1. 1
CHARACTERIZATION OF BIOLOGICAL MACROMOLECULES
FROM Chara vulgaris, Enteromorpha prolifera and
Microspora sp.
PROJECT SUBMITTED TO THE UNIVERSITY OF MUMBAI IN
REQUIREMENT OF PARTIAL FULFILMENT FOR DEGREE OF M.Sc.
(BY PAPER) IN LIFE SCIENCE (BIOTECHNOLOGY)
BY
JAY SUNIL PATHARE
M.Sc. PART II
YEAR 2012-13
UNDER THE GUIDANCE OF
PROF. Dr. Ganesh Iyer
DEPARTMENT OF LIFE SCIENCE
RAMNRAIN RUIA COLLEGE
MATUNGA, MUMBAI-400019

2. 2
Certificate
Department of Life Sciences,
Ramnarain Ruia College,
Matunga.
This is to certify that Mr. Jay Sunil Pathare, a student of this department,
bearing the seat no. 6677 has successfully completed his project on
“CHARACTERIZATION OF BIOLOGICAL MACROMOLECULES
FROM Chara vulgaris, Enteromorpha prolifera and Microspora sp.”,
towards the completion of MSc Part II examination in Life Sciences with a
specialization in Biotechnology, for the academic year 2012-2013 under my
guidance and supervision.
Dr. Ganesh Iyer
(Guide Supervisor)
(Head of the Department)
Place: Mumbai
Date:

3. 3
Acknowledgement.
The work presented in this thesis was carried out under the guidance of Dr Ganesh Iyer. I
express my gratitude to him for his motivation, encouragement and support during all
stages of this work. I take this opportunity to thank Mr. Yogesh Pawar, for providing me
the required facilities and an opportunity to undertake this unique and innovative project,
with special mention of Dr. Seema Menon. . I would also like to thank Ms Samruddha
Phadnis and Mr. Yash Gupte for their assistance and help offered to me at various stages
of my research work. I am also thankful to all my friends for their help and support. I
wish to thank all my colleagues, academic, scientific and administrative staff at
Ramnarain Ruia College for their ready help and willing cooperation at all times, with a
special mention to Mr. Pawar, Mr. Mahendra Pednekar, Mr. Baliram Adbal.
Last but not the least my parents the true sculptors of my life.

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Table of Contents
Table of figures...................................................................................................... 5
1. Abstract.............................................................................................................. 8
2. Introduction to Algae. ........................................................................................ 9
3. Introduction to Algae – Classification...............................................................12
PHYLUM CYANOPHYTA (Blue-Green
Algae/Cyanobacteria) ........................................................................................14
PHYLUM RHODOPHYTA................................................................................16
PHYLUM EUGLENOPHYTA ........................................................................18
PHYLUM CRYPTOPHYTA...............................................................................19
PHYLUM PYRROPHYTA .................................................................................21
PHYLUM RAPHIDOPHYTA .............................................................................22
PHYLUM CHRYSOPHYTA...............................................................................23
PHYLUM XANTHOPHYTA ..............................................................................25
PHYLUM CHLOROPHYTA..............................................................................27
PHYLUM EUSTIGMATOPHYTA .....................................................................29
PHYLUM PHAEOPHYTA ................................................................................31
PHYLUM PRASINOPHYTA .............................................................................33
PHYLUM GLAUCOPHYTA..............................................................................34
5. Purpose for the Project......................................................................................36
6. The Need for Research......................................................................................37
7. The Need for Research...................................................................................38
Work done by other Researchers ....................................................................39
8. Algae and their many uses ................................................................................44
9. Details about the Algal Species used in the Project. ..........................................48
Microspora species............................................................................................49
Enteromorpha Prolifera.....................................................................................52
Chara Vulgaris..................................................................................................56

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10. MATERIAL AND METHOD.........................................................................60
Extraction and Estimation of Lipids from Algal Culture. ...................................60
Transesterification of Lipids ..............................................................................64
Nile Red Staining...............................................................................................71
Extraction and Estimation of Protiens from the Algal Culture. ........................... 2
Folin Lowry’s Protein Assay .............................................................................. 3
Sodium Dodecyl Sulfate - Polyacrylamide Gel Electrophoresis (SDS – PAGE) . 7
Extraction and estimation of Carbohydrates using Phenol – Sulphuric Assay.14
11. Conclusion......................................................................................................16
12. Discussion.......................................................................................................17
13. Bibliography ...................................................................................................18

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Table of figures
Figure 1 Nostoc Pruniforme and Figure 2 Cylindrospermum _______________ 14
Figure 3 Polysiphonia ______________________________________________ 16
Figure 4 Some red algae ____________________________________________ 17
Figure 5 Some euglenoids ___________________________________________ 18
Figure 6 Rhodomonas salina_________________________________________ 19
Figure 7 Ceratium Furca____________________________________________ 21
Figure 8 Gonyostomum _____________________________________________ 22
Figure 9 Dinobryon divergens________________________________________ 23
Figure 10 Xanthophyceae ___________________________________________ 25
Figure 11 Xanthophyceae ___________________________________________ 26
Figure 12 Caulerpa racemosa________________________________________ 27
Figure 13 Caulerpa filiformis ________________________________________ 28
Figure 14 Nannochloropsis __________________________________________ 29
Figure 15 Brown algae _____________________________________________ 31
Figure 16 Macrocystis Pyrifera_______________________________________ 32
Figure 17 Glaucocystis Species_______________________________________ 34
Figure 18 World Population Growth Chart (1960- 2011) __________________ 36
Figure 19 Average Diesel Price in Dollars (World) _______________________ 37
Figure 20 Area under Agriculture (Sq. kms.) ____________________________ 38
Figure 21 Microspora at 100 X magnification ___________________________ 50
Figure 22 Microspora at 400 X magnification. Microscope: Labomed Lx 300 __ 50
Figure 23 ALgal bloom of Enteromorpha _______________________________ 52
Figure 24 Enteromorpha prolifera ____________________________________ 54
Figure 25 Area of collection for Enteromorpha prolifera___________________ 55
Figure 26 Chara in its natural habitat _________________________________ 56
Figure 27 Characteristic morphology of Chara vulgaris ___________________ 57
Figure 28 Light micrograph of a whole-mount slide of an oogonium and
antheridium of Chara_______________________________________________ 58
Figure 29 Transesterification reaction: 1- Glycerol, 2-Base, 3-Esters, 4 byproduct
________________________________________________________________ 65
Figure 30 Schematic representation of GC – MS _________________________ 67
Figure 31 Nile red staining in Microspora sp. ___________________________ 71

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Figure 32 Nile red staining in Chara vulgaris ___________________________ 71
Figure 33 Standard graph of proteins __________________________________ 5
Figure 34 Standard graph of molecular weight __________________________ 12

8. 8
1. Abstract.
Biofuels demand is unquestionable in order to cope up increasing demand of
fuel sources and to reduce gaseous emissions and their purported
greenhouse, climatic changes and global warming effects, to face the
frequent oil supply crises, Biodiesel is usually produced from oleaginous
crops, such as rapeseed, soybean, sunflower and palm. However, the use of
microalgae can be a suitable alternative feedstock for next generation
biofuels because certain species contain high amounts of oil, which could be
extracted, processed and refined into transportation fuels, using currently
available technology; they have fast growth rate, permit the use of non-
arable land and non-potable water, use far less water and do not displace
food crops cultures; their production is not seasonal and they can be
harvested daily. The present study was performed to make an attempt to
understand the characteristics of selected algal species in reference to lipids,
proteins and carbohydrates.
Keywords: Microspora sp., Chara vulgaris, Enteromorpha prolifera, GC
MS, Transesterification, Bligh and Dyer.

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2. Introduction to Algae.
Algae are a very large and diverse group of simple, typically autotrophic
organisms, ranging from unicellular to multicellular forms, such as the giant
kelps that grow to 65 meters in length. Most are photosynthetic like plants,
and "simple" because they lack the many distinct cell and organ types found
in land plants. The largest and most complex marine forms are called
seaweeds. Algae grow almost everywhere in the world. They are a vital part
of the aquatic ecosystem providing food and shelter to other organisms.
They play a crucial role in the ability of an aquatic ecosystem to absorb
nutrients and heavy metals.[5][9]
Though the prokaryotic cyanobacteria are informally referred to as blue-
green algae, this usage is incorrect since they are regarded as bacteria. The
term alga is now restricted to eukaryotic organisms. All true algae therefore
have a nucleus enclosed within a membrane and plastids bound in one or
more membranes. Algae constitute a paraphyletic and polyphyletic group, as
they do not include all the descendants of the last universal ancestor nor do
they all descend from a common algal ancestor, although their plastids seem
to have a single origin. Diatoms are also examples of algae.
Algae exhibit a wide range of reproductive strategies, from simple,
asexual cell division to complex forms of sexual reproduction.
Algae lack the various structures that characterize land plants, such as
the leaf-like phyllids of bryophytes, rhizoids in nonvascular plants and the
roots, leaves and other organs that are found in tracheophytes (vascular
plants). Many are phototrophic, although some groups contain members that
are mixotrophic, deriving energy both from photosynthesis and uptake of

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organic carbon either by osmotrophy, myzotrophy, or phagotrophy. Some
unicellular species rely entirely on external energy sources and have limited
or no photosynthetic apparatus.[8][9]
Nearly all algae have photosynthetic machinery ultimately derived from
cyanobacteria, and so produce oxygen as a by-product of photosynthesis,
unlike other photosynthetic bacteria such as purple and green sulfur bacteria.
Fossilized filamentous algae from the Vindhya basin have been dated back
to 1.6 to 1.7 billion years ago. Certain blue-green algal blooms are toxic and
algal toxins can seriously affect animals and humans. Toxic blue-green algal
blooms cause a rash known as “swimmer's itch”, while powerful
neuromuscular toxins released by other cyanobacteria (blue green Algae)
can kill fish living in the water or the animals that drink the water. They
are more closely related to the bacteria than to other algae and often
referred to as 'Cyanobacteria'. They are extraordinarily diverse and range
from solitary cells through to complex multicellular forms several meters in
length. The absence of a sterile layer of protecting cells surrounding the
reproductive organs was the main reason why those algal groups having
organelles (chloroplasts, nuclei) - the eukaryotic algae - were commonly
placed in the kingdoms Protista and Plantae. It is now apparent that the
evolutionary history of the plastids of these algae is exceedingly complex
and has involved several endosymbiotic events that have led to their
transmission from one group to another. There is beginning to emerge the
broad outlines of a global phylogenetic tree with the eukaryotic algae placed
in four of five supergroups or 'kingdoms', including the Plantae. The 'blue-
green algae' (Cyanophyta) lack membrane-bound organelles and are
therefore prokaryotic organisms.[8][5]

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Some of the most crucially important diagnostic characters seen with a
good quality light microscope are often lost or no longer visible when an
alga is preserved. Even when microalgae are mounted on glass slides they
can deteriorate in time and rarely retain useful diagnostic features; one of the
notable exceptions are diatoms whose silica walls normally provide all the
characters required for identification. As a result, many 'permanently
preserved' samples of freshwater algae provide little useful information. For
this reason, the type of many microscopic algae is frequently not a specimen
but an illustration, photograph or figure ('iconotype') and preserved voucher
material is of limited use for cross-checking identification.[8]

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3. Introduction to Algae – Classification.
The number of species of algae remains uncertain since there is no
authoritative inventory of names in current use. Various figures have been
given and these generally range from about 27,000-36,000 with less than one
third of the species occurring in marine or brackish water. In the United
Kingdom the number of known freshwater and terrestrial algal species is less
than 5,000 of which diatoms (Bacillariophyta) and green algae
(Chlorophyta) are by far the largest groups.
The phyla of algae are still distinguished on a combination of characteristics,
including chlorophyll pigments, accessory pigments, food reserve products,
cell covering, reproductive features and various aspects of cellular
organisation. Modern molecular phylogenetic studies are revealing
relationships within the 'algae' to be much more complicated than originally
believed. These studies are leading to a radical reorganisation of the
traditionally recognised algal groups although a consensus has still to
emerge.[4]

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There are fifteen recognised phyla:
 Cyanophyta (Cyanobacterial, bacteria / blue-green algae)
 Rhodophyta (red algae)
 Euglenophyta (euglenoids)
 Cryptophyta (cryptomonads)
 Pyrrophyta (dinoflagellates)
 Raphidophyta
 Haptophyta (Prymnesiophyta)
 Chrysophyta (golden/golden brown algae)
 Xanthophyta (=Tribophyta; yellow-green algae)
 Chlorophyta (green algae, including stoneworts)
 Eustigmatophyta
 Phaeophyta (Fucophyta, brown algae)
 Prasinopyta
 Bacillariophyta (diatoms)
 Glaucophyta

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PHYLUM CYANOPHYTA
(Blue-Green Algae/Cyanobacteria)
Cyanobacteria, also known as blue-green bacteria, blue-green algae, and
Cyanophyta, is a phylum of bacteria that obtain their energy through
photosynthesis. The name "cyanobacteria" comes from the color of the
bacteria.
The ability of cyanobacteria to perform oxygenic photosynthesis is
thought to have converted the early reducing atmosphere into an oxidizing
one, which dramatically changed the composition of life forms on Earth by
stimulating biodiversity and leading to the near-extinction of oxygen-
intolerant organisms. According to endosymbiotic theory, chloroplasts in
plants and eukaryotic algae have evolved from cyanobacterial ancestors via
endosymbiosis.[4]
Figure 1 Nostoc Pruniforme Figure 2 Cylindrospermum

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Cyanobacteria can be found in almost every terrestrial and aquatic
habitat: in oceans, fresh water - even bare rock and soil. They can occur as
planktonic cells or form phototrophic biofilms in fresh water and marine
environments, they occur in damp soil, or even on temporarily moistened
rocks in deserts. A few are endosymbionts in lichens, plants, various protists,
or sponges and provide energy for the host. Some live in the fur of sloths,
providing a form of camouflage. Aquatic cyanobacteria are probably best
known for the extensive and highly visible blooms that can form in both
freshwater and the marine environment and can have the appearance of blue-
green paint or scum.
Nitrogen Fixation in Cyanobacteria.
Cyanobacteria include unicellular and colonial species. Colonies may
form filaments, sheets or even hollow balls. Some filamentous colonies
show the ability to differentiate into several different cell types: vegetative
cells, the normal, photosynthetic cells that are formed under favorable
growing conditions; akinetes, the climate-resistant spores that may form
when environmental conditions become harsh; and thick-walled heterocysts,
which contain the enzyme nitrogenase, vital for nitrogen fixation.
Heterocysts may also form under the appropriate environmental conditions
(anoxic) when fixed nitrogen is scarce. Heterocyst-forming species are
specialized for nitrogen fixation and are able to fix nitrogen gas into
ammonia (NH3), nitrites (NO−)or nitrates (NO−)which can be absorbed by
plants and converted to protein and nucleic acids (atmospheric nitrogen is
not bioavailable to plants). Rice plantations utilize healthy populations of
nitrogen-fixing cyanobacteria (Anabaena, as symbiotes of the aquatic fern
Azolla) for use as rice paddy fertilizer.

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PHYLUM RHODOPHYTA
(Red Algae)
The red algae, or Rhodophyta, are one of the oldest groups of eukaryotic
algae, and also one of the largest, with about 5,000–6,000 species of mostly
multicellular, marine algae, including many notable seaweeds. Other
references indicate as many as 10,000 species; more detailed counts indicate
about 4,000 in about 600 genera (3,738 marine species in 546 genera and 10
orders (plus the unclassifiable); 164 freshwater species in 30 genera in eight
orders).
The red algae form a distinct group characterized by these attributes:
eukaryotic cells without flagella and centrioles, using floridean
polysaccharides as food reserves, with phycobiliproteins as accessory
pigments (giving them their red color), and with chloroplasts lacking
external endoplasmic reticulum and containing unstacked thylakoids. Most
red algae are also multicellular, macroscopic, marine, and have sexual
reproduction. They often have alternation of generations and may have three
generations rather than two. [4]
Figure 3 Polysiphonia

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Many of the coralline algae, which secrete calcium carbonate and play a
major role in building coral reefs, belong here. Red algae such as dulse
(Palmaria palmata) and laver (nori/gim) are a traditional part of European
and Asian cuisines and are used to make other products such as agar,
carrageenans and other food additives.
Most rhodophytes are marine, although freshwater species are found;
these generally prefer clean, running water, but with some exceptions.
One of the oldest fossils identified as a red alga is also the oldest fossil
eukaryote that belongs to a specific modern taxon. Bangiomorpha
pubescens, a multicellular fossil from arctic Canada, strongly resembles the
modern red alga Bangia despite occurring in rocks dating to 1200 million
years ago.
Red algae are important builders of limestone reefs. The earliest such
coralline algae, the solenopores, are known from the Cambrian period. Other
algae of different origins filled a similar role in the late Paleozoic, and in
more recent reefs.
Figure 4 Some red algae

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PHYLUM EUGLENOPHYTA
(Euglenoids)
Euglenoids (or euglena) are one of the best-known groups of flagellates,
commonly found in freshwater especially when it is rich in organic
materials, with a few marine and endosymbiotic members. Most euglenids
are unicellular. Many euglenids have chloroplasts and produce energy
through photosynthesis, but others feed by phagocytosis or strictly by
diffusion. They belong to the phylum Euglenophyta, and their cell structure
is typical of that group.
Euglenids are thought to descend from an ancestor that took up green algae
by secondary endosymbiosis.
Euglenids are distinguished mainly by the presence of a pellicle, which is
composed of proteinaceous strips underneath the cell membrane, supported
by dorsal and ventral microtubules. This varies from rigid to flexible, and
gives the cell its shape, often giving it distinctive striations. In many
euglenids the strips can slide past one another, causing an inching motion
called metaboly. Otherwise they move using the flagella.
Figure 5 Some euglenoids

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PHYLUM CRYPTOPHYTA
(Cryptomonads)
The cryptomonads (or cryptophytes) are a group of algae, most of which
have plastids. They are common in freshwater, and also occur in marine and
brackish habitats. Each cell is around 10-50 μm in size and flattened in
shape, with an anterior groove or pocket. At the edge of the pocket there are
typically two slightly unequal flagella.
Some may exhibit mixotrophy i.e. an organism that can use a mix of
different sources of energy and carbon.
Cryptomonads are distinguished by the presence of characteristic
extrusomes called ejectisomes or ejectosomes, which consist of two
connected spiral ribbons held under tension. If the cells are irritated either by
mechanical, chemical or light stress, they discharge, propelling the cell in a
zigzag course away from the disturbance. Large ejectisomes, visible under
the light microscope, are associated with the pocket; smaller ones occur
underneath the periplast, the cryptophyte-specific cell surrounding.
Figure 6 Rhodomonas salina

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Cryptomonads have one or two chloroplasts, except for Chilomonas, which
has leucoplasts and Goniomonas (formerly Cyathomonas) which lacks
plastids entirely. These contain chlorophylls a and c, together with
phycobiliproteins and other pigments, and vary in color (brown, red to
blueish-green). Each is surrounded by four membranes, and there is a
reduced cell nucleus called a nucleomorph between the middle two. This
indicates that the plastid was derived from a eukaryotic symbiont, shown by
genetic studies to have been a red alga.
A few cryptomonads, such as Cryptomonas, can form palmelloid stages, but
readily escape the surrounding mucus to become free-living flagellates
again. Some Cryptomonas species may also form immotile resting stages
with rigid cell walls (cysts) to survive unfavorable conditions. Cryptomonad
flagella are inserted parallel to one another, and are covered by bipartite
hairs called mastigonemes, formed within the endoplasmic reticulum and
transported to the cell surface. Small scales may also be present on the
flagella and cell body. The mitochondria have flat cristae, and mitosis is
open; sexual reproduction has also been reported. [4]

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PHYLUM PYRROPHYTA
(Dinoflagellates)
The dinoflagellates are a large group of flagellate protists. Most are
marine plankton, but they are common in fresh water habitats as well. Their
populations are distributed depending on temperature, salinity, or depth.
Many dinoflagellates are known to be photosynthetic, but a large fraction of
these are in fact mixotrophic, combining photosynthesis with ingestion of
prey. In terms of number of species, dinoflagellates form one of the largest
groups of marine eukaryotes, although this group is substantially smaller
than the diatoms. Being primary producers makes them an important part of
the aquatic food chain. Some species are endosymbionts of marine animals
and play an important part in the biology of coral reefs. Other dinoflagellates
are colorless predators on other protozoa, and a few forms are parasitic (see
for example Oodinium, Pfiesteria). Some dinoflagellates produce resting
stages, called dinoflagellate cysts or dinocysts, as part of their life cycles.
Figure 7 Ceratium Furca

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PHYLUM RAPHIDOPHYTA
Raphidophytes formerly referred to as Chloromonadophyceae and
Chloromonadineae are a small group of eukaryotic algae that includes both
marine and freshwater species. All raphidophytes are unicellular, with large
cells (50 → 100 μm) but no cell walls. Raphidophytes possess a pair of
flagella, organised such that both originate from the same invagination (or
gullet). One flagellum points forwards, and is covered in hair-like
mastigonemes, while the other points backwards across the cell surface,
lying within a ventral groove. Raphidophytes contain numerous ellipsoid
chloroplasts, which contain chlorophylls a, c1 and c2. They also make use of
accessory pigments including β-carotene and diadinoxanthin. In terms of
ecology, raphidophytes occur as photosynthetic autotrophs across a range of
aquatic systems. Freshwater species are more common in acidic waters, such
as pools in bogs. Marine species often produce large blooms in summer,
particularly in coastal waters. Off the Japanese coast, the resulting red tides
often cause disruption to fish farms, although raphidophytes are not usually
responsible for toxic blooms. [4]
Figure 8 Gonyostomum

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PHYLUM CHRYSOPHYTA
(Golden-Brown Algae)
Chrysophytes, or golden algae, are common microscopic chromists in fresh
water. Some species are colorless, but the vast majority are photosynthetic. As
such, they are particularly important in lakes, where they may be the primary
source of food for zooplankton. They are not considered truly autotrophic by some
biologists because nearly all chrysophytes become facultatively heterotrophic in
the absence of adequate light, or in the presence of plentiful dissolved food. When
this occurs, the chrysoplast atrophies and the alga may turn predator, feeding on
bacteria or diatoms. [4]
Figure 9 Dinobryon divergens
There are more than a thousand described species of golden algae, most of them
free-swimming and unicellular, but there are filamentous and colonial forms. Other
chrysophytes may spend part of their life as amoeboid cells. At the left and center

24. 24
of the above illustration is Dinobryon, a freshwater genus in which the individual
cells are surrounded by vase-shaped loricae, composed of chitin fibrils and other
polysaccharides. The colonies grow as branched or unbranched chains. A spherical
colonial form, Synura, is on the right; the surfaces of these cells are covered by
silica scales. Species which produce siliceous coverings may have bristles or scales
with quite complex structure. Some researchers group the chrysophytes with silica
scales in a separate taxon, the Synurophyceae.
The oldest known chrysophytes are from calcareous and siliceous deposits of
Cretaceous age, but they reached their greatest diversity in the Miocene. The group
actually has a fairly complete fossil record, because most freshwater chrysomonads
secrete resting cysts of silica, which may be abundant in certain rocks -- in some
Paleocene deposits, chrysophyte cysts outnumber the diatoms! The fossils of
chrysophytes, like those of diatoms and coccolithophorids, are often used as
paleoecological indicators to reconstruct ancient environments. [4]

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PHYLUM XANTHOPHYTA
(Yellow-Green Algae)
The yellow-green algae are photosynthetic species of organisms
belonging to the Xanthophyta Phylum, which is one of the phyla pertaining
to the Chromista Group in the Protista Kingdom. Xanthophyta encompasses
650 living species so far identified. Xanthophyta live mostly in freshwater,
although some species live in marine water, tree trunks, and damp soils.
Some species are unicellular organisms equipped with two unequal flagella
that live as free-swimming individuals, but most species are filamentous.
Filamentous species may be either siphonous or coenocytic. Coenocytes are
organized as a single-cell multinucleated thallus that form long filaments
without septa (internal division walls) except in the specialized structures of
some species. Siphonous species have multiple tubular cells containing
several nuclei.
Figure 10 Xanthophyceae
Xanthophyta synthesize chlorophyll a and smaller amounts of chlorophyll
c, instead of the chlorophyll b of plants; and the cellular structure usually
have multiple chloroplasts without nucleomorphs. The plastids have four
membranes and their yellow-green color is due to the presence of beta-

26. 26
carotene and xanthins, such as vaucheriaxanthin, diatoxanthin,
diadinoxanthin, and heretoxanthin, but not fucoxanthin, the brown pigment
present in other Chromista. Because of the presence of significant amounts
of chlorophyll a, Xanthophyceae species are easily mistaken for green algae.
They store polysaccharide under the form of chrysolaminarin and
carbohydrates as oil droplets. [4]
Figure 11 Xanthophyceae
One example of a relatively common Xanthophyta is the class Vaucheria
that gathers approximately 70 species, whose structure consists of several
tubular filaments, sharing its nuclei and chloroplasts without septa. They live
mainly in freshwater, although some species are found in seawater spreading
along the bottom like a carpet. Other Xanthophyceae Classes are Tribonema,
whose structure consists of unbranched filaments; Botrydiopsis, such as the
species Botrydium with several thalli, each thallus formed by a large aerial
vesicle and rhizoidal filaments, found in damp soil; Olisthodiscus, such as
the species Ophiocytium with cylindrical and elongated multinucleated cells
and multiple chloroplasts.

27. 27
PHYLUM CHLOROPHYTA
(Green Algae)
Chlorophyta is a division of green algae, informally called chlorophytes.
The name is used in two very different senses so that care is needed to
determine the use by a particular author. In older classification systems, it
refers to a highly paraphyletic group of all the green algae within the green
plants (Viridiplantae), and thus includes about 7,000 species of mostly
aquatic photosynthetic eukaryotic organisms. Like the land plants
(bryophytes and tracheophytes), green algae contain chlorophylls a and b,
and store food as starch in their plastids. [4]
In newer classifications, it refers to one of the two clades making up the
Viridiplantae, which are the chlorophytes and the streptophytes or
charophytes. In this sense it includes only about 4,300 species.
Figure 12 Caulerpa racemosa
Ecologically, green algae are incredibly important. For example, they
serve as a source of food for other aquatic organisms. Marine phytoplankton
is the first link in the great aquatic food chain. They are the producers which
are fed on by zooplankton (the other component of plankton comprised of

28. 28
protozoa, small crustaceans, jellyfish, etc.), which in turn are consumed by
larger animals such as fish and even the blue whale. Also, green algae
contribute largely to the world’s supply of oxygen. In fact, an estimated 90
percent of all photosynthesis and discharge of free oxygen occurs in the
oceans. However, chlorophytes can actually have a negative effect on the
environment, as when large populations of green algae produce an
unpleasant taste or smell in drinking water or as when algal populations in
freshwater lakes and ponds which are polluted by nitrates and phosphates
suddenly increase, forming an odorous scum and largely decreasing the
oxygen available to other organisms.
Figure 13 Caulerpa filiformis

29. 29
PHYLUM EUSTIGMATOPHYTA
Eustigmatophytes are a small group (7 genera; ~12 species) of
eukaryotic algae that includes marine, freshwater and soil-living
species. All eustigmatophytes are unicellular, with coccoid cells
and polysaccharide cell walls. Eustigmatophytes contain one or
more yellow-green chloroplasts, which contain chlorophyll a and
the accessory pigments violaxanthin and β-carotene.
Eustigmatophyte zoids (gametes) possess a single or pair of
flagella, originating from the apex of the cell. Unlike other
heterokontophytes, eustigmatophyte zoids do not have typical
photoreceptive organelles (or eyespots); instead, an orange-red
eyespot outside of a chloroplast is located at the anterior end of the
zoid. [4]
Figure 14 Nannochloropsis

30. 30
In terms of ecology, eustigmatophytes occur as photosynthetic
autotrophs across a range of systems. Most eustigmatophyte genera
live in freshwater or in soil, although Nannochloropsis contains
marine species of picophytoplankton (2 → 4 μm).

31. 31
PHYLUM PHAEOPHYTA
(Brown Algae)
The Phaeophyceae or brown algae (singular: alga), is a large group of mostly
marine multicellular algae, including many seaweeds of colder Northern
Hemisphere waters. They play an important role in marine environments,
both as food and for the habitats they form. For instance Macrocystis, kelp
of the order Laminariales, may reach 60 m in length, and forms prominent
underwater forests. Another example is Sargassum, which creates unique
habitats in the tropical waters of the Sargasso Sea. Many brown algae, such
as members of the order Fucales, commonly grow along rocky seashores.
Some members of the class are used as food for humans.
Figure 15 Brown algae
Worldwide there are about 1500–2000 species of brown algae. Some species
are of sufficient commercial importance, such as Ascophyllum nodosum, that
they have become subjects of extensive research in their own right.

32. 32
Brown algae belong to a very large group, the Heterokontophyta, a
eukaryotic group of organisms distinguished most prominently by having
chloroplasts surrounded by four membranes, suggesting an origin from a
symbiotic relationship between a basal eukaryote and another eukaryotic
organism. Most brown algae contain the pigment fucoxanthin, which is
responsible for the distinctive greenish-brown color that gives them their
name. Brown algae are unique among heterokonts in developing into
multicellular forms with differentiated tissues, but they reproduce by means
of flagellated spores and gametes that closely resemble cells of other
heterokonts. Genetic studies show their closest relatives to be the yellow-
green algae.
Figure 16 Macrocystis Pyrifera

33. 33
PHYLUM PRASINOPHYTA
The praesinophytes comprise a collection of organisms that generally
occur as unicells, but some are attached as filaments (Figures 1-4). They
occur in almost all aquatic environments. Indeed, there is a growing
realization that these might be significant contributors to marine plankton
primary production. The motile cells usually are covered with cellulosic
scales rather than a cell wall. In some the scales are quite complex.
However, the fusion of scales in taxa like Pedinomonas might indicate how
the more typical cell wall of the kingdom evolved.
Although they are photosynthetic, most obtain supplement their nutrition
by ingesting bacteria and other organic matter, a process called mixotrophy.
Likely, the earliest members of this line were completely heterotrophic, a
process by which they enslaved a chlorophyll a and b bearing
Cyanobacterium to become the first photobionts. [4]

34. 34
PHYLUM GLAUCOPHYTA
The glaucophytes, also known as glaucocystophytes or glaucocystids, are a
small group of freshwater microscopic algae. Together with the red algae
(Rhodophyta) and green algae plus land plants (Viridiplantae or
Chloroplastida), they form the Archaeplastida. However, the relationships
between the red algae, green algae and glaucophytes are unclear, in large
part due to limited study of the glaucophytes.
The glaucophytes are of interest to biologists studying the development of
chloroplasts because some studies suggest that they may be similar to the
original alga type that led to green plants and red algae.
Figure 17 Glaucocystis Species
The chloroplasts of glaucophytes are known as cyanelles. Unlike plastids
in other organisms they have a peptidoglycan layer that is believed to be a
relic of the endosymbiotic origin of plastids from cyanobacteria.
Glaucophytes contain the photosynthetic pigment chlorophyll a. Along with
red algae and cyanobacteria, they harvest light via phycobilisomes,
structures consisting largely of phycobiliproteins. The green algae and land
plants have lost that pigment.

35. 35
Glaucophytes have mitochondria with flat cristae, and undergo open mitosis
without centrioles. Motile forms have two unequal flagella, which may have
fine hairs and are anchored by a multilayered system of microtubules, both
of which are similar to forms found in some green algae. [4]

36. 36
5. Purpose for the Project
In today’s world uncontrolled population growth has become a big burden
on our society. The world population in last 50 years (1960-2011) has
doubled i.e. from 3.04 billion we have now touched 6.973 billion. The major
problems that arise due to this are energy crisis and hunger. Feeding a
population of nearly 7 billion people is a real task. Also increase in
population has led to a rise in the usage of fuels like petrol, diesel, C.N.G.
and L.P.G.
Figure 18 World Population Growth Chart (1960- 2011)
To keep up with the growing needs of our society countries all over the
world are now exploring new avenues in renewable sources of energy,
increasing per capita output of farms and new and simple sources of
proteins.

37. 37
6. The Need for Research.
The following graphs and statistics will show the current scenario of world
population demand.
Figure 19 Average Diesel Price in Dollars (World)
Diesel prices were under control till 2002 but since then there has been an
average 0.1$ increase every year due to increase in the demand for diesel.
Reasons being new factories in remote places had no electric supply increase
in demand for diesel vehicles in private sector. [2]

38. 38
7. The Need for Research
The agricultural sector also saw some changes in the past 50 years.
Figure 20 Area under Agriculture (Sq. kms.)
Although there has always been a steady growth in the area under
agriculture the years between 1992 – 94 saw phenomenal growth with upto 8
million sq. kms. were brought under agriculture.
Looking at all these statistics it is needless to say that more work is
needed in both these sectors. In this project we will find out how we can put
algae to use for productions of fuel and supplementary protein.[6]

39. 39
Work done by other Researchers
With the increase in research in alternative fuel sources, algal oil extraction
has also gained momentum. The past 10 years were very fruitful for this
research.
Studies performed by A. Demirbaşa on species of Cladophora and
Chlorella prove the presence of a diesel like fuel in the given species. This
fuel is said to be 20 times heavier than traditional diesel but trans-
esterification of the fatty ester is the promising solution. Given below is the
abstract from his book “Energy Sources, Part A: Recovery, Utilization, and
Environmental Effects” Volume 31, Issue 2, 2008.
Abstract
“A macroalga (Cladophora fracta) and a microalga (Chlorella
protothecoides) samples were used in this work. Most current research on oil
extraction is focused on microalgae to produce biodiesel from algal oil. The
biodiesel from algal oil in itself is not significantly different from biodiesel
produced from vegetable oils. Algal oils, as well as vegetable oils, are all
highly viscous, with viscosities ranging 10–20 times those of no. 2 diesel
fuel. Transesterification of the oil to its corresponding fatty ester is the most
promising solution to the high viscosity problem. Fatty acid (m)ethyl esters
produced from natural oils and fats is called biodiesel. Generally, methanol
has been mostly used to produce biodiesel as it is the least expensive
alcohol. The oil proportion from the lipid fractions of Chlorella
protothecoides is considerable higher than that of Cladophora fracta. The
higher heating value of Chlorella protothecoides (25.1 MJ/kg) also is higher

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than that of Cladophora fracta (21.1 MJ/kg). The average polyunsaturated
fatty acids of Chlorella protothecoides (62.8%) also are higher than those of
Cladophora fracta (50.9%).”
Ronald Halim, Brendan Gladman, Michael K. Danquah and Paul A.
Webley of the Bio Engineering Laboratory (BEL), Department of Chemical
Engineering, Monash University, Clayton, Australia have worked on
maximizing lipid extraction from Chlorococcum species by using super
critical carbon dioxide and hexane.
Abstract
“This study examines the performance of supercritical carbon dioxide
(SCCO2) extraction and hexane extraction of lipids from marine
Chlorococcum sp. for lab-scale biodiesel production. Even though the strain
of Chlorococcum sp. used in this study had a low maximum lipid yield (7.1
wt% to dry biomass), the extracted lipid displayed a suitable fatty acid
profile for biodiesel [C18:1 (∼63 wt%), C16:0 (∼19 wt%), C18:2 (∼4 wt%),
C16:1 (∼4 wt%), and C18:0 (∼3 wt%)]. For SCCO2 extraction, decreasing
temperature and increasing pressure resulted in increased lipid yields. The
mass transfer coefficient (k) for lipid extraction under supercritical
conditions was found to increase with fluid dielectric constant as well as
fluid density. For hexane extraction, continuous operation with a Soxhlet
apparatus and inclusion of isopropanol as a co-solvent enhanced lipid yields.
Hexane extraction from either dried microalgal powder or wet microalgal
paste obtained comparable lipid yields.”

41. 41
Luisa Gouveia, Ana Cristina Oliveira have written about their studies in
the “Journal of Industrial Microbiology & Biotechnology”, February 2009,
Volume 36, Issue 2, page 269-274. They write about the need for increasing
the production of biofuels and the harmful effects of fossil fuels.
Abstract
“Biofuels demand is unquestionable in order to reduce gaseous emissions
(fossil CO2, nitrogen and sulfur oxides) and their purported greenhouse,
climatic changes and global warming effects, to face the frequent oil supply
crises, as a way to help non-fossil fuel producer countries to reduce energy
dependence, contributing to security of supply, promoting environmental
sustainability and meeting the EU target of at least of 10% biofuels in the
transport sector by 2020. Biodiesel is usually produced from oleaginous
crops, such as rapeseed, soybean, sunflower and palm. However, the use of
microalgae can be a suitable alternative feedstock for next generation
biofuels because certain species contain high amounts of oil, which could be
extracted, processed and refined into transportation fuels, using currently
available technology; they have fast growth rate, permit the use of non-
arable land and non-potable water, use far less water and do not displace
food crops cultures; their production is not seasonal and they can be
harvested daily. The screening of microalgae (Chlorella vulgaris, Spirulina
maxima, Nannochloropsis sp., Neochloris oleabundans, Scenedesmus
obliquus and Dunaliella tertiolecta) was done in order to choose the best
one(s), in terms of quantity and quality as oil source for biofuel production.
Neochloris oleabundans (fresh water microalga) and Nannochloropsis sp.
(marine microalga) proved to be suitable as raw materials for biofuel
production, due to their high oil content (29.0 and 28.7%, respectively).

42. 42
Both microalgae, when grown under nitrogen shortage, show a great
increase (~50%) in oil quantity. If the purpose is to produce biodiesel only
from one species, Scenedesmus obliquus presents the most adequate fatty
acid profile, namely in terms of linolenic and other polyunsaturated fatty
acids. However, the microalgae Neochloris oleabundans, Nannochloropsis
sp. and Dunaliella tertiolecta can also be used if associated with other
microalgal oils and/or vegetable oils.”
Another important study by Laurent Lardon , Arnaud H lias, Bruno Sialve,
Jean-Philippe Steyer and Olivier Bernard from INRA, UR50 Laboratoire de
Biotechnologie de l’Environnement, France, and Comore, INRIA, France
write about the environmental impact of biodiesel production from micro-
algae.
Abstract
“This paper provides an analysis of the potential environmental impacts of
biodiesel production from microalgae. High production yields of microalgae
have called forth interest of economic and scientific actors but it is still
unclear whether the production of biodiesel is environmentally interesting
and which transformation steps need further adjustment and optimization. A
comparative LCA study of a virtual facility has been undertaken to assess
the energetic balance and the potential environmental impacts of the whole
process chain, from the biomass production to the biodiesel combustion.
Two different culture conditions, nominal fertilizing or nitrogen starvation,
as well as two different extraction options, dry or wet extraction, have been
tested. The best scenario has been compared to first generation biodiesel and
oil diesel. The outcome confirms the potential of microalgae as an energy

43. 43
source but highlights the imperative necessity of decreasing the energy and
fertilizer consumption. Therefore control of nitrogen stress during the
culture and optimization of wet extraction seem to be valuable options. This
study also emphasizes the potential of anaerobic digestion of oilcakes as a
way to reduce external energy demand and to recycle a part of the mineral
fertilizers.”

44. 44
8. Algae and their many uses
Earth’s early forms of life and first forms of food for subsequent species
now hold the potential to become the planet’s next major source of energy
and a vital part of the solutions to climate change and dependence on fossil
fuels. Cyanobacteria have caused more global environmental change than
humans could ever cause and are now poised to address many of society’s
greatest challenges. But given the fuel versus food problems associated with
other biofuels, the same issue and other issues relating to the scarcity of
resources and impacts on the environment must be considered when it comes
to algae.
Following are some of the other uses of algae:
 Fertilizer
Algae are used by humans in many ways. They are used as
fertilizers, soil conditioners and are a source of livestock feed.
Because many species are aquatic and microscopic, they are cultured
in clear tanks or ponds and either harvested or used to treat effluents
pumped through the ponds. Algaculture on a large scale is an
important type of aquaculture in some places.
 Energy source
Algae can be grown to produce hydrogen. In 1939 a German
researcher named Hans Gaffron, while working at the University of
Chicago, observed that the algae he was studying, Chlamydomonas
reinhardtii (a green-algae), would sometimes switch from the

45. 45
production of oxygen to the production of hydrogen. Algae can be
grown to produce biomass, which can be burned to produce heat and
electricity.
 Pollution control
Algae are used in wastewater treatment facilities, reducing the need
for greater amounts of toxic chemicals than are already used.
Algae can be used to capture fertilizers in runoff from farms. When
subsequently harvested, the enriched algae itself can be used as
fertilizer.
Algae Bioreactors are used by some powerplants to reduce CO2
emissions. The CO2 can be pumped into a pond, or some kind of tank,
on which the algae feed. Alternatively, the bioreactor can be installed
directly on top of a smokestack.
 Stabilizing substances
Chondrus crispus, (probably confused with Mastocarpus stellatus,
common name: Irish moss), is also used as "carrageen". The name
carrageenan comes from the Irish Gaelic for Chondrus crispus. It is an
excellent stabiliser in milk products - it reacts with the milk protein
caesin, other products include: petfoods, toothpaste, ice-creams and
lotions etc. Alginates in creams and lotions are absorbable through the
skin.
 Nutrition
1. Seaweeds are an important source of food, especially in Asia; They
are excellent sources of many vitamins including: A, B1, B2, B6,

46. 46
niacin and C. They are rich in iodine, potassium, iron, magnesium and
calcium.
2. Algae is commercially cultivated as a nutritional supplement. One of
the most popular microalgal species is Spirulina (Arthrospira
platensis), which is a Cyanobacteria (known as blue-green algae), and
has been hailed by some as a superfood. Other algal species cultivated
for their nutritional value include; Chlorella (a green algae),
and Dunaliella (Dunaliella salina), which is high in beta-carotene and
is used in vitamin C supplements. In China at least 70 species of algae
are eaten as is the Chinese "vegetable" known as fat choy (which is
actually a cyanobacterium). Roughly 20 species of algae are used in
everyday cooking in Japan.
 Other Uses of Algae
1. There are also commercial uses of algae as agar.
2. The natural pigments produced by algae can be used as an
alternative to chemical dyes and coloring agents.
3. Many of the paper products used today are not recyclable
because of the chemical inks that they use paper recyclers have
found that inks made from algae are much easier to break down.
4. There is also much interest in the food industry into replacing
the coloring agents that are currently used with coloring derived
from algal pigments.
5. Algae can be used to make pharmaceuticals.
6. Sewage can be treated with algae as well.
7. Some Cosmetics can come from microalgae as well.

47. 47
8. In Israel, a species of green algae is grown in water tanks, then
exposed to direct sunlight and heat which causes it to become
bright red in color. It is then harvested and used as a natural
pigment for foods such as Salmon.

48. 48
9. Details about the Algal Species used in the Project.
Three species of filamentous green algae were used:
1. Microspora species.
2. Enteromorpha prolifera.
3. Chara vulgaris.

49. 49
Microspora species.
Domain: Eukaryota
Kingdom: Viridiplantae
Phylum: Chlorophyta
Class: Chlorophyceae
Family: Microsporaceae
Genus: Microspora.
Description:
Microspora species are unbranched filamentous green algae. There is a
single dense net-like chloroplast, usually filling the cell, no pyrenoid. The
cells are frequently rather bulbous or barrel-shaped, but the chief diagnostic
character is the presence of H-shaped wall sections, which can usually be
seen in the filament by careful focusing under favorable lighting, and may be
most clearly seen at the ends of filaments. When the filament degenerates
they are frequently found free. The cell wall may be thin, with the H pieces
readily visible overlapping each other slightly or it may be thick and rather
gelatinous in appearance, in which condition it may be difficult to
distinguish from Binuclearia.

50. 50
Figure 21 Microspora at 100 X magnification
Figure 22 Microspora at 400 X magnification. Microscope: Labomed Lx 300
Microspora frequently show the presence of darkened, brown bands
between adjacent cells.
There are a number of species and mean cell diameter is a guide to species
discrimination. Diameters of 8, 11, 14, 17, 20 and 22 µm. are found in acid
waters. They are most frequently found tangled in moss or other vegetation,
although they initially attach to the substrate by means of a single holdfast
cell following settlement of the motile zoospore.

51. 51
Area of Collection:
The Microspora species used for this experiment was extracted from
Tansa Lake. This lake is located to the north east of Mumbai roughly a 120
kms away from Dadar on Mumbai-Nashik highway. It is one of Mumbai’s
major water sources and the area is protected by Brihanmumbai Municipal
Corporation

52. 52
Enteromorpha Prolifera
Domain: Eukaryota
Subkingdom: Viridaeplantae
Phylum: Chlorophyta
Class: Ulvophyceae
Order: Ulvales
Family: Ulvaceae
Genus: Enteromorpha
Description:
Enteromorpha prolifera is one of the dominant seaweeds in the littoral
zone of South East Asia, North America and some parts of Europe. It
distributes in a wide variety of coastal water, such as brackish-waters of
inner bays and estuaries and so on and greatly affects the carbon cycle and
recovery of the contaminant water body to border on the sea and often
contributes to the formation of the so-called ‘green tide’, which causes
ecological and indirect economic damages.
Figure 23 ALgal bloom of Enteromorpha

53. 53
It is better known as Enteromorpha prolifera, but Enteromorpha is now
considered to be part of the genus Ulva. The fronds are tubular, though often
more or less flattened, little too much-branched. The arrangement of the
cells, in longitudinal and transverse rows in the central part of the frond, is
characteristic of this species, as are the cylindrical chloroplasts seeming to
fill the cell and the usually single, central pyrenoids.
When Enteromorpha first begins growing, it forms a single row of cells,
this structure is monosiphonous. Soon after the monosiphonous filament is
formed, longitudinal division of cells creates a two layered filament.
Eventually, after more cell division the two cell layers separate to form a
tube, forming the adult morphology.
The thallus of Enteromorpha is tubular with the wall of the tube a single
cell layer thick. The thallus can be branched or unbranched, and there is a
wide variety of forms within the genus. Enteromorpha is attached to the
substrate by a disc-like holdfast. The holdfast is formed by the basal cell
dividing into three or four holdfast cells which elongate and undergo further
division.
The cells in Enteromorpha can vary in size and shape from species to
species, and sometimes they will form regular linear series in a frond, while
other times there is an irregular arrangement of the cells. Each cell contains
a single chloroplast, varying in size depending on the size of the cell.

54. 54
Figure 24 Enteromorpha prolifera
Area of Collection
The Enteromorpha prolifera sample for this project was collected from
Sasawne beach near Alibaug city in Raigad district. It is a coastal area with
dense population leading to a lot of pollution. Human settlements dump
nitrogen rich waste matter near the sea giving rise to algal bloom yearly.
This decreases the fish population of the area vital to the population

55. 55
Figure 25 Area of collection for Enteromorpha prolifera

56. 56
Chara Vulgaris
Kingdom: Plantae
Division: Charophyta
Class: Charophyceae
Order: Charales
Family: Characeae
Genus: Chara
Description:
Chara is a genus of green algae in the family Characeae. They are
multicellular and superficially resemble land plants because of stem-like and
leaf-like structures. They are found in fresh water, particularly in limestone
areas throughout the northern temperate zone, where they grow submerged,
attached to the muddy bottom. They prefer less oxygenated and hard water
and are not found in waters where mosquito larvae are present. They are
covered with calcium carbonate deposits.
Figure 26 Chara in its natural habitat

57. 57
The branching system of Chara species is complex with branches derived
from apical cells which cut off segments at the base to form nodal and
internodal cells alternately. They are typically anchored to the littoral
substrate by means of branching underground rhizoids. Chara plants are
rough to the touch because of deposited calcium salts on the cell wall. The
metabolic processes associated with this deposition often give Chara plants a
distinctive and unpleasant smell of hydrogen sulfide.
Figure 27 Characteristic morphology of Chara vulgaris
The plant body is a gametophyte. It consists of a main axis (differentiated
into nodes and internodes); dimorphic branches (Long Branch of unlimited
growth and short branches of limited growth), rhizoids (multicellular with
oblique septa) and stipulodes (needle shaped structures at the base of
secondary laterals.

58. 58
Figure 28 Light micrograph of a whole-mount slide of an oogonium and
antheridium of Chara

59. 59
Area of Collection:
Chara for this experiment was obtained from Vandri Lake, located 80 kms
north of Mumbai on the Mumbai-Ahmedabad highway. This lake is dammed
and supplies water to a dairy and a small village.

60. 60
10. MATERIAL AND METHOD
Extraction and Estimation of Lipids from Algal Culture.
Aim: To extract lipids from the given three algal cultures of
1. Microspora species.
2. Entromorpha prolifera.
3. Chara vulgaris.
Requirements:
Distilled water, Chloroform, Methanol, n-Hexane.
Principle:
The aim of all extraction procedures is to separate cellular or fluid lipids
from the other constituents, proteins, polysaccharides, small molecules
(amino acids, sugars...) but also to preserve these lipids for further analyses.
There is a great diversity of methodologies because biological tissues are not
similar when considering their structure, texture, sensitivities and lipid
contents. The ideal solvent for lipid extraction would completely extract all
the lipid components from a sample, while leaving all the other components
behind. In practice, the efficiency of solvent extraction depends on the
polarity of the lipids present compared to that of the solvent.
Polar lipids (such as glycolipids or phospholipids) are more soluble in polar
solvents (such as alcohols), than in non-polar solvents (such as hexane). On

61. 61
the other hand, non-polar lipids (such as triacylglycerols) are more soluble in
non-polar solvents than in polar ones. The fact that different lipids have
different polarities means that it is impossible to select a single organic
solvent to extract them all. Thus the total lipid content determined by solvent
extraction depends on the nature of the organic solvent used to carry out the
extraction: the total lipid content determined using one solvent may be
different from that determined using another solvent.
Ethyl ether and petroleum ether are the most commonly used solvents, but
pentane and hexane are also used for some foods.
Procedure:
1. 100 mg of dry biomass was homogenized in a mortar and pestle using
distilled water and made 10 ml final volume.
2. This was transferred to separating funnel and 37.5 ml of Chloroform:
Methanol in the ratio of 1:2 was added.

62. 62
3. The mixture was shaken thoroughly and kept standing for 30 mins.
4. 12 ml of chloroform was added to separate two layer from which
lower layer was collected in a preweighed beaker.
5. This was kept in water bath at 80 C for evaporation of chloroform.
6. The beaker was weighed again after complete evaporation of
chloroform.[11]
Observations:
Microspora species:
1. Weight of the sample – 100mg.
2. Weight of the beaker – 36.808gms.
3. Weight of the beaker and crude lipid – 36.819gms.
4. Weight of lipid – 0.011gms.
Entromorpha Prolifera:
1. Weight of the sample – 100mg.
2. Weight of the beaker – 35.445gms.
3. Weight of the beaker and crude lipid – 35.473gms.
4. Weight of lipid – 0.028gms.
Chara vulgaris:
1. Weight of the sample – 100mg.
2. Weight of the beaker – 35.995gms.
3. Weight of the beaker and crude lipid – 36.004gms.
4. Weight of lipid – 0.009gms.

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Inference:
Enteromorpha prolifera shows the highest amount of lipid content i.e.
0.028 gms. Chara and Microspora contain almost the same amount of lipid
i.e. 0.009-0.011 gms.

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Transesterification of Lipids
Lipids extracted from in the above given experiment cannot be directly
used. These lipids are tri-glycerides from which alcohol is deprotonated.
This process is called Trasnesterification.
‘Transesterification is the chemical process which replaces one type of
alcohol for another in an ester. An ester is made by combining an alcohol
with an acid.’
Principle:
Lipids are composed of triglycerides, which are esters containing three
free fatty acids and the trihydric alcohol, glycerol. In the
transesterification process, the alcohol is deprotonated with a base to
make it a stronger nucleophile. Commonly, ethanol or methanol are used.
The reaction has no other inputs than the triglyceride and the alcohol.
Under normal conditions, this reaction will proceed either exceedingly
slowly or not at all, so heat, as well as catalysts (acid and/or base) are
used to speed the reaction. It is important to note that the acid or base are
not consumed by the transesterification reaction, thus they are not
reactants, but catalysts. Common catalysts for transesterification include
sodium hydroxide, potassium hydroxide, and sodium methoxide.
Almost all biodiesel is produced from virgin plant oils using the base-
catalyzed technique as it is the most economical process for treating
virgin vegetable oils, requiring only low temperatures and pressures and
producing over 98% conversion yield (provided the starting oil is low in
moisture and free fatty acids).

65. 65
However, biodiesel produced from other sources or by other methods
may require acid catalysis, which is much slower. Since it is the
predominant method for commercial-scale production, only the base-
catalyzed transesterification process will be described below.
Triglycerides are reacted with an alcohol such as ethanol to give ethyl
esters of fatty acids and glycerol :
Figure 29 Transesterification reaction: 1- Glycerol, 2-Base, 3-Esters, 4 byproduct
Base-catalysed transesterification mechanism
The transesterification reaction is base catalyzed. Any strong base
capable of deprotonating the alcohol will do (e.g. NaOH, KOH, sodium
methoxide, etc.), but the sodium and potassium hydroxides are often
chosen for their cost. The presence of water causes undesirable base
hydrolysis, so the reaction must be kept dry.

66. 66
In the transesterification mechanism, the carbonyl carbon of the starting
ester (RCOOR1) undergoes nucleophilic attack by the incoming alkoxide
(R2O−) to give a tetrahedral intermediate, which either reverts to the
starting material, or proceeds to the transesterified product (RCOOR2).
The various species exist in equilibrium, and the product distribution
depends on the relative energies of the reactant and product.[2]

67. 67
Gas Chromatography Mass Spectroscopy
Gas chromatography–mass spectrometry (GC-MS) is a method that
combines the features of gas-liquid chromatography and mass spectrometry
to identify different substances within a test sample. Applications of GC-MS
include drug detection, fire investigation, environmental analysis, explosives
investigation, and identification of unknown samples. GC-MS can also be
used in airport security to detect substances in luggage or on human beings.
Additionally, it can identify trace elements in materials that were previously
thought to have disintegrated beyond identification.
Figure 30 Schematic representation of GC – MS
GC-MS has been widely heralded as a "gold standard" for forensic
substance identification because it is used to perform a specific test. A
specific test positively identifies the actual presence of a particular substance
in a given sample. A non-specific test merely indicates that a substance falls
into a category of substances. Although a non-specific test could statistically

68. 68
suggest the identity of the substance, this could lead to false positive
identification.
After Lipid extraction was carried out the samples were run in GC – MS.
The samples were analysed at Dr. P.S.Ramanathan Advanced Istrumentation
Centre Ramnarain Ruia College, Matunga, Mumbai-400019.
Date of Analysis:6th October 2012.
Following are the results of the same:
1. Microspora sps.

69. 69
2. Enteromopha prolifera.

70. 70
3. Chara Vulgaris.

71. 71
Nile Red Staining
Nile red (also known as Nile blue oxazone) is a lipophilic stain. It is
produced by boiling a solution of Nile blue with sulfuric acid. As can be
seen from the structural formulae, this process replaces an amino group with
a carbonyl group. Nile red stains intracellular lipid droplets red. In most
polar solvents Nile Red will not fluoresce, however when in a lipid-rich
environment can be intensely fluorescent, with varying colours from deep
red to strong yellow-gold emission. Whilst it generally excites at 485 nm,
and emits at 525 nm (552/636 nm in methanol), the fluorescence of the dye
is heavily dependent on the solvent used, and in some cases does not
fluoresce at all.
Since the reaction to generate Nile red does not usually completely exhaust
the supply of Nile blue, additional separation steps are required if pure Nile
red is needed. [7]
Observation:
Figure 31 Nile red staining in
Microspora sp.
Figure 32 Nile red staining in Chara
vulgaris

72. 2
Extraction and Estimation of Protiens from the Algal
Culture.
Aim: To extract Protiens from the given three algal cultures of
1. Microspora species.
2. Entromorpha Prolifera.
3. Chara Vulgaris.
Requirements:
Distilled water, 0.1 N NaOH, 0.5% beta Mercapto Ethanol, Bovine
Serum Albumin, Na2CO3, CuSO4, Sodium Potassium tartarate.
Procudure:
1. 100 mg of biomass was crushed in 4 ml distilled water and
centrifuged at 12000 rpm for 20 mins at 4 C.
2. The supernatant was collected and the pellet was treated with 2ml 2N
NaOH with 5% β-mercaptoethanol.
3. This was kept for 1 hr and centrifuged again at 12000 rpm for 20 mins
at room temperature.
4. The supernatant was mixed with the first supernatant and subjected to
Folin Lowry protocol for estimation of proteins and SDS PAGE.

73. 3
Folin Lowry’s Protein Assay
The Lowry protein assay is a biochemical assay for determining the total
level of protein in a solution. The total protein concentration is exhibited by
a color change of the sample solution in proportion to protein concentration,
which can then be measured using colorimetric techniques. It is named for
the biochemist Oliver H. Lowry who developed the reagent in the 1940s. His
1951 paper describing the technique is the most-highly cited paper ever in
the scientific literatures.
Principle:
The method combines the reactions of copper ions with the peptide bonds
under alkaline conditions (the Biuret test) with the oxidation of aromatic
protein residues. The Lowry method is best used with protein concentrations
of 0.01–1.0 mg/mL. and is based on the reaction of Cu+, produced by the
oxidation of peptide bonds, with Folin–Ciocalteu reagent (a mixture of
phosphotungstic acid and phosphomolybdic acid in the Folin–Ciocalteu
reaction). The reaction mechanism is not well understood, but involves
reduction of the Folin reagent and oxidation of aromatic residues (mainly
tryptophan, also tyrosine). Experiments have shown that cysteine is also
reactive towards to the reagent. Therefore, cysteine residues in protein
probably also contribute to the absorbance seen in the Lowry Assay. The
concentration of the reduced Folin reagent is measured by absorbance at 750
nm. As a result, the total concentration of protein in the sample can be
deduced from the concentration of Trp and Tyr residues that reduce the
Folin reagent.[12]

74. 4
Procedure:
1. Take 20 mg of B.S.A. and add 100ml Distilled water to it.
2. Reagent A – 2 gms of Na2CO3 in 100ml 0.1 N NaOH.
3. Reagent B – 0.5 gms CuSO4 in 1 gm of Na – K tartarate and make
the volume to 100 ml with distilled water.
4. Reagent C – mix Reagent A and B in the proportion of 50:1.
Conc.
Of BSA
(ug/ml)
Vol. of
BSA
(ml)
D/W
(ml)
Rgnt.C Folin’s
Soln.
O.D.
0 0 1 5 0.5 00
20 0.04 0.96 5 0.5 0.08
40 0.08 0.92 5 0.5 0.12
60 0.12 0.88 5 0.5 0.19
80 0.16 0.84 5 0.5 0.21
100 0.2 0.8 5 0.5 0.28
120 0.24 0.76 5 0.5 0.33
140 0.28 0.72 5 0.5 0.36
160 0.32 0.68 5 0.5 0.48
180 0.36 0.64 5 0.5 0.49
200 0.4 0.6 5 0.5 0.49
Microspora - 5 0.5 0.56
Ent. Proli. - 5 0.5 0.32
Chara
vulgaris
- 5 0.5 0.47
- Vortex and keep at room temperature for 10 mins.
- Keep at room temperature for 30 mins.

75. 5
Graph:
Figure 33 Standard graph of proteins
Ab
so
rb
an
ce
at
70
0
n
m
Scale:
X axis – 1 cm=20 µg
Y axis – 1 cm =0.025 units
Concentration µg

76. 6
Observations:
Sample Absorbance Concentration
(by graph in µg/ml)
Micorspora sps. 0.56 206
Enteromorpha
prolifera
0.32 118
Chara vulgaris 0.47 172
Dilution factor:
4 x 6 = 24.
4 = 2ml of D/W used for homogenizing 100mg of biomass x 2ml of (NaOH
+ β mercapto ethanol).
6 = 1ml of the above protein solution + 5 ml of reagent C.
Result:
Sample Concentration
(by graph in µg)
Concentration x
Dilution Factor
(µg)
Micorspora sps. 206 4944
Enteromorpha prolifera 118 2832
Chara vulgaris 172 4128
From the above extraction and estimation process for proteins it was found
that Microspora species contain 4.944%, Enteromorpha prolifera contains
2.832% and Chara vulgaris contains 4.128 % of proteins compared to dry
weight of biomass.

77. 7
Sodium Dodecyl Sulfate - Polyacrylamide Gel
Electrophoresis (SDS – PAGE)
SDS – PAGE is a simple, rapid and highly sensitive tool to analyze
proteins. The separation of proteins by electrophoresis is based on the fact
that charged molecules will migrate through a gel matrix upon application of
an electric field. In most proteins, the binding of SDS to the polypeptide
chain imparts an even distribution of charge per unit mass, thereby resulting
in a fractionation by approximate size during electrophoresis.
Principle:
When charged molecules are placed in an electric field, they migrate
toward either the positive or negative pole according to their charge. In
contrast to proteins, which can have either a net positive or net negative
charge, nucleic acids have a consistent negative charge imparted by their
phosphate backbone, and migrate toward the anode.
Proteins and nucleic acids are electrophoresed within a matrix or "gel".
Most commonly, the gel is cast in the shape of a thin slab, with wells for
loading the sample. The gel is immersed within an electrophoresis buffer
that provides ions to carry a current and some type of buffer to maintain the
pH at a relatively constant value.
As an electric field is applied across the gel, the negatively-charged proteins
to migrate across the gel towards the positive (+) electrode (anode).
Depending on their size, each protein will move differently through the gel

78. 8
matrix: short proteins will more easily fit through the pores in the gel, while
larger ones will have more difficulty (they encounter more resistance). After
a set amount of time (usually a few hours, though this depends on the
voltage applied across the gel; protein migration occurs more quickly at
higher voltages, but these results are typically less accurate than at those at
lower voltages) the proteins will have differentially migrated based on their
size; smaller proteins will have traveled farther down the gel, while larger
ones will have remained closer to the point of origin. Proteins may therefore
be separated roughly according to size (and thus molecular weight); however
certain glycoproteins behave anomalously on SDS gels.
Procedure:
1. Sample preparation.

79. 9
2.Preparing Acrylamide gels.
3. Preparing the plates.
4.Electrophresis.
[8]

80. 10
Given below is the molecular marker used for this experiment.
Observations:
Bands seen in the Molecular Marker:
o B1 – 0.5 cms
o B2 – 1.1 cms
o B3 – 1.8 cms
o B4 – 2.3 cms
kD
95
66
47
40
35
25
20
14
Mol.
Marker
E.
prolifera
Chara
vulgaris

81. 11
o B5 – 3.7 cms
o B6 – 4.5 cms
o B7 – 6.45cms
o B8 – 6.85 cms
Bands seen in Enteromorpha prolifera:
o EP1 – 1.5 cms
o EP2 – 1.9 cms
o EP3 – 2.3 cms
o EP4 – 6.45 cms
Bands seen in Chara vulgaris:
o CV1 – 1.97 cms
o CV2 – 1.38 cms
o CV3 – 1.23 cms
o CV4 – 1.18 cms
Protein degradation was observed in Microspora.

82. 12
Graph:
Figure 34 Standard graph of molecular weight
Scale:
On X axis: 1cm = 0.5 cms
On y axis: 1 cm = 0.1 log mol. wt.

83. 13
Observation:
Proteins extracted from Entromorpha prolifera and Chara vulgaris
showed 4 bands each. Proteins from Microspora sps. could not be detected
on silver stain due to storage related degradation.
Result:
Enteromopha prolifera showed proteins of 20, 45.71, 52.48 and 57.54 kD
mol.wt.
Chara vulgari showed proteins of 15.13, 16.98, 23.98 and 93.32 kD
mol.wt.

84. 14
Extraction and estimation of Carbohydrates using Phenol –
Sulphuric Assay
Aim: To extract carbohydrates from the given three algal cultures of
4. Microspora species.
5. Entromorpha Prolifera.
6. Chara Vulgaris.
Requirements:
Apparatus and Intruments:
1. Eppendoff tube
2. Motar Pestel
3. Standars flask
4. Beakers
5. Weighing balance
Chemicals:
1. Distilled water
2. Sodium Potassium tartarate
3. 2.5 N HCl
4. 96% Sulphuric acid
5. Phenol
6. Sodium Carbonate.

85. 15
Procedure:
 Hydrolyse 100 mg of dry biomass by adding 5ml of 2.5 N HCl and
keeping in boiling water bath for 3 hours.
 Cool to room temperature and neutralize with sodium carbonate
until the effervescence ceases.
 Make up the volume to 100 mL. Pipette out 1 mL of the working
standard into a test tube.
 Set a blank with 1 mL of distilled water. Add 1 ml phenol solution
to each tube and add 5 ml of 96% Sulphuric acid to each tube and
shake well.
 Shake well after 10 mins and keep in water bath at 20-30ᵒC for 20
mins. Orange colour develops for which absorbance is taken at
490 nm.
)bsevation:
Sr. no. Std.
Glucose
(µg)
Std.
Glucose
(ml)
D/W
(ml)
Phenol
(ml)
96%
H2SO4
(ml)
Wait
for
10
mins
then
shake
it and
keep
it in
the
water
bath
for
10
mins.
O.D.
@
490nm
1 Blank 0 1 1 5 0.0
2 200 0.2 0.8 1 5 0.35
3 400 0.4 0.6 1 5 0.58
4 600 0.6 0.4 1 5 0.67
5 800 0.8 0.2 1 5 0.89
6 1000 1 0 1 5 1.18
Microspora 1ml
Sample
- - 1 5 0.69
E. prolifera 1ml
Sample
- - 1 5 0.40
C. vulgaris 1ml
Sample
- - 1 5 0.52

86. 16
11. Conclusion
From above work on characterization of three filamentous green algae we
can conclude that though proteins are in less amount, lipids are are relatively
high. So we can use these lipids for production of biofuels whether diesel or
petrol.
Attempt to produce trans-esterification product using the extracted lipids
was made and inflammation test was performed which could give a crude
idea that fuel can be produced if more sophisticated methods are applied.
Lipid fluorescence technique using Nile red was also performed to check
lipid productivity in situ. This technique can be used in the starvation period
for obtaining greater yield of lipids in algae. We can calculate the harvest
time for these alga at which we can get highest lipid for the industrial use.
Carbohydrates were estimated and found that these algae are rich in
carbohydrates and can be used in diet. Many such algae are already in use
for diet purpose in few countries.

87. 17
12. Discussion.
The experiment conducted showed highest levels of lipid in Enteromorpha
prolifera (28 %). According to the findings of Mr. Alexis Baxter of the Florida
State University, show lipid levels of up to 40 - 45 % but after a growth period of
12 day under laboratory conditions. Whereas our sample was freshly analyzed and
there was no expenditure in growing the culture.

88. 18
13. Bibliography
 Barsanti Laura, Paolo Gualtieri.Algae: Anatomy, Biochemistry, and
Biotechnology.[1]
 Biodiesel: A Realistic Fuel Alternative for Diesel Engines.[2]
 Carl Branden (Author), John Tooze (Author).Introduction to Protein
Structure.[3]
 Classification of Algae: Simthsonian; National museum of Natural
History.[4]
 Graham James E. (Author), Lee W. Wilcox (Author), Linda E. Graham
(Author).Algae (2nd Edition).[5]
 Karen C. Timberlake. General, Organic, and Biological Chemistry:
Structures of Life(3rd Edition )
 Michael I. Gurr (Author), John L. Harwood (Author), Keith N. Frayn
(Author).Lipid Biochemistry: An Introduction. [7]
 Sameh Magdeldin: Gel Electrophoresis - Principles and Basicsmore . [8]
 Sharma. O.P..Textbook Of Algae.[9]
 Singh S. K. and Seema Srivastava, :A Textbook of Algae: Campus Books,
2008.[10]
 Standardized Protocols for Lipids, Proteins & Carbohydrates extraction &
estimation.
Lipids Ref: Bligh E G & Dyer W J. (1959) A rapid method of total lipid
extraction and purification. Can. J. Biochem. Physiol. 37: 911-917.[11]
Proteins Ref : Elisabete Barbarino & Sergio O. Louren¸co, 2005, An
evaluation of methods for extraction and quantification of protein from
marine macro- and microalgae, Journal of Applied Phycology 17: 447–
460.[12]

89. 19
Carbohydrates Ref: 1) Dubois, M, Gilles, K A, Hamilton, J K, Rebers, P
A and Smith, F (1956) Anal Chem 26: 350. 2) Krishnaveni, S, Theymoli
Balasubramanian and Sadasivam, S (1984) Food Chem 15: 229.
Modifications in the referred protocols were made as per requirements.[13]
 Westermeier. Reiner : Electrophoresis in Practice: A Guide to Methods and
Applications of DNA and Protein Separations.[14]