Nitrate and Nitrite Removal from Wastewater using Algae

Abstract

Background: Both land-based agriculture and aquatic algae culturing systems require a steady supply of macronutrients, primarily nitrogen (N) and phosphorus (P), in addition to a variety of micronutrients for biomass production. The use of commercial fertilizer for large-scale algae production significantly increases the cost of algae production. Microalgae have a high capability to remove combined nitrogen compounds, ammonia, nitrate and nitrite, from wastewaters. Methods: The algae assimilates inorganic nitrogen and converts nitrogen into biomass, thus providing an opportunity for efficient recycling of nutrients in wastewater. Furthermore, the microalgae can be a feedstock for biodiesel and other valuable by-products including pigments, proteins and lipids. Combined nitrogen is assimilated in different forms and at different rates that vary among the phylogenetically diverse strains of microalgae. Results: In this review, we summarize nitrate removal rates and biomass production of different microalgae species reported in the literature. Conclusion: A comparison of the literature suggests that Chlorella vulgaris, Neochloris oleoabundans and Dunaliella tertiolecta are able to remove nitrate more effectively than other strains studied. Moreover, important parameters influencing nitrate removal, including initial nitrate concentration, light intensity, pH and temperature, are discussed. Alternative culture methods, immobilization and biofilm formation for nitrate remediation, are introduced which are able to lower costs of the harvesting process.

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Nitrate and Nitrite Removal from Wastewater Using Algae
Mahboobeh Taziki1, Hossein Ahmadzadeh*,1, Marcia A. Murry2 and Stephen R. Lyon3
1Department of Chemistry, Ferdowsi University of Mashhad, Mashhad 1436-91779, Iran
2Department of Biological Sciences, California State Polytechnic University, Pomona, California 91768, USA
3Alga Xperts, LLC, Milwaukee, Wisconsin, USA
Abstract: Background: Both land-based agriculture and aquatic algae culturing systems require a
steady supply of macronutrients, primarily nitrogen (N) and phosphorus (P), in addition to a variety of
micronutrients for biomass production. The use of commercial fertilizer for large-scale algae
production significantly increases the cost of algae production. Microalgae have a high capability to
remove combined nitrogen compounds, ammonia, nitrate and nitrite, from wastewaters.
Methods: The algae assimilates inorganic nitrogen and converts nitrogen into biomass, thus providing
an opportunity for efficient recycling of nutrients in wastewater. Furthermore, the microalgae can be a
feedstock for biodiesel and other valuable by-products including pigments, proteins and lipids.
Combined nitrogen is assimilated in different forms and at different rates that vary among the
phylogenetically diverse strains of microalgae.
Results: In this review, we summarize nitrate removal rates and biomass production of different microalgae species
reported in the literature.
Conclusion: A comparison of the literature suggests that Chlorella vulgaris, Neochloris oleoabundans and Dunaliella
tertiolecta are able to remove nitrate more effectively than other strains studied. Moreover, important parameters
influencing nitrate removal, including initial nitrate concentration, light intensity, pH and temperature, are discussed.
Alternative culture methods, immobilization and biofilm formation for nitrate remediation, are introduced which are able
to lower costs of the harvesting process.
Keywords: Biomass concentration, microalgae, immobilization, nitrate removal rate.
1. INTRODUCTION
Population growth, industrialization and rapid
urbanization have led to excessive nitrogen (N) pollution,
often in the form of nitrate, presenting a water-quality
problem of growing concern [1]. Excessive fertilizer use in
urban and agricultural regions has caused serious problems
of nitrate and phosphate (P) pollution in surface waters,
groundwaters and the marine environments. Nitrate fertilizer,
not taken up by plants, is leached from soils and can
percolate into ground waters and/or be washed into
freshwater reservoirs and the ocean through urban storm
water systems. Municipal wastewater discharge [2], sewage
waste and septic tanks [3], livestock farms, processed food
plants, dairy and meat processing facilities and decomposit-
ion of decaying organic matter also release significant
amounts of N into aquatic environments [2]. While the
question of whether N or P input is the major factor in
eutrophication is questioned [4], and generally biologists
favor nitrogen as the limiting nutrient while geochemists
favor phosphate limitation [5], there is little debate that
increased N input into our waterways presents a major
*Address correspondence to this author at the Department of Chemistry,
Ferdowsi University of Mashhad, Mashhad 1436-91779, Iran.
Tel.: +985138797022ext375; fax: +9851387956416.
E-mail: H.Ahmadzadeh@um.ac.ir
perturbation to aquatic ecosystems [6]. Eutrophication of
surface waters including lakes, streams and drinking water
reservoirs has resulted in algae blooms. What were once
occasional algae blooms occurring as a regional phenomenon
are now appearing on a global basis with greater frequency.
The immediate consequences of these blooms include the
degradation of recreational lakes and total oxygen
consumption that result in major fish kills. Certain
cyanobacteria such as Microcycstis produce neurotoxins
(cyanotoxins) that can persist in the water column long after
the algae bloom has faded [7].
Nitrogen goes through a biogeological cycle producing
compounds with different oxidation states that are available
to plants, algae and microbes: Nitrate, nitrite, ammonium,
organic nitrogen including amino acids, urea and proteins.
While ammonium is energetically more favorable and is the
preferred nitrogen source when it is available [8, 9], in many
waterways nitrate concentrations are generally much higher
than ammonium concentrations. For example, many
industrial wastewaters often contain more than 200 mg NO3-
N while effluents from industries producing explosives,
fertilizers, pectin, cellophane, and metal finishing, contain
greater than 1000 mg NO3- N. The nuclear industry also
produces nitrate loaded wastes in extremely high
concentrations at many points during the nuclear fuel cycle
(up to 50,000 mg NO3- N/L). Therefore, in this review
Hossein Ahmadzadeh

2 Current Biotechnology, 2015, Volume 4, No. 3 Taziki et al.
article, we focus mainly on NO3- and NO2- removal. While
almost all algae can grow on low to high concentrations of
nitrate, many strains have acute sensitivity to high
concentrations of ammonium/ammonia. The review article
by Collos and Harrison [10] showed a ranking of sensitivity
to high levels of ammonium/ammonia (39-1.2 mM) where
the order of tolerance was: Chlorophyceae > Cyanophyceae,
Dinophyceae, Diatomophyceae, and Raphidophyceae.
Most wastewater treatment systems have two levels of
treatment, primary (physical settling of solids), secondary
(various forms of oxidation e.g. activated sludge or trickling
filters). Where regional or government regulations mandate
higher effluent quality, tertiary treatment is used for nutrient
removal and disinfection. In primary wastewater treatment
the major forms of nitrogen are organic-N and ammonium.
During secondary treatment the two major forms of nitrogen
are rapidly converted to nitrate by nitrifying bacteria, such as
Nitrosomonas, Nitrosococcus, Nitrobacter, and Nitrococcus
[11]. Nitrite is the most ephemeral form of nitrogen in the
environment. In both wastewater treatment systems and in
surface waters, it occurs as the least prevalent form of
inorganic nitrogen.
Although nitrate is one small component of the nitrogen
cycle, the focus of this review is nitrate assimilation by
algae. There are many biogeochemical and physical
processes within the nitrogen cycle. The daily shifts in pH in
surface waters due to algal photosynthesis facilitate the
conversion of ammonium ions to ammonia gas and its
subsequent volatilization into the atmosphere [12]. A
microscopic examination of almost any alga taken from an
oxidation pond will show hundreds of bacteria attached to
the outer surface of the alga. The bacterial-algal interactions
play a key role in nutrient processing. The main aspects of
this synergistic relationship include the photosynthetically
generated oxygen, which fuels bacterial mineralization of
organic material producing inorganic nutrients for algal
growth [13]. Table 1 below summarizes the forms of
nitrogen in surface waters and their impact on water quality
[14].
The removal of nitrate by bacterial dissimulatory nitrate
reduction plays a major role in the conversion of nitrate to
nitrogen gas in anaerobic sediments in lakes, ponds and
wetlands. Incomplete denitrification results in the release of
nitrous oxide, which has been shown to be 300 times more
potent a greenhouse gas than carbon dioxide [15] and is
considered the most active compounds in ozone depletion in
the 21st Century [16]. Nitrous oxide emissions into the
atmosphere are in part due to human activities including
agricultural fertilization and livestock feedlots [17]. The
contributions of nitrous oxide to the atmosphere by natural
and anthropogenic sources are a topic of the on-going
debates [18]. Recent studies by Guieysse et al. and Alcántara
et al. have shown that algae in high-rate production ponds
can also contribute to the production of nitrous oxide [19].
Many groundwater basins have been historically
underused for human consumption due to high nitrate
concentrations that leads to health consequences. Thus,
direct use of groundwater resources for human consumption
has been prohibited in many parts of the world. Nitrate may
be reduced to nitrosamines in the stomach which, as known
carcinogens, may be a factor causing gastric cancer [20, 21].
Nitrate reacts with hemoglobin in the blood to form
methemoglobin, leading to an overall reduced ability of the
red blood cells to release oxygen to the tissues. This lack of
oxygen results in methemoglobinemia (blue-baby syndrome)
[22].
Groundwaters contaminated with nitrate above the
United States Environmental Protection Agency (EPA) and
World Health Organization (WHO) maximum level (10 mg
L-1 NO3- N), must be treated before use as drinking water.
Europe also set a maximum of 12 mg L-1 NO3- N in drinking
water for the same concern [23]. Methods for nitrate and
nitrite removal in water resources are a controversial issue
that has attracted a good deal of attention.
Generally, there are two basic types of treatments for
removing nitrate from water or wastewater: physicochemical
and biological methods. Physicochemical methods include
reverse osmosis (RO) [24], ion exchange (IE) [25],
electrodialysis (ED) [26] and activated carbon adsorption in
conjunction with pH adjustment [27]. While IE and RO are
well developed, both are energy intensive processes and are
not highly efficient, producing brine waters that are
frequently discharged into adjacent waterways [28].
Recently, researchers have developed new methods for
nitrate removal, including metallic iron-aided abiotic nitrate
reduction (also known as zero-valent iron or ZVI) [29, 30].
Many have sought biological solutions to cost effective and
sustainable treatment processes that can be as effective as the
conventional physicochemical processes [20].
A variety of biological methods are available for the
denitrification of surface and ground waters based on plant
and microbial metabolic processes. The best described
mechanisms are assimilation of nitrate by plants, algae and
microbes and microbial respiratory denitrification where
nitrate and its reduction products serve as alternate electron
acceptors under anaerobic conditions resulting in the
conversion of nitrate to N2 gas (dissimilatory nitrate
reduction, DNR). Nitrate and ammonia assimilation, in
contrast to the respiratory nitrate reduction, results in N
being converted to biomass rather than being released to the
atmosphere as the relatively inert N2 gas. Other less known
microbial processes include a dissimilatory nitrate reduction
to ammonium [31], and anaerobic ammonium oxidation
(anammox) [32, 33]. In addition, denitrification can be
coupled to sulfide or iron oxidation [34-37].
Cyanobacteria and microalgae have been reported to be
more efficient for N bioremediation [38] than higher plants,
due in part to higher rates of biomass production but also
because algae lack the large stores of structural carbon (ie.
cellulose) characteristic of land plants. Thus, the C/N ratio of
higher plants ranges from 18-120 (by atoms) while
microalgae range from 5 to 20 [39] indicating that water
reclamation and nutrient recovery can be accomplished more
rapidly, and in a smaller area, using algae rather than
terrestrial plants. Mass-culture of algae on manure N and P is
an alternative to land spreading of manure effluents,
particularly in the case of confined animal feed operations
(CAFOs). Groundwater contamination is problematic in
these operations and many CAFOs do not have affordable
access to large tracts of land for manure application to soils.
A highly productive crop is needed to remove manure N and
P in smaller land areas than are required by crops such as

Nitrate and Nitrite Removal from Wastewater Using Algae Current Biotechnology, 2015, Volume 4, No. 3 3
corn. At least 70 percent of the cost of municipal wastewater
treatment can be attributed to secondary and tertiary
treatment. Much of this is due to the energy costs of oxygen
transfer in biological secondary treatment and to the
chemical requirements in tertiary treatment [40]. Microalgae
have been used for over 50 years in municipal wastewater
treatment where photosynthetically generated O2 is
consumed by bacterial populations that decompose organic
wastes to simple inorganic nutrients and in tertiary treatment
to remove inorganic nutrients before discharge to receiving
waters [41].
While there are approximately 4,000 known species of
microalgae and cyanobacteria [42], for this review, the
efficiency of N uptake and biomass production by selected
algal strains common to eutrophic waters were compared
(Section 3). Furthermore, experimental factors including
initial nitrate concentration and the ratio of ammonia to
nitrate (Section 4.1), light/dark cycle and light intensity
(Section 4.2), pH (Section 4.3), and temperature (Section
4.4), are also discussed. Finally, we provide a survey of
alternative culturing technologies, including immobilization
(Section 5.1) and biofilm formation (Section 5.2) aimed at
harvesting the biomass at a low cost.
2. PHOTOTROPHIC NITRATE ASSIMILATION
Phytoplankton are responsible for ~ 70% of global
nitrogen assimilation on earth with about 65% consumed as
reduced nitrogen (ammonia and organic nitrogen),
approximately 10% via nitrogen fixation and the balance as
nitrate [43]. Because N and P tend to be limiting nutrients for
algal growth and because phytoplankton grow in very dilute
nutrient solutions in natural waters, algae have developed
extremely efficient mechanisms for nutrient uptake. In many
waterways, especially estuarine and marine systems, nitrate
concentrations are generally much higher than ammonium
concentrations. Nitrate is tolerated at rather high levels by
both plants and algae while there are toxicity issues
associated with ammonium. Many algal strains show a high
tolerance for ammonium but others have a distinct sensitivity
to even low concentrations of ammonium [44]. This
Table 1. Overview of the primary forms of N found in surface waters and associated concerns.*
Nitrogen
Parameter
General Description
When Found
Sources to Surface Waters
Health and Environmental
Concerns
Nitrate-N (NO3-)
Main form of N in
groundwater and high-N
surface waters. Dissolved
in water and moves
readily through soil.
Present as a common form of
nitrogen, since most other N
forms can transform into
nitrate in N cycle.
Transformed
into nitrate from
other N forms found in
fertilizer, soil N, atmosphere
and human and animal waste.
Methemoglobinemia in infants
and susceptible adults. Toxic to
aquatic life, especially
freshwaters Eutrophication and
low oxygen (hypoxia), especially
in coastal waters.
Nitrite-N
(NO2-)
Low levels in waters –
typically measured in the
lab together with nitrate.
Less stable intermediary form
of N found during N
transforming processes.
Same as nitrate.
Methemoglobinemia
in infants and susceptible adults.
Toxic to aquatic life.
Ammonia-N
(NH3)
Unionized Ammonia –
low levels in most
waters.
Most of NH3 NH4+ is in
the NH4+ form. But NH3
increases with higher temps
and pHs (potential of
Hydrogen).
Human and animal waste
discharges.
Toxic to aquatic life.
Ammonium-N
(NH4+)
Measured in the lab
together with ammonia –
usually higher than
ammonia but less toxic
Usually found at low
levels compared to nitrate and
organic N. Found near waste
sources.
Human and
animal waste discharges.
Can convert to more
highly toxic ammonia in high pH
and temperature waters.
Organic-N
The main form of N in
low-N surface waters
(where nitrate is low).
Living and dead
organisms/algae. Found
naturally in water and is
supplemented by human
impacts.
Algae; soil; organisms; human
and animal waste.
Can convert to ammonium and
ultimately nitrate under certain
conditions.
Inorganic N
The sum of Nitrite,
Nitrate, ammonia,
and ammonium.
See separate
parameters above.
Total
Kjeldahl N (TKN)
Lab measurement which
includes organic-N,
ammonia and
ammonium.
Useful to determine
organic-N when ammonia
ammonium is also determined
separately and subtracted from
TKN.
See separate
parameters above.
Total N
Sum of TKN,
nitrite and
nitrate.
See separate
parameters above.
*Adapted from [14].

4 Current Biotechnology, 2015, Volume 4, No. 3 Taziki et al.
sensitivity/toxicity to ammonia is in part due to the pH shift
that occurs when carbon dioxide becomes limited and algae
begin to take up bicarbonate ions. The ion is broken down to
carbon dioxide and hydroxyl ions. The carbon dioxide is
used for photosynthesis and the hydroxyl ions are excreted
back into the water. This causes a rise in pH resulting in
ammonium ions (NH4
+) being converted to ammonia (NH3)
[45]. In a study of six classes of microalgae, Cyanophycea
(blue-green algae) had the highest tolerance to ammonium
while the Dinophyceae (dinoflagellates) had the least [46].
Ammonia is more toxic than nitrate to both plants and
animals because it dissipates transmembrane proton
gradients needed for both respiratory and photosynthetic
electron transport mechanisms. Thus, upon uptake, or
conversion of nitrate to ammonia, ammonia is incorporated
rapidly into amino acids. However, in polluted waters
including dairy [47-49], swine [50] and municipal
wastewaters [51, 52], ammonia and organic N are the
predominant forms of combined N, while nitrate and nitrite
are generally found at trace levels. Oxidation-reduction
potential can predict the oxidation state of N compounds and
their interconversion by nitrification and denitrification
reactions [53].
Nitrate reduction to ammonium takes place through
sequential reactions involving 2-electron and 6-electron
reductions catalyzed by nitrate reductase and nitrite
reductase. The ammonium produced is incorporated into
amino acids via glutamate dehydrogenase (at high
concentrations of ammonium) or the glutamine
synthetase/glutamate synthase cycle (at low levels). Plants
and algae assimilate nitrate and immediately reduce nitrate to
nitrite via the enzyme nitrate reductase (NR) using either
nicotinamide adenine dinucleotide (NADH) or nicotinamide
adenine dinucleotide phosphate (NADPH) as an electron
donor. Because nitrite is highly reactive and more toxic than
nitrate, in higher plants it is immediately transported from
the cytoplasm into the chloroplasts of the leaves or plastids
of root tissues. In higher plants, two different forms of nitrite
reductase both containing an iron-sulfur cluster and a
specialized heme prosthetic group [54], are found in the
chloroplasts and mitochondria where nitrite reductase
reduces nitrite to ammonia without intermediate forms of
varying redox levels. In the chloroplasts, reduced ferredoxin
produced by photosynthetic electron transport is used as
reducing agent for nitrite reductase while plastids use
NADPH derived from the oxidative pentose pathway [55].
Because of the high energetic cost of the process and
because nitrate reduction competes for reducing equivalents
with photosynthetic carbon fixation, nitrate reduction is
highly regulated [55]. While it is generally thought that the
presence of ammonium inhibits nitrate uptake, there is
evidence that in phytoplankton the uptake and assimilation
mechanisms are not as simple or as tightly coupled as
previously thought. Under various environmental conditions,
especially light and temperature, and among different
microalgal groups and even species, there is more flexibility
in the mechanisms regulating N assimilation [56].
Although many aspects of nitrate assimilation in
microalgae are similar to those of higher plants, differences
are seen due in part to the evolutionary diversity of algae and
to structural differences between these major taxonomic
divisions. In the cyanobacteria and unicellular algae there are
no storage vacuoles or transport systems seen in higher
plants. Thus, in some green algae, including Chlorella [57],
Chlamydomonas [58] and Monoraphidium braunii [59],
nitrate uptake and reduction is tightly coupled and stimulated
by blue light. However, nitrate uptake in Hydrodictyon, a
large vacuolated coenocytic alga (Characeae) is not as
closely coupled and is regulated more directly by energy
supply [60]. In algae, the major reductant used is NADPH of
photosynthetic origin and there are differences in the
structure, reducing agent and location of nitrate and nitrite
reductases [61]. Nitrate uptake and reduction to nitrite and
ammonium are driven in cyanobacteria by photosynthetically
derived ATP and reduced ferredoxin [62]. Nitrate reductase
(NR) from Chlorella sp. can utilize both NADH and
NADPH for nitrate reduction [63] presumably due to
light:dark cycles and because there is intense competition for
energy and reductant between photosynthetic carbon fixation
and other energy intensive processes. While nitrate reductase
is argueably [64, 65] localized in the cytoplasm of higher
plants, immuno-specific electron microscopy of NR
localized the enzyme in the pyrenoids, structures associated
with the chloroplasts of eukaryotic algae of Monoraphidium
braunii [66], Chlamydomonas reinhardii, Chlorella fusca,
Dunaliella salina, and Scenedesmus obliquus [66]. Starch
grains associated with the pyrenoids and enzymes, including
phosphoribulokinase, phosphoriboisomerase [67], and
ribulose bisphosphate carboxylase-oxygenase [68] suggest a
functional role for this structure in photosynthetic carbon
metabolism.
3. NITRATE BIOREMEDIATION BY MICROALGAE
3.1. Comparison of Various Algal Strains Toward Nitrate
Removal and Biomass Production
Previous studies have shown that species of microalgae
have different capabilities in N uptake and assimilation and
biomass production. However, variations in experimental
procedures make the data difficult to interpret and compare.
These include the use of different algae species and strains,
media composition including both defined inorganic media
or wastewater effluents, the ratios of reduced vs oxidized
forms of N and BOD levels, CO2 enrichment, light intensity,
diurnal light regimes and temperature. For example,
Sacristán de Alva et al. (2013), cultivated Scenedesmus
accutus in municipal wastewater after settling (primary
treatment) and after undergoing activated sludge treatment
(secondary treatment). The primary effluent had
approximately twice the levels of COD, nitrates and reduced
N and supported twice the biomass than effluent from
secondary treatment. They obtained a low nitrate removal
rate of 0.59 mg L-1 d-1 [69]. However, Doria et al. (2012)
used an outdoor photobioreactor with high light intensity
(from 100 to 1500 µmol m-2 s-1) under natural light/dark
cycles and improved nitrate removal efficiency for S. accutus
over 10-fold (6.26 mg L-1 d-1). The ten-fold difference in
nitrate uptake rate observed using the same species could be
attributed to the composition of wastewater and culture
methods. In their work [70], secondary treated municipal
wastewater also led to lower biomass concentration (0.74 g
L-1) than primary treated wastewater (1.1 g L-1) (Table 2)
[69]. Compared with secondary wastewater, the primary

Nitrate and Nitrite Removal from Wastewater Using Algae Current Biotechnology, 2015, Volume 4, No. 3 5
wastewater used in the Sacristán de Alva experiment had
higher levels of inorganic phosphate, organic nitrogen and
ammonia that potentially supported the higher algal growth.
Nunez et al. (2001) used artificial wastewater
(essentially, tap water, nitrate, ammonia and phosphate at
11.8, 40 and 4.5 mg L-1, respectively, and supplemented with
vitamins and trace minerals) for the cultivation of
Scenedesmus obliquus with 50% and 70% daily dilution in
continuous culture [71]. Nitrate uptake rates were 6.5 and 8.3
mg L-1 d-1, respectively, higher than the rate obtained by
Doria et al. (2012), and comparable to rates observed by
Sacristán de Alva et al. (2013) (Table 2). S. obliquus was
grown in continuous culture, replacing a fraction of growth
medium with fresh medium, which could account for higher
rates of nitrate uptake. But the influence of high BOD and
reduced N levels used in Sacristán de Alva’s work,
compared to the low BOD and relatively high levels of
nitrate to ammonia ratio of the Nunez study is unclear, along
with the question of whether the specific algae strains used,
also played a role in nitrate uptake.
Su et al. (2012a) compared three common green algae
species (Chlorella vulgaris, Scenedesmus rubescens,
Chlamydomonas reinhardtii and the cyanobacterium
Phormidium sp.) using effluent from a secondary clarifier
with a COD of 30 mg L-1, and total Kjeldahl nitrogen of 95.5
mg L-1 comprised of ammonia (95.5%) and the balance by
nitrate and nitrite. Cells were cultured in photobioreactors
with a light/dark cycle of 12:12 h, with 7000 lux. As
depicted in Fig. (1) and tabulated in Table 2, C. reinhardtii
removed nitrate in 4 days as compared with S. rubescens and
Phormidium sp that removed nitrate in 6 and 7 days,
respectively. In terms of removal capacity, C. reinhardtii had
the highest nitrate removal rate (0.16 mg L-1 d-1) while S.
rubescens had the highest biomass productivity (6.56 g m-2
d-1) (Table 2) [72]. Sydney et al. (2011) compared nitrate
uptake by 20 different strains of microalgae cultured under
identical conditions. They showed that Botryococcus braunii
and Chlorella vulgaris had the maximum nitrate removal
efficiency with an uptake rate of 22.2 and 20.28 mg L-1 d-1,
respectively (Table 2) [73]. While Botryococcus brauniiis is
known for its unusually high lipid content, its potential as a
source for biofuel production is limited due to its slow
growth rate [74], yet in this study showed a very high rate of
nitrate uptake.
Complete nitrate removal from primary treated sewage
was observed with the growth of Haematococcus pluvialis
[75]. Initial nitrate concentration was 42.4 mg L-1 and an
uptake rate of 8.48 mg L-1 d-1 was observed (Table 2). The
significant improvement in nitrate removal rates observed for
the same strain (40 mg L-1 d-1) [76] may be due to the higher
light intensity utilized (100 µmol photon m2 s-1 compared to
the 50 µmol photon m2 s-1) in the earlier study. Therefore, as
Sacristán de Alva suggested [69], light intensity can be
considered an important factor for nitrate removal efficiency
in some species (see section 3.2).
Another key to nitrate uptake variation in the literature
may reflect the fact that nitrate uptake is influenced by the
presence of other nitrogen sources especially ammonium ion.
For example, Chlorella vulgaris removed 62.5% of NO3-
with a removal rate of 2.65 mg L-1 d-1 and nitrite uptake at a
rate of 0.01 mg L-1 d-1 (Table 2) from effluent without
ammonium ion [77]. However, in wastewater containing 205
mg L-1 NH4+ in combination with initial nitrate
concentrations ranging from 1.5 to 198.3 mg L-1 NO3-, lower
rates of nitrate removal by C. vulgaris were observed [78]. In
these experiments, ammonium ion was the preferred nitrogen
source and nitrate uptake did not begin until the ammonium
ion was consumed [78]. This has been attributed to the
observation that ammonium ion assimilation does not
involve a redox reaction and it requires less energy [79].
Corey et al. (2013) measured nitrate uptake using five
different ratios of nitrate and ammonium in cultures of
Palmaria palmate and Chondrus crispus. Total N
concentration was 300 µM (with NO3-/NH4+ ratios of 300:0,
270:30, 150:150, 30:270, 0:300). P. palmate showed the
highest NO3- uptake (4.39 µmol NO3- gDW-1 h-1) at 270:30
NO3-/NH4+. For C. crispus nitrate uptake was equivalent at
300:0 NO3-/NH4+, 270:30 NO3-/NH4+, and 150:150 NO3-
/NH4+ with a mean uptake rate of 6.57 µmol NO3- gDW-1 h-1
(Table 2) [80]. Therefore, determining strain-specific
optimal ratios of ammonium and nitrate in the medium can
result in more efficient nitrate removal.
While clearly environmental parameters can affect nitrate
removal efficiency, there is little in the literature that
assesses the importance of each parameter and compares
their relative effect on nitrate removal efficiency. However,
finding species with significantly higher rate of nitrate
uptake relative to other species in comparable conditions can
provide researchers with valuable information. For example,
Neochloris oleoabundans was able to completely remove
nitrate with an initial concentration of 452 mg L-1 with an
uptake rate of 150 mg L-1 d-1 [81]. Dunaliella tertiolecta and
Chlorella vulgaris also have high nitrate uptake rates of 155
and 103.3 mg L-1 d-1, respectively [82]. Therefore,
Neochloris oleoabundans, Dunaliella tertiolecta and
Chlorella vulgaris are excellent candidates for nitrate
bioremediation.
4. EXPERIMENTAL PARAMETERS AFFECTING
REMOVAL OF NITRATE
The principle parameters affecting nitrate and nitrite
removal include, but are not limited to, initial nitrate
concentration, light intensity, pH and temperature. Data for
nitrate uptake by algae related to these parameters is
summarized in Tables 2-4.
4.1. Initial Nitrate Concentration
Initial nitrate concentrations reported in the literature for
algae growth experiments range from 45 to 1914 mg L-1
(summarized in Table 3) producing contradictory results for
the effect of initial nitrate concentration on biomass
production and nitrate removal rates. For example, Wang
and Lan (2011) grew Neochloris oleoabundans in media
containing 45 to 218 mg L-1 of NO3-. Their data showed that
increasing initial nitrate concentration increased nitrate
uptake rates, reaching a maximum of 1.82 mg L-1 h-1 at 140
mg NO3-. However, further increase in nitrate concentration
to 218 mg L-1 resulted in reduced cell growth [83]. It has

6 Current Biotechnology, 2015, Volume 4, No. 3 Taziki et al.
been suggested that increasing nitrates in the medium
stimulates NR activity leading to NH4+ accumulation and
toxicity [84]. A significant increase in biomass concentration
(from 4.5 to 6.4 g L-1) was also observed for Chlorella
vulgaris, when initial nitrate concentration was increased
from 124 to 1798 mg L-1 and nitrate uptake rates also
increased (Table 3) [84]. Haematococcus pluvialis, a
freshwater Chlorophyta, grown with nitrate concentrations
ranging from 32 to 1600 mg L-1, showed growth inhibition at
nitrate concentrations higher than 80 mg L-1 [76]. Continuous
feeding of nitrate at 40 mg L-1 to high density cultures of H. pluvialis
alleviated growth inhibition. Observations that increasing nitrate
concentrations result in an increase in nitrate removal rates
suggests that nitrate stimulates cellular nitrate reductase
activity at moderate nitrate levels [84]. Nannochloropsis
gaditana, a heterokont in the family Eustigmataceae, uses
nitrate as the sole nitrogen source at low concentrations (54
mg L-1) [85]. Nitrate removal decreased by 60% when initial
nitrate concentration was increased to 1914 mg L-1 (Table 3).
Nitrogen limitation is a key factor that initiates lipid
accumulation in many groups of algae. The final lipid
content in Nannochloropsis gaditana decreased 15% when
nitrate levels were significantly increased [85]. Ogbonna et
al. (2000) showed no significant effects on growth or nitrate
uptake by a photosynthetic bacterium Rhodobacter sphaeroides,
a green algae Chlorella sorokiniana and Spirulina platensis,
a cyanobacteria, with nitrate concentrations of 700 mg L-1
[86]. Thus, variation in initial nitrate concentrations can have
different effects on nitrate removal efficiency, assimilation
and growth in a taxa specific fashion.
Fig. (1). Nitrate removal of four different unicellular microalgae. (Chlamydomonas reinhardtii, Chlorella vulgaris, Scenedesmus rubescence
and phormidium sp). (Reproduced from [72] with permission).

Nitrate and Nitrite Removal from Wastewater Using Algae Current Biotechnology, 2015, Volume 4, No. 3 7
4.2. Light/Dark Cycle and Light Intensity
Light intensity and diurnal light cycles are important
factors affecting nitrate uptake [79]. In the natural
environment, these are not controllable, however, under
laboratory conditions, continuous light increases the rate of
nitrate uptake in some algae species. Data on nitrate removal
under light/dark cycles and light intensities is summarized in
Table 4. Comparisons of Chlorella kessleri grown under 12
h (L/D) lighting and continuous illumination [87],
demonstrated that continuous illumination lead to a higher
nitrate uptake rate (10.5 mg L-1 d-1) than 12 h (L/D) light
cycle (4.6 mg L-1 d-1) (Table 4). In contrast, no significant
difference in the nitrate and nitrite uptake was reported
between alternating (12:12 light/dark) and continuous
illumination (24 h light) by a mixed algae culture
(Chlamydomonas reinhardtii, Scenedesmus rubescens and
Chlorella. vulgaris), although a higher biomass production
capability was achieved for the continuous illumination
(Table 4) [88]. Biomass generation rates with 0, 12 and 24 h
illumination per day were 0.93, 7.51 and 9.38 g m-2 d-1,
respectively (Table 4). Lower productivity under a 12:12
hour light regime could be attributed to the loss of biomass
through respiration in the dark [88].
Light intensity also affects microalgae growth and nitrate
removal efficiency. Increasing light intensity is usually
accompanied by an increase in nitrate removal rates in
microalgal systems. Increasing light intensity from 400 to 1000
µmol photons m-2 s-1 led to an increase in nitrate uptake rate
from 2.2 to 6.3 mg L-1 d-1 by Chlamydomonas reinhardtii (Table
4) [89]. Increasing light intensity to the point of light saturation,
the point which photosynthetic activity reaches its maximum,
increases microalgae growth rates. However, at light intensities
above the saturation point, photoinhibition occurs, the
photosynthetic capacity decreases and growth is inhibited [90].
For example, in light intensities ranging from 5 to 50 µmol
photons m-2 s-1, maximum removal of nitrate was found at 10
µmol photons m-2 s-1 for Trentepohlia aurea. Nitrate removal
rates measured in this system were 0.94, 1.10, 1.02, and 0.80
mg L-1 d-1 when grown with 5, 10, 20, and 50 µmol photons m-2
s-1, respectively (Table 4) [91]. The data showed that nitrate
Table 2. Nitrate removal rate by microalgae species extracted from original references.
Algal Species
Days for
Assimilation
PH
T(˚C)
Light
Intensity
(µMOL M-2S-1)
Removal
Ratio
(%)
Uptake Rate
(MG L-1D-1)
Biomass
Concentration
(GDWL-1)
Reference
NO3
-
NO2
-
NO3
-
NO2
-
Chlorella. Sp
4
-
25
200
62
82
2.65
-
-
[77]
Scenedesmus accutus pvuw12
3
7.4
25
50
100
-
6.26
-
0.74
[70]
Phormidium sp
7
-
-
112
100
100
0.1
0.02
2.71G/M2/D
[72]
Chlamydomonas reinhardtii
4
-
-
112
92.7
22.2
0.16
0.005
6.06G/M2/D
[72]
Chlorella vulgaris
6
-
-
112
100
100
0.13
0.008
6.28G/M2/D
[72]
Scenedesmus rubescence
6
-
-
112
97.5
-
0.13
-
6.56G/M2/D
[72]
Rhodobacter sphaeroides
3
7
30
100
0
-
0
-
-
[86]
Chlorella sorokiniana
3
6
30
100
45.4
-
3.3
-
-
[86]
Spirulina platensis
3
9.2
30
100
100
-
7.2
-
-
[86]
Palmaria palmate
1
-
10
125
26
-
4.96
-
-
[80]
Chondrus cripus
1
-
10
125
56.35
-
11.21
-
-
[80]
Haematococcus pluvialis
5
7.5
23
50
100
-
8.48
-
-
[75]
Scenedesmus obliquus
50%DILLUTION 1
9.71
23
92
-
6.56
6.56
-
0.003
[71]
70%DILLUTION 1
9.35
23
88.9
-
8.3
8.3
-
0.17
[71]
Botryococcus braunii
14
7.2
25
56
79.74
-
22.21
-
-
[73]
Chlorella vulgaris
14
7.2
25
56
73.7
-
20.28
-
-
[73]
Haemotoccus pluvialis
2
7.5
23
100
-
-
40
-
0.6
[76]
Neochloris oleobundans
28
8
21.5
147
-
-
150
-
0.68
[81]
Chlorella vulgais
6
-
26
350
100
-
103.3
-
4
[82]
Dunaliella tertiolecta
4
-
26
350
100
-
155
-
3.3
[82]
Neochloris oleobundans
3
6.8
-
1280 (LUMEN)
100
-
43.7
-
3.15
[83]
Scenedesmus accutus
16
8.3
27
592
71
-
0.59
-
1.1
[69]

8 Current Biotechnology, 2015, Volume 4, No. 3 Taziki et al.
removal decreased at 20 and 50 µmol photons m-2 s-1, light
levels, which may exceed the saturation point for this green alga
adapted to growth on the trunks and branches of Monterey
cypress.
4.3. pH
The pH in microalgal cultivations media is altered by
uptake of inorganic carbon from the medium, nitrification in
ammonium treatment processes, and microalgal uptake of
nitrogen compounds [82]. Eustance et al. (2013), showed pH
increases in unbuffered growth media with nitrate as the
nitrogen source and air as carbon source during growth of
two Chlorophyte strains. Enriched CO2 concentrations can
help to prevent pH increases. For example, Scenedesmus sp.
131 and Monoraphidium sp. 92 exhibited no significant
fluctuation in pH when they were grown under 5% CO2
while growth under ambient air increased pH from 8.5 to 11.
Without CO2 enrichment, nitrate removal rates were 26.3 mg
L-1 d-1 and 17 mg L-1 d-1 for these strains, respectively, while
enrichment with 5% CO2 resulted in higher nitrate uptake
rates which reached 33 and 49.5 mg L-1 d-1 for the
Scenedesmus strain and Monoraphidium, respectively (Table
5) [92].
pH fluctuations can also be minimized by the high
buffering capacity of saline media or by use of organic
buffers. Buffers, including HEPES (pKa 7.4), CHES (pKa
9.3), and CAPS (pKa 10.4) have been used to stabilize pH in
culture media. Gardner et al. (2011) compared several
buffers on nitrate removal rates by Scenedesmus sp. and
Coelastrella sp. The unbuffered system had the highest
nitrate removal rate with 22.5 mg L-1 d-1 for Scenedesmus sp.
while Coelastrella sp. in media buffered by CHES had a
maximum nitrate uptake rate with 8.75 mg L-1 d-1 (Table 5)
[93]. Buffering solutions are not favored for large-scale algal
production because of the high cost of commercial organic
buffers and are thus confined to laboratory scale
experiments.
4.4. Temperature
The growth of microalgae is influenced by temperature
via effects on enzyme kinetics, changes in catalytic rate and
also unfolding/inactivation of enzymes [94, 95].
Additionally, temperature influences metabolite degradation
and biosynthesis and changes in conformation of vital
structures such as cell membranes [96]. As the temperature
drops, kinetic movement of phospholipids in the membrane
decelerate making the membranes more rigid; but as the
temperature increases, movements accelerate and
membranes become more fluid [94]. While most microalgae
can adapt to short-term as well as long-term changes in
temperature, each strain has a characteristic optimum
temperature [97, 98]. For example, the optimum growth
temperature for polar microalgae is usually below 10°C [99],
for temperate algae is around 10-25°C [100], for tropical
strains is around 25°C [97] and for desert algae is between
Table 3. Effect of initial nitrate concentration on nitrate removal efficiency extracted from original references.
Algal Species
Initial NO-
3
(mg L-1)
Biomass
Concentration
(g L-1)
Residue
NO-
3
(mg L-1)
Removal
Time
(d)
NO-
3
Removal
Rate
(mg L-1 d1)
Removal
Ratio
(%)
pH
T(˚C)
Light
Intensity
(µmol
m-2s-1 )
Reference
Neochloris
oleabundans
45
1.85
0
2
22.6
100
6.8
-
1280 lumen
[83]
70
2.37
0
2
34.8
100
6.8
-
1280
[83]
144
3.15
0
3
43.7
100
6.8
-
1280
[83]
218
2.91
1.4
5
42.5
99.3
6.8
-
1280
[83]
Nannochloropsis
gaditana
54
0.72
0
7
7.71
100
7
24
220
[85]
674.6
-
434
9
26.73
45
7
24
220
[85]
1294.68
-
806
9
54.22
44
7
24
220
[85]
1914.68
-
1240
9
74.88
40
7
24
220
[85]
Chlorella
vulgaris
124
4.5
94
17
0.45
24.19
-
25
600
[84]
248
6.1
186
17
0.57
25
-
25
600
[84]
372
11.5
298
17
0.60
19.89
-
25
600
[84]
744
10.5
653
17
0.85
12.23
-
25
600
[84]
1798
6.4
1667
17
0.95
7.28
-
25
600
[84]
Neochloris
oleabundans
135.67
1.85
0
1
135.67
100
-
30
360
[81]
226.11
2.37
0
2
113
100
-
30
360
[81]
452.23
3.15
0
3
150.74
100
-
30
360
[81]
678.35
2.91
4.52
6
112.30
99.3
-
30
360
[81]
904.47
2.70
224.76
6
113.28
75.15
-
30
360
[81]

Nitrate and Nitrite Removal from Wastewater Using Algae Current Biotechnology, 2015, Volume 4, No. 3 9
25 and 35°C [101]. Fluctuations in optimal temperatures can
affect microalgae growth. For example, the growth rate of C.
vulgaris decreased at 35°C by 17% when compared with
growth at 30°C. Further increase in temperature (38°C)
resulted in a cell death [102]. In another study, 20°C was
reported as optimal for growth of Nannochloropsis oculata.
At temperatures below 20°C, the growth rate dropped 50%,
falling from 0.13 to 0.06 day−1. A rapid decrease in the
microalgae growth rate was also found at higher
temperatures (25°C) [102].
Table 4. Effect of Light/Dark cycle and light intensity on nitrate removal efficiency and biomass production extracted from
original references.
Algal Species
L/D
Cycle
Light
Intensity
(µmol m-2s-1)
Initial NO-
3
(mg L-1)
Final NO-
3
(mg L-1)
Nitrate
Removal
Rate
(mg L-1 d1)
Removal
Time
(d)
Biomass
Concentration
(g L-1)
T(˚C)
pH
Reference
Chlorella
kessleri
12h(L+D)
-
168.1
154.1
4.6
3
-
30
-
[87]
24h(L)
168.1
136.5
10.5
3
-
30
-
[87]
Mixed algae
(Chlamydomonas
reinhardtii,
Scenedesmus
rubescence,
Chlorella
vulgaris)
12h(L+D)
112
7
0
0.7
10
7.51 g/m2/d
22.3
-
[88]
24h(L)
7
0
0.7
10
9.38 g/m2/d
22.3
-
[88]
Trentepohlia
urea
-
5
182
-
0.94
4
0.001
25
8
[91]
10
182
-
1.1
4
0.002
25
8
[91]
20
182
-
1.02
4
0.001
25
8
[91]
50
182
-
0.8
4
0.002
25
8
[91]
Chlamydomonas
reinhardtii
-
400
-
-
2.2
4h
-
25
6.5-7.5
[89]
800
-
-
5.8
4h
-
25
6.5-7.5
[89]
1000
-
-
6.3
4h
-
25
6.5-7.5
[89]
Table 5. Effect of pH on nitrate removal rate extracted from original references.
Species
pH
Buffer
Sparge
pH
Initial
NO3
-
(mg L-1)
Nitrate
Uptake
Rate
(mg L-1 d-1)
Biomass
Concentration
(g L-1)
T(˚C)
Light
Intensity
(µmol m-2 s-1)
Removal
Time
(Day)
Reference
Monoraphidium sp. 92
-
5% CO2
-
198
49.5
1.89
24
350
6
[92]
-
Air
-
204.6
17.05
1.13
24
350
12
[92]
Scenedesmus sp. 131
-
5% CO2
-
198
33
2.41
24
350
6
[92]
-
Air
-
210.8
26.35
1.23
24
350
8
[92]
Scenedesmus sp.
Unbuffered
-
8.4–11.1
90
22.5
0.83
27
75
4
[93]
Unbuffered
-
-
180
22.5
0.72
27
75
8
[93]
Unbuffered
-
-
360
25.7
1.56
27
75
14
[93]
HEPES
-
7.5
180
12.85
0.97
27
75
20
[93]
CHES
-
9.4
180
15
1.08
27
75
20
[93]
CAPS
-
9.4-10.5
90
22.5
1
27
75
4
[93]
CAPS
-
-
180
21.25
0.54
27
75
8
[93]
CAPS
-
-
360
25.7
1.60
27
75
14
[93]
Coelastrella saipanensis
Unbuffered
-
6.5–10
180
7.83
0.71
27
75
8
[93]
HEPES
-
7.5
180
7.5
0.86
27
75
14
[93]
CHES
-
9.0
180
8.75
0.96
27
75
12
[93]
CAPS
-
9.7–10.0
180
7.23
0.71
27
75
8
[93]

10 Current Biotechnology, 2015, Volume 4, No. 3 Taziki et al.
Adaptation to changes in temperature involves a variety
of responses in microalgae. Rising temperature increases
photosynthetic carbon fixation [95], although not the light
dependent reactions of photosynthesis. As photosynthetic
rates are enhanced, nutrient assimilation and other energy
and reductant requiring processes, including nitrate uptake,
also increase with temperature [103]. A maximum nitrate
uptake rate of 7.2 µmol mg-1 Chl per h in Chlamydomonas
rein hardtii was reported at 30°C [89] while arctic species
have optimal temperatures for both growth and nitrate
assimilation at temperatures near freezing [104]. Specific
affinity for inorganic N was studied in several algae and
bacterial strains in chemostat cultures. While specific affinity
for nitrate was strongly dependent on temperature (Q10 = 3,
where Q10 is the proportional change with a 10°C
temperature increase) and decreased below the optimum
temperature, the specific affinity for ammonium exhibited no
clear temperature dependence. This work implies that at low
temperatures, there is an increased dependence on ammonia
rather than nitrate as a N source [104].
4.5. Other Parameters
There are many other parameters affecting nitrate
removal rate and growth, including mixing velocity which
can act to optimize light exposure and nutrient availability in
culture media. Different mixing velocities of 0, 100 and 300
rpm were used for mixed algal cultures (Chlamydomonas
reinhardtii, Scenedesmus rubescens and Chlorella vulgaris)
[88]. The reactor with 300 rpm mixing velocity had the
maximum nitrate and nitrite removal efficiency. The initial
concentrations of nitrate and nitrite were 7.1 and 1.2 mg L-1
respectively, for the three mixing velocities. Both NO3- and
NO2- were removed above 99% at the end of the each
experiment [88]. Provision of both macro- and micro-
elements are also critical for growth and nitrate uptake. An
increase in nitrate removal rates by Scenedesmus accutus,
was observed by adding FeSO4 to wastewater. All of the
nitrate in the wastewater was removed after 48 h in
comparison to control media (with no FeSO4 addition) that
consume nitrate in 72 h [70] suggesting that Fe limited algal
growth in the wastewater used.
5. ALTERNATIVE TECHNOLOGIES FOR NITRATE
BIOREMEDIATION
Cell immobilization and biofilm systems have been
suggested as cost effective systems for wastewater treatment
including nitrate bioremediation [105].
5.1. Cell Immobilization
Chevalier and Noue (1985) were among the first to
immobilize microalgae in carrageenan beads for nutrient
removal. Since then, the entrapment of microalgae in gel
beads has been explored for nutrient removal from
wastewater as a method providing ease of harvest, one of the
major technical problems constraining algal systems [106].
Entrapment of microalgae in alginate or carrageenan beads is
the most common immobilization techniques [107], while
chitosan and polyvinyl foams are inexpensive polymers with
a long term performance [108]. Nitrate and nitrite removal
rates by Chlamydomonas reinhardtii in alginate beads were
reported at 5.3 and 4 µmol mg-1 chl h-1 respectively. A
sequential consumption of nitrite and then nitrate occurred
after ammonium was completely consumed, suggesting that
nitrite inhibits nitrate uptake in this system. The authors
suggested that photorespiration in the entrapped cells, due to
an in increased O2/CO2 ratio, lead to the local accumulation
of ammonium [109]. In freely suspended cells of C.
reinhardtii, nitrate and nitrite were consumed simultaneously
and at higher rates (6.1 and 5.8 µmol mg-1 chl h-1
respectively) [109].
Several studies have reported higher nitrate removal
efficiencies by immobilized cells in comparison with free
living microalgae. Chitosan immobilization of Scenedesmus
sp. cells resulted in a 70% nitrate removal within 12 h, at a
rate significantly higher than free living cells (20% nitrate
removal within 36 h of treatment) (Fig. 2) [110]. However,
the percentage of nitrate removal was lower than that
reported by Lau et al. (1998a), where complete consumption
of nitrate from its initial value of 11.5 mg L-1 by
immobilized C. vulgaris was observed [111]. The higher
initial nitrate concentration (44 mg L-1) of the Scenedesmus
sp. experiments may have influenced the rate [110].
Variations in immobilization methods, specific algae
strains and culture conditions, appear to have led to the
inconsistent rates found in the literature. The type of
immobilized bead was important parameter influencing
nitrate and nitrite removal efficiency in a study by Mallick
and Rai (1994). They compared nitrate and nitrite uptake
rates of Anabaena doliolum and Chlorella vulgaris in
immobilized beads composed of chitosan, agar, alginate,
carrageenan and free-living cells. They reported chitosan
immobilized cells had the maximum efficiency in term of
nitrate and nitrite removal (Table 6). The nitrate uptake rates
were 3.66 and 2.86 [µg NO3- (per mg dry wt-1) h-1] for
Anabaena doliolum and Chlorella vulgaris, respectively,
while nitrite removal rates were lower at 1.3 and 1.6 [µg
NO2- (mg dry wt-1) h-1], respectively [112].
A major problem in immobilization technology is that
microalgae may be released from the beads when the
maximum holding capacity is surpassed [113]. A twin layer
system was developed to separate microalgae from their
growth medium and allow diffusion of nutrients from the
media to the cells. A twin layer system was used to remove
nitrate from municipal wastewater by two green microalgae
(Chlorella vulgaris and Scenedesmus rubescens). Nitrate
concentrations at day 4 were 0.09 mg L-1 and 0.10 mg L-1 for
C. vulgaris and S. rubescens, respectively with initial value
of 4 mg L-1 by both algae [114].
5.2. Biofilm formation
Microalgal biofilm systems have some advantages
allowing short hydraulic retention times [115, 116] and
requires less energy input because stirring is not needed
compared to suspended microalgal systems. Nitrate removal
using a microalgal biofilm investigated at low, intermediate,
and high nutrient loads, showed that the higher loading rates
lead to lower nitrate removal. In the minimum loading rate
(0.18 g m-2 d-1), nitrate was completely removed from initial
concentration of 9 mg L-1 in 6 days [117].

Nitrate and Nitrite Removal from Wastewater Using Algae Current Biotechnology, 2015, Volume 4, No. 3 11
CONCLUSION
The use of microalgae for nitrate and nitrite removal
from wastewater has potential as an alternative technique to
conventional nutrient removal methods when coupled to
biomass production. Wastewater can be considered as a cost
effective and available medium for microalgae. By
assimilation of nitrate and nitrite and conversion into
biomass, algae growth provides an efficient means of
reclaiming the nutrients in wastewater and purifying water.
The conclusions from this review are that: a) The most
favorable strains for nitrate removal and biomass production
are Neochloris oleoabundans, Duniella tertiolecta and
Chlorella vulgaris, b) Higher initial nitrate concentrations
result in higher nitrate removal rates in many, but not all
species, c) Increasing light intensity to the saturation point
leads to maximum nitrate removal and when light intensity
surpasses that point, photosynthetic efficiency along with
nitrate uptake decreases, d) Optimal growth temperatures of
10-35˚C have been reported for different species and nitrate
uptake but ammonia uptake is not strongly influenced by
temperature, e) Using buffers or enriched CO2 concentrations
help to prevent major fluctuations of pH when nitrate is used
as the nitrogen source, f) Entrapment of microalgae in
alginate or carrageenan beads for wastewater treatment
shows promise and avoids the high cost of harvesting free-
living microalgae, and g) In addition to their bioremediation
capabilities, the microalgae feedstock can be used to produce
Fig. (2). Nitrate uptake from medium as percentage removed by free-living cells (!) and immobilized Scenedesmus sp. cells (▲).
(Reproduced from [110] with permission).

12 Current Biotechnology, 2015, Volume 4, No. 3 Taziki et al.
a variety of products including feed, biofuels, nutraceuticals,
high value chemicals and hydrogen in an integrated system.
Therefore, algal-based biotechnology is an environmentally
and economically sound approach to reduce nitrate and
nitrite level in wastewater while generating valuable co-
products. In the near future, wastewater engineers and
scientists from the algae biomass industry will retrofit
wastewater treatment plants to integrate wastewater
treatment and CO2 mitigation, such that environmental water
quality standards are maintained and multiple sources of
revenue can be generated from the production of algal
biomass. All of this is dependent on a fundamental
understanding of the physiology and growth characteristics
of the individual stains of algae.
CONFLICT OF INTEREST
The authors confirm that this article content has no
conflict of interest.
ACKNOWLEDGEMENTS
H. A gratefully acknowledge the financial support from
ATF Committee.
REFERENCES
[1] Aslan S, Turkman A. Nitrate and pesticides removal from
contaminated water using biodenitrification reactor. Process
Biochem 2006; 41(4): 882-86.
[2] Altintas O, Tor A, Cengeloglu Y, Ersoz M. Removal of nitrate
from the aqueous phase by Donnan dialysis. Desalination 2009;
239(1-3): 276-82.
[3] Calderer M, Jubany I, Pérez R, Martí V, de Pablo J. Modelling
enhanced groundwater denitrification in batch micrococosm tests.
Chem Eng J 2010; 165(1): 2-9.
[4] Dodds WK. Nutrients and the “dead zone”: the link between
nutrient ratios and dissolved oxygen in the northern Gulf of
Mexico. Front Ecol Environ 2006; 4(4): 211-17.
[5] Flynn KJ, Butler I. Nitrogen sources for the growth of marine
microalgae: role of dissolved free amino acids. Mar Ecol Prog Ser
1986; 34(3): 281-304.
[6] Burgin AJ, Hamilton SK, Have we overemphasized the role of
denitrification in aquatic ecosystems? A review of nitrate removal
pathways. Front Ecol Environ 2007; 5(2): 89-96.
[7] Florczyk M, Łakomiak A, Woêny M, Brzuzan P. Neurotoxicity of
cyanobacterial toxins. Environ Biotechnol 2014; 10(1): 26-43.
[8] McCarthy J, Carpenter E. Nitrogen cycling in near-surface waters
of the open ocean. In: Carpenter EM, Capone DG, Eds. Nitrogen in
the marine environment. New York: Academic Press 1983; pp.
487-512.
[9] Wheeler PA, Kokkinakis SA. Ammonium recycling limits nitrate
use in the oceanic subarctic Pacific. Limnol Oceanogr 1990; 35(6):
1267-78.
[10] Collos Y, Harrison PJ. Acclimation and toxicity of high ammonium
concentrations to unicellular algae. Mar Pollut Bull 2014; 80(1-2):
8-23.
[11] Mancinelli RL. The nature of nitrogen: an overview. Life Support
Biosph Sci 1995; 3(1-2): 17-24.
[12] Kang J. Mitigation of ammonia gas from animal house using
microalgae. Graduate Theses and Dissertations. Paper12782. Ames,
Iowa: Iowa State University 2012.
[13] Medina M, Neis U. Symbiotic algal bacterial wastewater treatment:
effect of food to microorganism ratio and hydraulic retention time
on the process performance. Water Sci Technol 2007; 55(11): 165-
71.
[14] Wall D, Mulla D, Weiss S, et al. Nitrogen in Waters: Forms and
Concerns. Nitrogen in Minnesota Surface Waters. Minnesota
Pollution Control Agency 2013.
http://www.pca.state.mn.us/index.php/about-mpca/mpca-
news/featured-stories/report-on-nitrogen-in-surface-water.html
[15] Howarth RW, Bringezu S, Eds. Proceedings of the Scientific
Committee on Problems of the Environment. International Biofuels
Project Rapid Assessment; 2008 September 22–25; Gummersbach,
Germany 2009.
[16] Ravishankara AR, Daniel JS, Portmann RW. Nitrous oxide (N2O):
the dominant ozone-depleting substance emitted in the 21st
century. Science 2009; 326(5949): 123-25.
[17] Chapin III FS, Matson PA, Vitousek PM. Principles of terrestrial
ecosystem ecology, Springer Science & Business Media 2011.
Table 6. Removal of NO3
- and NO2
- by free and immobilized Anabaena doliolum and Chlorella vulgaris over three cycles (I. II and
III). (Copied from [112] with permission).
Cells and Method of Immobilization
NO3
- Uptake [µg NO3
- (mg dry wt-1) h-1]
NO2
- Uptake [µg NO2
- (mg Dry wt-1) h-1]
I
II
III
I
II
III
Anabaena doliolum
Alginate
3.4
2.4
2.0
1.3
1.1
0.6
Agar
3.9
2.5
1.8
1.5
1.1
0.7
Carrageenan
2.5
2.0
1.0
1.2
1.0
0.5
Chitosan
4.3
3.7
3.0
1.6
1.5
0.9
Free cells
2.5
1.9
0.9
1.1
0.9
0.5
Chlorella vulgaris
Alginate
3.5
2.1
1.7
1.5
1.0
0.4
Agar
3.7
2.3
1.7
1.6
1.2
0.4
Carrageenan
2.9
1.7
1.5
1.4
0.9
0.3
Chitosan
3.8
2.6
2.2
1.7
1.2
0.6
Free cells
2.9
2.1
1.2
1.4
1.1
0.4
Standard error
0.02- 0.08
0.01- 0.05

Nitrate and Nitrite Removal from Wastewater Using Algae Current Biotechnology, 2015, Volume 4, No. 3 13
[18] Law Y, Ye L, Pan Y, Yuan Z. Nitrous oxide emissions from
wastewater treatment processes. Philos Trans R Soc Lond B Biol
Sci 2012; 367(1593): 1265-77.
[19] Alcántara C, Muñoz R, Norvill Z, Plouviez M, Guieysse B. Nitrous
oxide emissions from high rate algal ponds treating domestic
wastewater. Bioresour Technol 2015; 177: 110-117.
[20] Shrimali M, Singh KP. New methods of nitrate removal from
water. Environ Pollut 2001; 112(3): 351-59.
[21] Blowes DW, Robertson WD, Ptacek CJ, Merkley C. Removal of
agricultural nitrate from tile-drainage effluent water using in-line
bioreactors. J Contam Hydrol 1994; 15(3): 207-21.
[22] El Midaoui A, Elhannouni F, Taky M, et al. Optimization of nitrate
removal operation from ground water by electrodialysis. Sep Purif
Technol 2002; 29(3): 235-44.
[23] Glass C, Silverstein J. Denitrification of high-nitrate, high-salinity
wastewater. Water Res 1999; 33(1): 223-229.
[24] Ayyasamy PM, Shanthi K, Lakshmanaperumalsamy P, Lee S-J,
Choi N-C, Kim D-J. Two-stage removal of nitrate from
groundwater using biological and chemical treatments. J Biosci
Bioeng 2007; 104(2): 129-134.
[25] Pintar A, Batista J. Improvement of an integrated ion-
exchange/catalytic process for nitrate removal by introducing a
two-stage denitrification step. Appl Catal B 2006; 63(1-2): 150-59.
[26] McAdam EJ, Judd SJ. A review of membrane bioreactor potential
for nitrate removal from drinking water. Desalination 2006; 196(1-
3): 135-48.
[27] Ghafari S, Hasan M, Aroua MK. Bio-electrochemical removal of
nitrate from water and wastewater-A review. Bioresour Technol
2008; 99(10): 3965-74.
[28] Samatya S, Kabay N, Yüksel Ü, Arda M, Yüksel M. Removal of
nitrate from aqueous solution by nitrate selective ion exchange
resins. React Funct Polym 2006; 66(11): 1206-14.
[29] Choe S, Chang YY, Hwang KY, Khim J. Kinetics of reductive
denitrification by nanoscale zero-valent iron. Chemosphere 2000;
41(8): 1307-11.
[30] Ginner JL, Alvarez PJ, Smith SL, Scherer MM. Nitrate and nitrite
reduction by Fe0: influence of mass transport, temperature, and
denitrifying microbes. Environ Eng Sci 2004; 21(2): 219-229.
[31] Tiedje JM. Ecology of denitrification and dissimilatory nitrate
reduction to ammonium. In: Sehnder AJB, Eds. Biology of
Anaerobic Microorganisms. New York: Wiley 1988; pp. 179–244.
[32] Jetten MSM, Wagner M, Fuerst J, van Loosdrecht M, Kuenen G,
Strous M. Microbiology and application of the anaerobic
ammonium oxidation (‘anammox’) process. Curr Opin Biotechnol
2001; 12(3): 283-88.
[33] Jetten MSM, Strous M, van de Pas-Schoonen KT, et al. The
anaerobic oxidation of ammonium. FEMS Microbiol Rev 1998;
22(5): 421-37.
[34] Dannenberg S, Kroder M, Dilling W, Cypionka H. Oxidation of H2,
organic compounds and inorganic sulfur compounds coupled to
reduction of O2 or nitrate by sulfate-reducing bacteria. Arch
Microbiol 1992; 158(2): 93-99.
[35] Fossing H, Gallardo VA, Jorgensen BB, et al. Concentration and
transport of nitrate by the mat-forming sulphur bacterium
Thioploca. Nature 1995; 374(6524): 713-15.
[36] Otte S, Schalk J, Kuenen JG, Jetten MSM. Hydroxylamine
oxidation and subsequent nitrous oxide production by the
heterotrophic ammonia oxidizer Alcaligenes faecalis. Appl
Microbiol Biotechnol 1999; 51(2): 255-61.
[37] Weber KA, Achenbach LA, Coates JD. Microorganisms pumping
iron: anaerobic microbial iron oxidation and reduction. Nat Rev
Microbiol 2006; 4(10): 752-64.
[38] Kumar K, Das D. Growth characteristics of Chlorella sorokiniana
in airlift and bubble column photobioreactors. Bioresour Technol
2012; 116: 307-13.
[39] Collos Y, Berges J. Nitrogen metabolism in phytoplankton. Oxford,
UK: Eolss Publishers 2003.
[40] Tucker, Young, Jackson, Tull, Inc. Detroit Water and Sewerage
Department Wastewater Master Plan, DWSD Project No. CS-1314.
Basis of Cost Estimates. Technical Memorandum 2003.
[41] Oswald WJ. Terrestrial approaches to integration of waste
treatment. Waste Manage Res
1991; 9(1): 477-84.
[42] Kumar A, Ergas S, Yuan X, et al. Enhanced CO2 fixation and
biofuel production via microalgae: recent developments and future
directions. Trends Biotechnol 2010; 28(7): 371-380.
[43] Giordano M, Beardall J, Raven JA. CO2 concentrating mechanisms
in algae: Mechanisms, environmental modulation, and evolution.
Annu Rev Plant Biol 2005; 56: 99-131.
[44] Stewart WDP. The effect of available nitrate and ammonium-
nitrogen on the growth of two nitrogen-fixing blue-green algae. J
Exp Bot 1964; 15(1): 138-145.
[45] Azov Y, Goldman JC. Free ammonia inhibition of algal
photosynthesis in intensive cultures. Appl Environ Microbiol 1982;
43(4): 735-739.
[46] Collos Y, Harrison PJ. Acclimation and toxicity of high ammonium
concentrations to unicellular algae. Mar Pollut Bull 2014; 80(1-2):
8-23.
[47] Woertz I, Feffer A, Lundquist T, Nelson Y. Algae grown on dairy
and municipal wastewater for simultaneous nutrient removal and
lipid production for biofuel feedstock. J Environ Eng 2009;
135(11): 1115-22.
[48] Craggs R, Heubeck S, Lundquist T, Benemann J. Algal biofuels
from wastewater treatment high rate algal ponds. Water Sci
Technol 2011; 63(4): 660-65.
[49] Wilkie AC, Mulbry WW. Recovery of dairy manure nutrients by
benthic freshwater algae. Bioresour Technol 2002; 84(1): 81-91.
[50] Mulbry W, Kondrad S, Pizarro C, Kebede-Westhead E. Treatment
of dairy manure effluent using freshwater algae: Algal productivity
and recovery of manure nutrients using pilot-scale algal turf
scrubbers. Bioresour Technol 2008; 99(17): 8137-42.
[51] Martınez ME, Sánchez S, Jiménez JM, El Yousfi F, Munoz L.
Nitrogen and phosphorus removal from urban wastewater by the
microalga Scenedesmus obliquus. Bioresour Technol 2000; 73(3):
263-72.
[52] Lundquist TJ, Woertz IC, Benemann JR. Algal high rate ponds
with CO2 addition for energy efficient nutrient recovery. Nutrient
Recovery and Management conference. Miami, Florida 2011.
[53] Gerardi M. Management in Wastewater as an Indicator of Process
Efficiency. England Intertate Water Pollution Control Commision.
Interstate Water Report 2007.
[54] Siegal LM, Wilkerson JO. Structure and function of spinach
ferredoxin-nitrite reductase. In: wray JL, Kinghorn JR, Eds.
Molecular and genetic aspects of nitrate assimilation. New York:
Oxford University Press 1989; pp. 263-83.
[55] Buchanan BB, Gruissem W, Jones RL. Biochemistry & molecular
biology of plants. the American Society of Plant Physiologists.
Maryland, USA2000.
[56] Dortch Q. The interaction between ammonium and nitrate uptake in
phytoplankton. Mar Ecol Prog Ser 1990; 61: 183-201.
[57] Calero F, Ullrich WR, Aparicio PJ, Regulation by Monochromatic
Light of Nitrate Uptake in Chlorella fusca. In: Senger H, Eds. The
Blue Light Syndrome. Springer Berlin Heidelberg 1980; pp. 411-
21.
[58] Azuara MP, Aparicio P. Effects of light quality, CO2 tensions and
NO3- concentrations on the inorganic nitrogen metabolism of
Chlamydomonas reinhardii, Photosynth Res 1984; 5(2): 97-103.
[59] Ullrich W. Uptake and reduction of nitrate: Algae and fungi. In:
Lauchli A, Eds. Inorganic Plant Nutrition. Verlag Berlin-
Heidelberg: Springer 1983; pp. 376-97.
[60] Ullrich WR, Lazarova J, Ullrich CI, Witt FG, Aparicio PJ. Nitrate
uptake and extracellular alkalinization by the green alga
Hydrodictyon reticulatum in blue and red light. J Exp Bot 1998;
49(324): 1157-62.
[61] Ullrich WR. Transport of nitrate and ammonium through plant
membranes. In: Mengel K, Pilbeam DJ, Eds. Nitrogen Metabolism
of Plants. Oxford: Clarendon Press 1992; pp. 121-37.
[62] Flores E, Frías JE, Rubio LM, Herrero A. Photosynthetic nitrate
assimilation in cyanobacteria. Photosynth Res 2005; 83(2): 117-33.
[63] Hipkin CR, Al-Bassam BA, Syrett PJ. Pyridine nucleotide
specificity and other properties of purified nitrate reductase from
Chlorella variegata. Planta 1979; 144(2): 137-41.
[64] Dalling MJ, Tolbert NE, Hageman RH. Intracellular location of
nitrate reductase and nitrite reductase. I. Spinach and tobacco
leaves. Biochim. Biophys. Acta, Bioenergetics 1972; 283 (3): 505-
12.
[65] Wallsgrove RM, Lea PJ, Miflin BJ. Distribution of the enzymes of
nitrogen assimilation within the pea leaf cell. Plant Physiol 1979;
63(2): 232-36.
[66] Lopez-Ruiz A, Roldan JM, Verbelen JP, Diez J. Nitrate Reductase
from Monoraphidium braunii immunocytochemical localization

14 Current Biotechnology, 2015, Volume 4, No. 3 Taziki et al.
and immunological characterization. Plant Physiol 1985; 78(3):
614-18.
[67] Holdsworth RH. The isolation and partial characterization of the
pyrenoid protein of Eremosphaera viridis. J Cell Biol 1971; 51(2):
499-513.
[68] Satoh H, Okada M, Nakayama K, Miyaji K. Purification and
further characterization of pyrenoid proteins and ribulose-1, 5-
bisphosphate carboxylase-oxygenase from the green alga Bryopsis
maxima. Plant Cell Physiol 1984; 25(7): 1205-14.
[69] Sacristán de Alva M, Luna-Pabello VM, Cadena E, Ortíz E. Green
microalga Scenedesmus acutus grown on municipal wastewater to
couple nutrient removal with lipid accumulation for biodiesel
production. Bioresour Technol 2013; 146: 744-48.
[70] Doria E, Longoni P, Scibilia L, Iazzi N, Cella R, Nielsen E.
Isolation and characterization of a Scenedesmus acutus strain to be
used for bioremediation of urban wastewater. J Appl Phycol 2012;
24 (3): 375-83.
[71] Nunez VJ, Voltolina D, Nieves M, Pina P, Medina A, Guerrero M.
Nitrogen budget in Scenedesmus obliquus cultures with artificial
wastewater. Bioresour Technol 2001; 78(2): 161-64.
[72] Su Y, Mennerich A, Urban B. Comparison of nutrient removal
capacity and biomass settleability of four high-potential microalgal
species. Bioresour Technol 2012; 124: 157-62.
[73] Sydney EB, da Silva TE, Tokarski A, et al. Screening of
microalgae with potential for biodiesel production and nutrient
removal from treated domestic sewage. Appl Energy 2011; 88(10):
3291-94.
[74] Cheng P, Ji B, Gao L, Zhang W, Wang J, Liu T. The growth, lipid
and hydrocarbon production of Botryococcus braunii with attached
cultivation. Bioresour Technol 2013; 138: 95-100.
[75] Kang CD, An JY, Park TH, Sim SJ. Astaxanthin biosynthesis from
simultaneous N and P uptake by the green alga Haematococcus
pluvialis in primary treated wastewater. Biochem Eng J 2006;
31(3): 234-38.
[76] Kang CD, Han SJ, Choi SP, Sim SJ. Fed-batch culture of
astaxanthin-rich Haematococcus pluvialis by exponential nutrient
feeding and stepwise light supplementation. Bioprocess Biosyst
Eng 2010; 33(1): 133-39.
[77] Wang L, Min M, Li Y, et al. Cultivation of green algae Chlorella
sp. in different wastewaters from municipal wastewater treatment
plant. Appl Biochem Biotechnol 2010; 162(4): 1174-86.
[78] Ruiz J, Álvarez P, Arbib Z, Garrido C, Barragán J, Perales JA.
Effect of nitrogen and phosphorus concentration on their removal
kinetic in treated urban wastewater by Chlorella vulgaris. Int J
Phytoremediation 2011; 13(9): 884-96.
[79] Cai T, Park SY, Li Y. Nutrient recovery from wastewater streams
by microalgae: Status and prospects. Renew Sust Energ Rev 2013;
19: 360-69.
[80] Corey P, Kim JK, Duston J, Garbary DJ, Prithiviraj B.
Bioremediation potential of Palmaria palmata and Chondrus
crispus (Basin Head): effect of nitrate and ammonium ratio as
nitrogen source on nutrient removal. J Appl Phycol 2013; 25(5):
1349-58.
[81] Li Y, Horsman M, Wang B, Wu N, Lan CQ. Effects of nitrogen
sources on cell growth and lipid accumulation of green alga
Neochloris oleoabundans. Appl Microbiol Biotechnol 2008; 81 (4):
629-36.
[82] Hulatt CJ, Lakaniemi AM, Puhakka JA, Thomas DN. Energy
demands of nitrogen supply in mass cultivation of two
commercially important microalgal species, Chlorella vulgaris and
Dunaliella tertiolecta. BioEnergy Research 2012; 5(3): 669-84.
[83] Wang B, Lan CQ. Biomass production and nitrogen and
phosphorus removal by the green alga Neochloris oleoabundans in
simulated wastewater and secondary municipal wastewater
effluent. Bioresour Technol 2011; 102(10): 5639-44.
[84] Jeanfils J, Canisius MF, Burlion N. Effect of high nitrate
concentrations on growth and nitrate uptake by free-living and
immobilized Chlorella vulgaris cells. J Appl Phycol 1993; 5(3):
369-74.
[85] Ren M, Ogden K. Cultivation of Nannochloropsis gaditana on
mixtures of nitrogen sources. Environ Prog Sustain Energy 2014;
33(2): 551-55.
[86] Ogbonna JC, Yoshizawa H, Tanaka H. Treatment of high strength
organic wastewater by a mixed culture of photosynthetic
microorganisms. J Appl Phycol 2000; 12(3-5): 277-84.
[87] Lee K, Lee CG. Effect of light/dark cycles on wastewater
treatments by microalgae. Biotechnol Bioprocess Eng 2001; 6(3):
194-99.
[88] Su Y, Mennerich A, Urban B. Coupled nutrient removal and
biomass production with mixed algal culture: Impact of biotic and
abiotic factors. Bioresour Technol 2012; 118: 469-76.
[89] Garbayo I, Vigara AJ, Conchon V, Dos Santos VA, Vı́lchez C.
Nitrate consumption alterations induced by alginate entrapment of
Chlamydomonas reinhardtii cells. Process Biochem 2000; 36(5):
459-66.
[90] Markou G, Georgakakis D. Cultivation of filamentous
cyanobacteria (blue-green algae) in agro-industrial wastes and
wastewaters: A review. Appl Energy 2011; 88(10): 3389-401.
[91] Abe K, Takahashi E, Hirano M. Development of laboratory-scale
photobioreactor for water purification by use of a biofilter
composed of the aerial microalga Trentepohlia aurea
(Chlorophyta). J Appl Phycol 2008; 20(3): 283-88.
[92] Eustance E, Gardner R, Moll KM, Menicucci J, Gerlach R, Peyton
BM. Growth, nitrogen utilization and biodiesel potential for two
chlorophytes grown on ammonium, nitrate or urea. J Appl Phycol;
2013 25(6): 1663-77.
[93] Gardner R, Peters P, Peyton B, Cooksey KE. Medium pH and
nitrate concentration effects on accumulation of triacylglycerol in
two members of the Chlorophyta. J Appl Phycol 2011; 23(6);
1005-16.
[94] Ruelland E, Zachowski A. How plants sense temperature. Environ
Exp Bot 2010; 69(3): 225-32.
[95] Davison IR. Environmental effects on algal photosynthesis:
temperature. J Phycol 1991; 27(1): 2-8.
[96] Fogg GE. Algal adaptation to stress—some general remarks. In:
Rai LC, Eds. Algal Adaptation to Environmental Stresses. Verlag
Berlin Heidelberg: Springer 2001; pp. 1-19.
[97] Renaud SM, Thinh LV, Lambrinidis G, Parry DL. Effect of
temperature on growth, chemical composition and fatty acid
composition of tropical Australian microalgae grown in batch
cultures. Aquaculture 2002; 211(1): 195-214.
[98] MJ Raeesossadati, Ahmadzadeh H, McHenry MP, Moheimani NR.
CO2 bioremediation by microalgae in photobioreactors: Impacts of
biomass and CO2 concentrations, light, and temperature. Algal Res
2014; 6: 78-85.
[99] Teoh ML, Chu WL, Marchant H, Phang SM. Influence of culture
temperature on the growth, biochemical composition and fatty acid
profiles of six Antarctic microalgae. J Appl Phycol 2004; 16(6):
421-30.
[100] Barsanti L, Gualtieri P. Algae: anatomy, biochemistry, and
biotechnology. CRC press 2014.
[101] Weissman J. Screening for lipid yielding microalgae: Activities for
1983. U.S. `Department of Energy: Solar Energy Research Institute
1984.
[102] Converti A, Casazza AA, Ortiz EY, Perego P, Del Borghi M.
Effect of temperature and nitrogen concentration on the growth and
lipid content of Nannochloropsis oculata and Chlorella vulgaris for
biodiesel production. Chem Eng Process 2009; 48(6): 1146-51.
[103] Jiménez C, Niell FX. Growth of Dunaliella viridis teodoresco:
effect of salinity, temperature and nitrogen concentration. J Appl
Phycol 1991; 3(4): 319-27.
[104] Reay DS, Nedwell DB, Priddle J, Ellis-Evans JC. Temperature
dependence of inorganic nitrogen uptake: reduced affinity for
nitrate at suboptimal temperatures in both algae and bacteria. Appl
Environ Microb 1999; 65(6): 2577-84.
[105] Olguín EJ. Dual purpose microalgae bacteria based systems that
treat wastewater and produce biodiesel and chemical products
within a Biorefinery. Biotechnol Adv 2012; 30(5): 1031-46.
[106] Chevalier P, de la Noüe J. Wastewater nutrient removal with
microalgae immobilized in carrageenan. Enzyme Microb Tech
1985; 7(12): 621-24.
[107] Mallick N. Biotechnological potential of immobilized algae for
wastewater N, P and metal removal: a review. Biometals 2002;
15(4): 377-90.
[108] Olguı́n EJ. Phycoremediation: key issues for cost-effective nutrient
removal processes. Biotechnol Adv 2003; 22(1-2): 81-91.
[109] Garbayo I, León R, Vigara J, Vı́lchez C. Inhibition of nitrate
consumption by nitrite in entrapped Chlamydomonas reinhardtii
cells. Bioresour Technol 2002; 81(3): 207-215.

Nitrate and Nitrite Removal from Wastewater Using Algae Current Biotechnology, 2015, Volume 4, No. 3 15
[110] Fierro S, del Pilar Sánchez-Saavedra M, Copalcúa C. Nitrate and
phosphate removal by chitosan immobilized Scenedesmus.
Bioresour Technol 2008; 99(5): 1274-79.
[111] Lau PS, Tam NFY, Wong YS. Effect of carrageenan
immobilization on the physiological activities of Chlorella
vulgaris. Bioresour Technol 1998; 63(2): 115-21.
[112] Mallick N, Rai LC. Removal of inorganic ions from wastewaters
by immobilized microalgae. World J Microbiol Biotechnol 1994;
10(4): 439-43.
[113] Lau PS, Tam NFY, Wong YS. Operational optimization of
batchwise nutrient removal from wastewater by Carrageenan
immobilized Chlorella vulgaris. Water Sci Technol 1998; 38 (1):
185-92.
[114] Shi J, Podola B, Melkonian M. Removal of nitrogen and
phosphorus from wastewater using microalgae immobilized on
twin layers: an experimental study. J Appl Phycol 2007; 19(5):
417-23.
[115] Roeselers G, Van Loosdrecht MCM, Muyzer G. Phototrophic
biofilms and their potential applications. J Appl Phycol 2008;
20(3): 227-35.
[116] Schumacher G, Blume T, Sekoulov I. Bacteria reduction and
nutrient removal in small wastewater treatment plants by an algal
biofilm. Water Sci Technol 2003; 47(11): 195-202.
[117] Boelee NC, Temmink H, Janssen M, Buisman CJN, Wijffels RH.
Nitrogen and phosphorus removal from municipal wastewater
effluent using microalgal biofilms. Water Res 2011; 45(18): 5925-
33.
Received: May 7, 2015 Revised: September 2, 2015 Accepted: September 9, 2015
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