Visualization and Quantification of Lipid and Protein in Single Microalgal Cell

United Arab Emirates University Scholarworks@UAEU

Electrical Engineering Theses Electrical Engineering

4-2018

VISUALIZATION AND QUANTIFICATION OF LIPID AND PROTEIN IN SINGLE MICROALGAL CELL

Hina Laghari Mohammad Shakeel

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Recommended Citation Shakeel, Hina Laghari Mohammad, "VISUALIZATION AND QUANTIFICATION OF LIPID AND PROTEIN IN SINGLE MICROALGAL CELL" (2018). Electrical Engineering Theses. 13. https://scholarworks.uaeu.ac.ae/electric_theses/13

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Abstract

Microalgae have the potential to be used as a main source of renewable energy in the future due to their lipid contents. Microalgae are comprised of several components including carbohydrates, lipids and proteins. This lipid and protein composition needs to be quantified so the amount of usable energy can be measured. Therefore, the main objective of this thesis is to develop an image processing technique for the quantification of lipid and protein contents in single microalgal cell.

This image processing technique has made lipid quantification an easier process as compared to previous methods used to quantify lipid content. The technique is based on staining microalgal cells with several dyes for the visualization of the lipid and protein content in a microalgal cell. The quantification is then carried out by applying image processing methods. To enhance the lipid accumulation and simultaneously reduce the protein, these microalgal cells are put under stress conditions such as nitrogen starvation. The nitrogen starvation stress condition results in alteration of the lipid and protein content inside the cells. This will significantly influence the ex vivo optimization of microalgal growth conditions for enhanced oil productivity. The image processing-based approach is then used to analyze the time course of lipid accumulation and protein reduction patterns inside the corresponding-stained microalgal cells. There are many different types of strains of microalgae. In this study, Nannochloropsis microalgal strain was used and cultivated in a nitrogen deficient medium. The intracellular Nannochloropsis cell content was estimated by mathematically evaluating the lipid and protein volume inside the corresponding-stained cell. This mathematical evaluation was made vii possible with the image processing technique that has been developed in this thesis.

This image processing approach allows the quick characterization of microalgal cells. It also takes precedence over any techniques or methods that have been developed in the past to quantify lipid content of microalgal cells. This will be explored further in the subsequent studies.

Keywords: Lipid, Microalgal cell, Protein, Quantification, Visualization. viii

Title and Abstract (in Arabic)

ﺗﺤﺪﻳﺪ ﻣﻜﻮﻧﺎﺕ ﺍﻟﻬﻦ ﻭ ﺍﻟﺒﺮﻭﺗﻴﻦ ﺑﺪﺍﺧﻞ ﺧﻠﻴﺔ ﻃﺤﻠﺒﻴﺔ ﺑﺎﺳﺘﺨﺪﺍﻡ ﺗﻘﻨﻴﺔ ﺍﻟﺘﺼﻮﻳﺮ

ﺍﻟﻤﻠﺨﺺ

ﺍﻟﻄﺤﺎﻟﺐ ﺍﻟﺪﻗﻴﻘﺔ ﺑﺸﻜﻞ ﻋﺎﻡ ﻫﻲ ﻭﺍﺣﺪﺓ ﻣﻦ ﺍﻫﻢ ﺍﻟﻤﺼﺎﺩﺭ ﺍﻟﺮﺋﻴﺴﻴﺔ ﺍﻟﻤﺜﻴﺮﺓ ﻟﻼﻫﺘﻤﺎﻡ ﻓﻲ

ﻣﺠﺎﻝ ﺍﻟﻄﺎﻗﺔ ﺍﻟﻤﺘﺠﺪﺩﺓ ﺍﻟﺨﻀﺮﺍء. ﻭﺑﺎﻟﺘﺎﻟﻲ ﻣﻦ ﺍﻟﻤﻬﻢ ﻭ ﺍﻟﻤﺘﻮﻗﻊ ﺃﻥ ﻳﻜﻮﻥ ﻟﺪﻳﻨﺎ ﺍﻟﻘﺪﺭﺓ ﻋﻠﻰ ﻓﻬﻢ

ﺩﻭﺭﻣﺤﺘﻮﻳﺎﺗﻬﺎ ﻓﻲ ﻋﻠﻢ ﻭﻇﺎﺋﻒ ﺍﻟﺨﻠﻴﺔ. ﻭﻟﺬﻟﻚ، ﻓﺈﻥ ﺍﻟﻬﺪﻑ ﺍﻟﺮﺋﻴﺴﻲ ﻣﻦ ﻫﺬﻩ ﺍﻷﻃﺮﻭﺣﺔ ﻫﻮ ﺗﻄﻮﻳﺮ

ﺗﻘﻨﻴﺔ ﻣﺒﻨﻴﺔ ﻋﻠﻰ ﻣﻌﺎﻟﺠﺔ ﺍﻟﺼﻮﺭ ﻟﺘﻘﺪﻳﺮ ﻣﺤﺘﻮﻳﺎﺕ ﺍﻟﺪﻫﻦ ﻭﺍﻟﺒﺮﻭﺗﻴﻦ ﻋﻠﻰ ﻣﺴﺘﻮﻯ ﺧﻠﻴﺔ ﻭﺍﺣﺪﺓ.

ﻭﻳﺴﺘﻨﺪ ﻫﺬﺍ ﺍﻷﺳﻠﻮﺏ ﺍﻟﺤﺎﻟﻲ ﻋﻠﻰ ﺗﻠﻮﻳﻦ ﺧﻼﻳﺎ ﺍﻟﻄﺤﻠﺒﻴﺔ ﺑﻤﻮﺍﺩ ﻣﺸﻌﺔ ﺧﺎﺻﺔ ﻟﺘﻘﺪﻳﺮﻧﺴﺒﺔ

ﺍﻟﺪﻫﻮﻥ ﻭﺍﻟﺒﺮﻭﺗﻴﻦ. ﺛﻢ ﻳﺘﻢ ﺍﻟﻘﻴﺎﺱ ﺍﻟﻜﻤﻲ ﻭ ﺍﻟﻨﺴﺒﻲ ﻣﻦ ﺧﻼﻝ ﻃﺮﻕ ﻣﻌﺎﻟﺠﺔ ﺍﻟﺼﻮﺭ ﺍﻟﻤﻠﺘﻘﻄﺔ ﻣﻦ

ﻣﺮﺑﻊ ﺍﻟﻤﺰﺭﻋﺔ ﻓﻲ ﺃﻳﺎﻡ ﻣﺨﺘﻠﻔﺔ ﻣﻦ ﺗﺤﺖ ﺣﺎﻟﺔ ﺍﻹﺟﻬﺎﺩ ﺍﻟﻨﻴﺘﺮﻭﺟﻴﻦ. ﺣﺎﻟﺔ ﺍﻹﺟﻬﺎﺩ ﻫﺪﻩ ﺗﻠﻌﺐ ﺩﻭﺭ

ﻛﺒﻴﺮ ﻓﻲ ﺗﻐﻴﻴﺮ ﻣﺤﺘﻮﻯ ﺍﻟﺪﻫﻮﻥ ﻭﺍﻟﺒﺮﻭﺗﻴﻦ ﺩﺍﺧﻞ ﺍﻟﺨﻼﻳﺎ.

ﺍﻥ ﺇﻣﻜﺎﻧﺎﺕ ﺍﻟﻨﻬﺞ ﺍﻟﺘﺼﻮﺭ ﺍﻟﺤﺎﻟﻲ ﻳﺴﻤﺢ ﺗﻘﺪﻳﺮ ﻓﻲ ﺧﻠﻴﺔ ﺣﻴﺔ ﻭﻫﺬﺍ ﺳﻮﻑ ﻳﺆﺛﺮ ﺑﺸﻜﻞ

ﻛﺒﻴﺮ ﻋﻠﻰ ﺗﺤﺴﻴﻦ ﻇﺮﻭﻑ ﺍﻟﻨﻤﻮ ﺍﻟﺠﺰﺋﻲ ﻟﺘﺤﺴﻴﻦ ﺇﻧﺘﺎﺟﻴﺔ ﺍﻟﺪﻫﻮﻥ ﻛﻤﺼﺪﺭ ﺑﺪﻳﻞ ﻟﻠﻄﺎﻗﺔ.

ﻣﻔﺎﻫﻴﻢ ﺍﻟﺒﺤﺚ ﺍﻟﺮﺋﻴﺴﻴﺔ: ﻃﺤﺎﻟﺐ.ﺍﻟﺪﻫﻮﻥ، ﺍﻟﺨﻠﻴﺔﺍﻟﻄﺤﻠﺒﻴﺔ، ﺍﻟﺒﺮﻭﺗﻴﻦ، ﺍﻟﺘﺤﺪﻳﺪ ﺍﻟﻜﻤﻲ، ﺗﻘﻨﻴﺔ

ﺍﻟﺘﺼﻮﻳﺮ.

Acknowledgements

First and foremost, I would like to thank my thesis advisor and supervisor

Dr. Mahmoud F. Al Ahmad of the Electrical Engineering Department at United Arab

Emirates University. Without his consistent support and help, I most certainly could not have successfully conducted my thesis. I am very grateful for his endless patience towards me!

I would also like to thank Mrs. Shaima Raji and Engineer Nisreen Al Naji for their enthusiastic input and help throughout the researching process.

Finally, I must express my gratitude to my family (Dr. Mohammad Shakeel

Laghari, Anila Shakeel, Anum Laghari, Saad Laghari), including my cats (Coco

Pixie, Nala, Luna, Gracie, Chottu) and especially to Pippin for providing me with constant support and encouragement throughout my years of studying engineering and throughout the process of researching and writing this thesis. This accomplishment would not have been possible without them and my supervisor

Dr. Mahmoud F. Al Ahmad. Thank you.

Dedication

To my treasured family and cats xi

Table of Contents

Title ...... i Declaration of Original Work ...... ii Copyright ...... iii Approval of the Master Thesis ...... iv Abstract ...... vi Title and Abstract (in Arabic) ...... viii Acknowledgements ...... ix Dedication ...... x Table of Contents ...... xi List of Tables...... xiii List of Figures ...... xiv List of Abbreviations...... xv Chapter 1: Introduction ...... 1 1.1 Aim………………………………………………………………………..1 1.2 Motivation ...... 1 1.3 Research Objectives ...... 2 1.4 Thesis Significance ...... 3 1.5 Thesis Organization ...... 3 Chapter 2: Literature Review ...... 5 2.1 Microalgae Biomass as an Energy Source ...... 5 2.2 Conventional Methods for Extraction of Biomolecules...... 9 2.3 Image Processing by Fluorescent Microscopy ...... 13 Chapter 3: Materials and Methods ...... 18 3.1 Microalgae Cultivation...... 18 3.1.1 Growth under Controlled Conditions ...... 19 3.1.2 Determination of the Cell Concentration ...... 19 3.2 Microalgae Fluorescent Staining...... 20 3.2.1 Protein Visualization by Fluorescent Microscopy ...... 20 3.2.2 Lipid Visualization by Fluorescent Microscopy ...... 21 3.3 Image Processing Tools ...... 22 Chapter 4: Results and Discussion ...... 24 4.1 Visualization of Lipids, Protein Islands in Microalgal Cells ...... 24 4.1.1 Lipid, Protein Determination Using Biochemistry Method ...... 25 4.1.2 Statistical Analysis ...... 27 xii

4.2 Lipid, Protein Area Determination from Image Processing...... 29 4.3 Discussion ...... 31 Chapter 5: Conclusions and Future Work ...... 36 5.1 Conclusions ...... 36 5.2 Future Work ...... 37 References ...... 40

xiii

List of Tables

Table 1: Several extracted parameters per day using Aphelion Dev ...... 30

xiv

List of Figures

Figure 1: Microscopic picture of a single microalgal cell ...... 2 Figure 2: Microalgal cultivation tank ...... 7 Figure 3: Microalgae to biofuel conversion process ...... 8 Figure 4: Microalgae composition (a) Single cell showing carbohydrate, pigments, lipid and protein (b) Schematic showing percentage contribution of the composition in a general algae biomass ...... 10 Figure 5: Fluorescence microscopy images of Scenedesmus sp. cells stained with Nile Red and showing lipid accumulation after 1, 11, 14, 20 and 23 days of nitrogen starvation in cultures ...... 14 Figure 6: Microalgal cultivation process (a) First day (b) First day of starvation (c) Last day of starvation...... 18 Figure 7: Cell count over days of growing...... 20 Figure 8: Microscopic picture for (a) Protein staining (b) Lipid staining...... 21 Figure 9: Aphelion Dev program screenshot ...... 23 Figure 10: Captured fluorescence microscopy pictures over the growth period ...... 25 Figure 11: Extracted lipid and protein content using conventional biochemical methods and techniques...... 27 Figure 12: Histogram for protein versus level for several starvation days...... 28 Figure 13: Image processing and biochemical extraction results...... 31 Figure 14: Proposed in vivo system for real time measurements of cell content using thermal camera and electrical conditions...... 37

List of Abbreviations

CO2 Carbon Dioxide

DAPI 4',6-diamidino-2-phenylindole

DMSO Dimethyl Sulfoxide

DNA Deoxyribonucleic Acid

FITC Fluorescein Isothiocyanate

FTIR Fourier-transform Infrared Spectroscopy

GC Gas Chromatography

HCL Hydrochloric Acid

HEPES Organic Chemical Buffering Agent

HPLC High Pressure Liquid Chromatography

LC Liquid Chromatography

N2 Nitrogen

Nannochloropsis sp. Nannochloropsis Species

NIR Near Infrared Spectroscopy

NR Nile Red

SNR Signal-to-Noise Ratio

NaCl Sodium Chloride (Salt) xvi

NaNO3 Sodium Nitrate

NaOH Sodium Hydroxide

TAG Triacyl Glycerol

TCA Trichloroacetic Acid

Chapter 1: Introduction

1.116B Aim

The aim of this thesis is to develop an image processing based technique to quantify the amount of lipid and protein inside a single living microalgal cell. The quantification is based on estimating the volume of lipid and protein inside stained cells.

1.217B Motivation

Microalgae are one of the most promising sources for biodiesel production

[1]. A microscopic image of a single microalgal cell is shown in Figure 1.

Microalgal biodiesel production requires high lipid productivity that demands

continuous enhancement for biomass growth rate and lipid content [2], [3]. To do

so effectively, a continuous monitoring method at the level of single cell growth is

highly desired. Mainly such a method should be able to monitor and quantify the

amount of lipids and proteins inside living cells. Unfortunately, the existing

conventional cell bio-composition determination methods require the involvement

of several invasive steps such as Soxhlet [4] and Bligh & Dyer methods [5].

Drying the biomass based method includes disruption of the cell walls, for the

extraction of the cell components [2], [6], [7], [8], [9].

To avoid such invasive processes and develop faster and more accurate non-

invasive techniques, the use of Raman spectroscopy for the determination of lipid

content has been proposed [7]. Furthermore, an NMR based approach has been

developed and has been utilized to extract the lipid content [8]. In line with these 2

efforts, a radio frequency-(RF)-based novel technique has also been developed to

determine the oil content [2]. However, all the previous mentioned methods have

been carried out on bulky suspensions with predefined cells count.

Figure 1: Microscopic picture of a single microalgal cell

1.3 Research Objectives

The main research objective of this thesis is the contribution to the development of noninvasive techniques and methods for quantifying the composition inside a single microalgal cell. Therefore the main research objectives of this thesis are:

1) Develop an image processing algorithm for microalgae lipid and protein single

cell quantification.

2) Validate the results with conventional methods.

3) Compare the efficiency and sensitivity of the image based approach with the

other proposed methods in this domain. 3

1.4 Thesis Significance

It is currently accepted that biodiesel is a great alternative to petroleum fuel.

The reason being that biodiesel is proven to reduce dangerous emissions and thus it will be more environmentally friendly and safer for people. Since microalgae is a favorable source of oil for biodiesel production and has been utilized worldwide, this research topic is extremely important and could be beneficial for today’s society. It could benefit the world in countless ways. Another factor to consider is that the

U.A.E. is an ideal environment for microalgae cultivation. This is due to the fact that the U.A.E. receives abundant sunshine and it has readily available saltwater. This is an ideal situation for microalgae cultivation. Thus, this research will allow users to produce microalgae on a massive scale and estimate the oil content with the help of image processing. This type of oil estimation method could be very beneficial if it proves to be accurate and reduces the time and effort required to measure oil content of a single microalgal cell.

1.5 Thesis Organization

The thesis is organized as follows:

Chapter 1 (Introduction): This chapter presents the motivation and objectives of this work. It starts with explaining the demand for lipid from microalgae and the technologies used to extract it. The chapter ends by stating the objective and the significance of this research.

Chapter 2 (Literature Review): This chapter reviews the previous research work related to the topic of the thesis. It discusses the different techniques used to 4 determine lipid and protein volume content. It also presents the theory of characterization technique proposed in this work.

Chapter 3 (Materials and Methods): This chapter demonstrates the materials and tools used, in addition to the methodology and experimental procedures conducted in order to perform the desired measurements and analysis.

Chapter 4 (Results and Discussion): This chapter presents the results gained from performing the experiments, the processing and analysis of data, followed by a discussion of the results and the validation using a different technique.

Chapter 5 (Conclusions and Future Works): This chapter is the last one in the thesis that wraps up all the research findings, and relates them to the objectives presented in the first chapter.

Chapter 2: Literature Review

This chapter presents an overview of the use of microalgae as an energy source, and then it gives an overview of the conventional methods used for lipid and protein content determination. Furthermore, it summarizes the utility of image processing techniques for microalgae visualization and characterization.

2.1 Microalgae Biomass as an Energy Source

Microalgae are essentially microscopic algae typically found in freshwater and marine systems, mostly unicellular and can exist individually or in groups.

According to the species, their sizes differ in range from a few micrometers to a few hundred micrometers. They are capable of performing photosynthesis and produce just about half of the atmospheric oxygen and simultaneously consume carbon dioxide. Microalgae are found at the base of the food web, which highlights their importance on earth, and provide energy for all the trophic levels above them.

Microalgae are attaining increasing attention due to their wide range of applications in biology and biotechnology industries [10]. The demands of microalgae as a biodiesel source are on a leap in the modern era due to the increasing demand for fuels and environmental safety [11]. The main features that make the biofuels a superior alternative to fossil fuels are that they are renewable sources of energy, environmentally friendly and are more economic [12]. They possess high biomass productivity, speedy lipid accumulation and can survive in saline water.

Biofuels are expected to take the place of fossil fuels in the future due to the demand for clean energy, economic potential and renewable nature [13]. 6

In addition, microalgae are rich sources of carbohydrates, lipids and proteins, which make them important sources of feedstock [14]. The presence of long chain fatty acids, proteins and carbohydrates makes them important as components of health food supplements as well as various pharmaceutical industries [10].

Oleaginous microalgae like the Chlorella, Nannochloropsis, and

Scenedesmus species are particularly important sources of biofuels as well as feedstock due to the high levels of proteins and polyunsaturated fatty acids [15], [16],

[20]. The biomolecule components can be enhanced by manipulating the various parameters by subjecting them to stress, both genetic and environmental such as light, temperature, nutrient depletion like carbon or nitrogen, light intensity, salinity or harvesting procedure [17], [18], [19].

The high biomass growth rate coupled with high nutrient content is difficult to achieve simultaneously which can be achieved by the advancement of molecular biology and genetic engineering techniques [21], [22], [23]. Engineering the growth medium, proper selection of the algal strains, use of genetic engineering and use of suitable extraction methods can optimize nutrient content [24], [25], [26]. The studies involving continuous monitoring of the food, biofuel and other co-product production requires continuous monitoring of the biochemical components [18].

This work is aimed to study the increase in lipid content of microalgae by manipulating the nitrogen content of the medium in which it is cultured. The studies involving the cultivation of microalgae rich in lipids for biodiesel production and as valuable nutrient rich products for feedstock is on the increase in the recent times

[27]. Under conditions of stress, such as nitrogen depletion, studies have shown that a substantial increase in the fatty acid content, especially the triacyl glycerols (TAGs) 7 of the cells has been documented [28]. The fatty acid biosynthesis has a high energy requirement, which is mostly supported by photosynthetic process and also by the degradation of the carbohydrates that has accumulated during the nitrogen sufficient conditions [29]. An image of microalgal cells in a cultivation tank is shown in

Figure 2.

Figure 2: Microalgal Cultivation Tank

The nitrogen depletion on the other hand, is seen to affect the protein and carbohydrate contents in the microalgae [30]. It has been found that the growth of the algal biomass reduced sufficiently while the lipid content escalated considerably during nitrogen depletion [31]. The depletion of NaNO3, which is the nitrogen source for the microalgal culture, reduces the rate of protein biosynthetic pathways [32].

The biosynthesis of carbohydrates is affected by nitrogen depletion, which results in a breakdown of the carbohydrates rooting for the lipid biosynthetic pathway [33].

Pigments like lutein and astaxanthin accumulation was noticed in several microalgal species [34]. 8

Proteins make up a large portion of the cellular membrane and are important components of a number of enzyme systems that carry out the various cellular activities [35]. Some microalgal species like Nannochloropsis sp, Dunaliela sp. and

Chlorella sp. are able to accumulate 25 - 60% of lipids/dry weight [36]. Nitrogen depletion was used as the stress signal to induce lipid accumulation in

Nannochloropsis oculata for this study [42]. The Nannochloropsis species was found to contain a high oil content, around 33.3 - 37.8%, of dry cell weight [37].

Nannochloropsis is a motile, unicellular, green microalga with a simple cell morphology and diameter of 2 – 8 µm [38], [39]. They are found to have a high growth rate, ability to accumulate high amounts of lipids and a higher resistance to contamination [39]. The process steps of converting microalgae to biofuel is shown in Figure 3.

Figure 3: Microalgae to Biofuel Conversion Process

As the cell division slows down, the production of new cellular components is not required and the cells in turn will be involved in the formation of TAG (triacyl glycerols), which is a protection mechanism against the stress caused by nitrogen depletion [40]. This is mainly because the lipid-synthesizing enzymes are less susceptible to degradation than other enzymes involved in the biosynthetic pathway 9

[40]. It has also been reported that lipid accumulation can be attributed to the fact that under N2-depletion the cells continue to produce lipids and carbohydrates but affects protein and nucleic acid synthesis [41].

The biochemical methods commonly used for the extraction of proteins, carbohydrates and lipids use strong alkali, acid and organic solvents [25]. The microalgal cells have strong cell walls and the alkali treatment used for protein extraction lyses the cell walls [26]. The ethanol precipitation helps the carbohydrates to be extracted completely [25]. Several methods designed for lipid extraction turned out to be less satisfactory as there was loss or incomplete extraction of lipids and some of them were time consuming [43]. The Bligh and Dyer method used for this study gave optimum results [43]. Hemocytometer and spectrophotometer was used to determine the cell concentration [44].

2.2 Conventional Methods for Extraction of Biomolecules

Microalgal cells are made up of biomolecule components such as carbohydrates, lipids, proteins, nucleic acids and vitamins. The major biomolecule components of microalgal cells are shown in Figure 4. A vast majority of the conventional techniques that are used to determine the biomolecule components of microalgae are labor intensive and time consuming. These procedures make use of various methods and reagents that are disparaging and hazardous to health as well as the environment.

Therefore, the accurate, continuous and rapid detection of the amounts of carbohydrates, lipids and proteins in microalgal cells are extremely crucial and highly demanded while engineering the microalgae for the production of distinctive type of biofuels [50]. Several concerns arise while extracting the biomolecules from the microalgae, mostly due to the presence of resilient cell walls in a large number of 10 the species [51]. A substantial impedance encountered during the extraction of biomolecules from microalgae is the disintegration of the strong cell walls and extraction of the biomolecules both of which are rather laborious and time- consuming [52]. Most methods use dry biomass as the starting material for the extraction techniques of carbohydrates and lipids. This can avoid the inaccuracies that are subject to manifest due to the liquid carry over in these methods.

a) Microalgae Composition

Figure 4: Microalgae Composition (a) Single cell showing carbohydrate (CARBS), lipid and protein (b) Schematic showing the percentage contribution of the composition in a general algae biomass. 11

. Microalgal protein extraction is still in its infancy, which means that large- scale research is required to develop methods that provide optimum results [53].

Protein extraction and estimation are conventionally done using chromatographic techniques like High Performance Liquid Chromatography (HPLC) and Liquid

Chromatography (LC), which uses freeze-dried samples of microalgae [54]. Despite the fact that these techniques are time-consuming and labor-intensive, they present meticulous results. Alternatively, other methods that are largely sought after comprises atomic absorption spectrophotometry and mass spectrometry which provides information in regards to the amino acid levels present in the proteins. The

Dumas-based combustive method is largely implemented in the determination of the

Nitrogen content in organic samples. The renowned Kjeldahl method determines the protein content, mostly in organic contents by titration methods [55]. Dye-based procedures such as: Bradford, Biuret, Lowry and Microbiuret methods do provide high throughput procedures because of the use of colorimetry/spectrophotometry techniques but requires additional processing time of the fresh samples to release the protein content [56]. Pre-treatments involving chemical or physical methods are necessary for the complete release of proteins [57]. Heat extraction using strong alkali is a highly effective technique for the extraction of proteins but is compatible only with reduced number of microalgal strains [58].

Chromatographic techniques like Gas Chromatography (GC) are high throughput methods used in the quantitation of sugars and carbohydrates present in microalgae. Sample preparation time is sufficiently low for Liquid chromatography

(LC) compared to Gas Chromatography (GC) but affects the resolution of simple sugars [58]. Some of the commonly used methods that provide promising outputs for 12 the quantitation of sugars present in microalgae include phenol-sulphuric acid method or anthrone method [59]. The carbohydrates/sugars present in the sample get reduced to colored compounds called furans. The amounts of carbohydrates are estimated by measuring the color intensity of the furans/furfurals spectrophotometrically [58]. Once again, the presence of rigid cell walls makes the extraction of carbohydrates a time demanding method, which calls for the need of pre-treatment (chemical, enzymatic or physical) procedures like microwave-assisted, hot alkali, or ultrasonic-assisted extraction [60].

Numerous non-traditional, highly expensive and labor-intensive techniques are employed in the extraction of oils from microalgae [61]. Many of the lipid extraction and estimation techniques involve the use of organic and highly volatile solvents like butanol, chloroform, methanol etc. and methods like electroporation, supercritical CO2 or ultrasonic methods [62]. Conventional and highly acclaimed methods like the Folch method and the Bligh and Dyer methods are promising but use highly volatile solvents like chloroform and methanol for the total lipid extraction [63]. The lipids that get released into the chloroform layer are separated by various methods. Several mechanical methods like bead-beating, homogenization, sonication etc. are also used for the lipid extraction followed by gravimetric methods of measurement [64]. The most novel and recent advancements for the estimation of total lipids uses techniques like Raman spectroscopy as well as the measurement of fluorescence intensity using fluorescent dyes like Nile Red or BODIPY, where a standard curve correlating fluorescence intensity to oil content is constructed [49],

[65]. 13

2.3 Image Processing by Fluorescent Microscopy

The labor-intensive, time-consuming methods available currently for the determination of microalgal biomolecules such as lipids and proteins have highlighted the need for the development of more approachable methods for monitoring the biomolecules. Several quantification methods have been introduced for detection and monitoring of biomolecules that combines several traditional methods [66]. In the most recent times, the most suitable method is still the staining method that has modification superiority and great value for application [67]. The fluorescence microscopy can be used to visualize the various biomolecules (lipids, proteins, nucleic acids) in cells using appropriate staining dyes and techniques.

Fluorescence imaging relies on the illumination of fluorescently labeled biomolecules present in cells [68]. The fluorescence images can be acquired using an optical microscope equipped with a transmission and epi-fluorescence illumination sources [69]. The illumination source produces a defined wavelength of light considerably near to the excitation wavelength of the fluorophore. This causes the emission of a light of a longer wavelength that causes the fluorophores to fluoresce and enable visualization of biomolecules of interest [70]. A major challenge in confocal microscopy is the determination of the wavelength to cause the excitation of the fluorophores without causing photo bleaching [71]. Each time a fluorescently- labelled sample is illuminated, a portion of the fluorophores will be subjected to photo damage irreversibly [71]. An example of microalgal cells stained with Nile

Red, which illuminate lipid biomolecules fluorescently, is shown in Figure 5. 14

Figure 5: Fluorescence microscopy images of Scenedesmus sp. cells stained with Nile Red, showing lipid accumulation after 1, 11, 14, 20 and 23 days of nitrogen starvation

The filters for the excitation and emission wavelengths should be conveniently chosen to match each fluorophore to prevent excessive damage caused due to photo bleaching [72]. The magnification of the sample increases the irradiance significantly. An effective way to reduce photo bleaching and photo damage is by reducing the exposure time of the sample to the excitation light [73]. At the same time, it is compulsory to retain the signal-to-noise (SNR) ratio for the specific sample

[71]. The use of appropriate filters (blue, green and red) with appropriate wavelengths of the fluorophores used to stain the samples promises ample information using fluorescence microscopy.

Sample processing for fluorescence microscopy is an important step in fluorescence microscopy [74]. The samples, which are mostly cells or tissues, must be subjected to a number of pre-treatment processes for the effective staining of the cells. Some of the necessary steps include fixation, permeabilization, staining, washing, counterstaining if necessary, mounting, visualization and storage depending on the application [74].

Fixation of the cells is done by aldehydes or alcohols to maintain the cellular structure. Aldehydes like formaldehydes/paraformaldehydes are commonly used to cross-link the proteins whereas acetone and methanol allow the permeabilization of 15 the stains through the cell walls and membranes [75]. The stains like FITC used for protein staining are resuspended in acetone for the effective staining of amino acid units [76]. Washing is an important procedure between each step to remove the excess or unbound reagents. Physiological pH buffers or double distilled water is usually used for the washing steps [77]. Counter staining, using stains like those that

DAPI is commonly done while staining the proteins. Counterstains like DAPI,

Evan’s blue stains the nucleic acids, thus reducing the background fluorescence [78].

The final steps include mounting and storage of the slides [74].

The presence of thick and rigid cell walls in various species and strains of microalgae have made the development of standardized protocol for fluorescence microscopy quite difficult [79]. Therefore, the development and use of a common stain or staining protocol in staining the various strains is an almost impossible juncture. The biomolecule contents in the cells fluoresce when excited fluorescently.

This requires staining the biomolecules of interest with specific dyes and stains along with the suitable treatment procedures. Each strain requires specific physical and chemical treatment methods for the effective and efficient staining of the different biomolecules [80]. The treatment methods enable the fluorescent stains to penetrate the otherwise rigid cell wall without affecting the morphology or cellular make up

[79]. Chemical treatment of the cells with reagents such as DMSO, ethanol, acetone, depending on the strains, facilitates the uptake of different dyes by the cells as well as maintaining the cell structure at the same time [81].

Nitrogen deprivation, which is a stress condition, causes an increase in the lipid accumulation while the biomass and cell growth diminished [82]. The

Nannochloropsis sp. when subjected to Nitrogen starvation by decreasing the amount 16 of nitrogen content in the medium, showed an increase in the lipid content while the protein content reduced.

The rapid and simple measurement of neutral lipids is highly desired while studying the lipid metabolism. Nile red (9-diethylamino-5Hbenzo[a]phenoxazine-5- one), is a high throughput, intensely fluorescent dye under appropriate spectral ranges in most species of microalgae for visualizing the cytoplasmic lipid droplets, especially TAGs (triacyl glycerols) [83]. Nile red is scarcely soluble in water but highly soluble in organic solvents like acetone, ethanol, methanol, DMSO or xylene, which can be chosen depending on the application and the sample under test [84].

The Nile red stained lipid droplets can be observed under a fluorescent microscope with 510-640 nanometer exciter filters [84]. The fluorescent colors produced by Nile red ranges from golden yellow to deep red [85]. The Nannochloropsis samples were stained with Nile red dissolved in acetone and viewed for yellow gold and red fluorescence. The Nile red stained lipid droplets were seen as small distinct bodies scattered throughout the cytoplasm. Nile red can be considered as a general stain for lipids since it can stain phospholipids, cholesterols, cholesteryl esters, in addition to triacyl glycerols (TAGs) [86].

Bodipy 505/515 (4,4-difluoro-1,3,5,7-tetramethyl-4-bora-3a,4adiaza-s- indacene) is presently being used as an effective alternative to Nile Red [87]. They are widely used to stain lipid vesicles in microalgae. They are found to be more specific since it does not bind to cellular compartments other than lipids and chloroplasts [88]. A large number of solvents are used for resuspension of the

BODIPY 505/515 such as acetone, methanol, ethanol, DMSO depending on the application [85]. Lipid bodies are stained yellow to green and the chlorophyll bodies 17 appear red. The incorporation of BODIPY 505/515 into the cells is relatively fast since it can cross the cell and organelle membranes easily [89].

FITC (fluoresceinisothiocyanate isomer I) is used to stain the amino groups in the microalgal cells. The staining method involves rapid conjugation of proteins present in the microalgal cells with FITC [90]. A stable link forms between the protein and FITC while the non-protein components of the cells remain unstained

[91]. The FITC is made into a solution form using ethanol. The microalgal cells are fixed using formaldehyde to maintain the original structure of the cells and provide optimum staining [18]. A fluorochrome, DAPI (4’, 6 – diamino-2-phenylindole

2HCl) is used to counterstain the nucleic acids/DNA [91]. The nucleic acid contents emit a blue color under confocal microscope.

Chapter 3: Materials and Methods

The microalgal materials cultivation involved in this thesis experimental work along with the adopted methods for lipid and protein staining are summarized.

In addition to that, the utilized Dev. Aphelion software tools are also described and detailed.

3.1 Microalgae Cultivation

To obtain microalgal cells with different lipid and protein contents, the experiments are carried out using Nannochloropsis oculata species. The microalgae are cultivated locally in a sterile culture vessel of 10 L in Guillard F2 medium [45].

This sterile culture vessel is shown in Figure 6.

Figure 6: Microalgal Cultivation Process (a) First day (b) First day of starvation (c) Last day of starvation

The basal Guillard F2 medium is prepared using artificial seawater with the salinity adjusted to 3.5% NaCl in deionized water, enriched with 1 milliliter per liter of the following major nutrients (in mM); 0.880 NaNO3, 0.036 NaH2PO4.H2O and 19

0.106 Na2SiO3.9H2O and 1 milliliter per liter of trace metals solution consisting of

3 (10 milliliter); 0.08 ZnSO4.7H2O, 0.9 MnSO4.H2O, 0.03 Na2MoO4.2H2O, 0.05

CoSO4.7H2O, 0.04 CuCl2.2H2O, 11.7 Fe(NH4)2(SO4)2.6H2O and Na2EDTA.2H2O and followed by the addition of 1 milliliter per liter of Vitamins; B12, Biotin and

Thiamine prepared in HEPES at pH 7.4 is used for the growth of the microalgae.

3.1.1 Growth under Controlled Conditions

To alter the content of lipid and protein inside the living microalgal cell, the microalgal culture has been exposed to nitrogen starvation. The culture is started under optimal conditions in nitrogen sufficient medium. The cells were grown with optimum aeration and lighting. Light is provided in profusion, which helps the cells to undergo photosynthesis and operate in a linear growth phase. The microalgae is grown in nitrogen sufficient medium until the optimum number is obtained and then subjected to nitrogen depletion to induce lipid production. The nitrogen source and concentration in the medium itself causes the important changes in growth character and biochemical composition of algae, including oil and fatty acids [28].

3.1.2 Determination of the Cell Concentration

To ensure enough number of cells inside the microalgal culture, the concentration should be determined. The cell concentration is monitored using a UV- spectrophotometer (UV-1800, Schimadzu, Japan) [46]. The concentration is monitored daily at an optical density of 680 nm. The concentration (cells per milliliter) at any given cultivation time is calculated from a previously prepared calibration curve of optical density versus cell concentration determined using a 20

Neubauer Hemocytometer under a microscope. Figure 7 shows the cell count per milliliter throughout the days of growing.

Figure 7: Cell count per milliliter over days of growing

3.2 Microalgae Fluorescent Staining

The protein and lipid staining for visualization is detailed and discussed here.

3.2.1 Protein Visualization by Fluorescent Microscopy

Ten µL of the microalgal sample is immobilized on a slide under sterile conditions for 1 hour at RT and fixed. The fixation of the samples is performed by using 50 μL of 3.7% formaldehyde solution for 45 min. After 45 min at RT, the fixed sample is rinsed three times with sterile tap water and dried under sterile conditions for 30 min at RT. The amino groups in biofilm samples are stained with fluorescein isothiocyanate isomer I (FITC; Sigma-Aldrich, Germany). 30 μL of that working 21 solution is added to the samples for 60 min at RT. Nucleic acids are stained with 4,6- diamidino-2- phenylindoldihydrochloride (DAPI; Merck, Germany) by adding the stain to the microalgal samples for 40 min at RT. All of the procedures are conducted in the dark. The experiments are run in triplicate. After staining, the samples are dried under sterile conditions for 15 min at RT. The dried samples on the diagnostic slides are covered with immersion oil (ImmersolTM 518 F; Carl Zeiss AG,

Germany) and a cover glass. Microscopic analysis was performed using the fluorescence microscope with the appropriate filter blocs. At least 15 digital images are taken for each sample. Examples of these fluorescent digital images are shown in

Figure 8. This information and more details of protein staining can be found in [47].

Figure 8: Microscopic picture for (a) Protein staining (b) Lipid staining

3.2.2 Lipid Visualization by Fluorescent Microscopy

Here the lipid staining using different methods that used in this study are summarized.

3.2.2.1 Nile Red Staining

Microalgal cells were stained by 20-min incubation in the dark at room temperature in a 10 μg per milliliter solution of Nile red (Sigma, Saint Louis, USA). 22

In the presence of neutral lipids, Nile red emits a yellow-gold fluorescence (lmax =

580 nm) while in the presence of polar lipids, Nile red emits an orange-red fluorescence (lmax = 615 nm). Nile red fluorescence was observed using a fluorescent microscope at excitation of (475/40) and (510) optical filters. Images were captured with the Spot Insight 4 software (Diagnostic Instruments Inc., Sterling

Heights, USA). For more details, please check reference [48].

3.2.2.2 BODIPY Staining

Staining method: L g per milliliter of BODIPY in 1% DMSO for 30 min. Set of images were acquired with a fluorescent microscope. Cells were stained with each dye at a concentration of 10 μg per milliliter for 30 min before imaging. The green channel collected emissions between 510 and 560 nm, and the red channel collected emissions between 590 and 660 nm. For furthermore reading, a reference [49] is recommended.

3.3 Image Processing Tools

Aphelion Dev image processing tool has been employed through this thesis work to process the microscopic images. The tool imports the microscopic photos and its settings are adjusted and calibrated to extract the lipid and protein areas with high accuracy inside a single microalgal cell. Aphelion Dev tool also can display the lipid and protein distributions and accumulation patterns inside the living cells. The amount of lipid and protein present inside an individual cell can be estimated using images of their corresponding stained cells and the image processing steps summarized here. 23

To determine the area and height of the corresponding cell, the background of the images shown was first subtracted, which resulted in a clearer image of both lipid and protein bodies/islands. With the assistance of Aphelion Dev, it was possible to estimate the total amount of lipid and protein inside a single microalgal cell. Figure 9 presents a screenshot of the image processing tool that has been utilized for this thesis.

Figure 9: Aphelion Dev program screenshot 24

Chapter 4: Results and Discussion

To demonstrate the concept and working principle, Nannochloropsis sp. microalgae strains was used and to be tested and analyzed in this study. This chapter presents the results obtained from the conducted experiments and discusses them in relation to existing literature.

4.1 Visualization of Lipids and Protein Islands in Microalgal Cells

Set of selected captured fluorescence microscopy pictures over the growth period are displayed in Figure 10. According to the figure captions, the first column pictures (a), (c), (e), (g) and (i) visualize the protein content before starvation, day 4th

, day 10th , day 14th and day 18th after starvation. The second column pictures (b),

(d), (f), (h) and (j) visualize the lipid content before starvation, day 4th , day 10th , day

14th and day 18th after starvation. The selection of those pictures was based on their content that shows clearly the accumulation of a larger amount of lipid inside the cell as the starvation goes longer. Meanwhile the pictures also revealed that the amount of protein is reduced with starvation.

The fluorescence microscopy images of the microalgal cells induced for lipid and protein accumulations and reduction, respectively, were taken for cells in suspension concentration of 4×106 cells/ml from each time point (before starvation and after starvation on days 4, 10, 14, and 18) were stained with Nile red dye that emits a yellow fluorescence in the presence of lipid and visualized using a fluorescence microscope (Olympus). Meanwhile the cells were also stained with

FITC that emits a green fluorescence in the presence of protein. The detailed processes for staining have already been presented in Chapter 3. 25

Figure 10: Captured fluorescence microscopy pictures over the growth period. The first column pictures (a), (c), (e), (g) and (i) visualizes the protein content before starvation, day 4th , day 10th , day 14th and day 18th after starvations. The second column pictures (b), (d), (f), (h) and (j) visualizes the lipid content before starvation, day 4th , day 10th , day 14th and day 18th after starvations.

4.1.1 Lipid and Protein Content Determination Using Biochemistry Methods

The extraction of biomolecules from the microalgal cells is a laborious task due to their small size and resilient cell walls. Techniques for the release of proteins from 26 biomolecules require harsh treatment of the cells. The complete extraction of proteins requires labor-intensive disruption or homogenization steps by chemical or mechanical means. The hot-alkali method using NaOH or TCA is an effective and high throughput procedure for the complete release of proteins from most microalgal strains. Rausch in 1981 proposed the method of extraction of proteins using sodium hydroxide at a concentration of 0.5 N. Short incubation periods at 80 – 100°C facilitates the complete extraction of proteins without hydrolysis. Dye-based methods like the Microbiuret method for the estimation of proteins is a highly sensitive method and allows the detection of minute amounts of amino acid content in the cells. The microbiuret reagent, prepared from sodium hydroxide and copper sulphate develops color in the presence of proteins which can be measured colorimetrically.

Several conventional techniques exist for the extraction of lipids from microalgal cells. The most popular and commonly used ones are the Bligh and Dyer method and the Folch method. These highly efficient gravimetric methods use volatile solvents for the extraction of lipids from microorganisms. The Bligh and Dyer method is analogous to the Folch method but differs only in the ratio of the solvents used for the tissues. Methanol, chloroform and water are added to the microalgal samples in a two-step extraction procedure. The process starts with the homogenization of the microalgal cells using chloroform and methanol (1:2 ratio). This is followed by a liquid-liquid partitioning step. The mixture of biomolecules is separated into two homogenate phases. The chloroform-rich lower layer contains the non-polar and hydrophobic components that include mostly lipids. The upper (water-methanol) layer contains all the carbohydrates, proteins and polar lipids (mostly phospholipids). 27

The chloroform rich lower layer is allowed to evaporate leaving behind the lipids and the weight of the lipids can be determined.

Figure 11: Extracted lipid and protein content using conventional biochemical methods and techniques

Figure 11 shows both extracted lipid and protein biomass percentage over several starvations days. As previously revealed, the total volume of lipid increases meanwhile the total volume of the protein decreases with starvation.

4.1.2 Statistical Analysis

It is worth to mention that each day several photo fluorescence frames were taken. As the cell size and content is indeed varied within the same day, a statistical analysis is required. Here in this thesis; the principle of histogram is utilized. The histogram is considered to be an accurate representation of the distribution of numerical data. The histogram for the taken photo with different levels, should count for the relative stained composition. The histogram represents an estimate of the probability distribution of the relative stained composition. Figure 12 displays the 28 histogram curves versus different levels for several starvation days. With days, the level of protein corresponding histogram decreases. Thus, indicate that the protein level across the whole cells has been decreased. Comparing day 18 and day 4, the protein level has dropped dramatically.

Figure 12: Histogram for protein versus level for several starvation days

Throughout the experimental period, the microalgal cells were counted thrice to determine the cell numbers and the mean value was used for monitoring the cell growth. The fluorescent microscopy pictures of the microalgal cells were taken, starting from a day before starvation and continuing through the days of starvation.

It can be observed from the fluorescence microscopy images that the amount of protein in the cells reduces while advancing through the days of starvation. The amount of protein in the cells which appears as green due to the fluorescent stain

FITC, is observed to be more in the images of the cells taken before starting the starvation, they are found to occupy almost the entire area, approximately 90 percent of the cell. The amount of lipids (triglycerides, appearing as golden yellow islands) is 29 meagre to less than 10% of the cell area before starting the starvation. During the starvation period, the protein content of the cells, which accounts to 60% of the cells in the field, starts to diminish (Protein Day 4), whereas the lipid droplets started to occupy almost 20% of the cell surface area (Lipid Day 4). Further advancement into starvation, caused the loss of the protein content in 60-70% of the microalgal cells

(Protein Days 10, 14) in each field, whereas the lipid droplets in the cells began to form small islands and occupied nearly 60% of the cell area (Lipid Days 10, 14). The final days of starvation shows major loss of protein content in the cells due to the alteration in the metabolic pathway of the cells, occupying only 15 – 20% of the cell surface area whereas the lipid islands expanded occupying 80 – 90% of the cell area in 90% of the cells.

4.2 Lipid and Protein Area Determination from Image Processing

The captured fluorescence photos were imported to Aphelion software. The photos were processed to compute the lipid and protein total area inside a single microalgal cells. The extracted parameters are listed in Table 1.

Before After starvation starvation 1 Day 1st 4th 10th 14th 18th 2 Lipid area (a.u.) 0.005 0.095 1.766 0.6393 2.741 3 Cell area (a.u.) 0.258 1.317 8.815 2.663 6.812 4 Actual cell area in µm² 52 260 1750 540 1370 5 Lipid percentage (%) 2.0 7.0 20 24 40 6 Protein area (a.u.) 2.601 0.608 0.418 0.4491 0.103 7 Cell area (a.u.) 3.515 0.89 1.811 3.597 0.996 8 Actual cell area in µm² 710 185 350 715 207 9 Proteins percentage (%) 74 68 23 12 10

Table 1: Several extracted parameters per day using Aphelion Dev. (a.u. = arbitrary unit). Table 1 summarizes per day, the lipid and cell areas extracted using the software and the actual cell area measured using the microscope. The software can be calibrated directly to extract the right dimensions. Nevertheless, it is advisable not to change the imported photos settings, lest the quality becomes worst. As long as the extracted lipid and protein relative areas are corresponding to a relative extracted cell dimensions, their percentage can be computed directly by dividing their each total area within a single cell by its corresponding relative cell areas. Both lipid and protein percentages have been computed accordingly and listed in Table 1, rows 5 and 9, respectively. With nitrogen starvation and as displayed in Table 1, row 5 the total amount of lipid in a single cell increases, meanwhile as revealed by row 9, the protein percentage is decreased. This well corroborated with the biochemical extractions results reported in the previous section. Both results have been superimposed with each other and plotted in Figure 13. The biochemical method that has been applied is for the whole suspension that contains millions of cells. It is based on biomass extraction concept. On the contrary, the image processing has been carried at the level of single cell. Figure 13 revealed that single cell image results are validated as they follow the same trend of the biochemical analysis. 31

Figure 13: Image processing and biochemical extraction results

4.3 Discussion

A possible alternative to the fossil fuel is the biodiesel obtained from the microalgae, because they are renwable sources of lipids. Microalgae are photosynthetic organisms that accumulate significant amounts of neutral lipids in the form of TAGs (triacyl glycerols). Ever since the progression in the 1930’s, there has been a surge in the research to develop methods to analyze the microalgae oil content. There is a necessity to develop and improve the methods and techniques that provide accurate determination results of microalgae oil content. In addition, it is essential for the methods to require short preparation time. It is also essential to have low costs in the determination of oil content. Methods with these three particular necessities are those that are of real importance and have made valid contribution to this field.

Most of the previously used conventional techniques used in the determination of oil content can be either expensive or labor intensive or it can be a 32 combination of both. Another disadvantage would be that these conventional methods can be too invasive. One such example is the gravimetric oil determination.

The primary challenges in the production of biodiesel from microalgae are the selection of suitable strain with good oil productivity, their culture conditions, easy determination of the oil content and the determination of a feasible and suitable method for the complete extraction of the lipid content from the cells. In this research thesis, a novel, cost effective and entirely non-invasive image processing technique has been described. This saves the method from being labor-intensive and, consequently, is time effective especially when compared to the other popular techniques utilized worldwide.

Some of the commonly used techniques for the estimation of microalgal oil content have been reviewed and discussed previously. Some of the conventional methods of oil extraction and estimation are gravimetric quantification, liquid chromatographic techniques and infrared spectroscopy. Although the gravimetric methods are widely popular and commonly used ones, the use of a large sample volume and highly volatile and toxic reagents is a disadavntage in addition to the long processing times. TLC and HPLC provide highly accurate results but are expensive and require several processing steps [92]. The infrared spectroscopic methods like NIR and FTIR require minimal sample processing and accurate but need expertised people. The process is fast with minimal processing.

In this image processing method, fluorescent microscopy technique with the use of lipophillic and protein specific stains have been used to detect the lipids and proteins. The fluorescent microscopy is a gentle technique that stains the cellular components of interest due to the specificity of the stains while maintaining the 33 cellular structure. The optimization of a suitable pre-treatment method, maintains the cellular structure and provide optimum staining. Nannochloropsis strain used in the present study is an oleaginous microalgae that accumulate considerable amounts of lipids under conditions of stress like nitrogen starvation [28]. The fluorescent lipophillic dyes Nile Red and BODIPY are used due to their selective affinity to neutral lipid droplets in the cells. They specifically bind to the lipid particles which make them fluoresce under a fluorescent light which helps the user to visualize the lipid bodies. FITC specifically binds to the amino groups in proteins present in the cells.

Nile red staining has been used for the quantification of natural oils in various strains of microalgae like Chlamydomonas sp. and Scenedesmus strain. Studies have shown that the protocols for Nile red assays must be standardized for each strain of microalgae. Spectrophotometric protocols for the absolute quantification of the oil content using fluorescence are optimized. The NR assays were improved and modified for accuracy mostly by two approaches namely, bleaching of the cells prior to measurements and the use of milk as standard instead of other commonly used ones like triolein. The penetration of the stain for various species can be done using

DMSO, ethylene glycol or glycerol depending on the strains.

Another lipophillic dye, BODIPY, is highly promising for the staining of various strains of microalgae like the diatoms and dinoflagellates, due to its high photostability and fluorescent quantum yields. But the optimization of the staining technique requires several adjustments and modifications depending on the strains used due to the diverse algal taxa. 34

The amino groups in proteins are stained for fluorescent microscopy using

FITC. The FITC has affinity for the amino groups in the proteins present in the cells

[90]. The fixation of the microalgal cells with 3.7% formaldehyde preserved the structure of the cellular components. The optimal staining of the cells created a low background noise that reduced the exposure times, prevented photobleaching and produces high quality images. DAPI acts as a counterstain and stains the particles of

DNA [91].

The oil and protein monitoring technique using fluorescent microscopy can be used for the determination of oil and protein accumulation patterns in the cells. It provides information on the progression of oil accumulation under various conditions like environments and stresses, in this case, nitrogen starvation. Nitrogen is a critical macronutrient necessary for the growth and various cellular functions of the cells

[19]. The accumulation of lipids, mostly triglycerides, during the stationary phase of the culture, is due to a shift in the metabolic pathway of the cells. Due to the deficient nutrient conditions, the genetic make up of the cell prompts the cellular machinery to accumulate the lipids which is a method to overcome the stress. The fluorescent images show the progression of lipid islands as well as reduction in the protein levels while advancing through the days of nitrogen starvation. This is because the protein accumulated in the cells are used for the more necessary life-sustaining functions of the cells like photosynthesis. As a result, the rate of cell division becomes low and the cellular enzymatic machinery focuses on methods to overcome stress by accumulation of the biomass.

Chapter 5: Conclusions and Future Work

The summary of this work and future outlook are presented as follows:

5.1 Conclusions

Nannochloropsis microalgae strain has been cultured and cultivated for 66 days. The long period was due to the stain growth conditions. A growth period is needed to allow induction of metabolic alterations within the cells necessary under the experimental conditions. Nitrogen starvation process has been conducted to alter the lipid and protein contents inside the microalgal cells. The nitrogen starvation stress condition results in alteration of the lipid and protein content inside the cells.

Due to a lack of nitrogen needed to form the polypeptide bonds, the cells’ metabolism diverts towards lipid accumulation rather than protein synthesis. The intracellular Nannochloropsis cell content was estimated by mathematically evaluating each lipid and protein areas and volumes inside the corresponding-stained cell assisted by Aphelion Dev tools. The image processing based approach is then used to analyze the time course of lipid accumulation and protein reduction patterns inside the corresponding-stained microalgal cells.

From the images obtained, it can be observed that the image processing techniques indeed have been successful at measuring the lipid and protein content.

These image processing techniques, which are more modern and technical, have been proven to be a faster and more accurate method of obtaining the amount of lipid content available. As was predicted previously, the lipid content was found and proven to increase while at the same time the protein content was found and proven to decrease. This is a benefit as the main objective is to enhance and measure the 37 amount of lipid present in a microalgal cell. The results that were obtained corroborated with the results that were obtained from the previous methods used.

5.2 Future Work

In the current work, the staining and image processing techniques are being done on a single cell. Therefore, if the oil content needs to be known, the image processing techniques have to be performed on microalgae cell by cell. This subsection provides a future outlook to develop a new method for lipid and other biomolecules total content determination in a living microalgal single cell assisted by the use of thermal camera to capture the electrical current distribution propagated inside the cell. The method overview is depicted in Figure 14.

Figure 14: Proposed in vivo system for real time measurements of cell content using thermal camera and electrical conditions

The proposed future work allows the direct estimation of lipid and/or protein in real time in vitro measurements. Figure 14 provides a description of the future scope of this work. The microalgal cells are cultured under controlled (artificial) conditions in inside microalgae cultivation box, as shown in Figure 14 (a). Initially, a set of conditions are applied such as the addition of a nutrient medium that supports and maintains the growth of the microalgae, providing artificial conditions like aeration, illumination and temperature in conjunction to other physicochemical parameters and trace elements [1].

In addition to that, a micro pump is suggested to be used, as expressed in

Figure 14 (b). The micro pump is capable of allowing the passage of the culture suspension containing the cells and the medium at the required flow-rate. This passage is connected from the cultured system to a microfluidic system. As illustrated in Figure 14 (c), the micro pump is connected to the microfluidic system through an inlet that allows the flow of single microalgal cell suspension into the microfluidic system. The shown arrow denotes the direction of flow of the suspension (cells and media).

The microfluidic system is composed of a series of microfluidic channel that are etched or molded and allows the flow of liquid and cells inside along with several electrical electrodes. The result of this is that multiple single cells will be subjected to the same electrical condition. Therefore, the thermal camera will be able to monitor several single cells simultaneously. On the other hand, it could also be possible to have different microfluidic channels on the same board subjected to different set of electrical conditions. Such a scenario provides a wider scope and the ability to measure several sets of cells at different electrical conditions at the same time. 39

Furthermore, the outlet allows the discharge of the by-products such as used cells, medium or other liquids out of the system. The microfluidic system is connected to the Gamry Reference 600 electrical analyzer shown in Figure 14 (d) which is a high performance, stand alone, computer-controlled potentiostat/galvanostat/ZRA that interfaces with a computer illustrated in Figure 14

(e) via USB. The computer system is aimed at setting the electrical conditions to apply electrical signals of specific current and frequency to the microfluidic system with the help of the electrodes.

It is expected, based on the intracellular composition of the cells, the distribution of the electrical current varies in the cell. Thus, the magnitude of the electrical current and frequency differs according to the interaction with the biomolecule (lipids and proteins) composition inside the cells. When a current/voltage of defined magnitude and frequency passes through the cell and encounters lipids or protein, resistance or heat is generated.

Finally, a thermal camera as the one shown in Figure 14 (f) can be used to capture pictures of the microfluidic channel. A picture such the one shown in Figure

14 (g) can be obtained from the thermal camera. Based on our developed know how and expectation, the heat distribution produced by the propagation of the current signals will revealed a number of lipid and protein islands presented inside the cell.

Hence, the developed image processing method in this thesis can then be used to estimate the cell content accordingly.

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