ANALYSIS OF LIPIDS FROM CHROMULINA FREIBURGENSIS FOR
THEIR POTENTIAL USE AS BIOFUELS
by
Amanda Mondloch
A thesis submitted in partial fulfillment of the
requirements for the degree of
Interdisciplinary Master of Science
Montana Tech of The University of Montana
2012
ii
Abstract
Many uncertainties concerning energy availability exist for the future. Finding environmentally
friendly, alternative energy sources is a pressing issue, especially for countries that do not have
obtainable oil resources. Energy consumption is currently at an all-time high and is expected to
increase by 50% by 2020. The world consumes 84.2 million barrels of oil per day and
18.7 million barrels are used by the United States alone. Fifty-seven percent (57%) of the United
States’ crude oil is imported and 8.9 million barrels a day are used for motor vehicles. It is
uncertain how long the oil reserves will last and therefore it is imperative to find economically
viable, renewable energy sources with reduced waste emissions. Biofuels are renewable sources
that have received much attention in recent years because they have potential to be
environmentally friendly and readily available. Microalgae are a source of biofuels that have
many advantages over other biofuel sources. Algae do not require agricultural land to grow and
can grow in waste water. Algae fix atmospheric carbon which makes them nearly carbon neutral
and their fuel products do not contain sulfur or many other toxins that can be harmful to the
environment.
Chromulina freiburgensis is a species of algae that was isolated from the water of the Berkeley
Pit in Butte, Montana and has not yet been characterized in terms of biofuel potential. The goal
of this project was to isolate and grow C. freiburgensis, and to extract, characterize, and quantify
the lipids produced from this alga. Fatty acid methyl esters were extracted and quantified via
transesterification. Using the growth conditions presented in this study, C. freiburgensis did not
produce an adequate amount of lipids to be considered a potential biofuel source.
Keywords: algae, biofuels, lipids, Chromulina freiburgensis
iii
Dedication
I would like to thank my family, especially Dan, for their patience and support throughout my
education.
iv
Acknowledgements
Thank you, Dr. Doug Cameron, for all of your assistance and guidance. I also appreciate all of
the help from Dr. Marisa Pedulla and Dr. Grant Mitman. I would also like to say thanks to the
Chemistry and Geochemistry Department at Montana Tech and the National Center for Research
Resources for funding this project. “The project described was supported by Award Number
P20RR016455 from the National Center For Research Resources. The content is solely the
responsibility of the authors and does not necessarily represent the official views of the National
Center For Research Resources or the National Institutes of Health.”
v
Table of Contents
ABSTRACT ................................................................................................................................................ II
KEYWORDS: ALGAE, BIOFUELS, LIPIDS, CHROMULINA FREIBURGENSIS DEDICATION ...... II
ACKNOWLEDGEMENTS ........................................................................................................................... IV
LIST OF TABLES ...................................................................................................................................... VII
LIST OF FIGURES.................................................................................................................................... VIII
INTRODUCTION ....................................................................................................................................... 1
1.
2.
3.
BACKGROUND ................................................................................................................................... 1
1.1.
Biofuels ............................................................................................................................... 1
1.2.
Algae and its Evolution....................................................................................................... 3
1.3.
Lipid Metabolism in Algae .................................................................................................. 6
1.4.
Chromulina freiburgensis ................................................................................................... 7
METHODS ........................................................................................................................................ 7
2.1.
Isolation of Chromulina freiburgensis ................................................................................ 7
2.2.
Growth of Algae ................................................................................................................. 8
2.3.
Lipid Extraction................................................................................................................. 10
2.4.
Esterification of Lipids [26] ............................................................................................... 11
2.5.
Polysaccharide Extraction ................................................................................................ 12
2.6.
Characterization of Lipids ................................................................................................. 12
2.7.
Quantification of Lipids .................................................................................................... 14
2.8.
Characterization of Polysaccharides ................................................................................ 14
RESULTS AND DISCUSSION ................................................................................................................. 16
3.1.
Isolation of Chromulina freiburgensis .............................................................................. 16
3.2.
Characterization of Lipids ................................................................................................. 16
3.3.
Identification of Lipids ...................................................................................................... 23
vi
3.4.
Quantification of Lipids .................................................................................................... 38
4.
CONCLUSION .................................................................................................................................. 57
5.
FUTURE WORK................................................................................................................................ 58
REFERENCES CITED................................................................................................................................. 59
APPENDIX A: .......................................................................................................................................... 62
vii
List of Tables
Table 1: GC-MS Parameters ..............................................................................................13
Table 2: C4-C24 FAME Standard Analytes ........................................................................18
Table 3: C4-C24 Standard Mixture vs. EI Transesterification Extract Lipid Retention Times
................................................................................................................................23
Table 4: C4-C24 Standard FAME Mixture Areas and Concentrations ...............................39
Table 5: C4-C24 Standard FAME with Internal Standard Serial Dilutions ........................40
Table 6: C4-C24 Standard FAME with Internal Standard Response Factor Values ...........52
Table 7: Concentration of Fatty Acids in Transesterification Sample ...............................53
Table 8: Estimated Concentration of Fatty Acids in Transesterification Sample ..............53
Table 9: Ratio of Freeze Dry (FD) Peak Area and Height vs. Oven Dry (OD) Peak Area and
Height .....................................................................................................................54
viii
List of Figures
Figure 1: Chlamydomonas cell [17].....................................................................................5
Figure 2: Light Micrograph of Chromulina freiburgensis...................................................8
Figure 3: Petroff Hausser Counting Chamber ...................................................................10
Figure 4: C. frieburgensis colony ......................................................................................16
Figure 5: EI Chromatogram of C4-C24 Fatty Acid Methyl Ester Standard Mixture ........17
Figure 6: EI Chromatogram of C. freiburgensis extract- Bligh and Dyer Method............19
Figure 7: EI Chromatogram of C. freiburgensis extract- Ether Reflux Method ................20
Figure 8: EI Chromatogram of Transesterification Extract from C. freiburgensis............22
Figure 9: CI Chromatogram of Transesterification Extract from C. freiburgensis ...........22
Figure 10: EI Chromatogram for the Retention Time that Includes 45-50 minutes from C4-C24
FAME ....................................................................................................................24
Figure 11: EI Chromatogram for the Retention Time that includes 45-50 minutes from the
Transesterification Extract .....................................................................................25
Figure 12: CI Chromatogram for the Retention Time that includes 45-50 minutes from the
Transesterification Extract .....................................................................................26
Figure 13: EI Chromatogram for the Retention Time that includes 45-55 minutes from the
Transesterification Extract .....................................................................................27
Figure 14: Transesterification Mass Spectrum RT 49.19 minutes vs. C4-C24 FAME Standard
Mass Spectrum RT 49.23 minutes .........................................................................29
Figure 15: CI Mass Spectrum of Transesterification Extract at Retention Time 49.19 (C:18:0)
................................................................................................................................30
ix
Figure 16: Transesterification Mass Spectrum RT 48.31 vs. C4-C24 FAME Standard Mixture
Mass Spectrum Comparison of Peak at RT 48.33 .................................................31
Figure 17: Transesterification CI Mass Spectrum of Retention Time 48.31 Minutes (C:18:1)
................................................................................................................................32
Figure 18: Transesterification Mass Spectrum RT 47.97 minutes vs. C4-C24 FAME Standard
Mixture Mass Spectrum Comparison of Peak at RT 47.72 minutes......................33
Figure 19: Transesterification CI Mass Spectrum of Retention Time 47.97 (C:18:2) ......34
Figure 20: Transesterification Mass Spectrum RT 47.14 minutes vs. C4-C24 FAME Standard
Mixture Mass Spectrum Comparison of Peak at RT 46.99 minutes......................35
Figure 21: Transesterification CI Mass Spectrum of Retention Time 47.14 minutes (C:18:3)
................................................................................................................................36
Figure 22: EI Transesterification Mass Spectrum at Retention Time 47.39......................37
Figure 23: Transesterification CI Mass Spectrum of Retention Time 47.39 (C:18:4) ......37
Figure 24: EI Chromatogram of C4-C24 Fatty Acid Methyl Ester Standard Mixture ........38
Figure 25: C4-C24 Standard FAME C:12:0 Calibration Curve ..........................................42
Figure 26: C4-C24 Standard FAME C:13:0 Calibration Curve ..........................................42
Figure 27: C4-C24 Standard FAME C:14:0 Calibration Curve ..........................................43
Figure 28 :C4-C24 Standard FAME C:14:1 Calibration Curve ..........................................43
Figure 29: C4-C24 Standard FAME C:15:0 Calibration Curve ..........................................44
Figure 30: C4-C24 Standard FAME C:16:0 Calibration Curve ..........................................44
Figure 31: C4-C24 Standard FAME C:16:1 Calibration Curve ..........................................45
Figure 32: C4-C24 Standard FAME C:17:0 Calibration Curve ..........................................45
Figure 33: C4-C24 Standard FAME C:18:0 Calibration Curve ..........................................46
x
Figure 34: C4-C24 Standard FAME C:18:1 (RT 48.18 min) Calibration Curve ................46
Figure 35: C4-C24 Standard FAME C:18:3 Calibration Curve ..........................................47
Figure 36: C4-C24 Standard FAME C:18:2 (RT 47.72 min) Calibration Curve ................47
Figure 37: C4-C24 Standard FAME C:18:2 (RT 48.33 min) Calibration Curve ................48
Figure 38: C4-C24 Standard FAME C:18:3 Calibration Curve ..........................................48
Figure 39: C4-C24 Standard FAME C:20:0 Calibration Curve ..........................................49
Figure 40: C4-C24 Standard FAME C:20:5 Calibration Curve ..........................................49
Figure 41: C4-C24 Standard FAME C:21:0 Calibration Curve ...........................................50
Figure 42: EI Chromatogram of Transesterification Sample from Oven Dried C. freiburgensis
................................................................................................................................55
Figure 43: EI Chromatogram of Transesterification Sample from Freeze Dried C. freiburgensis
................................................................................................................................55
Figure 44: CI Chromatogram of Transesterification Sample from Oven Dried C. freiburgensis
................................................................................................................................56
Figure 45: CI Chromatogram of Transesterification Sample from Freeze Dried C. freiburgensis
................................................................................................................................56
1
Introduction
Finding alternative sources of energy has been an important topic since the oil crisis in
the 1970’s [1]. Not only is oil availability a concern for the future, but its combustion has a
negative impact on the environment in terms of greenhouse gas emissions [2-4]. Microalgae
have received much attention worldwide in recent years because they have many advantages
over other natural energy sources. Algae grow almost everywhere including sea water,
freshwater, sewage, and in waste lands [5]. Algae also grow especially well in areas where
carbon dioxide is present in abundance for example near coal fired power plants. Algae can also
grow in industrial settings where carbon is deficient but nitrogen and phosphorous are abundant
[6]. Algae are capable of multiplying very quickly and can produce high neutral lipid and
polysaccharide yields as byproducts of photosynthesis under the appropriate conditions
[2, 7].
Based on previous research, several species of algae have proven to be abundant
producers of oils and lipids and have been investigated as biofuel sources. Lipids comprise up to
50% of the dry mass of Chlorella sp., Chlamydomonas sp, and Neochloris oleoabundans [5, 6].
Some species of algae such as Caulerpa racemesa and Pterocladia capillacea also have high
polysaccharide contents [8]. Polysaccharides can be extracted from the biomass and used to
make ethanol for fuel or feedstock.
1. Background
1.1. Biofuels
Biofuels are solids, liquids, or gases that are made from recent biomass [9-11]. Biofuels
are originally produced from photosynthesis. Biofuels are alternative fuel sources made from a
2
variety of natural crops including corn, rapeseed, soybean, sunflower, flax, and hemp. They can
also be produced from waste vegetable oils and microalgae oils.
Biofuels are classified as 1st, 2nd, 3rd, or 4th generation [13, 14, 15, 16]. First generation
biofuels are made from feedstocks that have been used as food such as sugar, starch, or vegetable
oil. Second generation biofuels are fuels that are made from non-food feedstocks such as
lignocellulose from grasses, agricultural residues, and wood and forestry residues [12]. Third
generation biofuels are also made from non-food feedstocks, but the fuel is identical to their
petroleum counterparts and they are carbon neutral. Algae are a third generation biofuel source
and are considered carbon neutral because they use sunlight and carbon dioxide for
photosynthesis and therefore the amount of carbon used for growth essentially equals the amount
of carbon released into the atmosphere when these fuels are burned. Fourth generation biofuels
are made from genetically engineering third generation biofuels and are designed to be carbon
negative.
The major forms of biofuels today are biodiesel and bioethanol [12]. Bioethanol can be
used as an additive to gasoline to make mixtures of E15 (15% ethanol, 85% gasoline) and E85
(85% ethanol, 15% gasoline). Lignocellulosic ethanol is a second generation biofuel that is
produced from non-food residual biomass such as stems, leaves, wood chips, and non-food crops
including switchgrass and algae, however, the major feedstocks for bioethanol are sugarcane and
corn [13, 14, 15, 16]. Bioethanol is a particulate free burning fuel source that combusts cleanly
with oxygen [1]. Greenhouse gases may be reduced 12% by production and combustion of
ethanol versus conventional fuel sources [13]. Ethanol can also be used to make biodiesel.
3
Biodiesel is an alternative to diesel, and many engines can use 100% biodiesel. Biodiesel
is produced from vegetable oil, plant oil, and animal fats. The majority of biodiesel feedstock is
soybean, canola seed, rapeseed, sunflower, and palm oil.
1.1.1.
Advantages of Biofuels
The world’s petroleum reserves are expected to be depleted in less than 50 years at the
current rate of consumption [1, 14]. Biofuels are a biodegradable, renewable energy source that
can, in principle, be sustainably supplied. Biofuels are environmentally friendly; they create no
net increase in carbon dioxide released into the atmosphere, have no aromatic compounds, and
have no sulfur content which makes burning biofuels 90% less toxic compared to common diesel
sources [1]. Biofuels have better efficiency than conventional diesel fuel in terms of flash point
and biodegradability, and they have economic potential due to the ever increasing prices of nonrenewable fossil fuels.
When algal biofuels are produced via lipid extractions, a residue is left as waste. The
residue from these extractions can be used as aquaculture feed, animal food supplements, or a
source of pharmaceuticals [3].
1.2. Algae and its Evolution
The tree of life has three primary domains; bacteria, archaea, and eukaryotes. Bacteria
and archaea are prokaryotes. Eukaryotes and prokaryotes differ in size, complexity, structure and
function. First, eukaryotes are generally larger than prokaryotes with an average cell size of
10-100 microns and 0.1-0.2 microns, respectively [15]. Prokaryotes are mostly single celled
individuals while eukaryotes may be unicellular or multicellular organisms. Not only are
eukaryotes larger, they are much more complex than prokaryotes. Eukaryotes have membrane
bound organelles such as mitochondria, chloroplasts, Golgi apparatus, and endoplasmic
4
reticulum [15, 16]. Prokaryotes do not have membranous organelles, a cytoskeleton, or a nuclear
envelope that separates their DNA from the cytoplasm. Also, eukaryotes have multiple
chromosomes consisting of linear DNA molecules and prokaryotes contain a single circular
piece of DNA.
It is thought that mitochondria and chloroplasts in eukaryotes evolved from
endosymbiotic bacteria. Mitochondria were originally aerobic heterotrophic prokaryotes that
developed an endosymbiotic relationship with a bacterium which eventually became eukaryotic
cells [15, 16]. Plant and algal chloroplasts were originally photosynthetic prokaryotes and
evolved later, involving a second endosymbiotic event [15]. It is thought that there were several
symbiotic events thereafter, which accounts for the diversity of plastids in algal groups [15].
Algal groups are separated taxonomically by their 18S ribosomal RNA. 18S rRNA is part of the
small subunit of eukaryotic ribosomes. This particular piece of RNA is used as a target for
polymerase chain reaction and phylogenetic studies because it is easily accessible and highly
conserved flanking regions.
Algae are plant-like, photosynthetic, eukaryotic organisms. Algae are typically
autotrophic and have a membrane bound nucleus and plastids but do not have roots, stems,
leaves, or vascular tissue (Figure 1) [5]. They have relatively simple reproductive structures but
can reproduce via asexual cell division or through complex sexual reproduction.
Algae are separated taxonomically into seven different divisions based on several factors
such as the pigments they use for photosynthesis, storage carbohydrates, cell wall components,
and flagella. The seven divisions are Euglenophyta, Chrysophyta, Pyrrophyta, Chlorophyta,
Rhodophyta, Phaeophyta, and Xanthophyta [16]. They range in size from single celled
5
microorganisms to complex multi-cellular life forms such as giant kelps which can grow to
65 meters in length [16].
Figure 1: Chlamydomonas cell [17].
Algae are found in salt water, fresh water, and waste water around the world. They can be
cultivated under difficult climatic conditions and produce a wide variety of commercially
interesting byproducts including oils, fats, sugars, and functional bioactive compounds [5].
Algae are a very important species because they produce a large amount of the world’s
photosynthetic oxygen. Algae are at the base of the food chain and they are a food source for
many animals. Importantly, algae may also have the potential to become large producers of
biodiesel.
There are many advantages to using algae as a source of biodiesels such as:
6
1.
They fix carbon dioxide from the atmosphere and thus reduce the amount of
atmospheric CO2 which is currently considered a global environmental
problem [5, 9, 12].
2. Algal biofuels are non-toxic, biodegradable and contain no sulfur.
3. Algae can grow in conditions that may otherwise be considered unsuitable for
conventional plants.
4. They provide much higher yields of biodiesel fuel than other energy sources.
5. They have higher photosynthetic efficiencies and growth rates compared to
other crops.
6. Microalgal oils are more similar to conventional fuels than oils from other
biofuel sources and are more stable based on their flash point values.
7. Algae can be grown in photobioreactors (better growth control) or open ponds
(cheaper). The carbon source for algae growing in open ponds may come from
coal fired power plants; the CO2 and wastewater are therefore recycled.
1.3. Lipid Metabolism in Algae
Lipid metabolism has been extensively studied in plants and animals but little is known
about the key enzymes and biochemical pathways involved in the biosynthesis of lipids in
microalgae, especially Chromulina. Lipid metabolism is a complex process and many of the key
lipid metabolizing enzymes that have been identified in algae are based on orthology to fungi
and plants [18]. Most of this work has been done on the reference organism, Chlamydomonas
reinhardtii, which may or may not represent the lipid metabolizing enzymes or products found in
the same class, let alone division [19]. Genetic approaches have been used to determine the
7
essential genes involved in the synthesis of lipids [19]. Algal mutants have been used to
determine the functions and physiological roles of lipids and metabolizing enzymes in vivo.
1.4. Chromulina freiburgensis
Chromulina freiburgensis is a single-celled golden brown alga and is a member of the
class Chrysophyta [20, 21]. Members of Chrysophyta are mostly found in fresh water at a
slightly acidic pH. Their cell walls tend to have a high concentration of silica and their main
energy storage product is laminarin. Brown algae contain chlorophylls a and c in addition to
carotenes and xanthophylls [22]. Carotenes are unsaturated hydrocarbons that aid in
photosynthesis and protect algal tissues. Xanthophylls are similar in structure to carotenes but
they contain oxygen. Xanthophylls absorb light wavelengths that are not absorbed by
cholorophylls.
C. freiburgensis exhibits two life stages, a non-motile coccoid state and a flagellated state
[20, 21]. Cells in the coccoid life stage are round in shape with a diameter ranging from 5-10 µm.
Flagellated cells are 7-15 µm in length with a 3-5 µm flagellum [15, 20]. In uncontaminated
environments, C. freiburgensis is most commonly found in the flagellated state. The strain of
C. freiburgensis examined was isolated from the surface water of the Berkeley Pit in Butte,
Montana [14]. The Berkeley Pit is an acidic mine waste lake that contains heavy metals such as
iron, zinc, manganese, and copper.
2. Methods
2.1. Isolation of Chromulina freiburgensis
2.1.1.
Initial Algal Culture
All glassware, tubes, filters, and fertilizer were autoclaved. One tablespoon of Osmocote
fertilizer (ammonium nitrate, ammonium phosphate, calcium phosphate, and potassium
8
phosphate), 936 mL of Berkeley Pit surface water, and 10 mL of NaNO3 (5.0 g/200 mL) were
mixed together in an aspirator bottle [23]. A previously grown C. freiburgensis colony from a
1% modified acid medium (MAM) agar plate was used to inoculate the medium. The aspirator
bottle was connected to continuous air flow, a constant light intensity of 100 µmol/m2/s, and was
kept at a temperature of 9.8°C in the Percival Scientific culture chamber.
2.2. Growth of Algae
2.2.1.
MAM [24]
Ten mL of the initial algal culture (IAC) of C. freiburgensis (Figure 2) grown was used to
inoculate 1000 mL of MAM in an aspirator bottle. The bottle was placed in the Percival
Scientific culture chamber with constant air flow, a light intensity of 100 µmol/m2/s, and a
temperature of 9.8°C.
20 µm
Figure 2: Light Micrograph of Chromulina freiburgensis
9
As the algal culture was used for analysis, MAM was added to the aspirator bottles
allowing for a continuous supply of culture. Each culture was analyzed under the Nikon Eclipse
E800 microscope to make sure it was not contaminated.
2.2.2.
Berkeley Pit Water, NaNO3, Osmocote Fertilizer
One tablespoon of Osmocote fertilizer, 936 mL of Berkeley Pit water, and 10 mL of
NaNO3 (5.0 g/200mL) were combined in a 1000 mL aspirator bottle. The medium was
inoculated with 10 mL of IAC and placed in the Percival Scientific culture chamber with
constant air flow, a light intensity of 100 µmol/m2/s and a temperature of 9.8°C.
2.2.3.
Acidified Bold’s Basal Medium
Bold’s Basal Medium (BBM) was acidified to pH 2.5 with A.C.S. reagent grade sulfuric
acid obtained from JT Baker [25]. 1000 mL of Acidified Bold’s Basal Medium was placed in an
aspirator bottle. The medium was inoculated with 10 mL of IAC, placed in the Percival
Scientific culture chamber with constant air flow, a light intensity of 100µmol/m2/s, and a
temperature of 9.8°C.
2.2.4.
Algal Counts
Twenty µL of the culture was placed on a Petroff-Hausser counting slide (Figure 3) and
magnified at 20x under the Nikon Eclipse E800 microscope. Five boxes in the center grid were
arbitrarily selected and the algal cells were counted. The mean number of cells was calculated
and inserted into the following formula to determine the number of cells/mL (1).
(n1 + n2 + n3+ n4 + n5/5) x (25) x (50,000) = # cells/mL
N = the number of cells per small box
25 = the number of larger boxes in the center of the grid
50,000 = dilution factor
(1)
10
Figure 3: Petroff Hausser Counting Chamber
2.3. Lipid Extraction
2.3.1.
Ether/Water Reflux
The algal culture (~500 mL) was dried in a drying oven at 70° C for 24 hours. The dried
biomass (~1.0 g) was weighed and placed in a 250 mL glass round bottom flask. Thirty-five mL
of reagent grade 99.8% diethyl ether obtained from Acros Organics was added to the flask and
the mixture was refluxed for five hours. The solvent phase was decanted into a 45 mL glass vial
for esterification. Thirty-five mL of deionized water was added to the 250 mL round bottom
flask containing the solid residue and refluxed for another five hours. The aqueous solvent was
filtered and concentrated by gentle heating (60°C). The concentrated aqueous extract was
transferred to a glass 5 mL vial for analysis [26]. All extracts were stored in the refrigerator at a
temperature of 5°C.
2.3.2.
Bligh and Dyer Lipid Extraction [27]
The algal culture was dried in a drying oven at 70°C for 24 hours. The dried biomass
(~1.0 g) was massed and ground into a fine powder. Three mL of A.C.S. grade
chloroform:methanol (2:1) obtained from Fisher Scientific was added to the biomass. The
mixture was agitated for 20 minutes at room temperature. The solvent phase was recovered by
centrifugation and the whole solvent was evaporated. The organic material was re-dissolved in
11
98.5% hexane (Sigma Aldrich) for analysis via gas chromatography mass spectrometry
(GC-MS).
2.3.3.
Transesterification of Microalgae Lipids
C. freiburgensis was dried via a drying oven and a freeze dryer. The drying oven was set
at 80°C and the algae were placed in it for 24 hours [28]. A dry weight was obtained (~0.5 g).
The dried biomass was placed in a 50 mL glass round bottom flask where it was ground into a
fine powder. Two and a half mL of A. C. S. reagent grade 2.5% sulfuric acid in methanol was
added to the flask. The mixture was heated at 60°C for two hours with constant stirring. Then,
the mixture was vacuum filtered and the solution containing the supernatant was transferred to a
separatory funnel. Three mL of hexane was added and mixed well for 15 minutes and the
hydrophobic layer was transferred to a new vessel (repeated three times). The hydrophobic layer
was then washed with three mL of de-ionized water and transferred to a 45 mL vial. A.C.S.
reagent grade anhydrous sodium sulfate obtained from JT Baker was added to the hydrophobic
layer to remove all remaining water. Gentle heat was applied to the hydrophobic layer to
concentrate.
2.4. Esterification of Lipids [26]
2.4.1.
Method I-Esterification of Lipids from Bligh and Dyer
Extraction
Five mL of 2.5% sulfuric acid in methanol was added to the vial containing the Bligh and
Dyer lipid extract. The mixture was heated at 80°C for three hours with constant stirring. The
mixture was removed from the heat and an equal volume of de-ionized water and hexane was
added to the vial. The vial was mixed and the organic layer was recovered for further analysis.
12
2.4.2.
Method II-Esterification of Lipids from Ether Reflux
Five mL of 2.5% sulfuric acid in methanol was added to the ether reflux ether extract and
placed in the drying oven at 80°C for 80 minutes. Five mL of hexane were added and mixed
well. The mixture was centrifuged for two minutes and the top (organic) layer was recovered.
The process was repeated three times. The organic layer was further analyzed.
2.5. Polysaccharide Extraction
One liter of algae was dried in the drying oven at 80°C for 24 hours [29]. A dry weight
was obtained. The dried biomass was placed in a 250 mL round bottom flask. Fifty mL of ethyl
alcohol obtained from Aldrich was added to the flask. The mixture was heated for two hours at
80°C. The flask was cooled and the supernatant was removed. Fifty mL of 25°C de-ionized
water was added to the flask. The mixture was stirred for two hours. The supernatant was
transferred to a 150 mL beaker and all water was evaporated. Twenty mL of ethanol was added
to the beaker, mixed well, and transferred to a vial. The addition of ethanol was repeated four
times. The ethanol volumes were combined, centrifuged, and concentrated for analysis via liquid
chromatography mass spectrometry (LC-MS).
2.6. Characterization of Lipids
2.6.1.
GC-MS Method
One µL of a C4-C24 standard fatty acid methyl ester (FAME) mixture catalog number
18919-1AMP (100 mg/mL hexane) obtained from Sigma Aldrich was injected via the heated
needle technique into the Thermo Scientific Trace GC Ultra Model No. K24300000000080
coupled to an ITQ 1100 Mass Spectrometer (see Table 3.2.1). The FAME standard was analyzed
via electron ionization (EI) and chemical ionization (CI). The GC-MS parameters used for all
analyses are listed in Table 1.
13
Table 1: GC-MS Parameters
Trace GC Ultra Oven Parameters
Initial Temperature
100°C (hold 1.0 minute)
Ramp #1
2°C/min until 250°C (hold 1.0 min)
Ramp #2
15°C/min until 280°C (hold 1.0 min)
Carrier Gas
Constant Flow
PTV Mode
Helium
1.0 mL/min
Splitless Injection
PTV Inlet Parameters
Initial Temperature
280°C
Inject
0.1 min
Transfer Rate
14.5°C/sec to 280°C (hold 5 min)
ITQ 1100 MS Parameters (EI)
Source Temperature 280°C
Start Time
3 min
Damping Gas Flow
0.3 mL/min
Microscans
3
Maximum Ion Time
25 ms
Tune File
Autotune
Full Scan
100-550 amu
Electron Lens
10 V
Electron Energy
70 eV
Emission Current
200 μamp
ITQ 1100 MS Parameters (CI)
Source Temperature 280°C
Start Time
3 min
Damping Gas Flow
0.3 mL/min
Microscans
3
Maximum Ion Time
25 ms
Tune File
Autotune
Full Scan
100-550 amu
Electron Lens
10 V
Electron Energy
120 eV
Emission Current
200 μamp
*Acetonitrile was used as the reagent gas for all positive ion CI analyses.
Column- Restek Rxi-5MS, 30 m x 0.25 mm i. d. x 0.25 µm film thickness
14
The positive ion-source pressure was set by adjusting the ratio between the
1-methyleneimino-1-ethenylium ion and the adduct ion. The oven was heated to 280°C for one
hour between analyses to ensure no compounds adsorbed to the column. One µL of sample from
all extractions was injected, and the data were analyzed with XCalibur version 2.0 software.
2.7. Quantification of Lipids
2.7.1.
Internal Standard
102.8 mg naphthalene (obtained from Sigma Aldrich) /mL hexane was made as a stock
solution for an internal standard. The stock solution was diluted to make the internal standard
(ISS). Ten µL of the stock solution was diluted to one mL with hexane.
The transesterification method was performed on 1.3 g of dried algae. The final volume
of product dissolved in hexane after concentration was 2.5 mL. Ten µL of the ISS was added to
one mL of transesterification sample. One µL was analyzed via EI and CI GC-MS.
2.7.2.
Calibration Curve
Two hundred µL of the C4-C24 FAME standard mixture and five µL of ISS were
combined. One µL of this solution was analyzed via EI GC-MS. A series of standards were then
made by serially diluting the ISS (C4-C24 FAME standard mixture with internal standard) by a
factor of five with hexane. Calibration curves were developed to determine the concentration of
the lipids in the transesterification extract.
2.8. Characterization of Polysaccharides
2.8.1.
Standard Solutions
Standard solutions of glucose and sucrose obtained from Fisher Scientific and Sigma
Aldrich, respectively were made by adding 50 µg of each to 100 mL of de-ionized water. LC-MS
analyses were performed with an Agilent 1100 Series LC coupled with a Bruker Esquire 4000
15
APCI mass spectrometer. The column used was an Eclipse XDB-C18 column (5 µm x 4.6 µm x
150 mm). All LCMS analyses were performed using this instrument. Fifteen µL of each standard
solution was injected. An isocratic elution with 50:50 methanol:water as the solvent was
performed with a flow rate of 1 mL/min.
2.8.2. Polysaccharide Analysis
The concentrated sucrose standard was analyzed via negative ion atmospheric pressure
chemical ionization mass spectrometry (APCI MS/MS) with a vaporizer temperature of 400°C.
The standard was injected with a syringe pump.
16
3.
Results and Discussion
3.1. Isolation of Chromulina freiburgensis
C. freiburgensis was successfully isolated and grown in pure culture (Figure 4). The
MAM proved to be the best medium for the C. freiburgensis in terms of growth rate [20]. All
algal cultures reported in this thesis were grown with MAM.
20 µm
Figure 4: C. frieburgensis colony
3.2. Characterization of Lipids
Figure 4 is a representative chromatogram of an EI GC-MS analysis of the C4-C24 FAME
standard mixture. This chromatogram shows separation for most of the fatty acid methyl esters
present. High temperatures at the end of the analysis caused column bleed, and the resolution at
the tail end (retention time of 60 minutes and greater) was poor. Table 2 lists all of the analytes,
their weight percent, and their retention times (RT).
17
RT: 0.00 - 80.03
40.09
100
NL:
9.74E5
TIC MS
FAMESTD
_11032117
79.99 3049
90
49.23
80
57.68
48.02
70
60
65.56
50
30.25
40
20.14
30
35.01
56.55
44.74
61.70
73.06
11.02
20
55.54
29.59
5.06
15.32
10
5.43
0
0
10
20
30
40
Time (min)
50
60
70
Figure 5: EI Chromatogram of C4-C24 Fatty Acid Methyl Ester Standard Mixture
80
18
Table 2: C4-C24 FAME Standard Analytes
C4-C24 FAME Standard Mixture
Analyte
Wt %
Formula
RT
Analyte
Wt %
Formula
RT
Methyl butyrate
4.00
C:4:0
**
Methyl linoleate
2.00
C:18:2
*
Methyl hexanoate
4.00
C:6:0
**
Methyl linolenate
2.01
C:18:3
*
Methyl octanoate
3.98
C:8:0
5.06
Gamma-linolenic acid
methyl ester
2.02
C:18:3
*
Methyl decanoate
3.99
C:10:0
11.02
Methyl arachidate
4.02
C:20:0
57.68
Methyl
undecanoate
1.99
C:11:0
15.32
Methyl eicosenoate
2.04
C:20:1
*
20.14
Cis-11,14eicosadienoic acid
methyl ester
2.00
C:20:2
*
25.19
Cis-11,14,17eicosatrienoic acid
methyl ester
2.05
C:20:3
*
30.25
Cis-8,11,14eicosatrienoic acid
methyl ester
1.99
C:20:3
*
29.59
Methyl Cis-5,8,11,14Eicosatetraenoic acid
methyl ester
1.99
C:20:4
*
2.01
C:20:5
54.75
Methyl laurate
Methyl
tridecanoate
Methyl myristate
Myristoleic acid
methyl ester
Methyl
pentadecanoate
Cis-10Pentadecenoic acid
methyl ester
Methyl palmitate
Methyl
palmitoleate
Methyl
heptadecanoate
Cis-10heptadecenoic acid
methyl ester
Methyl stearate
Cis-9-oleic methyl
ester
3.98
2.01
3.98
1.99
C:12:0
C:13:0
C:14:0
C:14:1
2.00
C:15:0
35.24
Methyl Cis5,8,11,14,17Eicosapentaenoic acid
1.99
6.02
C:15:1
C:16:0
35.01
40.09
Methyl heneicosanoate
Methyl behenate
2.00
4.00
C:21:0
C:22:0
61.70
65.56
1.99
C:16:1
39.02
2.00
C:22:1
64.55
2.00
C:17:0
44.74
1.99
C:22:2
*
2.04
3.98
C:17:1
C:18:0
43.72
49.23
Methyl erucate
Cis-13,16docosadienoic acid
methyl ester
Cis-4,7,10,13,16,19Docosahexaenoic acid
methyl ester
Methyl tricosanoate
1.99
1.99
C:22:6
C:23:0
*
69.44
3.98
C:18:1
*
Methyl lignocerate
3.99
C:24:0
73.06
Trans-9-elaidic
methyl ester
2.05
C:18:1
48.33 Methyl nervonate
2.00
C:24:1
*
Linolelaidic acid
methyl ester
1.99
C:18:2
*
* C18-C24 unsaturates could not be resolved because a non-polar column was used. ** C:4:0 and C:6:0 fatty acid
methyl esters were not of interest because they were not in the transesterification extract. These compounds eluted
prior to acquisition of MS data using the chromatographic procedure described in Section 2.6.1.
19
Figure 6 is a representative chromatogram of an EI GC-MS analysis of the
C. freiburgensis extract from the Bligh and Dyer method. This method was repeated three times
and there were not any detectable free fatty acids present in any of the analyses. There are some
small peaks present (ex. RT 17.56, 40.37, and 70.68), but based on the NIST MS Library Search
2.0 the peaks were determined to be polysiloxanes from the column because these peaks were
also present in the control runs. The tail end of the chromatogram has poor resolution due to
column bleed from high temperatures similar to the FAME standard chromatogram.
RT: 0.00 - 80.03
79.35 NL:
2.69E8
TIC MS
FAMEChro
mulina1_10
080415342
5
100
90
80
70
78.80
60
50
40
77.35
30
20
70.68
10
3.27 8.06
0
0
10
17.56
20
66.56
59.63
55.29
40.37 50.09
30.09 32.37
30
40
Time (min)
50
75.65
69.77
60
Figure 6: EI Chromatogram of C. freiburgensis extract- Bligh and Dyer Method
70
80
20
Figure 7 is a chromatogram of an EI GC-MS analysis of the C. freiburgensis extract from
the ether reflux method. As in the analysis of the extract using the Bligh and Dyer method, there
were no detectable free fatty acids present. The few small peaks present are concluded to be
polysiloxanes.
RT: 0.00 - 80.03
78.77 NL:
7.08E4
TIC MS
09102010H
E_1009151
52612
100
90
80
70
60
50
75.40
73.91
40
30
3.34
5.48
20
10.99
18.87
11.26
27.47
46.21
52.58
58.62
64.72
61.77
72.55
67.39
10
0
0
10
20
30
40
Time (min)
50
60
70
80
Figure 7: EI Chromatogram of C. freiburgensis extract- Ether Reflux Method
Figures 8 and 9 are representative EI and CI chromatograms for the C. freiburgensis
transesterification extract. The retention times of the FAME’s in the EI and CI analysis were
reproducible. Tetradecanoic acid methyl ester, hexadecanoic acid methyl ester, octadecanoic acid
methly ester, and eicosanoic acid methyl ester and their corresponding unsaturates were the most
21
abundant and will therefore be the only lipids discussed. The other FAME EI and CI
chromatograms and mass spectra that had a relative abundance of at least 2% of the major peak
(C:18:1, RT 48.31) in the chromatogram from the transesterification extraction are listed in
Appendix A.
22
RT: 0.00 - 80.00
48.31
100
NL:
2.47E7
TIC MS
LipidQuantTr
ansesterificat
ion_1104291
30637
C:18:0
90
C:16:0
80
47.98
70
C:14:0
30.42
60
47.14
50
C:20:0
40
40.13
30
20
54.69
55.48
38.99
10
6.83
0
0
15.83
10
20
43.36
35.19
27.39
30
40
Time (min)
56.29
50
66.06
60
79.60
70
80
Figure 8: EI Chromatogram of Transesterification Extract from C. freiburgensis
RT: 0.00 - 80.03
48.31
100
NL:
7.38E5
TIC MS
CITranseste
rification
C:18:0
90
C:16:0
80
C:14:0
70
47.97
47.16
60
30.41
50
C:20:0
40
40.11
30
54.70
20
55.48
38.99
10
15.78 21.35
5.90
0
0
10
20
35.16
30
43.35
40
Time (min)
62.07
50
66.05
60
Figure 9: CI Chromatogram of Transesterification Extract from C. freiburgensis
79.60
70
80
23
3.3. Identification of Lipids
Retention times for each peak varied slightly with each analysis of the transesterification
extract. The volatile compounds were affected most. The variation was likely due to the
reproducibility achievable from manual sample injection. Table 3 compares the retention times
for the C4-C24 FAME (Figure 5) standard to the transesterification extract (Figure 8).
Table 3: C4-C24 Standard Mixture vs. EI Transesterification Extract Lipid Retention Times
Retention
Time
15.78
16.47
20.06
30.41
35.16
38.99
40.11
43.36
47.16
47.39
47.97
48.31
49.19
54.70
55.48
Transesterification
Compounds
C:8:2
C:9:1
C:12:0
C:14:0
C:15:0
C:16:1
C:16:0
C:17:0
C:18:3
C:18:4
C:18:2
C:18:1
C:18:0
C:20:5
C:22:4
Retention
Time
N/A
N/A
20.14
30.25
35.24
39.02
40.09
44.74
46.99
N/A
48.02
48.33
49.23
54.75
57.68
Standard
Compounds
N/A
N/A
C:12:0
C:14:0
C:15:0
C:16:1
C:16:0
C:17:0
C:18:3
N/A
C:18:2
C:18:1
C:18:0
C:20:5
C:20:0
The presence of peaks in the EI transesterification extract chromatogram at
approximately the same retention times as peaks in the EI C4-C24 FAME standard mixture
chromatogram is strong evidence that lipids are present in the transesterification extract. A
CI GC-MS analysis was performed on the transesterification extract in order to elucidate which
compounds were present. The mass spectra for each peak in the chromatograms were analyzed
using Xcalibur version 2.0 software. The mass spectra were used to identify the lipids in the
extracts and an NIST MS Routine Library Search 2.0 routine confirmed the presence of FAMEs.
24
Figures 10 and 11 are the EI chromatograms of retention time 45 to 50 minutes of the
C4-C24 FAME standard and transesterification extract, respectively. All of the C:18 unsaturates
could not be resolved because a non-polar column was used.
C:18:0
C:18:2
RT: 44.85 - 49.97
49.23
100
NL:
7.74E5
TIC MS
FAMESTD
_11032117
3049
48.02
90
80
C:18:1
70
60
C:18:3
50
48.33
40
30
47.72
20
46.99
10
44.95
45.13 45.67
46.24
49.39
48.51
47.24
46.75
0
45
46
47
48
49
Time (min)
Figure 10: EI Chromatogram for the Retention Time that Includes 45-50 minutes from C4-C24 FAME
Standard Mixture
25
C:18:2
C:18:1
RT: 44.97 - 49.97
48.31
100
NL:
2.47E7
TIC MS
LipidQuantTr
ansesterificat
ion_1104291
30637
48.28
90
C:18:4
80
C:18:3
47.98
70
60
47.14
50
47.07
40
C:18:0
47.39
30
20
46.92
10
45.16
0
45.48
45
45.66
46
48.47
46.48
47
48
49.19
49.50
49
Time (min)
Figure 11: EI Chromatogram for the Retention Time that includes 45-50 minutes from the
Transesterification Extract
Figures 12 and 13 are the CI and EI chromatograms of the transesterification extracts,
respectively. There is no significant difference in retention times for octadecanoic acid methyl
ester and its unsaturates.
26
C:18:4
C:18:2
C:18:1
RT: 44.97 - 49.86
48.31
100
C:18:3
NL:
7.38E5
TIC MS
CITranseste
rification
48.27
90
48.23
80
48.20
70
47.97
47.95
47.16
60
47.11
47.91
50
47.04
40
47.86
47.39
C:18:0
30
20
10
45.18 45.47
0
45.0
45.5
46.44
46.0
48.37
46.52
46.5
47.0
47.5
Time (min)
48.0
48.5
49.18
49.0
Figure 12: CI Chromatogram for the Retention Time that includes 45-50 minutes from the
Transesterification Extract
49.53
49.5
27
C:18:2
C:18:1
RT: 44.97 - 49.97
C:18:4
100
C:18:3
90
48.31
NL:
2.47E7
TIC MS
LipidQuantTr
ansesterificat
ion_1104291
30637
48.28
80
47.98
70
60
47.14
C:18:0
50
47.07
40
47.39
30
20
46.92
10
45.16
0
45
45.48
45.66
46
48.47
46.48
47
48
49.19
49.50
49
Time (min)
Figure 13: EI Chromatogram for the Retention Time that includes 45-55 minutes from the
Transesterification Extract
Figure 14 shows the EI mass spectra from the transesterification extract compound
eluting at retention time 49.19 minutes and the C4-C24 FAME standard mixture chromatogram
peak at retention time 49.23 minutes. In EI mass spectrometry, a 70 eV beam of electrons
interact with the sample in the ion trap and gives the sample molecules excess energy [30, 31].
An electron is knocked off the sample molecule forming a positive ion called the molecular ion
(M+). The molecular ion is energentically unstable and further fragments into smaller pieces. The
molecular ion in the figures below is at m/z=298 and the base peak is at m/z=143. The base peak
is the most abundant peak in the spectrum and it represents the most abundant fragmentation ion
(Note: the mass spectrometer was not set to trap and detect ions with m/z less than 100). The
following two spectra (Figure 14) are essentially identical which is strong evidence that there is
C:18:0 fatty acid methyl ester present in the transesterification extract. A library search was done
using NIST Xcalibur software version 2.0 to confirm the presence of C:18:0.
28
Figure 15 is the CI mass spectrum from the transesterification extract at retention time
49.19 minutes. Acetonitrile was used as the reagent gas for all of the CI analyses. When doing CI
analyses, ionized, gaseous acetonitrile is present in abundance in the ion trap and it collides with
the sample molecules producing the molecular ion plus H+ (M+H+), and characteristic adduct ion
(M+CH2CN+) that is equal to the molecular ion plus 40 mass units (M + 40)+. The abundance of
this ion in the spectrum varies with the reagent gas pressure.
Ionized acetonitrile also collides with neutral acetonitrile molecules to produce the
1-methyleneimino-1-ethenylium ion (H2C+NC=CH2) and creates an ion in the spectrum that is
equal to the molecular ion plus 54 mass units (M+54)+ in unsaturated fatty acids [31]. The
(M+H)+ and the (M+40)+ ions are found for the the saturated and unsaturated FAMES. The
(M+54)+ion is only found for the unsaturated FAMES. CI is a lower energy process than EI,
which results in less fragementation compared to the EI spectra. The mass spectrum for the
compound eluting at retention time 49.19 minutes is characteristic of C:18:0 with the M+H+ peak
at m/z=299. The peak at m/z=338 is the (M+40)+ adduct ion peak. CI spectra also produce
molecular weight information that alson confirms the identification of the various FAMES.
29
LipidQuantTransesterification_110429130637 #5689-5694 RT: 49.17-49.21 AV: 6 NL: 6.04E4
T: + c Full ms [100.00-550.00]
143
100
101
199
90
90
80
80
70
60
157
50
185
213
40
129
255
60
50
185
40
30
227
115
20
269
200
250
115
Molecular Ion
227
298
10
270 299
150
241
171
269
298
10
213
129
Molecular Ion
30
0
100
157
241
171
20
199
70
255
Relative Abundance
Relative Abundance
FAMESTD_110321173049 #5634-5648 RT: 49.17-49.27 AV: 15 NL: 5.77E4
T: + c Full ms [100.00-550.00]
101 143
100
300
350 383 415 441 462 489
350
m/z
400
450
500
532 544
550
0
100
270 299
327 354 372 404 430 460
150
200
250
300
350
400
450
502 535
500
550
m/z
Figure 14: Transesterification Mass Spectrum RT 49.19 minutes vs. C4-C24 FAME Standard Mass Spectrum
RT 49.23 minutes
30
CITransesterification #5497-5500 RT: 49.17-49.19 AV: 4 SB: 24 49.01-49.08 , 49.30-49.41 NL: 4.53E3
T: + c Full ms [100.00-550.00]
299
100
(M+H)+
90
80
70
60
50
(M+40)+
40
30
300
20
135
10
0
100
298
247 265
149
191 199 213
296
150
200
250
338
310
339
335
300
350
400
450
500
550
m/z
Figure 15: CI Mass Spectrum of Transesterification Extract at Retention Time 49.19 (C:18:0)
Figure 16 shows the mass spectra from the transesterification extract for the compound
eluting at retention time 48.31 minutes and the C4-C24 FAME standard compound eluting at
retention time 48.33 minutes. These mass spectra are equivalent indicating that there is C:18:1
fatty acid methyl ester present in the transesterification extract. A library search was performed
using Xcalibur software version 2.0 to confirm the presence of C:18:1. The molecular ion is at
m/z=296.
31
LipidQuantTransesterification_110429130637 #5561-5570 RT: 48.23-48.29 AV: 10 NL: 8.89E5
T: + c Full ms [100.00-550.00]
109
100
FAMESTD_110321173049 #5527-5535 RT: 48.30-48.35 AV: 9 NL: 1.09E4
T: + c Full ms [100.00-550.00]
109
100
90
90
121
121
Relative Abundance
70
60
80
123
135
137
50
264
151
60
166
221
30
265
50
166
200
265
222
266
180
250
253
10
296
267 298 343 371 393 412 464 483 501
150
221
30
20
246
Molecular Ion
264
40
180 207
193
10
151
152
235
20
137
207
Molecular Ion
40
0
100
123
70
Relative Abundance
80
300
350
m/z
400
450
500
539
550
0
100
281
150
200
250
296
327 354
300
350
400 428 450 467 503 535
400
450
500
550
m/z
Figure 16: Transesterification Mass Spectrum RT 48.31 vs. C4-C24 FAME Standard Mixture Mass Spectrum
Comparison of Peak at RT 48.33
32
Figure 17 shows the CI mass spectra for the compound eluting at retention time 48.31
minutes in the transesterification extract. The ion at m/z=297 is a result of the molecular ion plus
hydrogen (M+H)+. The ion at m/z=336 is due to the molecular ion plus the adduct ion
(M + CH2CN+). The ion at m/z=350 is the (M+54)+ peak (M+HC+NC=CH2), which is the result
of acetonitrile reacting with itself.31 The presence of these three ions is strong evidence C:18:1 is
present.
CITransesterification #5388-5395 RT: 48.25-48.30 AV: 8 SB: 24 49.01-49.08 , 49.30-49.41 NL: 5.18E4
T: + c Full ms [100.00-550.00]
265
100
M+H+
90
247
80
297
M+40
70
60
M+54
50
135
336
121
40
149
350
30
163
266
20
304
177
191
10
246
296
318
208 245
0
100
351
352 376
150
200
250
300
350
400
428
458
450
502
538
500
m/z
Figure 17: Transesterification CI Mass Spectrum of Retention Time 48.31 Minutes (C:18:1)
550
33
Figure 18 is a comparison of the mass spectrum for the compound eluting in the
transesterification extract at retention time 47.97 minutes to the mass spectrum for the compound
eluting in the C4-C24 FAME standard at retention time 48.02 minutes. The fragmentation pattern
in the two spectra are nearly the same indicating that C:18:1 is present. The molecular ion peak is
at m/z=294 and the base peak is at m/z=135. A library search was also performed to confirm the
presence of oleic acid.
LipidQuantTransesterification_110429130637 #5510-5527 RT: 47.86-47.99 AV: 18 NL: 8.54E5
T: + c Full ms [100.00-550.00]
135
100
FAMESTD_110321173049 #5445-5454 RT: 47.68-47.74 AV: 10 NL: 8.31E3
T: + c Full ms [100.00-550.00]
135
100
90
90
80
80
149
149
70
Relative Abundance
Relative Abundance
70
150
60
Molecular Ion
50
164
40
178
262
60
150
207
50
40
Molecular Ion
164
178
30
30
262
191
20
20
263
187
220
233
10
0
100
150
200
208
10
294
266 296
250
350
300
350
m/z
393 429 450 479 511 537 549
400
450
500
550
0
100
220
150
200
263
253
250
294 327
354 390 400 414 460 476 522 535
300
350
400
450
500
550
m/z
Figure 18: Transesterification Mass Spectrum RT 47.97 minutes vs. C4-C24 FAME Standard Mixture Mass
Spectrum Comparison of Peak at RT 47.72 minutes
34
Figure 19 is the CI mass spectrum for peak at retention time 47.97 minutes in the
chromatogram of the transesterification extract. The ion at m/z=348 is due to the molecular ion
plus 1-methyleneimino-1-ethenylium ion (M+54)+ and the ion at m/z=334 is the molecular ion
plus the adduct ion. The M+H+ ion at m/z=295, the adduct ion at m/z=348, and the
1-methyleneimino-1-ethenylium ion at m/z=348 suggests that C:18:2 is present.
CITransesterification #5346-5354 RT: 47.92-47.98 AV: 9 SB: 24 49.01-49.08 , 49.30-49.41 NL: 2.08E4
T: + c Full ms [100.00-550.00]
M+1
348
100
245
90
263
M+40
80
334
70
60
50
M+54
118
133
135
149
40
163
175
30
264
189
20
190
10
0
100
161
244
302
316
295
243
150
200
349
350
250
300
350
373
400
429 441
450
485
515 529
500
m/z
Figure 19: Transesterification CI Mass Spectrum of Retention Time 47.97 (C:18:2)
550
35
Below, the EI transesterification extract mass spectrum for the compound eluting at
retention time 47.14 minutes is compared to the EI C4-C24 FAME standard spectrum for the
compound eluting at retention time 46.99 minutes (Figure 20). The fragmentation pattern in each
spectrum is very similar. The molecular ion is present at m/z=292 and the base ion at m/z=121.
A library search was performed on the transesterification extract confirming the presence of
C:18:3.
FAMESTD_110321173049 #5358-5364 RT: 46.96-47.01 AV: 7 SB: 60 46.71-46.91 , 47.19-47.48 NL:
T: + c Full ms [100.00-550.00]
105
100
LipidQuantTransesterification_110429130637 #5390-5410 RT: 47.01-47.15 AV: 21 NL: 7.78E5
T: + c Full ms [100.00-550.00]
105
100
90
90
80
80
121
60
135
50
40
30
70
Molecular Ion
Relative Abundance
Relative Abundance
70
147
121
60
50
Molecular Ion
129
135
40
157
30
161
147
157
161
20
20
175
189
10
0
100
203
150
200
189
203
10
221
243
250
292
333 365 390 416 439 478 496 532 548
300
350
m/z
400
450
500
550
0
100
150
200
218 235
250
292 328
340 386 401 415 450
300
350
400
450
492 521 549
500
550
m/z
Figure 20: Transesterification Mass Spectrum RT 47.14 minutes vs. C 4-C24 FAME Standard Mixture Mass
Spectrum Comparison of Peak at RT 46.99 minutes
36
The mass spectrum in Figure 21 is the results of the CI analysis of the transesterification
extract at retention time 47.16 minutes. The M+H+ ion at m/z= 293, the adduct ion at m/z=332,
and the 1-methyleneimino-1-ethenylium ion at m/z=346 suggests that C:18:3 is present.
CITransesterification #5240-5252 RT: 47.07-47.16 AV: 13 SB: 24 49.01-49.08 , 49.30-49.41 NL: 1.93E4
T: + c Full ms [100.00-550.00]
346
100
M+40
M+H+
90
M+54
293
80
118
70
60
243
131
50
135
145
40
261
332
173
30
177
347
187
20
292
294
219
10
291
300
290
0
100
150
200
250
348
300
350
371 397
400
443
450
474 494
523 545
500
550
m/z
Figure 21: Transesterification CI Mass Spectrum of Retention Time 47.14 minutes (C:18:3)
Figures 22 and 23 are the EI and CI mass spectra for the compound eluting at retention
time 47.39 minutes. The M + H+ ion at m/z=291, the adduct ion at m/z=330, and the M+54 ion at
m/z=344 in the CI analysis confirms that there is C:18:4 in the transesterification extract. There
is not C:18:4 present in the C4-C24 FAME standard so a comparison between the EI spectra could
not be made.
37
LipidQuantTransesterification_110429130637 #5435-5446 RT: 47.33-47.41 AV: 12 NL: 8.62E5
T: + c Full ms [100.00-550.00]
105
100
90
80
Molecular Ion
70
119
60
50
40
133
147
30
161
20
171
189
10
197 215
0
100
150
200
261
290 311
250
350 369
300
397
350
400
424 440 472
524 536
450
500
550
m/z
Figure 22: EI Transesterification Mass Spectrum at Retention Time 47.39
CITransesterification #5274-5283
T: + c Full ms [100.00-550.00]
RT: 47.34-47.41
AV: 10
NL: 1.06E4
344
291
100
SB: 24 49.01-49.08 , 49.30-49.41
118
M+54
M+1
90
80
119
70
M+40
60
133
50
40
147
161
175
241
259
30
330
185
189
20
290
345
292
217
260
288
10
0
100
150
200
250
298
346
300
350
369
407
400
435
466
450
503
500
m/z
Figure 23: Transesterification CI Mass Spectrum of Retention Time 47.39 (C:18:4)
520
535
550
38
3.4. Quantification of Lipids
To quantify the lipids in the transesterification extract, an internal standard (naphthalene)
was added to the C4-C24 FAME standard and four serial dilutions were made (see Section 2.7).
The concentrations and areas of the peaks in the C4-C24 FAME standard and the areas of the
peaks in the transesterification extract were used to calculate the concentrations of the fatty acid
methyl esters in the transesterification sample. Below, the FAME standard chromatogram
(Figure 24) and a table (Table 4) corresponding to the pertinent analytes are shown.
RT: 0.00 - 80.03
40.09
100
NL:
9.74E5
TIC MS
FAMESTD
_11032117
79.99 3049
90
49.23
80
57.68
48.02
70
60
65.56
50
30.25
40
20.14
30
35.01
56.55
44.74
61.70
73.06
11.02
20
55.54
29.59
5.06
15.32
10
5.43
0
0
10
20
30
40
Time (min)
50
60
70
Figure 24: EI Chromatogram of C4-C24 Fatty Acid Methyl Ester Standard Mixture
80
39
Table 4: C4-C24 Standard FAME Mixture Areas and Concentrations
RT
(min)
Compound
20.14
29.59
30.25
35.24
39.02
40.09
44.74
46.99
47.72
48.02
48.18
48.33
49.23
54.75
57.68
61.7
C:13:0
C:14:1
C:14:0
C:15:0
C:16:1
C:16:0
C:17:0
C:18:3
C:18:2
C:18:2
C:18:1
C:18:1
C:18:0
C:20:5
C:20:0
C:21:0
Area of
Peak
1.34E+06
6.96E+05
2.18E+06
1.22E+06
8.55E+05
4.75E+06
1.64E+06
7.54E+05
8.98E+05
3.45E+06
1.33E+06
1.49E+06
3.91E+06
4.99E+05
3.96E+06
1.62E+06
Conc. in Amount
standard
on
(mg/ml Column
hexane)
(µg)
2.0
2.0
4.0
2.0
2.0
6.0
2.0
2.0
2.0
4.0
2.0
4.0
4.0
2.0
4.0
2.0
0.2
0.2
0.4
0.2
0.2
0.6
0.2
2.0
2.0
0.4
0.2
0.4
0.4
0.2
0.4
0.2
*The C:18 and C:20 unsaturates were unable to be resolved in the FAME standard. The
exactretention time for each unsaturate is unknown. The concentration of these species will be
estimated based on the RF values for the resolved analytes in the FAME standard.
40
Table 5: C4-C24 Standard FAME with Internal Standard Serial Dilutions
C:12:0
Conc.
Area
(mg/mL)
1.45E+07
0.8
2.45E+06
0.16
4.35E+05
0.032
n/a
0.0064
C:14:1
Dilution
Area
mg/mL
1
8.86E+06
0.4
2
1.43E+06
0.08
3
2.07E+05
0.016
4
n/a
0.0032
C:16:1
Dilution
Area
mg/mL
1
1.17E+07
0.4
2
1.82E+06
0.08
3
2.50E+05
0.016
4
n/a
0.0032
C:18:1 (RT 48.18)
Dilution
Area
mg/mL
1
1.03E+07
0.4
2
2.56E+06
0.08
3
3.47E+05
0.016
4
n/a
0.0032
C:18:3 (RT 46.99)
Dilution
Area
mg/mL
1
1.33E+07
0.4
2
1.86E+06
0.08
3
1.98E+05
0.016
4
n/a
0.0032
C:21:0
Dilution
Area
mg/mL
1
2.78E+07
0.4
2
4.61E+06
0.08
3
5.62E+05
0.016
4
2.29E+04
0.0032
Dilution
1
2
3
4
C:13:0
Conc.
Area
(mg/mL)
1.45E+07
0.4
2.45E+06
0.08
4.35E+05
0.016
n/a
0.0032
C:15:0
Dilution
Area
mg/mL
1
1.34E+07
0.4
2
2.28E+06
0.08
3
3.81E+05
0.016
4
n/a
0.0032
C:17:0
Dilution
Area
mg/mL
1
1.82E+07
0.4
2
3.06E+06
0.08
3
4.70E+05
0.016
4
n/a
0.0032
C:18:1 (RT 48.33)
Dilution
Area
mg/mL
1
1.17E+07
0.8
2
2.79E+06
0.16
3
5.29E+05
0.032
4
2.78E+04
0.0064
C:20:0
Dilution
Area
mg/mL
1
5.40E+07
0.80
2
9.75E+06
0.16
3
1.59E+06
0.03
4
6.97E+04
0.01
Dilution
1
2
3
4
C:14:0
Conc.
Area
(mg/mL)
2.29E+07
0.8
3.94E+06
0.16
7.30E+05
0.032
n/a
0.0064
C:16:0
Dilution
Area
mg/mL
1
5.02E+07
1.20
2
8.68E+06
0.24
3
1.56E+06
0.05
4
7.54E+04
0.01
C:18:0
Dilution
Area
mg/mL
1
4.39E+07
0.80
2
7.61E+06
0.16
3
1.35E+06
0.03
4
5.90E+04
0.01
C:18:2 (RT 47.72)
Dilution
Area
mg/mL
1
1.41E+07
0.4
2
2.21E+06
0.08
3
1.18E+06
0.016
4
n/a
0.0032
C:20:5
Dilution
Area
mg/mL
1
1.35E+07
0.4
2
1.76E+06
0.08
3
1.56E+05
0.016
4
n/a
0.0032
Dilution
1
2
3
4
*5 µL of napthalene was added to Dilution 2, 3, and 4 after the serial dilution.
41
The calibration curves (Figures 25-41) were made via Microsoft® Excel using the
method described in Section 2.7.2. The linearity over the range measured was selected based on
the areas of the peaks in the transesterification extracts. The limit of dectection (LOD) can only
be estimated based on the information obtained from the serial dilutions of the C4-C24 FAME
standard. The LOD for C:12:0 and C:14:0 is greater than 0.0064 mg/mL but less than
0.032 mg/mL. The LOD for C:13:0, C:14:1, C:15:0, C:16:1, C:17:0, and C:20:5 is less than
0.016 mg/mL but greater than 0.0032 mg/mL. The LOD for C:16:0, C:18:0, C:20:0, and C:21:0
is less than 0.0096mg/mL, 0.0064 mg/mL, 0.0064 mg/mL, and 0.0032 mg/mL, respectively.
The calibration curves shown in Figures 25-41 were used to estimate the concentration
of each fatty acid methyl ester (FAME) in the transesterification extact.
42
Figure 25: C4-C24 Standard FAME C:12:0 Calibration Curve
Figure 26: C4-C24 Standard FAME C:13:0 Calibration Curve
43
Figure 27: C4-C24 Standard FAME C:14:0 Calibration Curve
Figure 28 :C4-C24 Standard FAME C:14:1 Calibration Curve
44
Figure 29: C4-C24 Standard FAME C:15:0 Calibration Curve
Figure 30: C4-C24 Standard FAME C:16:0 Calibration Curve
45
Figure 31: C4-C24 Standard FAME C:16:1 Calibration Curve
Figure 32: C4-C24 Standard FAME C:17:0 Calibration Curve
46
Figure 33: C4-C24 Standard FAME C:18:0 Calibration Curve
Figure 34: C4-C24 Standard FAME C:18:1 (RT 48.18 min) Calibration Curve
47
Figure 35: C4-C24 Standard FAME C:18:3 Calibration Curve
Figure 36: C4-C24 Standard FAME C:18:2 (RT 47.72 min) Calibration Curve
48
Figure 37: C4-C24 Standard FAME C:18:2 (RT 48.33 min) Calibration Curve
Figure 38: C4-C24 Standard FAME C:18:3 Calibration Curve
49
Figure 39: C4-C24 Standard FAME C:20:0 Calibration Curve
Figure 40: C4-C24 Standard FAME C:20:5 Calibration Curve
50
Figure 41: C4-C24 Standard FAME C:21:0 Calibration Curve
51
The response factor was calculated for each analyte in all four serial dilutions with the
following equation (2). See Table 6.
RF=(AFA/CFA)/(AIS/CIS)
(2)
RF= Response Factor
AFA= Area of Fatty Acid Methyl Ester
CFA= Concentration of Fatty Acid Methyl Ester
AIS= Area of Internal Standard (Naphthalene)
CIS= Concentration of Internal Standard (Naphthalene)
The use of a response factor was necessary because the mass spectrometer generally will
have a different response to each component analyzed [32]. The concentration of the internal
standard and the FAME may be the same but the peak areas of each analyte may differ. The
known response factor was then used to calculate the concentration of the FAME in the
transesterification extract (Tables 7 and 8).
52
Table 6: C4-C24 Standard FAME with Internal Standard Response Factor Values
C:12:0
Dilution
1
2
3
4
Mean
SD
RF
5.24E-01
2.31E-01
3.69E-01
n/a
3.75E-01
1.47E-01
C:13:0
Dilution
1
2
3
4
Mean
SD
RF
1.05E+00
8.88E-01
7.87E-01
n/a
9.08E-01
1.32E-01
C:14:0
Dilution
1
2
3
4
Mean
SD
RF
8.29E-01
3.70E-01
6.20E-01
n/a
6.06E-01
2.30E-01
C:14:1
Dilution
1
2
3
4
Mean
SD
RF
6.41E-01
6.89E-01
3.75E-01
n/a
5.69E-01
1.69E-01
C:15:0
Dilution
RF
1
9.70E-01
2
8.23E-01
3
6.89E-01
4
n/a
Mean
8.27E-01
SD
1.40E-01
C:16:0
Dilution
RF
1
1.21E+00
2
5.44E-01
3
8.83E-01
4
6.52E-01
Mean
8.22E-01
SD
2.95E-01
C:16:1
Dilution
RF
1
8.46E-01
2
6.58E-01
3
4.53E-01
4
n/a
Mean
6.52E-01
SD
1.96E-01
C:17:0
Dilution
RF
1
1.32E+00
2
5.75E-01
3
7.98E-01
4
n/a
Mean
8.98E-01
SD
3.82E-01
C:18:0
Dilution
RF
1
1.59E+00
2
7.15E-01
3
1.14E+00
4
7.64E-01
Mean
1.05E+00
SD
4.04E-01
C:20:0
Dilution
RF
1
1.95E+00
2
9.16E-01
3
1.35E+00
4
9.03E-01
Mean
1.28E+00
SD
4.94E-01
C:20:5
Dilution
RF
1
9.78E-01
2
2.82E-01
3
1.02E+00
4
n/a
Mean
7.59E-01
SD
4.14E-01
C:21:0
Dilution
RF
1
2.01E+00
2
8.66E-01
3
9.55E-01
4
n/a
Mean 1.28E+00
SD
6.37E-00
C:18:1 (48.18)
Dilution
RF
1
7.44E-01
4.81E-01
2
5.89E-01
3
n/a
4
Mean
6.04E-01
SD
1.32E-01
C:18:1 (48.33)
Dilution
RF
1
4.23E-01
2.62E-01
2
4.49E-01
3
3.61E-01
4
Mean
3.73E-01
SD
8.31E-02
C:18:2 (48.02)
Dilution
RF
1
3.05E+00
4.81E-01
2
5.89E-01
3
7.21E-01
4
Mean
1.21E+00
SD
1.23E+00
C:18:2 (47.72)
Dilution
RF
1
1.02E+00
4.15E-01
2
2.01E+00
3
n/a
4
Mean 1.15E+00
SD
0.803975
53
Calculation of Concentration of Fatty Acid for C:18:0
RF=(AFA/CFA)/(AIS/CIS)
1.05=(3.70E6/CFA)/(1.73E5/5.01E-3)
CFA=9.39E-2
9.39E-2 mg/mL*1.0 ml of hexane =9.39E-2 mg C:18:0/1252.9 mg algae
Weight % = 0.00749 %
Table 7: Concentration of Fatty Acids in Transesterification Sample
Cmpd
C:14:0
C:15:0
C:16:1
C:16:0
C:17:0
C:18:3
C:18:4
C:18:2
C:18:1
C:18:0
C:20:5
C:21:0
Apex RT
30.43
35.17
38.99
40.13
43.35
47.15
47.4
47.99
48.32
49.19
54.71
61.84
Area
1.11E+08
1.84E+06
1.77E+07
5.91E+07
4.01E+06
1.28E+08
5.34E+07
1.54E+08
2.14E+08
3.70E+06
3.56E+07
2.60E+04
Conc.
%Area
%Height (mg/mL)
13.03
11.96 4.88E+00
0.22
0.35 5.92E-02
2.08
2.83 7.22E-01
6.94
8.24 1.92E+00
0.47
0.67 1.19E-01
15.03
12.72
*
6.28
8.07
*
18.07
15.26
*
25.15
22.47
*
0.44
0.69 9.39E-02
4.18
5.59 1.25E+00
0.00
0.10 5.42E-04
Wt %
3.89E-01
4.72E-03
5.77E-02
1.53E-01
9.51E-03
*
*
*
*
7.49E-03
9.98E-02
4.33E-05
Table 8: Estimated Concentration of Fatty Acids in Transesterification Sample
Cmpd
C:18:3
C:18:4
C:18:2
C:18:1
Apex
RT
47.15
47.4
47.99
48.32
Area
1.28E+08
5.34E+07
1.54E+08
2.14E+08
Conc.
%Area
%Height (mg/mL)
15.03
12.72 3.41E+00
6.28
8.07 1.43E+00
18.07
15.26 4.10E+00
25.15
22.47 5.71E+00
Wt %
2.72E-01
1.14E-01
3.27E-01
4.56E-01
*The concentrations for these compounds were estimated using an RF value of 1.00.
54
The composition of high quality microalgal biodiesels should have an equal mixture of
saturated and unsaturated fatty acids to maintain high oxidative stability and low temperature
property [4]. Also, according to European Standards for quality biodiesel, the linolenic acid
(C:18:3) content cannot exceed 12% of the total fatty acids [4]. The linolenic acid limit is not
met and unfortunately the amount of lipids produced using the growing conditions described in
Section 2.2 are too minute for Chromulina to be considered a good source of biodiesel.
Two different methods were used to dry the same batch of algae, a drying oven and a
freeze dryer. The transesterification method described in Section 2.2.3 was used to extract the
lipids from each sample. The oven dried (OD) and freeze dried (FD) transesterification extracts
were analyzed via GC-MS with the parameters described in section 2.2.1. Figures 42 and 43 are
the EI chromatograms for the OD and FD transesterification extracts, respectively. The retention
times (RT) of the FAME were the same in the GC-MS analysis but the relative abundance of
each compound differed. The relative abundance of C:14:0, C:15:0, C:16:0, and C:16:1 was
decreased but the C:18 unsaturates and C:20:5 relative abundance was increased in the FD EI
analysis. The CI analysis illustrated similar results (see Figure 44 and Figure 45). Table 9 is the
ratio of the area and height of each FAME from the freeze dry to the oven dry method.
Table 9: Ratio of Freeze Dry (FD) Peak Area and Height vs. Oven Dry (OD) Peak Area and Height
Compound
C:14:0
C:15:0
C:16:1
C:16:0
C:18:3
C;18:2
C:18:1
C:20:5
FD/OD RT
FDa/ODa FDh/ODh
30.32/30.23
0.91
0.78
35.16/35.37
0.28
0.26
38.95/38.93
0.87
0.77
40.10/40.05
0.81
0.66
47.09/46.95
1.23
0.87
47.36/47.21
2.03
1.67
47.75/47.67
1.40
0.78
54.67/54.60
1.55
1.35
55
RT: 0.00 - 80.03
40.05
100
NL:
2.11E6
TIC MS
EITranseste
rification2_
110712104
644
46.95
90
30.23
80
47.67
70
60
47.94
50
40
30
79.54
54.60
38.93
20
35.37
10
55.67
61.43
66.83
27.36
6.74
5.46
42.69
10.93
18.79
0
0
10
20
30
40
Time (min)
50
60
70
80
Figure 42: EI Chromatogram of Transesterification Sample from Oven Dried C. freiburgensis
RT: 0.00 - 80.01
47.09
100
NL:
3.66E6
TIC MS
EITranseste
rification3
47.36
90
80
40.10
30.32
70
47.75
60
50
48.01
54.67
40
30
20
38.95
46.53
10
4.59
0
0
6.75 15.77
10
20
62.08
64.38
55.69
35.16
24.56
30
40
Time (min)
50
60
70
Figure 43: EI Chromatogram of Transesterification Sample from Freeze Dried C. freiburgensis
79.58
80
56
RT: 0.00 - 80.03
30.25
100
NL:
1.03E3
TIC MS
CITranseste
rification3
46.94
90
80
70
40.05
60
50
47.67
40
47.95
30
20
54.63
10
38.95
6.75
27.38
35.39
42.66
77.71
63.82
66.83
75.02
0
0
10
20
30
40
Time (min)
50
60
70
80
Figure 44: CI Chromatogram of Transesterification Sample from Oven Dried C. freiburgensis
RT: 0.00 - 80.05
47.12
100
30.35
NL:
2.38E4
TIC MS
CITranseste
rification2
47.40
90
80
40.12
70
60
47.78
50
48.04
40
30
38.97
20
54.69
10
46.54
3.25
4.59
15.77
35.17
24.56
62.08
0
0
10
20
30
40
Time (min)
50
60
74.88
77.72
70
Figure 45: CI Chromatogram of Transesterification Sample from Freeze Dried C. freiburgensis
80
57
4. Conclusion
C. freiburgensis was isolated from the Berkeley Pit in Butte, Montana by Grant Mitman
and successfully grown in MAM in pure culture. Chromulina does not appear to produce any
free fatty acids under the growing conditions described in Section 2.2.1. This conclusion is based
on not finding any detectable FAMEs from the Bligh and Dyer and Ether extraction methods.
Methyl myristate (C:14:0), pentadecanoate (C:15:0), palmitate (C:16:0), palmitoleate (C:16:1),
heptadecanoate (C:17:0), stearate (C:18:0), oleate (C:18:1), linoleate (C:18:2), linoleneate
(C:18:3), arachidate (C:20:0), and eicosapentaenoate (C:20:5) were extracted using the
transesterification method. The FAMES were identified and quantified by comparison to a
C4-C24 FAME standard.
Using the growing conditions in this study, C. freiburgensis did not produce enough fatty
acid methyl esters to be used as a biofuel source. Microalgae that are currently being used to
make biodiesel have a lipid content that is at least 25 % of their dry weight [4]. The most
abundant lipids produced were methyl myristate (C:14:0), palmitic acid methyl ester (C:16:0),
and saturated and unsaturated octadecanoic acid methyl esters (C:18:0, C:18:1, C:18:2, C:18:3,
C:18:4). Most microalgal biodiesels are composed of unsaturated fatty acids such as
palmitoleate, linoleate, and linoleneate and saturated fatty acids such as palmitate and oleate
[33].
58
5. Future Work
The algal stocks used in this work were obtained from a mixture of organisms growing in
Berkeley Pit water. The date of sample water extraction from the Berkeley Pit is unknown. A
new sample must be collected from the waters of the Pit. Proper culturing techniques should be
employed and freezer stocks should be made to ensure the cells are not continually changing.
The genetics of the new Chromulina stock must to be compared to the previous sample to make
sure the organism has not mutated since its original extraction.
The preliminary data (not shown) obtained from the polysaccharide extraction was
promising. Further extraction, identification, and quantification of polysaccharides is crucial in
determining the potential of C. freiburgensis as a source for biofuels. Polysaccharides have a
high molecular weight; therefore, the polysaccharide extracts need to be analyzed via
electrospray ionization.
Varying the growth conditions of C. freiburgensis to determine any resulting change in
composition needs to be further explored. Changing the temperature and decreasing the nitrogen
and phosphorous concentrations in the growth media may produce a larger quantity or alter the
distribution of lipids [4]. Chromulina should also be grown in large scale and analyzed to
determine any differences in lipid and polysaccharide production.
Lastly, repeat analyses of the freeze dry method should be performed. The data obtained
thus far indicate that the drying method does not have a significant effect on lipid quantities but
should be further investigated.
59
References Cited
1.
Huang, G., et al., Biodiesel Production by Microalgal Biotechnology. Applied Energy,
2010. 87: p. 38-46.
2.
Hahn-Hagerdal, B., et al., Bioethanol-the fuel of tomorrow from the residues of today.
TRENDS in Biotechnology, 2006. 24 (12): p. 549-556.
3.
Hongjin, Q. and W. Guangce, Effect of carbon source on growth and lipid accumulation
in Chlorella sorokiniana. Chinese Journal of Oceanology and Linmology, 2009. 27:
p. 762-768.
4.
Mutanda, T., S. Karthikeyan, and F. Bux, The utilization of post-chlorinated municipal
domestic wastewater for biomass and lipid production by Chlorella spp. under batch
conditions. Applied Biochemistry and Biotechnology, 2011. 164(7): p. 1126-1138.
5.
Oilgae. 2011 [cited 2011 February 23]; Available from:
http://www.oilgae.com/algae/algae.html.
6.
Wu, L.F., et al., The feasibility of biodiesel production by microalgae using industrial
wastewater. Bioresource Technology, 2012. 113: p. 14-18.
7.
Algae Biofuels for the Future. [cited 2011 February 23]; Available from:
http://www.alternative-energy-news.info/algae-biofuels-future/.
8.
El-Sarraf, W.M. and G. El-Shaarawy, Chemical composition of some marine algae from
the Mediterranean Sear of Alexandria, Egypt. Bulletin of H.I.P.H., 1994. 24 (3):
p. 523-534.
9.
United States Energy Information.Biofuels:Ethanol and Biodiesel Explained. 2011
[cited 2011 June 10]; Available from:
http://www.eia.gov/energyexplained/index.cfm?page=biofuel_home#tab2.
10.
Smithsonian National Museum of Natural History: Algae Research. 2011 [cited 2011
February 23].
11.
Wigmosta, M.S., et al., National Microalgae Biofuel Production Potential and Resource
Demand. Water Resources Research, 2011. 47: p. 1-13.
12.
Cheng, J.J. and G.R. Timilsina, Status and barriers of advanced biofuel technologies: A
review. Renewable Energy, 2011. 36: p. 3541-3549.
13.
Hill, J., et al., Environmental, economic, and energetic costs and benefits of biodiesel and
ethanol biofuels. Proc Natl Acad Sci U S A, 2006. 103 (30): p. 11206-10.
60
14.
Mitman, G., Algal Bioremediation of the Berkeley Pit Lake. 1998, Montana Tech of the
University of Montana: Butte, Montana.
15.
Alberts, B., et al., Molecular Biology of the Cell. 4th ed. 2002, New York: Garland
Science. 1463.
16.
Campbell, N.A. and J.B. Reece, Biology. 6th ed. 2002, San Francisco: Pearson Education
Inc. 1249.
17.
Merchant, S.S., The Chlamydomonas Genome Reveals the Evolution of Key Animal and
Plant Functions. Science, 2007. 318 (245): p. 245-251.
18.
Merchant, S.S., et al., TAG, You're It! Chlamydomonas as a reference organism for
understanding algal triacylglycerol accumulation. Current Opinion in Biotechnology,
2011. 23: p. 1-12.
19.
Harwood, J.L. and I.A. Guschina, The versatility of algae and their lipid metabolism.
Biochimie, 2009. 91: p. 679-684.
20.
Dakel, S.M., Bioremediative Potential of Chromuline freiburgensis in Culture from the
Berkeley Pit, in Biological Sciences. 2001, Montana Tech of the University of Montana:
Butte.
21.
Mitman, G., Biological Survey of the Berkeley Pit Lake System, in Mine Waste
Technology Program. 1998, Montana Tech of the University of Montana: Butte,
Montana.
22.
Biology Reference: Algae. [cited 2011 February 23]; Biology Encyclopedia]. Available
from: http://www.biologyreference.com/A-Ar/Algae.html.
23.
Stewart, A., A. Mondloch, Editor. 2009, Montana Tech of the University of Montana:
Butte.
24.
Olaveson, M.M. and P.M. Stokes, Responses of the acidophillic alga Euglena mutabilis
(Euglenophyceae) to carbon enrichment at pH 3. Journal of Phycology, 1989. 25:
p. 529-539.
25.
Bold's Basal Medium. The Provasoli-Guillard National center for Culture of Marine
Phytoplankton [cited 2011 June 9]; Available from:
https://ccmp.bigelow.org/index.php?q=node/71.
26.
Xu, D.J., et al., Molecular weight and monosaccaride composition of Astragalus
polysaccharides. Molecules, 2008. 13: p. 2408-2415.
61
27.
Bligh, E.G. and W.J. Dyer, A rapid method of total lipid extraction and purification.
Canadian Biochem. Physiol., 1959. 37: p. 911-917.
28.
Ehimen, E.A., Z.F. Sun, and C.G. Carrington, Variables affecting the in situ
transesterification of microalgal lipids. Fuel, 2010. 89 (677-684).
29.
Yu, G., et al., A comparative analysis of four kinds of polysaccarides purified from
Furcellaria lumbricalis. Journal of Ocean University China, 2007. 6: p. 16-20.
30.
Pelt, C.K.V., J.T. Brenna, and B.K. Carpenter, Studies of structure and mechanism in
acetonitrile chemical ioniation tandem mass spectrometry of polyunsaturated fatty acid
methyl esters. J Am Soc Mass Spectrum, 1999. 10: p. 1253-1262.
31.
Pelt, C.V. and J.T. Brenna, Acetonitrile chemical ionization tandem mass spectrometry to
locate double bonds in polyunsaturated fatty acid methyl esters. Ananlytical Chemistry,
1999. 71: p. 1981-1989.
32.
Sobrino, F.H., C.R. Monroy, and J.L. Perez, Biofeuls and fossil fuels: Life cycle analysis
(LCA) optimisation through productive resources maximization. Renewable and
Sustainable Energy Reviews, 2011. 15: p. 2621-2628.
33.
Constenoble, O., et al., Improvements needed for the biodiesel standard EN 14214.
Bioscopes, 2008: p. 1-171.
62
Appendix A:
Table A.1: CI Transesterification Peaks
CI Transesterification
Apex
RT
15.78
21.35
30.41
35.16
38.99
40.11
43.35
54.70
55.48
66.05
Start RT
15.73
21.30
30.12
35.09
38.84
39.78
43.25
54.48
55.29
65.94
End
RT
15.92
21.49
30.47
35.26
39.18
40.19
43.45
54.80
55.59
66.16
Area
%Area
37888.02
0.14
20654.50
0.07
3687728.21 13.31
45987.22
0.17
681681.61
2.46
1682383.41
6.07
81426.06
0.29
1128524.73
4.07
700241.23
2.53
65359.31
0.24
Height
9532.55
4523.78
399801.24
9862.62
116305.02
230360.11
15295.98
181414.27
119225.03
12092.74
%Height
0.31
0.15
12.96
0.32
3.77
7.47
0.50
5.88
3.86
0.39
Table A.2: EI Transesterification Peaks
EI Transesterification
Apex
RT
15.83
21.59
30.42
35.19
38.99
40.13
43.36
54.69
55.48
66.06
Start
RT
15.79
21.50
30.16
35.12
38.85
39.80
43.26
54.52
55.32
65.94
End
RT
16.09
21.76
30.47
35.33
39.19
40.24
43.49
54.87
55.80
66.28
Area
%Area
184314.43
0.02
55766.07
0.01
126072218.41 16.44
1232407.67
0.16
14174312.10
1.85
54765377.26
7.14
3292439.66
0.43
25349494.14
3.31
16940300.10
2.21
2301768.79
0.30
Height
%Height
27069.38
0.03
5647.72
0.01
14375315.70
14.93
246290.08
0.26
2567944.60
2.67
8601021.65
8.93
588135.25
0.61
4177992.63
4.34
2799950.64
2.91
382243.54
0.40
63
FAMESTD_110321173049 #3322-3334 RT: 30.22-30.30 AV: 13 SB: 35 30.06-30.15 , 30.48-30.66 NL:
T: + c Full ms [100.00-550.00]
101
100
143
90
80
199
70
60
50
157
40
185
129
30
20
171
115
213
10
214 242
0
100
150
200
250
267 293 315
300
356 377 406 432
350
400
462 486
450
500
m/z
Figure A.1: FAME Standard EI Mass Spectrum at RT 30.25 minutes (C:14:0)
528 548
550
64
LipidQuantTransesterification_110429130637 #3306-3331 RT: 30.26-30.43 AV: 26 SB: 116
T: + c Full ms [100.00-550.00]
143
100 101
90
199
80
70
60
50
157
185
40
129
30
171
20
213
115
10
0
100
243
214
244 271
150
200
250
320 344
300
350
389
400
432 445 473 502
450
500
m/z
Figure A.2: Transesterification EI Mass Spectrum at RT 30.14 minutes (C:14:0)
533
550
65
CITransesterification #3234-3256 RT: 30.21-30.38 AV: 23 SB: 279 28.71-29.80 , 30.59-31.84 NL: 6.51E4
T: + c Full ms [100.00-550.00]
243
100
90
80
70
60
50
40
30
20
103
117
10
0
100
244
135
149
150
191
242
209
282
254
281
157
200
250
283
285
329
300
368 382
350
425 442
400
450
477
518 535
500
m/z
Figure A.3: Transesterification CI Mass Spectrum at RT 30.41 minutes (C:14:0)
550
66
FAMESTD_110321173049 #3934-3945 RT: 35.21-35.29 AV: 12 SB: 47 35.01-35.16 , 35.44-35.66 NL:
T: + c Full ms [100.00-550.00]
101
100
90
80
143
70
60
213
157
50
199
40
171
129
30
20
115
227
10
0
100
256
258
150
200
250
329 341
300
350
384 401 430
400
460 476
450
512
500
m/z
Figure A.4: FAME Standard EI Mass Spectrum at RT 35.24 minutes (C:15:0)
541
550
67
LipidQuantTransesterification_110429130637 #3906-3914 RT: 35.15-35.21 AV: 9 SB: 44
T: + c Full ms [100.00-550.00]
101
100
90
143
80
70
213
157
60
199
50
40
171
129
30
20
115
227
10
0
100
256
258
150
200
250
304 328
300
368 388 404
350
400
455 487 517 529
450
500
m/z
Figure A.5: Transesterification EI Mass Spectrum at RT 35.19 minutes (C:15:0)
550
68
CITransesterification #3821-3828 RT: 35.14-35.20 AV: 8 SB: 38 34.85-34.97 , 35.33-35.52 NL: 2.48E3
T: + c Full ms [100.00-550.00]
257
100
90
80
70
60
50
40
30
20
258
103
10
0
100
135
149
163
150
205
199
200
256
255
236
250
268
296
295
297
298
300
350
400
450
500
m/z
Figure A.6: Transesterification CI Mass Spectrum at RT 35.16 minutes (C:15:0)
550
69
FAMESTD_110321173049 #4521-4531 RT: 40.04-40.11 AV: 11 SB: 43 38.81-38.93 , 39.17-39.38 NL:
T: + c Full ms [100.00-550.00]
101
100
90
143
80
70
227
60
171
50
185
199
40
30
129
157
115
213
241
20
10
0
100
270
271 294 331 343
255
150
200
250
300
350
389
400
428 450 476 502 516 538
450
500
m/z
Figure A.7: FAME Standard EI Mass Spectrum at RT 39.02 minutes (C:16:0)
550
70
LipidQuantTransesterification_110429130637 #4376-4387 RT: 38.95-39.03 AV: 12 SB: 44
T: + c Full ms [100.00-550.00]
109
100
123
90
80
70
60
137
148
50
152
236
40
30
165
193
194
237
20
218
10
0
100
238
268
239
295 327 349
150
200
250
300
350
384 401 427 454
400
450
489 505 529 543
500
m/z
Figure A.8: Transesterification EI Mass Spectrum at RT 38.99 minutes (C:16:1)
550
71
CITransesterification #4271-4282 RT: 38.93-39.02 AV: 12 SB: 110 38.24-38.78 , 39.17-39.54 NL: 7.84E3
T: + c Full ms [100.00-550.00]
219 237
100
90
80
269
70
60
135
50
121
40
30
149
308
163
322
20
238
177 218
180
10
0
100
268
276
290
323
347
150
200
250
300
350
384
400
422 439
450
505 528
500
m/z
Figure A.9: Transesterification CI Mass Spectrum at RT 38.99 minutes (C:16:1)
550
72
FAMESTD_110321173049 #4521-4531 RT: 40.04-40.11 AV: 11 SB: 43 38.81-38.93 , 39.17-39.38 NL:
T: + c Full ms [100.00-550.00]
101
100
90
143
80
70
227
60
171
50
185
199
40
30
129
157
115
213
241
20
10
0
100
270
271 294 331 343
255
150
200
250
300
350
389
400
428 450 476 502 516 538
450
500
m/z
Figure A.10: FAME Standard EI Mass Spectrum at RT 40.09 minutes (C:16:0)
550
73
LipidQuantTransesterification_110429130637 #4520-4534 RT: 40.05-40.14 AV: 15 SB: 48
T: + c Full ms [100.00-550.00]
101
100
90
143
80
70
227
171
60
185 199
50
157
129
40
213
30
115
241
20
10
0
100
270
243
150
200
250
272 312 333
300
350
380 402 414
400
453
450
499 521 532
500
m/z
Figure A.11: Transesterification EI Mass Spectrum at RT 40.13 minutes (C:16:0)
550
74
CITransesterification #4401-4417 RT: 40.02-40.14 AV: 17 SB: 179 39.24-39.59 , 40.30-41.44 NL: 4.61E4
T: + c Full ms [100.00-550.00]
271
100
90
80
70
60
50
40
30
20
10
272
103
0
100
219
135 149
171 199
150
200
270
250
250
310
282
311
308 313
300
367 381
350
400
439 451
450
490 529 540
500
m/z
Figure A.12: Transesterification CI Mass Spectrum at RT 40.11 minutes (C:16:0)
550
75
LipidQuantTransesterification_110429130637 #4929-4934 RT: 43.33-43.37 AV: 6 SB: 28
T: + c Full ms [100.00-550.00]
101
100
143
90
185 199
80
227
70
241
255
60
50
115
40
129
157
30
171
20
213
256
10
0
100
284
269 285
150
200
250
340 365
400
350
400
300
450 462 485 519 538
450
500
m/z
Figure A.13: Transesterification EI Mass Spectrum at RT 43.36 minutes (C:17:0)
550
76
CITransesterification #4796-4798 RT: 43.34-43.36 AV: 3 SB: 179 39.24-39.59 , 40.30-41.44 NL: 4.38E3
T: + c Full ms [100.00-550.00]
285
100
90
80
70
60
50
40
30
20
10
0
100
286
135 149
150
163
233
227
199
200
283
255
250
324
296
325
300
350
400
450
500
m/z
Figure A.14: Transesterification CI Mass Spectrum at RT 43.35 minutes (C:17:0)
550
77
FAMESTD_110321173049 #6296-6307 RT: 54.71-54.79 AV: 12 SB: 49 54.45-54.64 , 54.91-55.11 NL:
T: + c Full ms [100.00-550.00]
104.89
100
90
80
70
60
118.88
50
132.85
40
30
146.85
20
174.85
202.84
10
214.82
0
100
150
200
266.64
250
315.65 345.48 376.58
300
350
400
450.45
450
503.02 538.18
500
m/z
Figure A.15: FAME Standard EI Mass Spectrum at RT 54.54 minutes (C:20:5)
550
78
LipidQuantTransesterification_110429130637 #6368-6381 RT: 54.63-54.72 AV: 14 SB: 60
T: + c Full ms [100.00-550.00]
105
100
90
80
70
60
119
133
50
40
147
30
20
175
187
10
0
100
201
247 261 289 318
341
150
200
250
300
350
382
427 442 460
400
450
496 517
500
m/z
Figure A.16: Transesterification EI Mass Spectrum at RT 54.69 minutes (C:20:5)
545
550
79
CITransesterification #6140-6158 RT: 54.59-54.74 AV: 19 SB: 116 53.63-54.29 , 54.93-55.24 NL: 6.11E3
T: + c Full ms [100.00-550.00]
372
100
319
90
80
70
60
50
118
119
131
40
269
287
147
358
171
30
203
189
20
373
320
245
235
10
318
288
316
0
100
150
200
250
326
300
350
374 397
400
441 471 503 515
450
500
m/z
Figure A.17: Transesterification CI Mass Spectrum at RT 54.70 minutes (C:20:5)
550
80
RT: 38.44 - 53.01
49.45
100
48.29
40.33
90
NL:
2.51E7
TIC MS
LipidStdNa
phthalene
80
70
60
44.84
50
40
47.07
43.81
30
39.10
20
10
41.91
0
40
42
49.78 51.16
45.69
44
46
Time (min)
48
Figure A.18: EI Chromatogram of C:14-C:20
50
52
81
Figure A.21: C. freiburgensis in three different media; Left- BP water & NaNO3, Center- MAM, RightAcidified Bold Basal Medium
82
Figure A.22: C. freiburgensis in MAM in culture chamber.
83
Table A.3: Modified Acid Medium Recipe
Modified Acid Medium (MAM)
Chemical
Formula
M.W.
Macronutrients
Stock
Solution
weight
(g/L)
Final
Concentration
addition
(mL/L)
mg/L
mM
Ammonium sulphate
(NH4)SO2
131.70
20.00
10.00
200.46
1.52
Calcium chloride
CaCl2 *2(H2O)
146.99
0.80
10.00
7.99
0.06
Magnesium sulphate
MgSO4*7(H2O)
246.36
50.00
10.00
499.91
2.03
Potassium phosphate
KH2PO4
139.09
30.00
1.00
29.99
0.22
Sodium chloride
NaCl
58.44
2.40
10.00
23.99
0.41
372.24
1.60
10.00
16.00
0.04
174.25
19.20
9.00
172.80
1.10
1145.57
10.00
1.00
10.22
0.01
61.83
10.00
1.00
10.00
0.16
Na-EDTA
Potassium sulphate
K2SO4
Micronutrients
Ammonium molybdate
(NH4)6Mo7O24*4(H2O)
Boric Acid
H3BO3
Colbalt chloride
CoSO4*6(H2O)
237.90
30.00
1.00
29.89
0.13
Copper sulphate
Ferrous ammonium
sulphate
CuSO4*5(H2O)
249.81
60.00
1.00
59.99
0.24
Fe(NH4)2(SO4)2*6(H2O)
392.04
1170.00
1.00
1169.42
2.93
Manganese sulphate
MnSO4*4(H2O)
223.05
690.00
1.00
689.66
3.09
Sodium vanadate
NaVO3
121.93
2.00
1.00
2.00
0.02
Zinc sulphate
ZnSO4*7(H2O)
287.54
370.00
1.00
370.07
1.29
337.27
50.00
0.67
100.00
0.30
1.30
1.00
Vitamins
Thiamine-HCl B1
Cyanocobalamin B12
*Combine 10 ml of each micronutrient to make a stock solution. Add 10 ml of stock micronutrients
solution to a 9476 ml of de-ionized water. Add 10 ml of each macronutrient except KH2PO4 and K2SO4.
Add 10 ml of KH2PO4 and 90 ml of K2SO4. Weigh 0.03 g of B1 and add to solution. Add 13.0 ml of B 12 to
bottle and mix well.
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