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Morphology, composition, production, processing and applications
of Chlorella vulgaris: A review
Carl Sa
a,b,
n
, Bachar Zebib
a,b
, Othmane Merah
a,b
, Pierre-Yves Pontalier
a,b
,
Carlos Vaca-Garcia
a,b,c
a
Université de Toulouse, INP-ENSIACET, LCA (Laboratoire de Chimie Agro-industrielle), F-31030 Toulouse, France
b
INRA, UMR 1010 CAI, F-31030 Toulouse, France
c
King Abdulaziz University, Jeddah, Saudi Arabia
article info
Article history:
Received 8 March 2013
Received in revised form
9 March 2014
Accepted 6 April 2014
Available online 25 April 2014
Keywords:
Chlorella vulgaris
Algo-renery
Growth conditions
Morphology
Primary composition
Production
abstract
Economic and technical problems related to the reduction of petroleum resources require the
valorisation of renewable raw material. Recently, microalgae emerged as promising alternative feedstock
that represents an enormous biodiversity with multiple benets exceeding the potential of conventional
agricultural feedstock. Thus, this comprehensive review article spots the light on one of the most
interesting microalga Chlorella vulgaris. It assembles the history and a thorough description of its
ultrastructure and composition according to growth conditions. The harvesting techniques are presented
in relation to the novel algo-renery concept, with their technological advancements and potential
applications in the market.
&2014 Elsevier Ltd. All rights reserved.
Contents
1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266
2. Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266
2.1. Cell wall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267
2.2. Cytoplasm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267
2.2.1. Mitochondrion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267
2.2.2. Chloroplast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267
3. Reproduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267
4. Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267
4.1. Autotrophic growth. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268
4.1.1. Open pond systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268
4.1.2. Closed photo-bioreactor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268
4.2. Heterotrophic growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268
4.3. Mixotrophic growth. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268
4.4. Other growth techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268
4.5. Harvesting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269
4.5.1. Centrifugation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269
4.5.2. Flocculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269
4.5.3. Flotation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269
4.5.4. Filtration. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269
5. Primary composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269
5.1. Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269
5.2. Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270
Contents lists available at ScienceDirect
journal homepage: www.elsevier.com/locate/rser
Renewable and Sustainable Energy Reviews
http://dx.doi.org/10.1016/j.rser.2014.04.007
1364-0321/&2014 Elsevier Ltd. All rights reserved.
n
Corresponding author at: Université de Toulouse, INP-ENSIACET, LCA (Laboratoire de Chimie Agro-industrielle), F-31030 Toulouse, France. Tel.: þ33650452965.
E-mail address: csa@me.com (C. Sa).
Renewable and Sustainable Energy Reviews 35 (2014) 265278
Author's personal copy
5.3. Carbohydrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271
5.4. Pigments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271
5.5. Minerals and vitamins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271
6. Cell disruption techniques. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272
7. Applications and potential interests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272
7.1. Biofuels. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272
7.2. Human nutrition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273
7.3. Animal feed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273
7.4. Wastewater treatment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274
7.5. Agrochemical applications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274
8. Algo-renery concept . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275
9. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275
Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275
1. Introduction
Microalgae have an ancient history that left a footprint 3.4 bil-
lion years ago, when the oldest known microalga, belonging to the
group of cyanobacteria, fossilised in rocks of Western Australia.
Studies conrmed that until our days their structure remains
unchanged and, no matter how primitive they are, they still
represent rather complicated and expertly organised forms of life
[1]. Nevertheless, other reports estimated that the actual time of
evolution of cyanobacteria is thought to be closer to 2.7 billion
years ago [2,3]. Hence, evolutionary biologists estimate that algae
could be the ancestors of plants. Thus, through time algae gave rise
to other marine plants and moved to the land during the
Palaeozoic Age 450 millions years ago just like the scenario of
animals moving from water onto land. However, evolutionists
need to overcome multiple obstacles (danger of drying, feed,
reproduction, and protection from oxygen) to denitely conrm
this scenario complemented with more scientic evidence.
Like any other phytoplankton, microalgae have a nutritional
value. The rst to consume the blue green microalga were the
Aztecs and other Mesoamericans, who used this biomass as an
important food source [4]. Nowadays, these microscopic organisms
are still consumed as food supplement such as Chlorella vulgaris and
Spirulina platensis [5] and their products are also used for different
purposes like dyes, pharmaceuticals, animal feed, aquaculture and
cosmetics. For the last two decades, microalgae started to take a
new course with increasing applications motivated by the depletion
of fossil fuel reserves, the consequent increase in oil prices and the
global warming concern. These dramatic thresholds are forcing the
world to nd global strategies for carbon dioxide mitigation by
proposing alternative renewable feedstocks and intensifying
researches on third-generation biofuels. In this context, microalgae
are regarded nowadays as a promising sustainable energy resource
due to their capacity to accumulate large quantities of lipidssuitable
for biodiesel production that performs much like petroleum fuel
[6,7]. They also proved to be a source of products such as proteins,
carbohydrates, pigments, vitamins and minerals [8]. In addition,
microalgae capture sunlight and perform photosynthesis by produ-
cing approximately half of atmospheric oxygen on earth and
absorbing massive amounts of carbon dioxide as a major feed.
Therefore, growing them next to combustion power plants is of
major importance due to their remarkable capacity to absorb carbon
dioxide that they convert into potential biofuel, food, feed and
highly added value components [914].
Microalgae can grow in both fresh and marine water as well as
in almost every environmental condition on earth from frozen
lands of Scandinavia to hot desert soils of the Sahara [15].If
production plants were installed in an intelligent way, microalgae
would not compete with agricultural lands, there would be no
conict with food production [16] and especially would not cause
deforestation.
Microalgae represent an enormous biodiversity from which
about 40.000 are already described or analysed [17]. One of the
most remarkable is the green eukaryotic microalga C. vulgaris,
which belongs to the following scientic classication: Domain:
Eukaryota, Kingdom: Protista, Divison: Chlorophyta, Class: Tre-
bouxiophyceae, Order: Chlorellales, Family: Chlorellaceae, Genus:
Chlorella, Specie: Chlorella vulgaris. Hence, Martinus Willem Bei-
jerinck, a Dutch researcher, rst discovered it in 1890 as the rst
microalga with a well-dened nucleus [18]. The name Chlorella
comes from the Greek word chloros (
Χλωρός
), which means
green, and the Latin sufxella referring to its microscopic size. It
is a unicellular microalga that grows in fresh water and has been
present on earth since the pre-Cambrian period 2.5 billion years
ago and since then its genetic integrity has remained constant [1].
By the early 1900s, Chlorella protein content (455% dry weight)
attracted the attention of German scientists as an unconventional
food source. In the 1950s, the Carnegie Institution of Washington
[19] took over the study and managed to grow this microalga on a
large scale for CO
2
abatement. Nowadays, Japan is the world leader
in consuming Chlorella and uses it for medical treatment [20,21]
because it showed to have immune-modulating and anti-cancer
properties [2226]. After feeding it to rats, mice and rabbits in the
form of powder, it showed protection properties against haema-
topoiesis [27] age-related diseases like cardiovascular diseases,
hypertension and cataract; it lowers the risk of atherosclerosis and
stimulates collagen synthesis for skin [28,29]. Furthermore, C.
vulgaris is also capable of accumulating important amounts of
lipids, especially after nitrogen starvation with a fatty acid prole
suitable for biodiesel production [30,31].
The available reviews have focused so far on evaluating micro-
algae as an important source of lipids for biofuel production
[32,33] and also explained in details the different production
processes and harvesting techniques. The following review covers
greater information about C. vulgaris, including not only produc-
tion and harvesting techniques already conducted on this micro-
alga, but also detailed information about its ultrastructure and
chemical composition accompanied by cell wall breaking techni-
ques and extraction processes. The last section focuses on the
multiple applications and potential interests of this microalga in
different areas and not only on the production of fatty compounds.
2. Morphology
C. vulgaris is a spherical microscopic cell with 210
μ
m dia-
meter [3335] and has many structural elements similar to plants
(Fig. 1).
C. Saet al. / Renewable and Sustainable Energy Reviews 35 (2014) 265278266
Author's personal copy
2.1. Cell wall
The rigidity preserves the integrity of the cell and is basically
a protection against invaders and harsh environment. It varies
according to each growth phase. During its early formation in its
autosporangia, the newly formed cell wall remains fragile, forming
a 2 nm thin electron-dense unilaminar layer [33,36]. The cell wall
of the daughter cell gradually increases in thickness until it
reaches 1721 nm after maturation [33,35], where a microbrillar
layer is formed representing a chitosan-like layer composed of
glucosamine [36,37], which accounts for its rigidity. In the mature
stage, cell wall thickness and composition are not constant because
they can change according to different growth and environmental
conditions. Furthermore, some reports [38,39] explained the rigidity
of the cell wall by focusing on the presence of a sporopollenin layer,
even though it is generally accepted that C. vulgaris has a unilaminar
cell wall that lacks sporopollenin, which is an extremely resistant
polymerised carotenoid found on the cell wall of Haematococcus
pluvialis [40] and Chlorella fusca [41]. However, a contradictory study
conducted on C. vulgaris by Martinez et al. [42] reported the presence
of sporopollenin by observing an outer trilaminar layer and by
detecting resistant residues after being submitted to acetolysis.
2.2. Cytoplasm
It is the gel-like substance conned within the barrier of the
cell membrane and it is composed of water, soluble proteins and
minerals. It hosts the internal organelles of C. vulgaris such as
mitochondria, a small nucleus, vacuoles [43], a single chloroplast
and the Golgi body [44].
2.2.1. Mitochondrion
Every mitochondrion contains some genetic materials, the
respiratory apparatus and has a double-layer membrane; the outer
membrane surrounds the whole organelle and is composed of an
equal ratio of proteins and phospholipids. Nevertheless, the inner
membrane is composed of thrice more proteins than phospholi-
pids; it surrounds the internal space called the matrix, which
contains the majority of mitochondrial proteins [44].
2.2.2. Chloroplast
C. vulgaris has a single chloroplast with a double enveloping
membrane composed of phospholipids; the outer membrane is
permeable to metabolites and ions, but the inner membrane has a
more specic function on proteins transport. Starch granules,
composed of amylose and amylopectin, can be formed inside the
chloroplast, especially during unfavourable growth conditions. The
pyrenoid contains high levels of ribulose-1,5-bisphosphate carbox-
ylase oxygenase (RuBisCO) and is the centre of carbon dioxide
xation. The chloroplast also stores a cluster of fused thylakoids
where the dominant pigment chlorophyll is synthesised masking
the colour of other pigments such as lutein. During nitrogen stress,
lipid globules mainly accumulate in the cytoplasm and the
chloroplast [15,45].
3. Reproduction
C. vulgaris is a non-motile reproductive cell (autospore) that
reproduces asexually and rapidly. Thus, within 24 h, one cell of
C. vulgaris grown in optimal conditions multiplies by autosporula-
tion, which is the most common asexual reproduction in algae.
In this manner, four daughter cells having their own cell wall
are formed inside the cell wall of the mother cell (Figs. 2 and 3)
[33,35]. After maturation of these newly formed cells, the mother
cell wall ruptures, allowing the liberation of the daughter cells and
the remaining debris of the mother cell will be consumed as feed
by the newly formed daughter cells.
4. Production
Annual production of Chlorella reached 2000 t (dry weight) in
2009, and the main producers are Japan, Germany and Taiwan
[46]. This microalga has a rapid growth rate and responds to
each set of growth condition by modifying the yield of a specic
component. C. vulgaris is ideal for production because it is
remarkably resistant against harsh conditions and invaders. On
the one hand, lipid and starch contents increase and biomass
productivity ceases or decreases [47] during unfavourable growth
conditions such as nitrogen and phosphorus limitation, high CO
2
concentration, excessive exposure to light [30,4850], excess of
iron in the medium [51] or increase in temperature [52]. On the
other hand, protein content increases during normal and managed
growth conditions (nitrogen supplementation). Therefore, many
growth techniques have been tested in order to voluntarily target
biomass productivity, lipid, proteins, carbohydrates and pigments
content.
Fig. 2. Drawings showing the different phases of daughter cell-wall formation in Chlorella vulgaris: (a) early cell-growth phase; (b) late cell-growth phase; (c) chloroplast
dividing phase; (d) early protoplast dividing phase; (e) late protoplast dividing phase; (f) daughter cells maturation phase and (g) hatching phase [35].
Fig. 1. Schematic ultrastructure of C. vulgaris representing different organelles.
C. Saet al. / Renewable and Sustainable Energy Reviews 35 (2014) 265278 267
Author's personal copy
4.1. Autotrophic growth
4.1.1. Open pond systems
Open ponds are the most common way of production and are
the cheapest method for large-scale biomass production. These
systems are categorised into natural waters (lakes, lagoons and
ponds) or wastewater or articial ponds or containers. They are
usually built next to power plants or heavy industry with massive
carbon dioxide discharge where the biomass absorbs nitrogen
from the atmosphere in the form of NO
x
. In order to allow easy
exposure of all the cells to sunlight, especially at the end of the
exponential growth phase, the optimal pond depth is 1550 cm
[46,52]. On the other hand, open pond systems have some
limitations because they require a strict environmental control to
avoid the risk of pollution, water evaporation, contaminants,
invading bacteria and the risk of growth of other algae species.
In addition, temperature differences due to seasonal change cannot
be controlled and CO
2
concentration and excess exposure to sun-
light are difcult to manage. Moreover, near the end of the
exponential growth phase, some cells are not sufciently exposed
to sunlight because other cells oating near the surface cover them,
leading to lower mass yields. Therefore, stirring of the medium is
preferable and is currently practiced.
4.1.2. Closed photo-bioreactor
This technology was implemented mainly to overcome some
limiting factors in the open pond systems, thus growing the
biomass in a managed environment (pH, light intensity, tempera-
ture, carbon dioxide concentration) to obtain higher cell concen-
tration as well as products that are more suitable for the
production of pure pharmaceuticals, nutraceuticals and cosmetics.
In addition, these systems are more appropriate for sensitive
strains that cannot compete and grow in harsh environment.
Feeding the biomass with CO
2
comes by bubbling the tubes.
Fluorescent lights are used in case the tubes are not or not
sufciently exposed to sunlight. The tubes are generally 20 cm or
less in diameter [32] and the thickness of their transparent walls is
few millimetres, allowing appropriate light absorption. Hence,
multiple designs have been used and tested: at-plate photo-
bioreactor [53,54], tubular photo-bioreactor [55] and column
photo-bioreactor [56]. Degen et al. [57] achieved 0.11 g L
1
h
1
dry biomass productivity after growing the cells of C. vulgaris in a
at panel airlift photobioreactor under continuous illumination
(980
μ
Em
2
s
1
). Nonetheless, the main disadvantages of a
closed system are the cost of the sophisticated construction, small
illumination area and sterilising costs [58].
4.2. Heterotrophic growth
This technique does not require light and the biomass is fed
with organic carbon source. Thus, microalgae are grown in a
stirred tank bioreactor or fermenter where a higher degree of
growth are expected as well as low harvesting cost due to the
higher dry biomass productivity achieved (up to 0.25 g L
1
d
1
)
and high accumulation of different components such as lipids
2254 mg L
1
d
1
[42,59,60]. The carbon sources used for C.
vulgaris are glucose, acetate, glycerol and glutamate with max-
imum specic growth rate obtained with glucose. Nevertheless,
the major disadvantage of this system is the price and availability
of sugars, which compete with feedstocks for other uses such as
food and biofuel productions.
4.3. Mixotrophic growth
C. vulgaris is capable of combining both autotrophic and
heterotrophic techniques by performing photosynthesis as well
as ingesting organic materials such as glucose, which is the most
appropriate for C. vulgaris[5963]. Hence, the cells are not strictly
dependent on light or organic substrate to grow. This technique
competes favourably with autotrophic systems and according to
Yeh and Chang [63] mixotrophic conditions showed high dry
biomass productivity (25gL
1
d
1
) and lipids productivity
(67144 mg L
1
d
1
). The main advantages of mixotrophic meta-
bolism are limiting the impact of biomass loss during dark
respiration and reducing the amount of organic substrates used
for growing the biomass.
4.4. Other growth techniques
Growth of C. vulgaris can take an additional dimension by co-
immobilising it with plant growing bacterium Azospirillum brasi-
lense in alginate beads [64,65]. This technique has been extra-
polated to C. vulgaris and other microalgae from the hypothesis
that A. brasilense promotes terrestrial plant growth performance
by interfering with the host plant hormonal metabolism and
provides O
2
for the bacteria to biodegrade pollutants and then
the microalga consumes CO
2
released from bacterial respiration
[66]. Consequently, depending on the strain of C. vulgaris [67] this
technique has an impact on prolonging its life span, enhancing
biomass production, increasing cell size (62% larger) and accumu-
lating pigments and lipids. Simultaneously, uptake of zinc, cad-
mium, phosphorus, nitrogen and other heavy metals from
wastewater increases. On the other hand, growing C. vulgaris with
its associative bacterium Phyllobacterium myrsinacearum also has a
different impact by ceasing its growth or cell death [68]. Further-
more, mixing and shear stress have an effect on increasing the
photosynthetic activity and growth of C. vulgaris. Thus, optimal
conditions (tip speed of 126 cm s
1
and friction velocity
2.06 cm s
1
) increased the photosynthetic activity by 45% with
4871% stronger growth compared to null tip speed or friction
velocity. Nevertheless, higher tip speed and friction velocity
decreased both photosynthetic activity and growth to the value
of the unstirred condition and even lower [69].
Fig. 3. Newly formed cells emerging outside the cell wall of the mother cell after
hatching [33].
C. Saet al. / Renewable and Sustainable Energy Reviews 35 (2014) 265278268
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4.5. Harvesting
4.5.1. Centrifugation
This process contributes to 2030% of the total biomass
production cost [55]. The most common harvesting technique for
C. vulgaris is centrifugation (5000 rpm, 15 min) [30,70] because it
is highly efcient (95% recovery), not time consuming, and treats
large volumes. In addition, the morphology of C. vulgaris permits
high centrifugal stress without damaging its structure during the
process. Other techniques are also applied such as occulation,
otation and ltration or by combining two techniques to max-
imise recovery of the biomass.
4.5.2. Flocculation
During the exponential growth phase, the algal cells have
high negative surface charge and are difcult to neutralise, and
thus the cells remain dispersed. After reaching the stationary
or the declining phase, the negative charge decreases, allowing
the cells to aggregate and to form lumps, thereby resulting
in a process called auto-occulation. This phenomenon is asso-
ciated with elevated pH due to CO
2
, nitrate and phosphate assimi-
lation [71]. Moreover, auto-occulation can occur by interactions
between algae and bacteria or excreted organic molecules or by
simply cutting CO
2
supply; this method is less expensive but time-
consuming. In general, culture of microalgae is very stable and
auto-occulation probability is negligible and sometimes mislead-
ing. In order to accelerate coagulation, it is necessary to increase
the pH by adding a base. The most effective is sodium hydroxide,
which induces more than 90% occulation at pH 11 and requires
less quantity (9 mg of NaOH per gram of dry biomass) [71,72]. But
on an industrial scale, lime seems to be the most cost-efcient.
This mechanism is associated with Mg
2þ
from hydrolysed Mg
(OH)
2
, which precipitates attracting with it the negatively charged
microalgal cells. Chitosan is also an interesting occulating agent
[73], which showed maximum efciency at pH 7 with 90%
microalgal recovery. Further on, using bioocculants like Paeniba-
cillus sp. with the presence of a co-occulant (CaCl
2
) also showed
an efcient occulation (83%) at pH 11 [74]. Flocculation is some-
times considered as a pre-harvesting step in order to facilitate
or complement other harvesting methods like centrifugation or
ltration [75,76].
4.5.3. Flotation
To ourknowledge, there is very limited evidence ofits feasibility,
but this method consists of trapping the cells using dispersed
micro-air bubbles. Flotation can also occur naturally when the lipid
content in microalgae increases. Cheng et al. [77] induced effective
otation on C. vulgaris by using dispersed ozone gas (0.05 mg g
1
biomass). Thus, unlike occulation, this method does not require
synthetic chemicals, but its economic viability is not yet known,
especially on an industrial scale.
4.5.4. Filtration
This method involves continuous passing of the broth with the
microalga across a lter on which algal cells will concentrate
constantly until it reaches a certain thickness. Due to the small size
of C. vulgaris, conventional ltration is not an adequate method
to be applied. Instead, ultraltration or microltration is more
efcient. Fouling generated by soluble compounds like exopoly-
saccharides of some microalgae such as Porphyridium is one of
the major limitations during the ultraltration process, but with
Chlorella this phenomenon is negligible, and thus its structure
provides more important permeation ux without the need of an
additional unit operation like swirling while ltering [78,79].More-
over, microltration and ultraltration are affected by different
parameters such as lter type; transmembrane pressure, ow
velocity, turbulent cross-ow and growth phase, and therefore
a compromise that takes into consideration these parameters
should be made. Furthermore, they can be accompanied by another
harvesting technique (otation or occulation) that improves the
process [75,76,80].
5. Primary composition
5.1. Proteins
Proteins are of central importance in the chemistry and
composition of microalgae. They are involved in capital roles such
as growth, repair and maintenance of the cell as well as serving as
cellular motors, chemical messengers, regulators of cellular activ-
ities and defence against foreign invaders [44].
Total proteins content in mature C. vulgaris represents 4258%
of biomass dry weight [8185], and varies according to growth
conditions. Proteins have multiple roles, and almost 20% of the
total proteins are bound to the cell wall, more than 50% are
internal and 30% migrate in and out of the cell [86]. Their
molecular weight revealed by SDS-PAGE comprises between 12
and 120 kDa, with the majority between 39 and 75 kDa after
growing C. vulgaris under autotrophic or heterotrophic conditions.
Nevertheless a higher intensity peak is observed for cells grown in
autotrophic conditions [82,87].
Protein nutritional quality is determined by its amino acid
prole [81,88], and like the majority of microalgae, the amino acid
prole of C. vulgaris compares favourably and even better with the
standard prole for human nutrition proposed by World Health
Organisation (WHO) and Food and Agricultural Organisation
(FAO), because the cells of C. vulgaris synthesise essential and
non-essential amino acids (Table 1). Furthermore, regardless of the
extraction procedure, C. vulgaris proteins showed excellent emul-
sifying capacity [89] that is comparable and even better than the
commercial ingredients. Results showed that the emulsifying
capacity of C. vulgaris proteins extracted at pH¼7 reached
3090750 mL oil/g protein with a stability of 7971%. Therefore,
proteins of C. vulgaris open the gate for additional valorisation
options of this microalga in the market, especially in the food
sector.
Protein extraction is technically the same for all microalgae and
is mainly conducted by solubilisation of proteins in alkaline
solution [83,90,91]. Further purication can be followed by pre-
cipitating the solubilised proteins with trichloroacetic acid (25%
TCA) [92,93] or hydrochloric acid (0.1 N HCl) [94]. Another
separation method could be applied by means of ultraltration.
Indeed, this method is usually applied for harvesting the cells
but considering the study conducted by Saet al. [95], a two-stage
ultraltration process was applied on the aqueous extract of
Tetraselmis suecica containing solubilised molecules (starch, pro-
teins and low molecular weight polysaccharides). The rst phase
of the process completely retained starch molecules, and then the
second phase completely retained proteins, allowing only small
polysaccharides to be present in the ltrate of the second phase
of the process. This process could be extrapolated to C. vulgaris
with minor modications of the cut-off of the ultraltration
membranes [95].
Quantication is carried out by elemental analysis, Kjeldahl,
Lowry assay, Bradford assay or the dye binding method. However,
the rst two analyses take into consideration total nitrogen
present in the microalga, and multiplying it by the standard
nitrogen to protein conversion factor (NTP) 6.25 may lead to
overestimation or underestimation of the true protein quantity.
Therefore, several studies calculated from an amino acid prole
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recommended a new NTP lower than the standard 6.25 [9610 0] .
Nevertheless, a study conducted by Saet al. [83] correlated the
evaluation of the NTP to the rigidity of the cell wall by evaluating
the NTP of ve crude microalgae including C. vulgaris and their
protein extract, and concluded that no universal conversion factor
could be recommended for multiple reasons such as cell wall
rigidity, growth conditions, growth media and environmental
uncertainty. Gonzalez-Lopez et al. [97] determined the NTP using
a different technique that correlates protein content (Lowry assay)
to total nitrogen content (Kjeldahl and elemental analysis) and
also estimated that the Kjeldahl method correlates better with the
Lowry assay. In addition, Servaites et al. [84] quantied proteins of
12 different microalgae including C. vulgaris by staining the
protein isolate with Coomassie brilliant blue R-250 (CBB) on a
paper and then eluting the remaining stained proteins in 1%
sodium dodecyl sulphate (SDS) followed by measuring the absor-
bance at 600 nm. This method gave almost similar results com-
pared to the Dumas method. On the other hand, the colorimetric
method of Lowry [101] was also considered as one of the most
accurate methods to quantify proteins [102], but with time this
method showed to only quantify hydro-soluble proteins
[83,88,101105], which represents the major part of proteins.
The Lowry assay is more acceptable than the Bradford assay
because the latter does not react with all the amino acids present
in the extract, thus giving lower protein concentrations [92].
5.2. Lipids
Lipids are a heterogeneous group of compounds that are
dened not by their structure but rather by the fact that they
are soluble in non-polar solvents and relatively insoluble in water
[90]. During optimal growth conditions C. vulgaris can reach 540%
lipids per dry weight of biomass [81], and are mainly composed of
glycolipids, waxes, hydrocarbons, phospholipids, and small
amounts of free fatty acids [15,17]. These components are synthe-
sised by the chloroplast and also located on the cell wall and on
membranes of organelles (chloroplast and mitochondrion mem-
branes). Nevertheless, during unfavourable growth conditions,
lipids content (mainly composed of triacyglycerols) can reach
58% [8,81,106]. Unlike other lipids, triacylglycerols do not perform
a structural role but instead accumulate as dense storage lipid
droplets in the cytoplasm and in the inter-thylakoid space of the
chloroplast [17].
Liu et al. [51] optimised a method that detects the accumula-
tion of lipid droplets inside the cells of C. vulgaris after each
growth phase. The method relies on staining the cells with Nile
red dye and then observing the accumulation of lipids with
uorescence microscope by emitting blue light that reveals the
lipid droplets, especially neutral lipids. This technique showed a
correlation between the quantity of neutral lipids accumulated
and uorescence intensity. However, according to Chen et al. [107]
without cell disruption, this method could be ineffective due to
the presence of a thick cell wall of some microalgae that can
prevent complete access of the reagent inside the cell. Thus, cell
disruption is a necessity to prevent wrong measurements and
quantication.
The extraction process of total lipids from C. vulgaris is
generally conducted by the method of Bligh and Dyer (a mixture
of chloroform and methanol), or by hexane, or petroleum ether
[31,49,51,58,108110]. Quantication of total lipids is conducted
gravimetrically after evaporating the extracting solvent; in addi-
tion, column chromatography is carried out in order to separate
different lipid constituents followed by evaporating the solvent
and then weighing the remaining lipid extract [111]. Indeed, these
solvents are not used on an industrial scale because they are
harmful for the environment, toxic, highly ammable and con-
taminate the extract [109]. Total lipids are composed of three
major fractions phospholipids (PL), glycolipids (GL) and neutral
lipids (NL). These fractions are fractionated by sequential elution of
chloroform and acetic acid for NL, acetone and methanol for GL,
and methanol for PL recovery [111]. Supercritical carbon dioxide
(SC-CO
2
) extraction has been identied as an alternative for a
greener extraction since it gives pure extracts free of contamina-
tion. Moreover, in order to increase the yield of extraction, a co-
solvent to SC-CO
2
such as ethanol can be used or a preliminary cell
disruption technique can be performed [112]. It is noteworthy that
the addition of ethanol increases the extraction yield of total
lipophilic molecules (lipids and pigments), but it could also bypass
the energetic yet efcient cell disruption technique, and therefore
the production cost could be signicantly reduced [113].
Table 1
Amino acid prole of Chlorella vulgaris compared to other resources expressed in grams per 100 g of protein.
Amino acids C. vulgaris
b
C. vulgaris
a
C. vulgaris
c
Recommendation from FAO/WHO
b
Eggs
b
Soya
b
Aspartic acid 9.30 10.94 9.80 N/A 11.00 1.30
Threonine 5.30 6.09 5.15 4.00 5.00 4.00
Serine 5.80 7.77 4.32 N/A 6.90 5.80
Glutamic acid 13.70 9.08 12.66 N/A 12.60 19.00
Glycine 6.30 8.60 6.07 N/A 4.20 4.50
Alanine 9.40 10.90 8.33 N/A n.d 5.00
Cysteine n.d 0.19 1.28 3.50 2.30 1.90
Valine 7.00 3.09 6.61 5.00 7.20 5.30
Methionine 1.30 0.65 1.24 N/A 3.20 1.30
Isoleucine 3.20 0.09 4.44 4.00 6.60 5.30
Leucine 9.5 7.49 9.38 7.00 7.00 7.70
Tyrosine 2.80 8.44 3.14 6.00 4.20 3.20
Phenylalanine 5.50 5.81 5.51 N/A 5.80 5.00
Histidine 2.00 1.25 1.97 N/A 2.40 2.60
Lysine 6.40 6.83 6.68 5.50 5.30 6.40
Arginine 6.90 7.38 6.22 N/A 6.20 7.40
Tryptophan n.d 2.21 2.30 1.00 1.70 1.40
Ornithine n.d 0.13 n.d N/A n.d n.d
Proline 5.00 2.97 4.90 N/A 4.20 5.30
n.d: not detected; N/A: not available.
a
[83].
b
[192,193].
c
[194].
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The fatty acid prole changes with respect to growth condi-
tions and is suitable for different applications. For instance,
according to Yeh and Chang [63], the fatty acid prole of C. vulgaris
grown under mixotrophic growth conditions can accumulate
6068% saturated and monounsaturated fatty acids composed of
palmitic acid C16:0, stearic acid C18:0 fatty acids, palmitoleic acid
C16:1 and oleic acid C18:1 [31]. Such a prole is more suitable for
biodiesel production [114]. On the contrary, if it is grown under
favourable growth conditions, its fatty acid prole is unsuitable
for biodiesel [106] but more suitable for nutritional uses because
it is more concentrated in polyunsaturated fatty acids such as
linoleic acid C18:2, linolenic acid C18:3, and eicosapentaenoic acid
C20:5 [107].
5.3. Carbohydrates
Carbohydrates represent a group of reducing sugars and poly-
saccharides such as starch and cellulose. Starch is the most
abundant polysaccharide in C. vulgaris. It is generally located in
the chloroplast and is composed of amylose and amylopectin, and
together with sugars they serve as energy storage for the cells.
Cellulose is a structural polysaccharide with high resistance, which
is located on the cell wall of C. vulgaris as a protective brous
barrier. In addition, one of the most important polysaccharides
present in C. vulgaris is the
β
1-3 glucan [115], which has multiple
health and nutritional benets.
Total carbohydrates are generally quantied by the sulphuric
phenol method [116,117], yielding simple sugars after hydrolysis at
110 1C, then quantication of the latter by HPLC (especially HPIC).
Starch quantication is much better using the enzymatic method
compared to the acidic method [118,119]. During nitrogen limita-
tion, total carbohydrates can reach 1255% dry weight.[120,121].
Moreover, C. vulgaris has a remarkably robust cell wall [122],
mainly composed of a chitosan like layer, cellulose, hemicellulose,
proteins, lipids and minerals [123125].
The sugar composition (Table 2) of the cell wall is a mixture of
rhamnose, galactose, glucose, xylose, arabinose and mannose
[126130], rhamnose being the dominant sugar [128,131,132].
5.4. Pigments
The most abundant pigment in C. vulgaris is chlorophyll, which
can reach 12% dry weight and is situated in the thylakoids. C.
vulgaris also contains important amounts of carotenoids (Table 3)
that act as accessory pigments by catching light;
β
-carotene for
instance is associated with the lipid droplets in the chloroplast, and
primary carotenoids are associated with chlorophyll in thylakoids
where they trap light energy and transfer it into the photosystem.
However, as in terrestrial plants, some pigments act as photo-
protectors by protecting chlorophyll molecules from degradation
and bleaching during strong exposure to radiation and oxygen [44].
These pigments have multiple therapeutic properties, such as
antioxidant activities [133], protective effect against retina degen-
eration [134,135], regulating blood cholesterol, prevention from
chronic diseases (cardiovascular and colon cancer) and fortifying
the immune system [136,137]. Pheophytins are biochemically
similar to chlorophyll but lacking Mg
þþ
ion; they can form after
chlorophyll degradation during the growth of microalgal cells or
during harsh extraction conditions. In addition, these pigments are
lipophilic and their extraction is generally associated with lipid
extraction.
Many studies worked on optimising the extraction process of
pigments using solvents (dimethyl formamide, dichloromethane,
acetone, hexane, and ethanol), soxhlet, ultrasound-assisted extrac-
tion [70,138141], and pressurised liquid extraction (PLE) that
showed useful simultaneous extraction of carotenoids and chlor-
ophyll, and also minimised the formation of pheophytins [70,142]
at high temperature (4110 1C). Moreover, SC-CO
2
extraction was
also carried out to enhance carotenoids recoveries, and the best
conditions were 35 MPa and 4055 1C on crushed cells, and under
these conditions the extract was golden and limpid unlike solvents
extraction; thus by using SC-CO
2
, higher selectivity can be
achieved [139,142]. This hypothesis was conrmed by Kitada
et al. [20], using different optimum conditions (50 MPa and
80 1C) because the study was conducted on whole cells; thus
stronger conditions were required. In addition, co-solvent such as
5% ethanol has been added as a booster to increase the extraction
yield. Analyses and quantication of pigments are conducted by
high performance liquid chromatography (HPLC) and spectro-
photometry using specic equations [143] or by plotting the
calibration curve for each pigment.
5.5. Minerals and vitamins
Minerals are determined after incinerating the biomass and
then analysis by atomic absorption spectrophotometry (Table 4).
They play important functional roles in humans [44]. For instance,
potassium cation is principal for human nutrition; it is associated
with intracellular uid balance, carbohydrate metabolism, protein
synthesis and nerve impulses. In addition, it is used as chemical
fertilizer in agriculture in the form of chloride (KCl), sulphate
(K
2
SO
4
) or nitrate (KNO
3
). Magnesium is important in maintaining
normal and constant nervous activity and muscle contraction;
hence magnesium deciency in human organism can lead to
depression and symptoms of suicidal behaviour. Zinc is an essen-
tial component of enzymes, which participates in many metabolic
processes including synthesis of carbohydrates, lipids and proteins
and it is also a cofactor of the superoxide dismutase enzyme,
which is involved in the protection against oxidative processes and
reducing the severity of strong diarrhoea.
Table 3
Potential pigments content in C. vulgaris under different growth conditions.
Pigments μgg
1
(dw) References
β-Carotene 712,000 [20,65,70,139,170]
Astaxanthin 550,000 [170,195,196]
Cantaxanthin 362,000 [139,140,170,195]
Lutein 523830 [20,65]
[67,70]
[139,170]
Chlorophyll-a2509630 [65]
[20,67]
[68,139]
Chlorophyll-b725770 [65]
[20,67]
[70,139]
Pheophytin-a23105640 [70]
Pheophytin-bN/A [70]
Violoxanthin 1037 [65]
[67]
N/A: not available.
Table 2
Simple sugars composition of the cell wall poly-
saccharides [128].
Neutral sugars Percentage (%)
Rhamnose 4554
Arabinose 29
Xylose 719
Mannose 27
Galactose 1426
Glucose 14
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Vitamins are classied as water-soluble (C and B) and fat-
soluble (A, D, E, and K). C. vulgaris has an important vitamin prole
(Table 5) that are key elements for cell growth and differentiation
in the human body (Vitamin A), and have antioxidant activity that
acts as radical scavenger together with improving blood circula-
tion and controlling muscle functions (Vitamins E and C) [144].
Vitamin B complex occupies the largest number in living organ-
isms and is a major actor for enzymes activity in metabolism [145],
promotes red blood cells growth, reduces the risk of pancreatic
cancer, and maintains healthy skin, hair and muscles. Vitamins
prole is sensitive to growth conditions; thus the best concentra-
tion was achieved after 24 h autotrophic growth with 10% CO
2
, but
during heterotrophic conditions vitamins content was higher than
autotrophic due to the presence of glucose in the medium and
used as carbon source to produce organic compounds [87].
Another possible explanation for the high content of vitamins
may be the alterations in the ultrastructure of the photosynthetic
apparatus which were found to be associated with changes in
cellular components [146].
6. Cell disruption techniques
C. vulgaris has a resistant cell wall, which is a major barrier for
digestibility and extraction process of all internal components.
Breaking the cell wall is an important challenge and a costly unit
operation. Multiple techniques have been carried out on C. vulgaris
(Table 6). Cooling the system during mechanical cell breaking is
always required because the high-energy input overheats the
broken microalga and jeopardises the integrity of target compo-
nents by damaging or oxidising them. Enzymatic treatment is a
promising technique that requires a deep understanding of the
ultrastructure and composition of the cell wall in order to select
the appropriate enzyme and to reduce the enzyme concentration
required to hydrolyse the cell wall. According to Lee et al. [108]
and Zheng et al. [31] the best cell disruption techniques with 30%
dry weight lipid recovery of C. vulgaris grown under autotrophic
conditions were autoclaving, microwave, enzymatic and grinding
with liquid nitrogen. Nonetheless, the quality of the target mole-
cules is susceptible to be different with respect to the cell disruption
method applied. Thus, the amino acid prole of proteins obtained
after conducting an alkaline treatment on C. vulgaris is different
from the amino acid prole obtained after high-pressure homo-
genisation [147].
The success of cell disruption techniques is generally assessed
by conducting microscopic observations or by comparing the
extracted yield of a component before and after applying the cell
disruption.
7. Applications and potential interests
7.1. Biofuels
Dependency on energy sources is growing faster, especially
with the exponential increase in demand, which is leading to more
dramatic consequences for the environment. Third generation
biofuel form algae or microalgae is considered as one of the
alternatives to current biofuel crops such as soybean, corn, rape-
seed and lignocellulosic feedstocks because it does not compete
with food and does not require arable lands to grow [16]. However,
biofuel from microalgae is promising in the long term because it is
now accepted that the production cost is still high and cannot yet
compete with conventional fuel. But it competes favourably with
crops by their potential of producing 1020 times more oil [148]
within a shorter period of time. As mentioned previously, C.
vulgaris has the potential to accumulate high amounts of lipids,
especially while growing it under mixotrophic conditions. Its fatty
acid prole showed to be suitable for biodiesel production with an
oxidative stability after transforming it to biodiesel, and has
properties [149] that comply with the US Standard (ASTM 6751),
European Standard (EN 14214), Brazilian National Petroleum
Agency (ANP 255) and Australian Standard for biodiesel [150]
and also compared favourably with (ASTM and EN) an Indian
biodiesel standard [61]. After lipid extraction the remaining
residue is rich in proteins, carbohydrates and minor amounts of
lipids. Thus, Wang et al. [149] applied fast pyrolysis on C. vulgaris
remnants using an atmospheric-pressure uidised bed reactor at
500 1C and obtained bio-oil and biochar representing 94% of
energy recovery from the remnant, without forgetting the small
amount of biogas recovered. However, the quality of bio-oil was
poor due to the presence of nitrogen in signicant amounts (12.8%
dry weight). Besides, C. vulgaris has high starch content and algal
starch proved to be a good source for bioethanol production.
Hirano et al. [151] extracted starch from C. vulgaris and achieved
65% ethanol-conversion rate after saccharication and fermenta-
tion with yeast. Hydrothermal liquefaction is another alternative
route for biofuel production from microalgae. It involves the
reaction of biomass in water at high temperature with or without
the presence of a catalyst to obtain bio-crude [152]. The main
advantage of this method is that it improved 1015% the energetic
Table 5
Vitamins prole of C. vulgaris.
Vitamins Content (mg 100 g
1
)
Maruyama
et al. [203]
Yeh
et al. [114]
Panahi
et al. [198]
B1 (Thiamine) 2.4 N/A 1.5
B2 (Riboavin) 6.0 N/A 4.8
B3 (Niacin) N/A N/A 23.8
B5 (Pantothenic acid) N/A N/A 1.3
B6 (Pyridoxine) 1.0 N/A 1.7
B7 (Biotin) N/A N/A 191.6
B9 (Folic acid) N/A N/A 26.9
B12 (Cobalamin) tr N/A 125.9
C (Ascorbic acid) 100.0 39.0 15.6
E (Tocopherol) 20.0 2787.0 N/A
A (Retinol) N/A 13.2 N/A
tr: traces; N/A: not available.
Table 4
Minerals prole of C. vulgaris.
Minerals Mineral content (g 100 g
1
)
Maruyama et al. [203] Tokusoglu and Unal [197] Panahi et al. [198]
Microelements
Na N/A 1.35 N/A
K 1.13 0.05 2.15
Ca 0.16 0.59 0.27
Mg 0.36 0.34 0.44
P N/A 1.76 0.96
Macroelements
Cr N/A tr tr
Cu N/A tr 0.19
Zn N/A tr 0.55
Mn N/A tr 0.40
Se N/A tr N/A
I N/A N/A 0.13
Fe 0.20 0.26 0.68
tr: traces; N/A: not available.
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value of C. vulgaris by acting on the whole biomass, suggesting that
oil is also derived from carbohydrates and proteins [153], and thus
no need to stress the microalgae to increase lipid content. Hence,
the best conditions applied on C. vulgaris in a batch reactor were
300350 1C, with 150200 bar in water or with the presence of an
organic acid or heterogeneous catalysts, and the results indicate
that bio-oil formation follows the trend lipids4proteins 4carbo-
hydrates [152154].
Nowadays, algal biofuel is suffering from several drawbacks,
jeopardising its commercialisation on an industrial scale due to
high production cost that is far from being competitive with fossil
fuel, and also questioning the sustainability of this production.
Hence, different studies considered life cycle assessment analysis
as an effective tool to identify the reasons leading to production
decit and exploring its environmental impact [155162]. There-
fore, it was agreed that the major costs come from infrastructure,
production set-up, fertilizers, harvesting, drying the biomass,
transportation, water footprints, cell disruption and oil extraction
process. For instance, Lardon et al. [163] performed an analysis by
taking into account all the energetic debt for 1 MJ biodiesel
production from C. vulgaris. The only positive balance obtained
was 0.57 MJ for wet oil extraction with low nitrogen for cell
growth (Table 7), and all the other revealed negative balance.
Hence, microalgal biofuel production still needs efcient improve-
ment to reduce energy input needed in order to reach competitive
prices with petroleum in the market, and more important to be an
overall sustainable production.
7.2. Human nutrition
C. vulgaris is one of the few microalgae that can be found in the
market as a food supplement or additive [5,140], colourant (C.
vulgaris after carotenogenesis) and food emulsion [119]. These
products come in different forms such as capsules, tablets, extracts
and powder [164,165]. Nevertheless, despite all the healthy
benets that C. vulgaris and other microalgae can provide, with
their remarkable richness in proteins, lipids, polysaccharides,
pigments and vitamins, they are rather considered as nutraceu-
ticals instead of food products due to the lack of clear common
ofcial legislations in terms of quality and requirements regarding
microalgae [166,167]. Moreover, C. vulgaris extract proved to have
preservative activity higher than those obtained synthetically, i.e.,
butylated hydroxyanisole (BHA) and butylated hydroxytoluene
(BHT) [168].
7.3. Animal feed
It is estimated that about 30% of microalgal production is sold
for animal feed purposes [169] due to the increasing demand for
Table 7
Cumulative energy demand and energy production associated with the production
of 1 MJ of biodiesel from C. vulgaris [159].
Oil
extraction
Nitrogen for
culture
Energy
production (MJ)
Cumulative energy
demand (MJ)
Yield
(MJ)
Dry Sufcient 2.7 5.29 2.59
Wet Sufcient 3.84 3.99 0.15
Dry Low 1.57 2.32 0.75
Wet Low 2.23 1.66 0.57
Table 6
Different cell disruption techniques carried out on C. vulgaris.
Cell disruption Time Experimental set-up References
Acid treatment 25 min Hot Ac
2
OþH
2
SO
4
(9:1, v-v) [70]
Alkaline treatment 60 min 2 N NaOH [83]
Autoclaving 5 min 125 1Cþ1.5 MPa [106]
Bead milling 20 min Beads: 0.40.6 mm [31]
Rotational speed: 1500 rpm
5 min Beads: 0.1 mm, [106]
Rotational speed: 2800 rpm
2 min Beads: 1 mm [59]
Electroporation N/A Electric eld: 3 kV/cm [73]
Electrode: 2 cm
Enzymatic lysis 60 min Snailase (5 mg L
1
), 37 1C[31]
10 h Cellulase or lysozyme (5 mg L
1
), 55 1C
N/A 4% Cellulaseþ1% others (w/v) [199]
25 mM sodium phosphate buffer
pH 7.0
0.5 M mannitol
10 h 4% Cellulase þ1% macerozyme R10 þ1% pectinase (w/v) [90]
pH 6.0
25 mM phosphate buffer
0.6 M sorbitol/mannitol (1:1)
24 h Cellulase 0.5 mg L [200]
0.5 M mannitol
French press N/A 138 MPa [201]
N/A N/A [78]
Manual grinding 110 min With liquid nitrogen or quartz [31]
N/A With dry ice [169]
High pressure homogeniser N/A N/A [202]
Microwaves 5 min 100 1C, 2450 MHz [31,106]
5 min 4050 1C, 2450 MHz [107]
Osmotic shock 48 h 10% NaCl [106]
60 min 2 N NaOH [83]
Ultra-sonication 6 min 10 W [84]
20 min 600 W [31]
5 min 10 kHz [106]
1560 min N/A [50]
N/A: not available.
C. Saet al. / Renewable and Sustainable Energy Reviews 35 (2014) 265278 273
Author's personal copy
food with natural composition instead of synthesised ingredients.
This has triggered intensive research into nding natural ingre-
dients that improve the quality of animal food products [119].
Thus, while stressing C. vulgaris, it accumulates important amount
of carotenoids and after feeding it to animals such as sh and
poultry it showed interesting pigmentation potential for sh esh
and egg yolk in poultry, together with enhancing health and
increasing life expectancy of animals [165,169174]. Moreover, C.
vulgaris showed a protective effect against heavy metals and other
harmful compounds (lead, cadmium, and naphtalene) by reducing
signicantly the oxidative stress induced by these harmful com-
pounds, and increasing the antioxidant activity in the organisms of
tested animals [175177].
7.4. Wastewater treatment
Many studies demonstrated the remarkable potential of C.
vulgaris in xating up to 74% carbon dioxide when grown in a
photobioreactor [178], and in absorbing 4597% nitrogen, 2896%
phosphorus and in reducing the chemical oxygen demand (COD)
by 6186% from different type of wastewater such as textile,
sewage, municipal, agricultural and recalcitrant [179185]. Micro-
algae provide a pathway for the removal of vital nutrients (nitro-
gen and phosphorus), carbon dioxide, heavy metals and pathogens
present in wastewaters and necessary for their growth. In addi-
tion, saving and requirements for chemical remediation and
possible minimisation of fresh water use for biomass production
are the main drivers for growing microalgae as part of a waste-
water treatment process [46]. Thus, a faster growth rate accom-
panied by an elimination of water-contamination level is a
promising and advantageous process. Furthermore, performance
of C. vulgaris in synthesised wastewater was improved when co-
immobilised in alginate beads with microalgae growth-promoting
bacteria, and removed 100% of ammonium (NH
4þ
) during four
consecutive cycles of 48 h, and 83% for phosphorus after one cycle
of 48 h [186]. Thus, C. vulgaris is considered as one of the best
microalga for bioremediation of wastewater with an impressive
potential to completely remove ammonium and sometimes mod-
est potential to eliminate phosphorus present in the medium
[187].
7.5. Agrochemical applications
Blue-green algal extract excretes a great number of substances
that inuence plant growth and development [188]. These micro-
organisms have been reported to benet plants by producing
growth promoting regulators, vitamins, amino acids, polypeptides,
antibacterial and antifungal substances that exert phytopathogen
biocontrol, and polymers such as exopolysaccharides that improve
plant growth and productivity [189].
The bio-fertilisation effect using algae extract are recom-
mended for increasing the growth parameters of many plants
[190,191]. This is due to the biochemical prole of algae extract
rich in nitrogenase, nitrate reductase, and minerals, which are
Fig. 4. Algo-renery concept from production to valorisation.
C. Saet al. / Renewable and Sustainable Energy Reviews 35 (2014) 265278274
Author's personal copy
essential nutrients for plant growth. The effect of the aqueous
extract of C. vulgaris as foliar feeding on nutrients status, growth,
and yield of wheat plant (Triticum aestivum L. var. Giz 69) has been
investigated [192]. Thus, this study found that a concentration of
50% (v/v) algae extract as one time foliar spray (25 days after
sowing) increased the growth yield and weight gain by 140% and
40%, respectively. Moreover, another study showed the bio-
fertilisation impact of C. vulgaris on growth parameters and
physiological responses of Lactuca sativa germination seeds in
culture medium containing microalga grown for 3, 6, 9, 12 and 15
days [193]. As a result, the addition of C. vulgaris to the culture
medium or soil signicantly increased fresh and dry weight of
seedlings as well as pigments content. The best treatments were
2 and 3 g dry alga kg
1
soil. All these studies were conducted on
the liquid extract of C. vulgaris as bio-fertilizer for plant growth.
Therefore, further studies should be carried out to estimate costs
on a large scale of the algae cell extract as foliar fertilizer,
compared to other commercial foliar fertilizers present in the
market.
8. Algo-renery concept
The concept of biorenery has been inspired from the petro-
leum renery concept. It reects a platform that integrates a
process to fractionate the components of a biomass [194,195] to
produce multiple products, and thus a biorenery takes advantage
of the various components in the biomass in order to improve the
value derived from each component and also generating its own
power, which maximises protability and preserve the environ-
ment. Hence, C. vulgaris with all its potential and richness in
proteins, carbohydrates, lipids, pigments, minerals and vitamins
described previously deserves to be completely rened (Fig. 4)
without forgetting that every operation unit should take into
account the next stage and preserve the integrity of all compo-
nents of interest in the downstream process.
9. Conclusion
This review reects a broader image about the potential
benets of C. vulgaris, and gives an insight about the technological
advancements already conducted. C. vulgaris can easily be cultured
with inexpensive nutrient regime and has faster growth rate as
compared to terrestrial energy crops and high biomass produc-
tivity. However, production-processing cost remains too high to
compete in the market. Indeed, this is the major problem facing
the microalgal industry nowadays, but it should be recognised that
much improvements have been achieved during the last decade
and expectations are estimating that the nearest future of micro-
algal industry will be strongly competitive on different levels in
the market. The remarkable values of C. vulgaris set the ground-
work to additional research for futuristic applications where it will
be represented as a strong candidate for tomorrows bio-industry.
Acknowledgements
TheauthorswouldliketothankAgenceNationaledelaRecherche
(ANR) for the nancial support, and Laboratoire de Chimie Agro-
industrielle (LCA) for providing all the necessary tools and require-
ments to dress this review.
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... C. vulgaris, 2 ila 10 µm boyutlarında ve yüksek bitkilere benzer bir yapıya sahip mikroskobik bir organizmadır. Basit bir şekilde bir hücre duvarı, çekirdek, mitokondri ve fotosentez gerçekleştirmek için kloroplasta sahiptir [6]. ...
... Hücre bölünmesi 6 adımdan oluşur; 1) hücre boyutunda artış, 2) yavru hücre için iç hücre duvarının oluşumu, 3) kloroplastın 2'ye bölünmesi, 4) kloroplastların ikinci bölünmesiyle 4, 5) yavru hücre duvarının oluşumu ve olgunlaşması. 4 yeni hücrenin bölünmesi ve 6) 4 yavru hücrenin serbest bırakılması için eski hücre duvarının yırtılması [6]. Bu işlemin, hücre uygun büyüme koşullarında olduğu sürece gerçekleştirildiğini belirtmek önemlidir, çünkü algler strese maruz kalırsa daha uzun bir büyüme süresine sahip olabilmektedir. ...
Conference Paper
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Bu çalışmada Chlorella vulgaris mikroalgi kullanılarak belirli çevrim hızlarına sahip dinamik sistem ile statik sistem arasındaki popülasyon, konsantrasyon ve pH değişimleri incelenmiştir. Statik ve dinamik sistemlerde 14 günlük kültür süresince mikroalgin üreme süreci gözlemlenmiştir. Dinamik sistemde 1,5/3/6 mL/dk olmak üzere üç farklı çevrim hızı ile çalışılmıştır. 3000 lux ışık şiddeti ve 25 ℃ sabit ortam sıcaklığında, 12 saatlik periyotlarla ölçülen absorbanslar ile konsantrasyon miktarları (g/L) hesaplanmıştır. Sedgwick-Rafter yöntemiyle yapılan hücre sayımları ile her ölçüm saatindeki canlı hücre popülasyonları hesaplanmıştır. Konsantrasyon ve popülasyon ile birlikte sistemlerin pH değişimleri de gözlemlenmiştir. Hücre konsantrasyonu arttıkça pH’nın da arttığı gözlemlenmiştir. Elde edilen verilere göre dinamik sistemde statik sisteme oranla daha yüksek konsantrasyon, hücre popülasyonu ve popülasyon artış hızı elde edilmiştir. Sisteme kazandırılan dolaşımla birlikte statik sisteme kıyasla biyokütle birikmesinde bir azalma görülmüştür. 1.5 mL/dk çevrim hızında statik sisteme oranla aynı kültür süresi içerisinde, daha yüksek popülasyon ve konsantrasyon elde edilmiştir. 3 mL/dk çevrim hızı ile yapılan kültür süresince statik sisteme ve 1.5 mL/dk çevrim hızına sahip sisteme göre daha da yüksek popülasyon ve konsantrasyon elde edilmiştir. En yüksek konsantrasyon (0,39 g/L), pH artış hızı ve canlı hücre popülasyonu (1,76 × 106) 6 mL/dk çevrim hızına sahip dinamik sistemde elde edilmiştir. Böylelikle C. vulgaris mikroalginin dinamik bir sistemde kültür edilmesiyle daha hızlı bir konsantrasyon artışı ve daha yüksek bir popülasyon sağlanmaktadır.
... Therefore, considering the potential for dazomet to reach the aquatic systems, the main objective of this study was to evaluate the influence of soil pH in the toxicity of eluates obtained from soils contaminated by Basamid® on two freshwater producers species that might be directly exposed to leachates of this agrochemical: the microalgae Raphidocelis subcapitata and the duckweed Lemna minor (EFSA, 2010;Vryzas, 2018). Both species are well-known and characterized by rapid growth rates, low maintenance in the lab and can provide multiple endpoints as chlorophyll pigments, cell damage, growth rates, and biomass content to assess adverse effects (Safi et al., 2014). ...
... Also, L. minor has a great tendency to bioaccumulate compounds such as metals and pesticides, which could be an additional explanation for its great sensitivity in comparison to the algae (Mkandawire et al., 2014). Additionally, the cell wall of R. subcapitata can be the main mechanism of defence of algae from the surround medium as it is observed for C. vulgaris (Safi et al., 2014) while Lemna sp. present a potential higher exposure directly through the leaves and systemically throughout the exposed roots (Mkandawire et al., 2014). ...
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Intensive agriculture along with the use of agrochemicals has been associated with soil infertility, erosion, and soil acidity. Management of soil pH through liming is a common practice in agriculture to increase soil fertility and nutrient availability. When altering soil pH, different chemical reactions occur depending on soil composition and agrochemicals presence. Basamid® is a fumigant used worldwide targeting soil nematodes, fungi, and weeds in diverse crops, that can reach freshwater ecosystems by leaching through the soil layers. The major goal of this work was to assess the influence of soil pH in the toxicity of Basamid® eluates to the microalgae Raphidocelis subcapitata and the duckweed Lemna minor. For this, eluates were prepared from soils with different pH (5.5, 6.5 and 7.5), contaminated with the recommended dose of Basamid® corresponding to 145.7 mg of dazomet/Kg soil. Soil was amended with calcium carbonate (CaCO3). Raphidocelis subcapitata and L. minor were exposed to the eluates during 72 h and 7 days respectively, and multiple endpoints were assessed: growth rate, biomass, pigment as chlorophyl content and cell damage. Results showed that soil pH can influence the performance of the tested species and also be a major factor in influencing Basamid®'s toxicity. However, a clear pattern of the influence of soil pH on Basamid®'s toxicity was not observed and was species dependent. For R. subcapitata lower soil pHs induced higher toxicity of Basamid®'s to the algae [ED50 for growth rate: 30 % (confidence limits-CL: 22.8–37.2) for soil pH 5.5; >100 % for soil pH 6.5 and pH 7.5], while for L. minor the opposite was observed [ED50 for number of fronds: 27.2 % (CL: 22.8–31.6) for pH 5.5; 20.3 % (CL: 10.0–30.6) for pH 6.5 and 10.7 % (CL: 6.3–15.1)]. Overall, these results showed that leachates of Basamid® through soils, at recommended doses, can have a severe impact on aquatic systems, with or without the influence of abiotic factors.
... Algal biomass either in the form of dry biomass or living cells can be used as an anodic substrate Xiao & He, 2014). Chlorella vulgaris biomass has high nutritional value comprising of carbohydrates (12-55%) and proteins (42-55%) that can be mineralized by electrogenic bacteria to generate electricity Safi et al., 2014). In a previous study, Chlorella vulgaris and Ulva lactuca feedstocks have tested in dry powder as a substrate for MFC operation, the power density produced was 0.98 Wm-2 from the Chlorella vulgaris, and 760 mWm −2 for Ulva lactuca-based MFC (Velasquez-Orta et al., 2009). ...
... In this study we used Chlorella vulgaris, a freshwater green microalga, which is broadly used in the food, pharma, and water industry (21). The main purpose was to investigate the effect of different nitrate and phosphate concentrations and two different environmental conditions on the production of MAAs, as well as to determine the optimum conditions, using an experimental design method. ...
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