remaining

biomass (100 mL) was cultivated again with fresh media and glucose, with the goal to obtain more

biomass. However, due to a problem which lead to an increase of the temperature of the room, the

algae were heavily contaminated with bacteria and foamed out of the PBR. The cultivation was stopped,

and the biomass harvested (late harvesting) with the idea of using it as positive control in the LPS assay.

To harvest, the content of the PBR’s was pumped (Carl Roth GmbH SN 1206 - 081) to a Schott

bottle (900 mL). Then it was transferred to plastic containers (200 mL) centrifuged for 10 min at 3000

rpm. The biomass was then washed with distilled water. The cycle of centrifugation/washing was

repeated two times and the biomass was transferred to 50 mL Falcon Tubes and kept in the fridge at -

20 °C until it was freeze dried.

The biomass was dried by a vacuum freeze-dryer (CHRIST ALPHA 1-4 LD plus) for 27 hours,

24 hours of main drying and 3 h of final drying. The lid of the freeze dryer was covered to prevent

pigment degradation by light. After freeze drying, the dried biomass was stored in the fridge at -20°C

until it was used.

(2.2)

(2.3)

26

2.2.3.2 Harvested Biomass Labeling Code

The code used to label the harvested biomass for further procedures is as follows in table 2.6:Table 2. 6 Identification of

the biomass from each PBR cultivated and harvested.

First 3 letters (Photobioreactor)

First digit (Number of photobioreactor)

Time of harvesting (stop and second digit)

PBR

1 .1

Early harvesting (before the 30 days of cultivation) 2

3 .2

Normal harvesting (at 30 days of cultivation) 4

5 .3

Later harvesting (after the 30 days of cultivation) 6

7 -

In the case of PBR that were only harvested once the stop and the

last digit is omitted. 8

Example: PBR 3 .1

First 3 letters: PBR Photobioreactor

First Digit: 3 Number of photobioreactor

Stop and second digit: .1 Time of harvesting (Early harvesting)

2.2.4 Lipid Content Analysis

To study the possibility of an interference of the lipid content in carotenoid extraction and the

difference of the lipid content in cultivation with and without glucose, samples of the first cultivation set

(PBR 1-4) were sent to FH Wels Upper Austria to be analyzed regarding the total lipid content and the

percentage of each fatty acid inside the samples. From each photobioreactor of this cultivation set were

sent two samples that correspond to two replicates. In the results the replicates from each

photobioreactor were treated as A and B.

2.2.5 LPS Assay

2.2.5.1 Procedure

To study the toxicity/safety of the samples in future applications an LPS assay was performed.

This LPS assay was performed twice with samples of extracts, liquid and biomass of the PBR’s

harvested. The protocol followed was based on the SOP “Chromogener LAL Test zur Bestimmung des

Endotoxingehalts in Extraken (LPS)” (Annex 6.2).

The treatment gave to the LPS assay samples is different in the case of liquid and dried samples.

In the case of liquid samples, 1 mL was collected from the photobioreactor for further processing while

for dried samples, biomass or extracts were dissolved in 50% (v/v) ethanol: sterile water in order to

obtain a concentration of 10 μg/mL of sample. Then, the samples were briefly vortexed and putted in

the ultrasonic bath for 30 minutes to aid the dissolution of the extract/biomass. After ultrasonic bath the

samples were taken to a Thermomixer (Eppendorf Thermomixer R) for 10 min at 300 rpm and 70°C.

After this step, the samples were allowed to cool until room temperature.

27

Depending on the condition of testing, the samples were filtered through a 0.22 μm sterile filter

(Carl Roth GmbH, Rotilabo®-PVDF sterile). To test different concentrations the samples were then

diluted into: 1.0; 0.2; 0.02; 0.002 and 0.0002 mg/mL with 1% (v/v) ethanol: sterile water.

In the case of the standard it was used the lyophilized endotoxin standard and 1 mL of 1% (v/v)

ethanol: sterile water that was then vortexed for 15 min.

To have the calibration curve the standard was diluted into 1.0; 0.5; 0.25; 0.1 and 0 mg/mL.

Then the Chromogenic substance was reconstituted with 6500 μL of sterile water along with the LAL

enzyme mix (1500 μL sterile water).

Both plates were then constructed following the layout represented in figure 2.4. The samples

applied in the wells in yellow in figure 2.4 were applied according to the dilution, that is to say, the higher

concentration (0.2 mg/mL) in line E and the lowest concentration in line B according to the sample.

Figure 2. 4 Layout of the 96 well plate used in both LPS assays. The wells in blue were filled with sterile water, the ones in yellow were used to samples, the green line was used to test the standards and the orange line was filled with the blank.

After the samples were placed in lines B to E, the standards in line F and the blank in line G the

plate was thermomixed for 15 minutes at 37°C. Then the LAL enzyme mix was added to the plate (B2

to F11) and the plate was shaken at 300 rpm (10 s at 37°C) and then incubate (10 min for 37°C. After

the chromogenic substance was added to the plate followed by a shaking again at 300 rpm (10 s at

37°C) and an incubation (4.5 min for 37°C), 25% (v/v) acetic acid solution was added to the plate to stop

the reaction with a final shake at 300 rpm (10 s at 37°C).

The plate was then analyzed by a spectrophotometer (Mithras LB 940, Berthold Technologies)

at 405 nm with the aid of the program MikroWin 2000.

Std 0.00

Std 0.00

Std 0.10

Std 0.10

Std 0.25

Std 0.25

Std 0.50

Std 0.50

Std 1.0

Std 1.0

28

2.2.5.2 LPS Assay Code

The LPS assay samples were labeled according to table 2.7:

Table 2. 7 Identification of the LPS assay samples.

First 3 letters (Photobioreactor)

First digit (Number of

photobioreactor)

Time of harvesting (stop and second digit)

First letter (Type of sample)

Second letter (Treatment of the

sample)

PBR

1 .1 Early harvesting L

Liquid Sample

F Filtered Sample

2

3 .2 Normal harvesting B Biomass

4

5 .3 Later harvesting

E Extract

U Unfiltered Sample

6

7

-

In the case of PBR that were only

harvested once the stop and the last digit are omitted.

8

Example: PBR 3 .1 E F

First 3 letters: PBR Photobioreactor

First Digit: 3 Number of photobioreactor

Stop and second digit: .1 Time of harvesting (Early harvesting)

Second letter: E Type of sample (Extract)

Third letter: F Treatment of the sample (Filtered)

2.2.5.3 List of LPS assay samples

The samples tested in both LPS assays are listed below:

1st LPS assay:

o PBR 1 (extract unfiltered)

o PBR 2 (extract unfiltered))

o PBR 3.1 (extract unfiltered)

o PBR 3.2 (extract unfiltered)

o PBR 3.3 (extract and dried biomass unfiltered)

o PBR 4 (extract and dried biomass unfiltered)

2nd LPS assay:

o PBR 1 (biomass unfiltered)

o PBR 3.3 (extract filtered, dried biomass filtered and unfiltered)

o PBR 4 (extract and biomass filtered)

o PBR 5 (liquid filtered and unfiltered)

o PBR 6 (liquid filtered)

29

2.2.6 Extraction

2.2.6.1 Standard Procedure

The extraction method used was based on a Standard Operating Procedure (SOP) “Microalgal

Pigment and lipids extraction for bioassay and for high performance liquid chromatography” (Annex 6.3).

The procedure for standard extraction of pigments is as follows, approximately 25 mg of

lyophilized algae was weighed with 1 g of glass beads 0.75 – 1.0 mm into a 5 mL Eppendorf. The glass

beads should be washed previously and the solvent mixture, Dichloromethane: Methanol made in a

ratio of 1:1. Then, 4 mL of Dichloromethane: Methanol mixture was added to the 5 mL Eppendorf

containing the biomass and the beads. After, the suspension was vortexed for 3 minutes and

subsequently centrifuged at the maximum power for 15 min. After centrifugation, the supernatant was

placed in a glass tube previously weighed and to the pellet was added 2 mL of fresh solvent and the

vortex and centrifugation cycles repeated. After, the process is repeated one more time.

If the solvent mixture includes organic solvents, the extract is left open in the laminar flow to

evaporate. If this is not the case, the extract is placed in an evaporator (Eppendorf Concentrator 5301)

for 2 hours at 45°C and then the temperature is raised to 60 °C until the solvent is evaporated.

To optimize the extraction procedure, one approach was performed: Change the size of the

glass beads to improve the cell disruption.

2.2.6.2 Experiment with different glass beads size

The extraction procedure remained the same with the exception of the glass beads size. It was

study four types of cell disruption with glass beads: Big (4.0 ± 0.3 mm), Medium (0.75-1.0 mm), small

(0.25-0.50 mm) and a mix with small and big glass beads. Biomass from PBR 3.2 was used to perform

the experiment.

2.2.6.3 Extraction Results Analysis

To compare the results of the extractions performed the extraction yield was calculated

according to equation 2.4.

𝐸𝑥𝑡𝑟𝑎𝑐𝑡𝑖𝑜𝑛𝑦𝑖𝑒𝑙𝑑(%) =𝑚𝑎𝑠𝑠𝑜𝑓𝑒𝑥𝑡𝑟𝑎𝑐𝑡(𝑔)

𝑚𝑎𝑠𝑠𝑜𝑓𝑏𝑖𝑜𝑚𝑎𝑠𝑠(𝑔)× 100

2.2.7 Fractionation

2.2.7.1 Procedure

Fractionation is a technique with the goal of purifying the pigments, to do so, the process of

fractionation obtains different fractions that should contain different pigments. To obtain different

pigments in different fractions it is necessary to look to the principle of polarity. Different pigments have

different polarities what makes the solubility in solvents/solvents mixtures also different. Researching

the targeted pigments in this project (astaxanthin and canthaxanthin), showed that both pigments have

different polarities and solubilities in different solvents, which means that they can be fractioned, in

theory.

(2.4)

30

For the fractionation, approximately 70-90 mg of freeze-dried biomass and 0.5 g of glass beads

0.75-1.0 mm were weighed and added to a 5 mL Eppendorf with 4 mL of the first solvent/solvent mixture

studied. Then, the sample was vortexed for 3 minutes and centrifuged for 15 min at 9000 rpm. After

centrifugation, the supernatant was collected to a glass tube previously weighed and the pellet was left

open for 10 min, in order to evaporate the remaining solvent.

The process was repeated, according to the number of solvents/solvent mixtures studied and

the supernatant was always collected to a different glass tube to have the different fractions of the

process. All the fractions are left open in the laminar flow to evaporate the solvent and allow the extract

to dry. In order to preserve the pigments, the glass tubes are covered with aluminum foil and placed in

the dark.

The methods and solvents used in fractionation are described in Table 2.8.

Table 2. 8 Methods used for fractionation and sequence of the solvents/solvent mixtures used in each case.

Fractionation Method Sequence of Solvents/Solvent Mixtures

A: Standard Method Ethanol: Water (8:2) > Ethanol: Water (8:2) > Dichloromethane:

Methanol (1:1) > Dichloromethane: Methanol (1:1) > Ethanol: Water (8:2) > Dichloromethane: Methanol (1:1)

B: Apolar Scheme Ethanol > Acetonitrile > Acetone > Dichloromethane > Chloroform

C: Polar Scheme Hexane > Ethyl Acetate > Ethanol > Methanol > Water

D: Astaxanthin / Canthaxanthin

Detection

Acetonitrile: Dichloromethane (2:1) > Isopropanol: Dichloromethane (2:1) > Ethyl Acetate: Acetone (8:2) > Ethyl Acetate: Dichloromethane (2:1) >

Methanol: Hexane (1:1)

E: Canthaxanthin / Astaxanthin Detection

Ethyl Acetate: Acetone (8:2) > Ethyl Acetate: Dichloromethane (2:1) > Acetonitrile: Dichloromethane (2:1) > Isopropanol: Dichloromethane (2:1)

> Methanol: Hexane (1:1)

F: Polarity Scale Water: Ethanol (1:1) > Ethanol: Isopropanol (1:1) > Isopropanol:

Acetonitrile (1:1) > Acetonitrile: Dichloromethane (1:1) > Dichloromethane: Ethyl Acetate (1:1) > Ethyl Acetate: Hexane (1:1)

2.2.7.2 Fractionation’s sample code

Each extract from the fractionation was labeled according table 2.9.

Table 2. 9 Identification of the extracts obtain in each fractionation method.

First letter (Fractionation)

Second letter (Type of fractionation)

First digit (Number of

fraction) Suffix (Replicate)

F

A Standard Method 1

A First

Replicate B Apolar Scheme 2

C Polar Scheme 3

D Astaxanthin/Canthaxanthin 4

B Second

Replicate E Canthaxanthin/Astaxanthin 5

F Polarity Window 6

31

Example: F A 1 B

First letter: F Fractionation

Second letter: A Type of fractionation (Standard Method)

First Digit: 1 Number of fraction (Fraction number 1)

Second letter: B Replicate (Second replicate)

2.2.8 Quantitative and Qualitative Analysis of high valuable pigments

2.2.8.1 Procedure

For the quantitative and qualitative analysis of the pigments a RP-HPLC method was chosen.

In this type of chromatography, the analytes are separated based on their hydrophobicity, where the

stationary phase is less polar than the mobile phase. The analytes that are more hydrophobic create

stronger bonds with the stationary phase and have longer retention times. The HPLC system (Agilent

1100 series) is constituted by different modules: the pump, the auto sampler, the column compartment

and the diode array detector. While the pump pushes the solvents through the column, the auto sampler

injects the analytes to be analyzed thorough the column after being eluted from the column according

to their hydrophobicity, they are then detected by a UV detector and the chromatogram is plotted.

The analyses were performed injecting 5 μL of the sample in a C18 column. They were eluted

with a flow of 1mL/min, following a specific gradient elution profile according to the method used. The

data was analyzed with the software ChemStation for LC 3D systems.

Both the standards and the extracts were diluted in DMSO and then filtered through a 0.2 µm

filter (sartorius Minisart® Syringe Filter) and added to the HPLC vials. While for the standards the volume

of DMSO added was calculated according to the final concentration of study, for the extracts a

concentration of 10 mg of extract/mL was targeted and the volume of DMSO calculated with the g of

extract obtained. The methods and columns used in this procedure are described in table 4.

Table 2. 10 Columns used and methods tried in each case.

Column Method

Phenomenex C18 3 μm (100 x 2 mm)

General Method

Solvent A: Acetone: Water (1:1)

Solvent B: Acetone

Flow: 1 mL/min

Temperature: 60°C

Injection Volume: 5 μL

Detection wavelength: 450 nm

Time of the running: 30 min

Method description:

0 min 100% A

0-60 min 100-0% A

C18 5 μm (250 x 4 mm)

+

Guard Column LiChrodpher®

100RP-18 (5μm)

General Method

(GM)

Solvent A: Acetonitrile : Methanol

(74:6)

Solvent B: Methanol : Hexane (5:1)

Flow: 1 mL/min

Temperature: 30°C

Injection Volume: 5 μL

Detection wavelength: 440 nm

Time of the running : 27 min

Method description:

4 min at 100% A

4-9 min 0-100% B

9-25 min at 100% B

25-27 min at 100-0% B

32

Astaxanthin Detection

(AD)

Solvent B: Methanol : Hexane (5:1)

Flow: 1 mL/min

Temperature: 30°C

Injection Volume: 5 μL

Detection wavelength: 440 nm

Time of the running : 25 min

Method description:

15 min at 100% B

15-20 min 100-0% B

20-25 min at 100% B

Canthaxanthin Detection

(CD)

Solvent B: Methanol : Hexane (5:1)

Flow: 1 mL/min

Temperature: 30°C

Injection Volume: 5 μL

Detection wavelength: 440 nm

Time of the running : 15 min

Method description:

5 min at 100% B

5-10 min 100-0% B

10-15 min at 100% B

2.2.8.2 HPLC Results Analysis

The limit of detection (LOD) and limit of quantification (LOQ) were calculated with software Data

Analysis from MS Excel and based on the standard deviation of y-interception of the regression line

(SD) and the slope (S), using the equation 2.5 and 2.6, respectively.

𝐿𝑂𝐷 = 3.3 ×𝑆𝑡𝑎𝑛𝑑𝑎𝑟𝑑𝐷𝑒𝑣𝑖𝑎𝑡𝑖𝑜𝑛

𝑆𝑙𝑜𝑝𝑒

𝐿𝑂𝑄 = 10 ×𝑆𝑡𝑎𝑛𝑑𝑎𝑟𝑑𝐷𝑒𝑣𝑖𝑎𝑡𝑖𝑜𝑛

𝑆𝑙𝑜𝑝𝑒

To compare the results of HPLC measurements in terms of quantification of astaxanthin and

canthaxanthin between different samples a relation was made obtaining the results in mg of carotenoid

per g of biomass (DW) tested, equation 2.7.

[𝑝𝑖𝑔𝑚𝑒𝑛𝑡](𝜇𝑔/𝑚𝐿) × 𝑣𝑜𝑙𝑢𝑚𝑒𝑜𝑓𝐷𝑀𝑆𝑂𝑎𝑑𝑑𝑒𝑑(𝑚𝐿) = 𝑚𝑎𝑠𝑠𝑜𝑓𝑝𝑖𝑔𝑚𝑒𝑛𝑡(𝜇𝑔)

𝑚𝑎𝑠𝑠𝑜𝑓𝑝𝑖𝑔𝑚𝑒𝑛𝑡(𝜇𝑔)

𝑚𝑎𝑠𝑠𝑜𝑓𝑒𝑥𝑡𝑟𝑎𝑐𝑡(𝑔)×

𝑚𝑎𝑠𝑠𝑜𝑓𝑒𝑥𝑡𝑟𝑎𝑐𝑡(𝑔)

𝑚𝑎𝑠𝑠𝑜𝑓𝑏𝑖𝑜𝑚𝑎𝑠𝑠(𝑔)=𝜇𝑔𝑜𝑓𝑝𝑖𝑔𝑚𝑒𝑛𝑡

𝑔𝑜𝑓𝑏𝑖𝑜𝑚𝑎𝑠𝑠

(2.5)

(2.6)

(2.7)

33

3. Results and Discussion

3.1 Cultivation

The aim of this experiment was to cultivate Chromochloris sp. (MCI-31) with different conditions

and therefore observe the difference of carotenoid content in each condition. Since it has been

documented that inducing stress in microalgae cultivations enhances the synthesis of carotenoids, as

astaxanthin and canthaxanthin, it was decided to study the following conditions: mixotrophy by

supplementation of glucose, two-stage cultivation, nitrogen and phosphorus deficiency, high salinity and

different light regimes.

Another aim was to study the influence of a co-culture in carotenoid synthesis. Even though in

general, lack of purity is problematic with bacteria overtaking microalgae cultures, some reports

identified symbiotic relations between bacteria and microalgae that during long term cultivation achieve

a state of equilibrium and ultimately live in co-culture that enhances microalgae growth and lipid

accumulation. However, it is still unknown if a co-culture prejudices or enhances the synthesis of

pigments.

Furthermore, the first set of cultivation was performed to test the influence of the purity in the

carotenoid content, therefore, two PBR were cultivated with a pre-culture of a co-culture of microalgae

and uncharacterized bacteria while the other two PBR were cultivated with a starting culture of

Chromochloris sp. (unialgae). In this first experiment it was also tested a two stage strategy, where in

the first stage algae follow a photoautotrophic cultivation and in the second stage, glucose is added and

the PBR has a mixotrophic cultivation.

The second set of cultivation was performed to test the influence of nitrogen/phosphorus

deficiency, high salinity and mixotrophy in the biosynthesis of astaxanthin/canthaxanthin. Moreover, a

comparison between the two sets of cultivation studies the influence of light regimes in algae growth

and carotenoid content.

3.1.1 Study of the influence of bacterial co-culture in cell growth of Chromochloris sp. and carotenoid content

To study the influence of purity in carotenoid content PBR 1 and 2 were cultivated with a

bacterial co-culture of Chromochloris sp. while PBR 3 and 4 were cultivated with Chromochloris sp.

unialgae. Figure 3.1 shows the evolution of the PBR’s from the first setup of cultivation along the

cultivation period.

34

Figure 3. 1 Pictures illustrating the evolution of the first set of cultivation with four PBR’s. A: day 7 of cultivation for PBR 1, 2 and 3 and day 0 for PBR 4. B: day 19 of cultivation for PBR 1, 2 and 3 and day 11 for PBR 4. C: Harvested biomass at day 30 of cultivation for PBR 1, 2 and 3.

3.1.1.1 Proof of purity

To ensure the validity of the study, samples were taken regularly from the PBR’s and visualized

under the microscope. In the case of PBR’s cultivated with a bacterial co-culture of Chromochloris sp.

(PBR 1 and 2), the samples were regularly visualized under the microscope to ensure that the bacteria

didn’t take over the culture. In the case of PBR’s inoculated with a pre-culture of Chromochloris sp.

unialgae, microscope pictures were taken to ensure the purity of the culture. Figure 3.2 shows the

evolution of the microalgae in PBR 1 and 4 during the cultivation period.

A B

C

PBR 2 PBR 1 PBR 3 PBR 4 PBR 2 PBR 1 PBR 3 PBR 4

PBR 3 PBR 2 PBR 1

35

Figure 3. 2 Microscope pictures of the first cultivation setup, PBR 2 and 4. In PBR 2 Chromochloris sp. appears in co-culture with bacteria at cultivation day 6 (A) and 21 (B). In both pictures, the black arrow indicates the bacteria contamination. In PBR 4 showing the purity ensured with no apparent contamination visible at day 0 of cultivation (C) and at day 13 (D). (Magnification: 1000x)

Once ensured the purity of PBR’s 3 to 8 and the co-culture in PBR 1 and 2 the results of the

growth curves were plotted and analyzed.

3.1.1.2 Growth Curve and Rate Analysis

To study the influence of bacterial co-culture in cell growth, OD measurements were performed

every 2 days while the cultivation was running. The growth curves of all four PBR’s from the first

cultivation set can be observed in Figure 3.3.

Furthermore, the growth rate and the duplication time were calculated for the exponential phase

of all four PBR’s in the first set of cultivation, in order to compare the growth of microalgae in the

presence of bacteria and how that affects the cell growth.

A B

C D

36

Figure 3. 3 Graphic representation of growth curves for PBR 1 bacterial co-culture (), PBR 2 bacterial co-culture (), PBR 3 two-stage cultivation () and PBR 4 control () based on OD measurements taken during the 30 days of cultivation.

Table 3. 1 Exponential growth rates (µ) and duplication times (tD) of the first cultivation set.

Photobioreactor PBR 1 PBR 2 PBR 3 PBR 4

Growth Rate (day-1) 0.171 0.167 0.213 0.195

Duplication Time (day) 4.1 4.2 3.3 3.6

3.1.2 Study of Stress Conditions: N and PO4- Deficiency and High Salinity in cell

growth

In this experiment, 4 PBR´s were cultivated with a pre-culture of Chromochloris sp. unialgae.

While PBR 5, 6 and 7 were started with glucose 5 % (m/v) in the media following a mixotrophic

cultivation, PBR 8 was started with 3N BBM following a phototrophic cultivation. After attaining stationary

phase, nitrogen/phosphorus deficiency stress was induced in PBR 5 and high salinity stress in PBR 6.

The growth curves of all 4 PBR’s are shown in Figure 3.4 and the associated growth curves and

duplication times in table 3.2.

Figure 3. 4 Graphic representation of growth curves for PBR 5 high salinity stress (), PBR 6 N and P deficiency (), PBR 7 mixotrophic cultivation () and PBR 8 control () based on OD measurements taken during the 30/45 days of cultivation.

0

2

4

6

8

10

12

14

16

18

0 5 10 15 20 25 30 35

OD

75

0 n

m

Time (days)

PBR 1 PBR 2 PBR 3 PBR 4

0

1

2

3

4

5

6

7

8

9

0 5 10 15 20 25 30 35 40 45 50

OD

75

0 n

m

Time (days)

PBR 5 PBR 6 PBR 7 PBR 8

37

Table 3. 2 Exponential growth rates (µ) and duplication times (tD) of the second cultivation set.

PBR PBR 5 PBR 6 PBR 7 PBR 8

Growth Rate (day-1) 0.176 0.182 0.175 0.071

Duplication Time (day) 3.9 3.8 4.0 9.8

3.1.3 Comparison mixotrophic with phototrophic conditions in cell growth

For studying both mixotrophic and phototrophic conditions of cultivation it was established a

comparison between PBR 3, PBR 7 and PBR 8. PBR 7 was supplemented with glucose from day 0 of

cultivation while for PBR 8 the carbon source was exclusively CO2. In the case of PBR 3, before attaining

stationary phase this PBR followed a photoautotrophic cultivation and after the glucose supplementation

a mixotrophic cultivation, with cells retrieving C from CO2 and glucose.

Furthermore, PBR 7 can be identified as a purely mixotrophic type of cultivation, PBR 8 as a

purely phototrophic cultivation while PBR 3 has a phototrophic type of cultivation before glucose

supplementation and a mixotrophic cultivation after the addition of this nutrient.

Figure 3.5 plots the growth curves of PBR 3, 7 and 8 to establish a comparison between a

phototrophic and a mixotrophic type of cultivation. Besides OD and DW measurements, microscope

pictures (Figure 3.6) followed the cell morphology and color.

Figure 3. 5 Graphic representation of growth curves for PBR 3 two stage cultivation (), PBR 7 mixotrophic cultivation () and PBR 8 photoautotrophic cultivation (). The arrow targets the time point when glucose was added to PBR 3 turning the phototrophic cultivation in mixotrophic cultivation.

0

2

4

6

8

10

12

14

16

18

0 5 10 15 20 25 30 35 40 45 50

OD

75

0 n

m

Time (days)

PBR 7, mixotrophic PBR 3, phototrophic and mixotrophic PBR 8, phototrophic

38

Figure 3. 6 Microscope pictures of PBR 3: (A) at day 6 of cultivation (before adding glucose), (B) at day 21 of cultivation (after the addition of the C source). (Magnification: 1000x)

Microscope pictures of PBR 3 before and after glucose supplementation show differences in

color, with cells changing from green to orange and in size with cells in mixotrophic conditions becoming

enlarged. Differences in cell shape were not observed during cultivation period. The color change was

also observed in PBR 7 that had a purely mixotrophic state from the start of the culture, with cells

changing color 2 days after the exposure to the media with glucose. The difference in cell color was also

observed macroscopically, with the culture color changing from green to orange after exposure to

glucose. Previous studies reported similar results concluding that the addition of glucose to the media

turns cells bigger in size and orange.[49]

The reason behind these changes is still unknown. One possible explanation could be that it

happens due to an increase of carotenoid accumulation. It can also be explained by the fact that, a

decrement in the amount of photosynthetic activity causes a decrease on the amount of chlorophyll

synthetized which, gives the green color to the cultivation that causes a enhancing of the green color of

the pigments.

3.1.4 Study of the impact of photoperiod regime in cell growth

To study the influence of photoperiod regime in cell growth and in the accumulation of

carotenoids, the first set of cultivation was exposed to 24 of light while the second set was exposed to

cycles of 16:8 h of L:D. Since other conditions were studied, it is possible to directly compare in terms

of the impact of light, only PBR 4 to PBR 8. Both bioreactors were started with Chromochloris sp.

unialgae and both follow a photoautotrophic type of cultivation during the experiment. The growth curves

and the calculated growth rates can be observed in figure 3.7.

A B

39

Figure 3. 7 Graphic representation of growth curves of PBR 4 – control, continuous light () and 8 – control, 16:8 (L:D) () based on OD measurements taken during the 30/45 days of cultivation.

3.1.5 Cultivation Summary and Discussion

Table 3.3 summarizes the cultivation studies performed and compiles other information not

present in figures and tables shown above.

Table 3. 3 Photobioreactors cultivated, type of cultivation followed, strain and media tested, time of glucose addition, type of stress induced and pH at the time, as well as, end measurements (OD, DW and pH).

PBR PBR 1 PBR 2 PBR 3 PBR 4 PBR 5 PBR 6 PBR 7 PBR 8

Assay 1 1 1 1 2 2 2 2

Type of strain

MCI-31 co-culture with

bacteria

MCI-31 co-culture with

bacteria

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

MCI-31 unialgae

Type of Cultivation

Phototrophic Phototrophic Phototrophic Phototrophic Mixotrophic Mixotrophic Mixotrophic Phototrophic

Media 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM 3-N BBM

Glucose addition

- - Day 16 - Day 0 Day 0 Day 0 -

Type of stress

- - - - Salt Stress

(0.3 M NaCl) N and PO4

-

depletion - -

Photoperiod Regime

24:0 hours 24:0 hours 24:0 hours 24:0 hours 16:8 hours 16:8 hours 16:8 hours 16:8 hours

Growth Rate (day-1)

0.171 0.167 0.213 0.195 0.176 0.182 0.175 0.071

Duplication Time (day)

4.1 4.2 3.3 3.6 3.9 3.8 4.0 9.8

Stress pH - - - - 4.89 4.95 - -

End OD 750 nm

11.18 9.695 17.04 13.62 1.713 2.830 6.585 8.090

End pH 9.082 9.284 8.685 9.251 4.976 6.012 5.876 9.129

End DW (g/L)

5.2 4.5 19.36 - 2.35 4.3 6.8 3.1

0

2

4

6

8

10

12

14

16

0 5 10 15 20 25 30 35 40 45 50

OD

75

0 n

m

Time (days)

PBR 4, 24 h Light PBR 8, 16:8h (L:D)

40

Figure 3.3 shows the growth curves of the first set of cultivation with all the PBR’s obtaining

similar growth patterns. While PBR 1 and 2 show the presence of lag phase with both algae and bacteria

adapting to the media, in PBR 3 and 4 there is the absence of the lag phase and microalgae start to

grow almost immediately. Regarding the exponential phase, results show a pattern of the growth rates

with PBR 1 and 2 obtaining similar rates as well as PBR 3 with PBR 4, with the cultivation with

Chromochloris sp. unialgae (PBR 3 and 4) obtaining higher growth rates than the co-culture strategy

cultivation.

In addition, end OD measurements show a higher cell density in PBR 4 than PBR 1 and 2.

Bacterial co-culture lower growth rates and low end OD measurements indicate that in photoautotrophic

cultivation a strategy of enhancing microalgae growth with a bacterial co-culture is not beneficial.

Relations of mutual symbiosis between microalgae and bacteria have been reported with

bacteria supplementing vitamins and preventing other bacteria to invade the culture, while algae induce

the synthesis of photosynthetic metabolites and lipids. One particular study demonstrated an increasing

effect in pigment and lipid accumulation with a bacterial co-culture of Chlorella spp. with bacteria

Azospirillum brasilense. [67] Further studies should be conducted investigating the impact of co-culture

in mixotrophic and heterotrophic cultivations of Chromochloris sp., as well as, the characterization of the

bacteria used to prove and fully understand the impact of bacteria in culture and the possibility of benefic

microalgae-bacteria interactions.

Second set of cultivation (Figure 3.4) shows a more uneven pattern of growth curves. While

PBR 5, 6 and 7 have a similar shape of growth curves with no evidence of lag phase and a similar

exponential phase only differing when stress is induced (day 24 of cultivation), PBR 8 has a long lag

phase, with the exponential phase only starting on day 10 of cultivation.

PBR 5 and 6 growth curves show a decaying effect in OD measurements when stress is

induced. This indicates that both stress conditions have a similar effect in the culture. While for N and P

deficiency condition (PBR 6), the cell decaying is moderate, high salinity induces a drastic decaying

effect in cell density. The results also indicated that after inducing the stress a slight increase of cell

density is observed, but it is not possible to conclude if cells overcame the negative impact of the salt

during the cultivation time.

These results are consistent with a previous study that showed evidence of a reduction effect

in cell density when high salinity stress is induced. In this study with Chlorella zofingiensis salt stress

was induced with NaCl concentrations of 0.2 and 0.4 M and results show a decrease in cell density of

30 and 50%, respectively. [57]

In PBR 6 the shift from an N-rich medium to an N-depleted medium causes a decaying in

biomass at the stress inducing moment followed by a recovery until the end of cultivation. Previous

studies reported that nitrogen depletion causes the cells to look for other sources of this nutrient with

chlorophyll being the primary candidate as it is a nitrogen-rich compound and easily accessible. The

loss of chlorophyll reduces the efficiency of energy collection affecting photosynthesis and promotes the

41

accumulation of other pigments. [43] Therefore, a decrease in cell growth is observed while algae are

adapting to the new media followed by the recovery observed in Figure 3.4.

The decaying effect on cell growth observed in PBR 5 and 6 can also be explained by the pH

of the media. When inducing the stress, the pH of PBR 5 and 6 decreases to 4.89 and 4.94, respectively.

A pH lower than 5.5 was reported to slow cell growth since algae are not tolerant to such acidic

environments. [49] The end pH values suggest that the algae are slowly recovering which, is reflected

in the growth curve by an increment in OD measurements with the increasing of the pH.

Figure 3.5 shows the growth curves of PBR 7 to PBR 8, these two PBR have different cultivation

types, with PBR 7 following a mixotrophic cultivation while PBR 8 follows a photoautotrophic cultivation.

A clear distinction can be observed in the growth curve profiles, as pointed before; PBR 8 shows a long

lag phase with 8 days of duration while PBR 7 proceeded almost immediately to exponential phase with

no lag phase observed. Moreover, PBR 7 attains stationary phase more rapidly than PBR 8. These

results are concordant with the calculated growth rates that show a faster growth for PBR 7 (0.175 day-

1), and a huge discrepancy to PBR 8 with a growth rate of 0.071 day-1.

Therefore, results of growth rates and the growth curves profile suggest that glucose stimulates

the biomass production with algae attaining faster stationary phase but does not have an increasing

effect on biomass concentration with results of OD showing similar measurements for phototrophic and

mixotrophic cultivations.

However, cell density results obtained by DW measurements reveal a different pattern of

biomass concentration with PBR 7 obtaining 6.8 g/L while PBR 8 only 3.1 g/L, which represents an

increment of 2-fold with mixotrophic cultivation.

From another point of view, an interesting pattern is seen when observing pH results of the end

of cultivation. While PBR 8 that followed photoautotrophic cultivations shows a pH value of 9.129

mixotrophic PBR 7 has a pH value of 5.876. This result suggests that glucose supplementation lowers

the pH of the cultivation. A previous study investigated the impact of pH in Chlorella zofingiensis growth,

with results showing that this alga is capable of tolerating pH range of 5.5-8.5 and as a pH optimum for

astaxanthin yield of 5.5. This suggests than in these conditions a pH value of 9 slows cell growth, with

algae not capable of tolerating a high basic media.

Consulting the literature for microalgae growth in these conditions, it is documented that the

addition of glucose to the media enhances its growth not only enabling the biomass to grow faster but

also to attain higher cell densities. [48,86] Replicates of both cultivations are needed to fully understand

and evaluate the effect of glucose in Chromochloris sp. growth.

A singular result was obtained when comparing PBR 7 with PBR 3. The two stage cultivation

PBR started with a photoautotrophic cultivation and was then supplemented with glucose turning it into

a mixotrophic cultivation. Figure 3.5 shows that at the starting point (day 0) of PBR 3 and 7 OD is similar,

however unexpectedly, right from day one of cultivation PBR 3 still in phototrophic state has a higher

growth than PBR 7 that was supplement with glucose at day 0 and PBR 8 that also followed a

42

photoautotrophic cultivation. This growth is then enhanced by the glucose supplementation of PBR 3 at

day 16 of cultivation and culminates in end OD measurements two times higher than PBR 7 or 8.

These results are also observed in DW measurements with PBR 3 exhibiting a cell density of

19.4 g/L which translates in an increase of 2.85 times the biomass yield of PBR 7. The growth rate

results show that PBR 3 grows faster, with a rate of 0.213 day-1 higher than PBR 7 and PBR 8. The

results indicate that a two-stage cultivation enhances biomass yield and the growth rate and that should

be further investigate to optimize it.

Available literature shows that these results were previously reported in a study with

Haematococcus pluvialis. This study concluded that a two-stage cultivation, followed by PBR 3, enables

a much higher cell number to be obtained when compared to a pure mixotrophic cultivation. Moreover,

has the advantage of minimizing the contamination risk owing to the shorter exposure time of the cells

to any organic carbon source.[87]

Considering that the only difference between PBR 3 and 7 besides type of cultivation is the light

regime, results also suggest a high impact of light regime in Chromochloris sp. growth. In fact, all the

PBR’s of the first cultivation set exhibit higher cell densities than PBR’s from second cultivation set.

Light regime impact in algal growth was studied comparing PBR 4 with PBR 8; both PBR’s

followed a photoautotrophic cultivation and were set under different light cycles. Figure 3.7 suggests

that a high photoperiod regime (24 h) enhances algal growth with no photo-inhibition effect observed.

These results are supported by the calculated growth rates where PBR 4 obtained a value of 0.195 day-

1 while PBR 8 obtained 0.071 day-1. While some studies defend the use of continuous light in cultivation

since they achieve the maximal growth rate, others suggest the use of light/dark cycles instead. L:D

cycles mimics the cells natural environment where algae are not exposed to light 24h per day and hints

the importance of a dark phase necessary for cell division and specific metabolite synthesis. [88]

Moreover, most studies reported that the use of light/dark cycles were benefic for algal growth

and pigment accumulation in Chlorella zofingiensis emphasizing on the importance of the dark phase

and in the fact that cell growth is also affected by the amount of energy offered per cycle, and not only

by the duration of the photoperiod.[33,54,88]

At this light intensity, results suggest that Chromochloris sp. growth is highly dependent on the

photoperiod regime, but replicates are needed to validate this theory as well as higher light intensities

to understand the light saturation limit of Chromochloris sp. and its impact in algal growth and pigment

content. Furthermore, future studies should be performed with mixotrophic cultivations, other stress

factors and different light regimes to fully comprehend the effect of light in Chromochloris sp. growth.

3.2 Lipid Content in Chromochloris sp. (1st Cultivation Set)

Knowing that Chromochloris sp. is a strain with high lipid content, which could interfere in the

extraction and analysis of carotenoids it was performed a total lipid content analysis to the biomass

harvested in the first set of cultivation. The samples included one sample from each PBR with exception

to PBR 3. In this case, three samples were taken: early, normal and late harvest. The results are shown

in Figure 3.8.

43

Figure 3. 8 Total lipid content in percentage of Dry Weight obtained for biomass harvested of Chromochloris sp. in the first cultivation setup. The data obtained is the result of a mean between the results of two samples and the deviation is represented be the error bars.

Results showed that total lipid content in PBR 1 to 4 varies in a range of 28 to 70%. The highest

lipid content was found in the photobioreactor supplemented with glucose, PBR 3.1, .2 and .3 and the

lowest lipid content in PBR 4, the control condition. In addition, mixotrophic cultivation (PBR 3) obtained

a higher lipid content than the photoautotrophic cultivation (PBR 1, 2 and 4). In fact, the supplementation

of glucose to the media increased the lipid accumulation, with total lipid content increased by 2-fold over

the control condition.

The addition of a carbon source to the medium has been widely reported to increase the lipid

content in microalgae, with glucose being the most commonly used source of these metabolites. It can

be argued that in a mixotrophic cultivation, glucose provides additional energy and material for

biosynthesis, with intermediate metabolites such as acetyl-CoA and NADPH playing important roles in

lipid metabolism. [48]

Furthermore, focusing on PBR 3 and in the role of glucose in lipid accumulation, PBR 3.1 has

the highest lipid content while PBR 3.2 the lowest, this could indicate an accumulation of lipids in an

early stage of stationary phase (PBR 3.1) and a minor decrease on lipid content towards the end of

stationary phase (PBR 3.2). In a similar study, results showed early-stationary phase as the ideal harvest

time for microalgae Chlorella sorokiniana GXNN01, a potential strain for lipid production. [89]

While analyzing results of PBR 3.3, the combination of glucose re-supplementation and bacterial

contamination had an increasing effect in lipid accumulation. In fact, bacterial co-culture has been

reported as a factor that enhances lipid accumulation in microalgae.[90] Likewise, PBR 1 and 2

Unialgae + + + - - +

Glucose + + + - - -

0.0

10.0

20.0

30.0

40.0

50.0

60.0

70.0

80.0

90.0

100.0

PBR 3.1 PBR 3.2 PBR 3.3 PBR 1 PBR 2 PBR 4

Tota

l Lip

id c

onte

nt

[% o

f D

W]

Total Fatty Acids in 1st setup of cultivation

44

cultivated with a co-culture of microalgae and bacteria also demonstrated the same enhancing effect

when compared with PBR 4 (control).

Furthermore, to understand the impact of cultivation conditions in lipid composition, the fatty

acid profile should be studied. The results of the major fatty acids found in the profile as well as the

amount of saturated and unsaturated fatty acids found in the biomass tested can be observed in table

3.4.

Table 3. 4 Fatty acid profile of PBR 1 – 4 expressed as major fatty acids, quantity of saturated FA and unsaturated fatty acids present in the samples. The data is expressed as a mean of two independent measurements in percentage of DW.

PBR 3.1 PBR 3.2 PBR 3.3 PBR1 PBR2 PBR4

Co-culture - - - + + -

Glucose + + + - - -

18:0 (%) 3.6 ± 0.1 3.1 ± 0.0 3.6 ± 0.0 1.8 ± 0.0 1.8 ± 0.05 1.3 ± 0.2

18:1 (%) 34.3 ± 0.3 34.9 ± 0.2 41.9 ± 1.3 29.7 ± 0.8 27.1 ± 0.1 14.1 ± 0.4

18:2 (%) 14.8 ± 0.1 13.7 ± 0.1 12.7 ± 0.4 12.9 ± 0.3 10.7 ± 0.3 5.4 ± 0.1

18:3 (%) 4.5 ± 0.5 3.9 ± 0.1 3.7 ± 0.1 3.5 ± 0.1 3.2 ± 0.02 2.4 ± 0.0

Saturated fatty acids (%)

4.4 ± 0.4 3.7 ± 0.1 4.2 ± 0.0 2.2 ± 0.0 2.2± 0.03 1.9 ± 0.3

Unsaturated fatty acids (%)

53.7 ± 0.0 52.7 ± 0.3 58.3 ± 1.8 46.3 ± 1.2 41.1 ± 0.4 21.9 ± 0.6

The fatty acid profile showed a predominance of C18 fatty acids over fatty acids of longer and

shorter chains in all PBR’s tested, with C18:1 having the highest content of all FA analyzed. Smaller

(12:0, 14:0 and 16:0) and longer chain (20:0) FA were detected in trace amounts and are not shown in

table 3.4. One similar study that investigated the lipid and astaxanthin content in C. zofingiensis under

mixotrophic conditions also obtained C18 fatty acids as the major constituents of lipid content and C18:1

with the highest proportion of all FA.[91]

Saturation has a high importance when searching for an ideal biodiesel, with saturated FA being

considered excellent for this purpose. In this project, unsaturated fatty acids were found in higher

quantity than saturated fatty acids in all conditions tested, which could lead to the conclusion that the

lipids synthetized by Chromochloris sp. have no commercial value in biodiesel industry.

However, previous studies also reported that a high proportion of 18:1 in total fatty acids has

potential in biodiesel industry since it has both the oxidation- and low-temperature- stability necessary.

In this project, results show that glucose supplementation and bacteria co-culture did not induce

the synthesis of other FA, changing the diversity of the fatty acid profile but influenced the quantity of

fatty acids synthetized.

Between cultivation conditions, a trend can be observed in table 3.4 with C18 fatty acids

saturated and unsaturated exhibiting a higher proportion in mixotrophic cultivation rather than in

photoautotrophic cultivation. This corroborates what has been said before, as glucose supplementation

increases the lipid accumulation as well as a bacteria co-culture with the algae. Individually 18:0, 18:1,

18:2 and 18:3 FA had an increase of 2-fold in mixotrophic cultivation when compared to control

conditions (PBR 4).

45

Bacterial co-culture also had an enhancing effect in C18 content, with 18:1 and 18:2 being

synthetized 1.5 times more than in control conditions (PBR 4) and 18:0 and 18:3 doubling its synthesis

in co-culture with bacteria.

Overall, mixotrophic cultivation conditions and a bacterial co-culture stimulated the lipid

production with highlight for C18 fatty acids but did not alter the FA profile, with the same fatty acids

being synthesized only in higher quantity. Since it has been reported that stimulating the cells for

astaxanthin accumulation has the same effect in lipid accumulation, when talking about an economic

sustainable project for astaxanthin production with Chromochloris sp., lipids in special 18:1 has the

potential to be a co-product. [91]

3.3 LPS assay: Study of Toxicity

High valuable pigments are currently approved by FDA as a color pigment in animal feed, with

potential applications in cosmetic and pharmaceutical industries being in development but not

implemented. The production of carotenoids from a microalgae source entails risks, like bacterial

contamination along the process. Therefore, it is necessary to ensure that the downstream process

eliminates these risks and the final product is considered safe and non-toxic for human consumption.

Furthermore, in this project two LPS assays were performed to characterize the endotoxin activity of the

different samples, understand how the purification process impacts on endotoxin levels and the

necessity of the addition of a process that ensures the safety of the samples.

3.3.1 Impact of Extraction procedure in Sample Toxicity

The first LPS assay was performed to understand the effect of the extraction procedure in the

endotoxin level of the samples. As control conditions 2 samples were chosen: PBR 3.3 and PBR 4

before extraction, PBR 4 as negative control with its purity ensured by microscope pictures, low

endotoxin levels are expected and PBR 3.3 as positive control with heavily contamination detected by

microscope pictures and high endotoxin levels expected.

The test conditions included then samples of extracts of PBR 1-3.2, without any type of filtration

being applied in the extraction process. The results are shown in Figure 3.9.

46

Figure 3. 9 Endotoxicity level assay performed with biomass (B) and extract (E) samples of PBR 1, 2, 3 and 4. The sample concentration described is 0.2 mg/mL and all the samples were analyzed without filtration (U). The results are expressed as endotoxin units per mililiter, (EU/mL).

Ten samples were tested in the first endotoxin assay with the results showing presence of

endotoxins in all samples tested. Extract of PBR 2 showed the highest endotoxin content (2.51 EU/mL)

while the extract of PBR 3.3 showed the lowest endotoxin level (0.52 EU/mL). The positive control (PBR

3.3 BU) obtained a high endotoxin level as expected but negative control (PBR 4 BU) contradict the

expected results having a high endotoxicity.

Results demonstrated that the extraction procedure decreases the endotoxin level of samples,

opposing PBR 3.3 and PBR 4 samples before and after extraction. While for PBR 4 the reduction in

endotoxin level is of 1.6 times, for PBR 3.3 endotoxin level decreases 5 times with the extraction

procedure. The disparity of the results for both samples indicates that the extraction procedure is not a

trustable procedure to minimize the endotoxin level. Moreover, it suggests the need for the

implementation of other procedures in the downstream process to ensure the safety of the final product.

Extracts of PBR 1 and 2 had high endotoxin levels after the extraction procedure, of

approximately 2.5 EU/mL. The lack of samples of biomass before extraction does not allow a conclusion

on the impact of this procedure in PBR’s cultivated with a bacterial co-culture of Chromochloris sp.

While analyzing the results of PBR 3.1 and 3.2 it is possible to observe low levels of endotoxins

in both cases and no significant difference in the level of endotoxin of both harvestings. In addition, lower

endotoxin levels of extracts of PBR 3 when compared to extract PBR 4 can indicate that the type of

cultivation have impact in the endotoxin level. However, replicates are needed to prove this hypothesis.

0.0

0.5

1.0

1.5

2.0

2.5

3.0

PBR 1EU

PBR 2EU

PBR3.1EU

PBR 3.2EU

PBR 3.3EU

PBR 4EU

PBR 3.3BU

PBR 4BU

Endoto

xin

Level (E

U/m

L)

Endotoxin Level Assay(Sample concentration = 0.2 mg/mL)

1st replicate

2nd replicate

Unialgae - - + + + + + +

Glucose - - + + + - + -

47

When comparing the results obtained in this LPS assay with the proper regulation, all samples

are still above the FDA limit of 0.5 EU/mL, defined for medical devices. The limits defined by the FDA

for cosmetic products are not quantified in terms of Endotoxin units, with the limits depending on the

dosage of use, the patient weight, among other factors. Moreover, theorizing the implementation of

astaxanthin/ canthaxanthin in industry it is necessary to improve the knowledge of the correct dosage,

the formula of the final product before taking further conclusions on the safety of the product.

A comparison with axenic samples, as well as, more replicates should be performed to further

investigate the effect of the extraction procedure in the endotoxin level of the samples and allow more

reliable conclusions.

3.3.2 Impact of Filtration procedure in Sample Toxicity

The second LPS assay was focused on the relevance of a filtration process in the safety of the

samples. Therefore, samples from extracts, biomass and liquid from different PBR’s were chosen to

apply the filtration procedure. Also, to have the possibility to compare the results of the first assay with

the second, a common sample was chosen: PBR 3.3 BU, before extraction and unfiltered. The results

from the second LPS assay are presented in figure 3.10.

Figure 3. 10 Endotoxin level asay of biomass (B), extract (E) and liquid (L) samples of PBR 1, 3.3, 4, 5 and 6. The samples are also classified as filtered (F) or unfiltered (U). The sample concentration chosen was 0.2 mg/mL and the results are expressed as endotoxin units per mililiter, (EU/mL).

Results of the second LPS assay demonstrated that a filtration process has influence on the

endotoxin level in biomass and liquid samples. PBR 3.3 exhibits a decrease in endotoxin level by

Unialgae + + - + + + + + +

Glucose + - - + - + + + +

Filtered + + + + - - - + +

0.0

0.5

1.0

1.5

2.0

2.5

PBR 3.3EF

PBR 4EF

PBR 1BF

PBR 3.3BF

PBR 4BF

PBR 3.3BU

PBR 5LU

PBR 6LU

PBR 5LF

Endoto

xin

Level (E

U/m

L)

Endotoxin Level Assay (Sample concentration = 0.2 mg/mL)

48

approximately half with the filtration process in biomass sample while PBR 5 also decreases the

endotoxicity by applying the filtration process to the sample.

Sample of biomass of PBR 1 filtered obtained the lowest endotoxin level (0.4 EU/mL) of all the

test samples, along with PBR 3.3 and 4 which obtained similar endotoxin levels as the extract samples

of these PBR’s. Moreover, it can be speculated that the filtration process has a similar effect in endotoxin

level as the extraction procedure.

By direct comparison with the first endotoxin assay, it is possible to investigate the influence of

filtration in the endotoxicity of sample PBR 3.3 extract. The results of both assays indicate that a filtration

procedure does not affect endotoxicity in extracts, since PBR 3.3 extract sample have a higher endotoxin

level filtered (0.74 EU/mL) than unfiltered (0.52 EU/mL).

However, comparing the results of PBR 3.3 BU, a sample tested in both assays, the results

show a decrement in endotoxicity in the second assay, with endotoxin level decreasing from 2.5 EU/mL

to 1.0 EU/mL. This result invalidates a direct comparison between the two assays, with further assays

needed to understand the effect of a filtration procedure in extract samples.

The introduction of high valuable pigments with cosmetic and pharmaceutical applications in

these industries is currently a trending topic causing an urgent need to ensure the safety and no-toxicity

of the final product. [92]

3.4 Extraction

To optimize the extraction procedure the first step was to understand which part of the standard

SOP could be improved. Since the limiting steps of an extraction are the cellular disruption and the

carotenoid recovery it was decided to study the influence of the glass beads size in the disruption of the

cells and the solubility of the carotenoids in different solvent mixtures. However, since previous studies

indicated that the mixture of DCM and Methanol (1:1) was efficient in recovering pigments in other

strains, the focus of the optimization of the extractions was moved to the impact of the glass beads size.

3.4.1 Impact of glass beads size

To study the impact of the cell disruption in the extraction, four different type of condition were

tested. These conditions were focused on the size of the glass beads used to disrupt the cells. This is a

really important step since that, in theory, the higher the number of the cells disrupted the higher the

carotenoid content. Furthermore, there were three different types of glass beads and a mix of glass

beads with different sizes, with the effectiveness of the cell disruption methods being determined using

extraction yield percentages.

In this case, it was used the biomass from PBR 3 (.2) at the normal harvesting time since this is

a biomass that is expected to have a high content of carotenoids. The results of this test can be observed

in figure 3.11. Microscope pictures were taken at various stages of the extraction process to investigate

the effectiveness of cell disruption. The cell disruption in each experiment can be seen in figure 3.12

with pictures taken while doing this experiment.

49

Figure 3. 11 Effect of different glass bead sizes in cell disruption. The average extraction yield is reported as a percentage of Dry Weight and it is the result of the mean between two independent samples.

Four different glass beads size were tested to extract high valuable pigments from

Chromochloris sp. All four methods successfully rupture the cells and release the intracellular

components, (Figure 3.11 and 3.12) From the four methods tested, maximum yield of extraction was

obtained using glass beads with a diameter of 0.25 - 0.50 mm (73.1%) while minimum yield of extraction

was obtained using 4.0 ± 0.3 mm glass beads (50.9%).

One similar study investigated the influence of glass beads size in cell disruption and extraction

of lipids and proteins in Chlorella vulgaris with results indicating that glass beads with approximately

0.40 mm of diameter were the most effective in extracting the components.[93]

While looking at the error bars in Figure 3.11 it is possible to observe that the small and mix

condition have the highest error associated, which means that the replicates made were distant from

each other. The use of smaller beads for the extraction of carotenoids enhances the cell breakage into

smaller pieces; these are difficult to settle in the pellet in the centrifugation step, which causes the

supernatant to have a turbid appearance and the collection of it for further evaporation difficult.

In the case of 4.0 mm glass beads, the usage of two beads with the vortex for 3 minutes is not

enough to disrupt the cells. Unlike smaller beads, this type of beads induces not the breakage of the

cells into small pieces but one big opening that allows the inside content to be released. Furthermore,

more cycles of vortex/centrifugation are necessary to achieve the same level of disruption as the smaller

beads.

0.0

10.0

20.0

30.0

40.0

50.0

60.0

70.0

80.0

90.0

100.0

0.25-0.50 mm 0.75-1.0 mm 4.0± 0.3 mm 0.25-0.50 mm + 4.0±0.3mm

Extr

action Y

ield

[P

erc

enta

ge o

f D

W (

%)]

Glass Beads Size Tested

Glass Beads Size Test

50

Figure 3. 12 Microscope pictures of samples from the extraction procedure with different glass beads size after the first centrifugation cycle (Magnification: 1000x) A: 0.25 – 0.50 mm; B: 0.75 – 1.0 mm; C: 4.0 ± 0.3 mm; D: 0.25 – 0.50 mm + 4.0 ± 0.3 mm

Overall, bead milling can be considered a good process for extracting intracellular components

from Chromochloris sp., with the effectiveness on extracting high valuable pigments being studied

further in this project by an HPLC procedure. Other approaches can be further investigated in an effort

to improve the extraction yield of this specific strain since available literature suggests that cell disruption

strongly depends on microalgae species, age of the culture and composition of cell wall. Therefore,

results obtained from one species cannot be generalized to all other species.

Other approaches can be further investigated in an effort to improve the extraction yield of this specific

strain since available literature suggests that cell disruption strongly depends on microalgae species,

age of the culture and composition of cell wall. Therefore, results obtained from one species cannot be

generalized to all other species. [1], [68,70]

3.5 Fractionation

The purpose of a fractionation procedure is to extract different carotenoids in different fractions.

Doing so, this procedure can be seen as a purifying step in the production of one specific carotenoid.

The main goal of the purification step in this project was to obtain astaxanthin and canthaxanthin isolated

from other carotenoids and if possible also separated from each other in different fractions.

While looking for the best approach to separate both pigments, the first thought was to see the

difference between the two standard methods of extraction that are currently used in MCI to extract

components in microalgae: ethanol: water (8:2) and dichloromethane: methanol (1:1) and compare the

extraction rate and carotenoid content.

A B

D C

51

The second approach focus on polarity and its connection with carotenoid affinity and solubility.

In this case two fractionation procedures were developed, an apolar and a polar fractionation. These

two procedures were based in the same principle: the polarity of the solvents that constitute them. In

the polar approach, 5 different solvents were chosen and ordered by polarity, from the least polar to the

most polar. The apolar approach was similar, but in this case with the solvents being ordered from the

most polar to the least polar. The solvents were chosen according to their relative polarity and their

position in the polarity scale, so that a big portion of this scale would be covered.

The third approach has the aim to develop a fractionation procedure that was based on the

relation of carotenoid solubility with solvent relative polarity. Based on literature astaxanthin is soluble

in solvents like acetone and acetic acid while canthaxanthin is soluble in solvents like ethanol, acetone

and ethyl acetate.[24,37,94] Arranging the solvents in the polarity scale, it was found a relation between

carotenoid solubility and solvent polarity. So, it was thought that astaxanthin prefers solvents with a

polarity between 0.20 and 0.55 while canthaxanthin is soluble in solvents with a polarity between 0.10

and 0.355. Positioning these values on a scale, as it can be seen in figure 3.13, it is possible to observe

that there is zone where both pigments are soluble and two zones where, in theory, only

canthaxanthin/astaxanthin are soluble.

Figure 3. 13 Relative Polarity Scale of solvents, with the interval of solubility of astaxanthin in orange and canthaxanthin in blue with the region where the solubility of the two carotenoids intersect.

In this case, two procedures were considered to separate astaxanthin from canthaxanthin.

Following again the principle of 5 successive extractions with 5 different solvents/ solvent mixtures, in

both of these procedures the first two solvent mixtures were chosen according to the interval where only

one carotenoid was soluble and the next two solvents on the interval where the other carotenoid was

soluble. For instance, one procedure focused first on the interval where only astaxanthin was soluble,

choosing two solvent mixtures with a relative polarity between 0.355 and 0.55 and the third and fourth

fractions were focused on canthaxanthin choosing solvent mixtures with a polarity between 0.20 and

0.30.

The last approach was to consider all the polarity scale and investigate which part of this scale

corresponds to the highest yield of extraction and highest carotenoid content. To do so, five solvent

mixtures were chosen so that all the polarity scale would be covered.

3.5.1 Comparison between different methods

Six methods of fractionation were evaluated to understand the efficiency of a purification step

to separate different carotenoids and understand the affinity of the pigments with different solvent/

solvent mixtures. The effectiveness in extracting intracellular components from Chromochloris sp. is

analyzed as percentage of dry weight in Figure 3.14.

Astaxanthin

Canthaxanthin

0 1 0.50 0.55 0.355 0.10 0.20

52

Figure 3. 14 Extraction yields of fractionation methods tested. All approaches are based in one replicate with exception to method A, the standard method, where it was performed two replicates.

0.0

10.0

20.0

30.0

40.0

50.0

60.0

Ethanol Acetronitrile Acetone DCM Chloroform Chloroform

Tota

l Y

ield

of

Extr

action (

% o

f D

W)

B: Apolar Scheme

0.0

10.0

20.0

30.0

40.0

50.0

60.0

Hexane EthylAcetate

Ethanol Methanol Water Water

Tota

l Y

ield

of

Extr

action (

% o

f D

W)

C: Polar Scheme

0.0

10.0

20.0

30.0

40.0

50.0

60.0

Acetonitrile:DCM (2:1)

Isopropanol:DCM (2:1)

EthylAcetate:

Acetone (2:1)

EthylAcetate:

DCM (2:1)

Methanol:Hexane (2:1)

Tota

l Y

ield

of

Extr

action (

% o

f D

W)

D: Astaxanthin/Canthaxanthin

0.0

10.0

20.0

30.0

40.0

50.0

60.0

Ethyl Acetate:Acetone (2:1)

Ethyl Acetate:DCM (2:1)

Acetonitrile:DCM (2:1)

Isopropanol:DCM (2:1)

Methanol:Hexane (2:1)

To

tal Y

ield

of

Extr

actio

n (

% o

f D

W)

E: Cantaxanthin/Astaxanthin

0.0

10.0

20.0

30.0

40.0

50.0

60.0

Water:Ethanol (1:1)

Ethanol:Isopropanol

(1:1)

Isopropanol:Acetonitrile

(1:1)

Acetonitrile:DCM (1:1)

DCM : EthylAcetate (1:1)

EthylAcetate:

Hexane (1:1)

Tota

l Y

ield

of

Extr

action (

% o

f D

W)

F: Polarity Scale

-10.0

0.0

10.0

20.0

30.0

40.0

50.0

60.0

Ethanol:Water(8:2)

Ethanol:Water(8:2)

DCM:Methanol

(1:1)

DCM:Methanol

(1:1)

Ethanol:Water(8:2)

DCM:Methanol

(1:1)

Tota

l Y

ield

of

Extr

action (

% o

f D

W)

A: Standard Method

1st Replicate 2nd Replicate

53

All the fractionation methods tested resulted in the extraction of intracellular components.

Regarding the extraction yield, from all the procedures tested, individually methanol obtained the highest

yield of components extracted while as a method polar scheme obtained the higher extraction yield.

Results of apolar and polar scheme methods show different patterns. While in the apolar

scheme ethanol show an extraction rate of 10%, in polar scheme solvent hexane exhibits an extraction

yield of 1%. This may be explained by the inability of hexane to aid the cell disruption and to solubilize

the intracellular components. The low polarity of this solvent along with its inability to extract components

from Chromochloris sp. indicates the preference for more polar solvents on the extraction of high

valuable pigments.

Moreover, results show a high extraction rate with chloroform (22.2%), however the addition of

this solvent to the pellet caused the dissolution of it and further centrifugation cycles were incapable of

a proper separation of phases. The high dissolution power, the rapid evaporation and its toxicity are

aspects that make chloroform a poor extraction solvent in this case.

Results of the polarity scale method (F) show a higher extraction rate of intracellular components

in solvents mixtures: Ethanol: Isopropanol (1:1) and Isopropanol: Acetonitrile (1:1). The relative polarity

of these mixtures is concentrated in the middle of the polarity scale with a slight deviation to the apolar

side. This result suggests that pigments and other intracellular compounds prefer solvents with a

moderate polarity instead of the extremes of the polarity scale. Other studies have shown the preference

of non-polar solvents when extracting high valuable pigments. [70,71] The structure of these molecules

provides them a high degree of hydrophobicity and therefore a preference for non-polar solvents.

Regarding the standard extraction method approach, similar results are described. While the

mixture DCM: Methanol obtain high extraction yields, the mixture Ethanol: Water (8:2) obtained low

extraction yields, with even negative yields being found. These results indicate the inability of the polar

extraction procedure in extracting pigments, with the high degree of polarity of this mixture does not

allow the dissolution of pigments and other metabolites.

In the attempt to create a process of fractionation able to separate astaxanthin from

canthaxanthin, two approaches were tested. In both approaches the results showed a higher extraction

rate in the first two solvent mixtures applied. In the approach to separate canthaxanthin from astaxanthin

the solvent mixture ethyl acetate: acetone (2:1) obtained the highest extraction rate, while in the

astaxanthin approach, Acetonitrile: DCM had the highest extraction rate.

A similar pattern of results was obtained in a previous study that investigated the influence of

various solvent mixtures in the extraction of astaxanthin synthetized by Phaffia rhodozyma, with the

results showing that acetone and isopropanol were efficient in extracting astaxanthin.[94]

Overall, results showed a preference for non-polar solvents on the extraction of intracellular

components, except for methanol on polar scheme method. This solvent obtained an extraction rate of

almost 60%. Since the affinity of pigments for non-polar solvents has been reported by previous studies

and in this project, this can be explained by an error committed while collecting the supernatant.

54

Therefore, to prove the efficiency of methanol on extracting intracellular compounds from Chromochloris

sp. replicates of this method are needed.

The results also suggest that other attempts should be made while optimizing the extraction

procedure. As of now, the standard procedure uses a mixture of DCM: Methanol to extract the pigments

for further investigation. However, even though this mixture obtains high extraction rates, the selectivity

of products extracted can increase and new solvent mixtures should be tested. Furthermore, new

approaches can be pursued to optimize the carotenoid extraction rates from Chromochloris sp.

The development of a fractionation process able to purify canthaxanthin and astaxanthin is

incomplete, with the results needed to be validated introducing replicates to the study, further testing of

different solvents/ solvent mixtures and new approaches investigated. However, the time limitation

during the project made impossible a more detailed investigation of this procedure and further

conclusions.

3.6 Analytics

3.6.1 Method Development

A HPLC method was developed to quantify and identify the pigments recovered in the

extractions performed. The first approach was to use a C18 column allied with a mobile phase composed

by two solvents, A (Acetone: Water 1:1) and B (Acetone). Although this approach was implemented by

Austrian Drug Screening Institute (ADSI) in their projects that involve pigments identification, the use of

this elution method with a C18 column (3 μm (100 x 2 mm)) was not ideal in this project. Over pressure

problems (>400 bar) while running the standards showed the impossibility of using this method. A

primary solution was to reduce the flow in half (0.5 mL/min) which would double the run time from 30 to

60 minutes. Even though with these conditions the pressure problems were overcome it was impossible

to obtain peaks that would allow the identification of astaxanthin. Moreover, problems in runs with lower

concentrations of the standards were associated with the advanced age of the column. The long run

times, the inability to properly identify and differentiate astaxanthin esters allied to the age of the column

led to the decision of purchasing a new column and adopt a new method of elution.

The new column was also a C18 (5 μm (250 x 4 mm)) and it was also purchased a guard column.

The guard column and a step of filtration of the samples prior to the injection will protect the column of

some impurities present in the samples and preserve the column for a longer time. The new mobile

phase comprehend two solvents: A (Acetonitrile: Methanol (74:6)) and B (Methanol: Hexane (5:1)) and

is currently applied by the Botanical Institute of Innsbruck in the detection of pigments. One of the

advantages of using this method was the temperature. In this case the column was stabilized at 30 °C

and it was at this temperature that the elutions were performed thus the possibility of thermal

degradation is improbable since the pigments only start to degrade at T > 60ºC.

Moreover, the slightly higher particle size (5 μm) allows higher flows and shorter run times

without the existence of overpressure problems and without compromising the efficiency of the HPLC

analysis.

55

With the new approach, the standards of astaxanthin and canthaxanthin were successfully

eluted. While canthaxanthin exhibits a single peak at 9.92 min that is classified as the free form of

canthaxanthin, the astaxanthin standard is a mixture of esters from Haematococcus pluvialis and

likewise the chromatogram shows the elution of this mix between 11.192 and 21.038 min with the

highest peak at 12.574 min. (Figure 3.15)

Figure 3. 15 Chromatograms of canthaxanthin (150 µg/mL) (A) and astaxanthin (500 µg/mL) (B) standards and respective retention times. The black arrow shows the peaks chosen to quantify in the samples.

Good results in the elution of canthaxanthin and astaxanthin as well as a differentiation of

astaxanthin esters allowed for the next step of the method development.

To quantify the pigments a range of concentrations of both astaxanthin and canthaxanthin

standards were chosen to test. In the case of canthaxanthin, the concentrations were: 150, 100, 50, 10

and 5 µg/mL, while for astaxanthin concentrations of 500, 250, 100, 50, and 10 µg/mL were chosen. To

quantify the pigments in the samples a co-relation was made between the concentration of the pigment

and the integrated peak area of the eluted pigment. While this was a simple task for canthaxanthin that

has a single peak in its chromatogram, in the case of astaxanthin with a mixture of esters the highest

peak was chosen to integrate.

Furthermore, two replicates were made for each concentration and the peak area is an average

of both measurements for each concentration. The standards curves obtained for both canthaxanthin

and one astaxanthin-ester are shown in Figure 3.16.

A

B

56

Figure 3. 16 Calibration curve of canthaxanthin (left) and astaxanthin (right) using the general method for quantification of pigments in samples. The equations obtained were [canthaxanthin] = 0.33416 Area Peak – 0.05557, R² = 0.99998 for canthaxanthin and [astaxanthin] = 1.12697 Area Peak + 2.00414, R² = 0.999874 for astaxanthin.

The linearity of both curves is proved by the high values of squared r, 0.9998 for canthaxanthin

and 0.999874 for astaxanthin ester. Recurring to the Data Analysis software of Excel the limit of

detection (LOD) and limit of quantification (LOQ) were calculated. In canthaxanthin’s case the values

were 2.3 and 15.8 µg/mL, respectively while in astaxanthin case LOD was 4.9 µg/mL and LOQ was 32.9

µg/mL. LOD gives the perception of the lowest standard concentration that is reliably distinguished from

the basal line while LOQ is the lowest concentration of analyte that can be determined with an

acceptable level of precision without interferences. These results show that the method development

still has room to be optimize so lower concentrations can be analyzed as well as other pigments

quantified.

In terms of repeatability, since the pigments are light sensitive and measurements of the same

sample with one week apart showed loss of integrated area peak, a decision was made of: prior to the

analysis of the extracts, also analyze the standards so it was possible to correlate with the calibration

curve and introduce a possible correction.

Parallel to the development of the general method that allows the identification of both

astaxanthin and canthaxanthin, attempts were made to develop two specific methods that would allow

a rapid identification of both pigments separately. These methods are especially interesting in cases

where only one of the pigments is of interest. For canthaxanthin the attempt of method development

was successful with the method based on the general one having a runtime of 15 minutes that allows

the rapid identification of canthaxanthin.

However, for astaxanthin the runtime of the specific method was of 25 minutes only 2 minutes

less that the general method. Therefore, to allow a rapid identification of astaxanthin further optimization

is needed.

0

20

40

60

80

100

120

140

160

0 100 200 300 400 500

Canth

axanth

in C

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Canthaxanth in Standard Curve

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Astaxanth in Standard Curve

57

3.6.2 Influence of cultivation conditions in astaxanthin and canthaxanthin content

The success of the method development turned possible the quantification of the content in

astaxanthin and canthaxanthin in the biomass harvested and extracted.

For the first running, one of the samples that in theory was expected to have a higher carotenoid

content was chosen, PBR 5. The chromatogram obtained is shown in Figure 3.17.

Figure 3. 17 Chromatogram obtained after analysis of N and P deficiency cultivation of Chromochloris sp. (PBR 5) and retention times identified by the software ChemStation. The analysis was performed by injecting 5 µL of sample in a C18 5 μm (250x4 mm) column. The flow was kept at 1 mL/min during washing, injection and elution and the temperate at 30ºC.

The chromatogram shows a peak at 9.924 min identified as canthaxanthin and one at 12.590

min identified as an astaxanthin ester. Recurring to a chromatogram provided by Innsbruck Botanical

Institute (Annex 6.4) that shows the elution of a sample of Haematococcus pluvialis it is possible to

classify the astaxanthin esters. In this provided chromatogram free form astaxanthin elutes at min 5.512,

while monoesters forms elutes from min 8.572 to 9.679 and diesters from min 10.879 till min 13.383.

According to this classification the astaxanthin eluted in this sample is classified as a di-ester form of

astaxanthin.

An HPLC analysis was performed to study the influence of different cultivation conditions in

carotenoid content. Samples of the 8 PBR’s (PBR 1 to 8) were measured to quantify astaxanthin and

canthaxanthin. Replicates of each sample were performed with each sample and its duplicate run twice

to ensure the validity of the analysis.

Chromatograms of different cultivation conditions showed different carotenoid profiles.

Chromatograms of PBR 4, 3.1 and 3.2 are shown as examples in Figure 3.18 and 3.19. The results of

carotenoid content described as mg of pigment per g of biomass are shown in Figure 3.20.

Figure 3. 18 Chromatogram obtained after analysis of Chromochloris sp. in control conditions (PBR 4) with retention times identified by software ChemStation. The analysis was performed by injecting 5 µL of sample in a C18 (5 μm 250x4 mm) column. The flow was kept at 1 mL/min during washing, injection and elution and the temperate at 30ºC.Canthaxanthin and astaxanthin diester are marked by the black arrow and unidentified significant peaks are marked by the blue arrow.

58

Figure 3. 19 Chromatograms obtained after analysis of Chromochloris sp. in two-stage cultivation at A) early stationary phase (PBR 3.1) and B) end of stationary phase (PBR 3.2) with retention times identified by software ChemStation. The analysis was performed by injecting 5 µL of sample in a C18 (5 μm 250x4 mm) column. The flow was kept at 1 mL/min during washing, injection and elution and the temperate at 30ºC. Canthaxanthin and astaxanthin diester are marked by black arrows and unidentified significant peaks are marked by blue arrows.

Figure 3. 20 Carotenoid content in different cultivation conditions. An average of the carotenoid content is reported as mg of pigment per g of biomass.

Both astaxanthin and canthaxanthin were successfully identified in all samples measured.

Furthermore, all samples showed a higher content in astaxanthin than in canthaxanthin. PBR 6, that

was submitted to a N and P deficiency stress exhibit the highest content in astaxanthin while PBR 5 that

was put through a high salinity stress exhibit the highest content in canthaxanthin.

In the second set of cultivation comparing the cultivation conditions tested (PBR 5-7) to the

control (PBR 8) it is possible to observe that the impact of the stresses on astaxanthin accumulation is

reduced, in the order of 2.0 mg higher. N deficiency has been previously reported to enhance

astaxanthin accumulation in Haematococcus pluvialis and in Chlorella zofingiensis. [15], [53] It is worth

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PBR 1 PBR 2 PBR 3.1 PBR 3.2 PBR 3.3 PBR 4 PBR 5 PBR 6 PBR 7.1 PBR 7.2 PBR 8

Caro

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Canthaxanthin Content Astaxanthin Content

A

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59

noting that the implementation of the N deficiency stress factor is always accompanied by a high light

irradiance, which can explained the low astaxanthin yield in this project since high light irradiance was

not studied. Further investigation should focus on understanding the impact of higher light intensities

and its conjugation with an N deficiency stress in astaxanthin content.

In the case of canthaxanthin, high salinity induced the accumulation of this pigment with results

showing a canthaxanthin content 2 times higher in this condition in comparison with the control

conditions (PBR 8), while N and P deficiency did not influence canthaxanthin accumulation. High salinity

stress was previous reported to enhance canthaxanthin accumulation in Chlorella zofingiensis. [59]

Bacterial co-culture impact represented by PBR 1 and 2 shows uneven results between them

with PBR 1 displaying higher contents of astaxanthin and canthaxanthin than PBR 2. However, the error

bar of PBR 2 indicates that astaxanthin/canthaxanthin content has a significant error introduced in the

calculation of these pigments contents. Furthermore, while comparing PBR 1 and 2 with PBR 4, results

show no significant influence of bacterial co-culture in canthaxanthin and astaxanthin-di-ester content.

The analysis of the chromatograms (Figure 3.18 shows the example of PBR 4) showed an

interesting pattern with PBR 1, 2 and 4 displaying two unidentified single peaks at min 9.385 and at min

11.826. No difference in quantification and diversity of analytes between PBR 1, 2 and 4 suggests that

bacterial co-culture does not influence pigment production and accumulation.

However, the fact that the three PBR’s had the same chromatogram pattern and knowing that

all three followed a photoautotrophic cultivation, the study progressed into observing the influence of

different type of cultivations in pigment accumulation.

Therefore, a comparison of carotenoid content in PBR 7 and PBR 8, photobioreactors with

purely cultivation states, mixotrophic and phototrophic respectively, shows no influence of glucose and

mixotrophic cultivation in carotenoid accumulation. While chromatogram of PBR 8 showed the same

pattern of PBR 4 with two unidentified peaks around 9 and 11 minutes, chromatogram of PBR 7 shows

the absence of these peaks. Therefore, although mixotrophy shows no influence in astaxanthin and

canthaxanthin quantity, results show a clear influence in pigments diversity.

The identification of these peaks remains unclear. A comparison with Botanical Institute

reference chromatogram (Annex 6.4) shows that the peak at 9.385 min could potential be chlorophyll a

or a mono-ester form of astaxanthin while the peak at 11.826 min could be an astaxanthin di-ester form

or β-carotene. An argument can be made that 9.385 peak probably is chlorophyll a, hence the presence

in photoautotrophic cultivation and the absence in mixotrophic. This could be a possible explanation for

the change in cell color from green to orange when glucose is supplemented. However, to validate this

hypothesis it would be necessary to acquire chlorophyll a standard and compare the results with the

peak obtained, which due to time limitations it was not possible do.

PBR 3 obtained good results in terms of algal growth and biomass yield. Regarding carotenoid

accumulation, the two-stage cultivation strategy shows no influence in astaxanthin content and an

increment of canthaxanthin content when compared with PBR 7 (pure mixotrophy) and 8 (pure

phototrophy).

60

When opposing carotenoid content of PBR 3.1 and 3.2 that correspond to a harvesting in the

beginning and at the end of stationary phase, respectively, results show a higher carotenoid content in

PBR 3.2. The increment of the astaxanthin content from one stage to the other is not significant with the

majority of the carotenoid synthesis happening before stationary phase. The same trend is observed for

PBR 7, with the early (7.1) and normal (7.2) harvesting showing no significant difference in astaxanthin

content when compared. In canthaxanthin’s case the increment of the content from early to late

stationary phase is reduced but significant (approximately 4.0 mg/g) in PBR 3. In purely mixotrophic

cultivation (PBR 7), canthaxanthin content remains similar in both harvestings suggesting that its

accumulation is not induced by stationary phase but probably due to the glucose supplementation.

Furthermore, astaxanthin synthesis show the same pattern as biomass growth, which indicates

that the pigment is a growth-associate product. Others have demonstrated the same for astaxanthin,

showing that its synthesis and accumulation is correlated to an early exponential phase in Chlorella

zofingiensis.[95] In contrast, astaxanthin production by H. pluvialis requires a two-stage strategy

because it is synthesized during the secondary carotenoid induction phase.

The low accumulation of astaxanthin and canthaxanthin during stationary phase also suggests

that the color change of PBR 3 is mostly due to a decrease of photosynthetic activity and not derived

from an increment of astaxanthin and canthaxanthin accumulation. However, the reason behind this

change should be investigated with harvests before and after glucose supplementation.

When observing PBR 3.1 and PBR 3.2 chromatograms (Figure 3.19) a curious result was found.

Besides the astaxanthin di-ester and the canthaxanthin peaks, two unidentified peaks were found.

These peaks have the same run time of the ones found in PBR 4 and 8 chromatogram that followed a

photoautotrophic cultivation. Moreover, since these peaks were found in phototrophic cultivation and

PBR 3 followed this type of cultivation until day 16 of cultivation, it was expected that PBR 3 also showed

the presence of these unidentified peaks. Even though, PBR 3.1 and 3.2 chromatograms show the

presence of the unidentified peaks it is possible to see a decrement of the area of both peaks in PBR

3.2. This can represent a change that occurs during stationary phase and that may be associated with

the supplementation of glucose to the media and subsequent decrease of photosynthetic activity.

Regarding the impact of light regimes, while comparing both cultivation sets, PBR 1-4 and PBR

5-8, results show a higher astaxanthin content in the second set. However, the same observation cannot

be made for canthaxanthin, with the distribution of this pigment in both cultivation sets appearing random

and non-homogeneous.

Focusing on PBR 4 and 8 it is possible to relate the light regime with pigment accumulation.

Both studied pigments show a higher content in light regime of 16:8 h than in continuous light regime

(24 hours). This result suggests that pigment accumulation is influenced by photoperiod regime and that

the existence of a dark phase is important in pigment accumulation. Previous studies show no influence

of light in canthaxanthin accumulation, while high light irradiance is a stress factor commonly used to

enhance astaxanthin accumulation by microalga Chlorella zofingiensis. [33,51]

61

3.6.3 Comparison between different glass beads size in carotenoid content

An HPLC analysis was performed to understand the difference of different glass beads size in

the extraction of carotenoids. Figure 3.21 shows the carotenoid content of astaxanthin and

canthaxanthin for the different sizes of glass beads tested.

Figure 3. 21 Carotenoid content with different extraction procedures. An average of the carotenoid content is reported as mg of pigment per g of biomass.

All extraction procedures tested were able to extract astaxanthin and canthaxanthin from the

intracellular media. Glass beads of 0.75-1.0 mm obtained the highest content of astaxanthin and

canthaxanthin, contrary to glass beads of 4.0 ± 0.3 mm that obtained the lowest content in both

carotenoids.

The carotenoid content of the mix of glass beads result is similar to the smaller beads showing

no advantage in using this strategy to enhance cell disruption and carotenoid recovery. Furthermore,

the results indicate that even though glass beads of 0.25 - 0.50 mm obtained higher extraction yields,

glass beads of 0.75 - 1.0 mm are more effective in extracting carotenoids. However, the sensitivity

limitation of the HPLC analysis as well as the small difference of carotenoid content in both tests does

not allow the selection of one method over the other.

The results confirm that bigger glass beads (4.0 mm) are not the ideal method for cell disruption

of Chromochloris sp. cells showing lower extraction rates and lower carotenoid content than the other

methods. It is also the method that is more time consuming and leads to a higher energy consumption.

3.6.4 Comparison between different fractionation methods

The present study was directed towards the understanding the effect of the different solvent

mixtures in fractionating astaxanthin and canthaxanthin. Figure 3.22 shows the results obtained for

carotenoid content in five fractionation procedures. Standard method performed poorly in carotenoid

extraction and likewise due to time limitation, the results of the HPLC analysis were not interpreted.

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tenoid

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f caro

tenoid

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62

Figure 3. 22 Carotenoid content in the five fractionation procedures studied.

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E: Canthaxanthin/AstaxanthinCanthaxanthin Content Astaxanthin Content

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Ethyl Acetate :Hexane (1:1)

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F: Polarity Scale Scheme

Canthaxanthin Content Astaxanthin Content

63

Results of carotenoid content in fractionation procedure show the same trend observed in the

extraction yields, with non-polar solvents obtaining higher carotenoid contents than polar solvents.

Methanol in polar scheme obtained the highest extraction yield of all solvents and methods,

however, the same is not observed in terms of carotenoid content. One possible explanation is that the

supernatant collection of this fraction had an error evolved, with biomass fragments also being collected

or that methanol extracts other type of intracellular compounds that have a high affinity to polar

components.

The same conclusion can be reached in the case of chloroform (apolar scheme) that obtained

22.2% of extraction yield in the apolar scheme but performed poorly when extracting carotenoids with

contents bellow 1 mg/g. In chloroform’s case, the difference observed between extraction rate and

carotenoid content is possibly due to the high dissolution capacity of chloroform observed when

performing the supernatant collection. This is indicative that chloroform is not the adequate solvent when

extracting or fractionating carotenoids from Chromochloris sp.

In the same line of thinking, hexane performed poorly in extraction rate and in carotenoid content

which confirms the incapacity of solubilize carotenoids and extracting them.

Regarding acetone and ethyl acetate, from apolar and polar scheme, respectively, both these

solvents had high extraction rates and carotenoid contents. This suggests that the moderate polarity of

these solvents seems to be ideal for carotenoid extraction and therefore should be considered for further

extraction and fractionation purposes. However, it is necessary to point the inability to separate

canthaxanthin from astaxanthin with the content of both carotenoids being similar in both methods.

Moreover, when analyzing the results of astaxanthin/canthaxanthin approach the first two

solvents: acetonitrile: DCM (2:1) and isopropanol: DCM (2:1) obtained high carotenoid content

confirming the results showed before regarding extraction yields. In the case of canthaxanthin/

astaxanthin approach even though the first two solvents had high extraction rates, only mixture ethyl

acetate: acetone (2:1) obtained high carotenoid content. This indicates that mixture ethyl acetate: DCM

(2:1) is efficient in extraction intracellular compounds but not astaxanthin or canthaxanthin.

In terms of separation of astaxanthin from canthaxanthin and vice-versa, acetonitrile and mixture

of Acetonitrile: DCM (2:1) in astaxanthin fractionation procedure seems to be the only solvents that with

some degree of efficiency separates astaxanthin from canthaxanthin. In fact, the structure similarity of

these two molecules turns the isolation of one from the other based in solvent polarity, a difficult task.

Results of polarity scale scheme indicate again the higher affinity of carotenoids for solvents

with a moderate polarity with mixture water: ethanol (extremely polar) and mixture ethyl acetate: hexane

(extremely apolar) performing poorly when extracting carotenoids.

64

4. Conclusions and Future Work

The main goal of this project was to accomplish a successful method of cultivation of alga

Chromochloris sp., extract its intracellular compounds and develop an HPLC method able to identify

and quantify high valuable pigments, in particular, astaxanthin and canthaxanthin.

Regarding cultivation, several strategies were tried to enhance the accumulation of high-

valuable pigments: bacterial-co-culture, light regime, N and P depletion, high salinity, mixotrophic

cultivation and two-stage cultivation.

Bacterial co-culture strategy in photoautotrophic was proven not beneficial in enhancing cell

growth and carotenoid content. Further research of this strategy should involve the identification of the

bacteria in co-culture with Chromochloris sp. and focus on understanding possible symbiotic interactions

between both microorganisms.

High salinity stress and N and P deficiency induced a decaying effect in algal growth once the

stress was induced. This decrement of cell density is thought to be associated with the acidic pH values

that the culture achieves once the stress is induced which slows the algae growth. Regarding carotenoid

content, high salinity stress induced the accumulation of canthaxanthin but showed no influence in

astaxanthin content. N and P deletion had the inverse effect with astaxanthin content increasing in this

condition while canthaxanthin content remained similar to the control condition.

High salinity and N and P depletion are conditions that should be further studied since they

showed an enhancing effect of canthaxanthin and astaxanthin, respectively. Future work should focus

on optimizing the salt concentration that enables high levels of canthaxanthin but allows algae to

maintain its growth. In N and P deficiency it would be interesting to understand the effect of the depletion

of each nutrient in pigments content. Also, allying N depletion to high light intensity should be

investigated since others reported this strategy as one that enhances astaxanthin content.

Mixotrophic cultivation was studied as an alternative to photoautotrophic cultivation. Results

showed an increment of the growth rate but no change in biomass accumulation. In the presence of

glucose, cells turned orange, which it is suspect to be due to a decrease of the photosynthetic activity

and not due to a higher carotenoid accumulation since carotenoid content was similar to control

conditions.

Two stage cultivation condition enhanced algal growth and biomass yield showing the strategy

potential to mass cultivation of Chromochloris sp.. Regarding carotenoid content, the strategy did not

have an impact in astaxanthin accumulation and showed a reduced increment in canthaxanthin content.

Future work should focus on understanding the optimal glucose concentration that improves cell growth

and enhances canthaxanthin accumulation.

Light regime was studied in photoautotrophic cultivation, with results showing an increment of

algal growth in the presence of a continuous light source. An inverse trend was found in carotenoid

content results with PBR’s with a light regime of 16:8 (L:D) having higher carotenoid content than with

65

continuous light regime. It appears that a dark phase is important to astaxanthin and canthaxanthin

accumulation. To prove this theory other light regimes should be investigated as well as other light

intensities, since studies have reported the enhancing effect of high light intensities in astaxanthin

content.

In summary, Chromochloris sp. is a strain with potential in the high valuable pigment production

but further optimization is needed. It is worth noting that the results obtained for cultivation data are

dependent of one experiment for each condition furthermore, replicates are necessary to validate the

conclusions.

The lipid content of the first cultivation set was studied since lipids can interfere with the

extraction of high-valuable pigments. Results showed an increment of total lipid content when glucose

was supplemented to the media (PBR 3), the same effect was observed in PBR 1 and 2 that were

submitted to a bacterial co-culture cultivation. The study allowed for the association of lipid accumulation

to the early stationary phase with PBR 3.1 obtaining the highest lipid content. Results of the fatty acid

profile showed that stress conditions tested caused a difference in quantity of lipids but not in quality, as

the same profile was obtained in all PBR’s samples tested. Lipid profile shows 18:1 as the major fatty

acid in the cultivation tested. Further investigation is needed to understand the impact of other stress

conditions in lipid profile and to study the interest and applicability of 18:1 as a co-product on the

production of high-valuable pigments by Chromochloris sp.

Endotoxicity study was performed to understand the potential of Chromochloris sp. synthetized

pigments in future applications. Results showed that the extraction procedure decreases the endotoxin

level of the samples. Bacterial co-culture PBR’s performed poorly in the endotoxin test after extraction,

which can be a drawback in the usage of this strategy to enhance pigment production.

The second LPS assay focused on the implementation of a filtration process that in theory could

be applied in the pigment production process to ensure its safety. Results showed a decrement of

endotoxin level in biomass and liquid samples but no effect was found in extract samples. In conclusion,

a filtration process is a good idea but further improvement is needed. Furthermore, the impact of different

cultivation conditions, of the extraction procedure and other purification processes should be examined

by further research. The impending FDA approval of carotenoid from microalgal source in

cosmetic/pharmaceutical applications turns the search for a safe process to produce them an interesting

topic for future work.

The carotenoid content depends on the efficiency of the extraction procedure. In this case, the

main focus was the cell disruption method with 4 different glass beads size being tested. Results show

that smaller and medium glass beads have a high efficiency in cell disrupting which leads to a high

carotenoid content, on the contrary, big glass beads extraction procedure, is more time consuming,

needing more time to attain the same level of extraction, as the other methods.

Cell disruption strongly depends on microalgae species, age of the culture and composition of

cell wall, therefore, further improvement of the extraction procedure should focus on other approaches

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such as different physical disruption methods as well as other solvent mixtures that could obtain a higher

carotenoid content.

A fractionation procedure was developed to separate astaxanthin and canthaxanthin from other

pigments and intracellular compounds that are also extracted. Six different approaches were proposed

and tested, all based in polarity affinity and dissolution of carotenoids by different solvents. Results of

extraction yields and carotenoid content were distinct, with some solvents having high extraction rates

but low carotenoid content, as the case of methanol or chloroform.

Others solvents such as hexane, proved its incapacity of extracting metabolites from

Chromochloris sp. performing poorly in terms of extraction rate and carotenoid content. The study also

allowed the discovery of new solvent/solvent mixtures that obtained higher extraction rates and

carotenoid content. These solvents are potential agents to be used in a new extraction procedure with

a comparison with the standard method.

One limitation encountered was the inability to isolate astaxanthin from canthaxanthin, with

acetonitrile being the only solvent to show some evidence of a moderate separation of astaxanthin from

canthaxanthin. The high similarity of the structure of both molecules turns the separation difficult, with

the solvent affinity of both carotenoids also being similar.

Despite the limitations, this was the first step in the development of a fractionation procedure

that helped understanding the affinity of astaxanthin and canthaxanthin for different solvents and

indicated potential solvents/solvent mixtures to be tested in the optimization of the fractionation

procedure.

Future research should first focus on the analysis of replicates to validate the results obtained

in this project and on examining other solvents/solvent mixtures, other ratios of the solvent mixtures

tested, as well as other approaches of a fractionation process especially when the goal is to isolate

astaxanthin from canthaxanthin.

An HPLC method was developed to identify and quantify high valuable pigments in extracts.

The first approach was not successful with results showing problems on the separation of astaxanthin

and overpressure problems leading to an extension of the running time, which was not ideal. Moreover,

the inability of running lower concentrations of the standards and the advanced age of the column lead

to the decision of purchasing a new column. In the end of this project, the method developed could

identify two high valuable pigments: canthaxanthin and astaxanthin, and quantify canthaxanthin and one

astaxanthin di-ester.

The successful development of a general method that allows the identification of canthaxanthin

and astaxanthin simultaneously, lead to the development of two specific methods that allowed the

identification of only one pigment. These methods allow the rapid identification of the pigment and are

ideal in cases where only one of the pigments is of interest. By the end of this internship, the specific

method of canthaxanthin was developed and allowed the identification of canthaxanthin in a running

time of five minutes. The time limitation did not allow the quantification step of this method which is

desirable for future work. The development of astaxanthin specific method failed, as the runtime of the

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method was not optimized compared to the general method. Future work should explore other mobile

phases that would allow a rapid elution of astaxanthin standard maintaining the peak separation.

The general method developed in this project should be used in the future since it is able to

identify and quantify canthaxanthin and astaxanthin and aids the understanding of the different

conditions in the synthesis of both pigments. Future work should focus on increasing the sensitivity of

this method, optimizing the limit of detection and quantification and analyze the difference of carotenoid

content in different concentrations of the extract.

Recommendations for a close future include the purchase of standards of other carotenoids,

which will allow a broadening of the variety of high valuable pigments identified and quantified such as

chlorophyll, lutein, zeaxanthin, β-carotene, etc. Also, the purchase of other type of astaxanthin standard

that allows the evaluation of the presence or absence of astaxanthin free form in the extracts. On a long-

term vision, the purchase of a method, which will allow the proper identification of all the astaxanthin

esters found in the extracts, such as a mass spectrometry technique should be considered.

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6. Annex

6.1 Bold Basal Media Composition

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6.2 LPS Assay Procedure

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6.3 Extraction SOP

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6.4 Chromatogram of Haematococcus pluvialis and Hordeum vulgare samples

Figure 6. 1 Chromatograms obtained after the analysis of samples of Hordeum vulgare and Haematococcus pluvialis by RP-HPLC. The analysis was performed by injecting 5 µL of sample in a C18 column while the flow was kept at 1 mL/min during washing, injection and elution and the temperate at 30ºC. The chromatograms were obtained by Innsbruck Botanical Institute.