Fruit and Vegetable Waste Hydrolysates as Growth Medium for Higher Biomass and Lipid Production in Chlorella vulgaris
JEWM
Fruit and Vegetable Waste Hydrolysates as Growth
Medium for Higher Biomass and Lipid Production in
Chlorella vulgaris
Abhimanyu Pratap1, Mritunjay Kumar2 and Sibi Ganapathi3*
1,2
Department of Microbiology, Bangalore City College, Bengaluru, India
3*
Department of Biotechnology, Indian Academy Degree College-Autonomous, Bengaluru, India
Fruit and vegetable wastes include peels, pulp and seeds that constitute about 40% of the total
mass and constitute huge environmental problems. Cultivation of microalgae that utilizes fruit
and vegetable wastes as feedstock to produce value added products such as biomass and lipids
is a unique approach. Different concentrations of fruit waste hydrolysate (FWH) and vegetable
waste hydrolysate (VWH) were used for heterotropic cultivation of Chlorella vulgaris thereby
optimizing the suitable hydrolysate concentration for higher biomass and lipid production. FWH
in the ratio of 8:2 has produced maximum specific growth rate of 1.92 µ d-1
. Higher biomass was
recorded in growth medium supplemented with FWH (0.16 mg L-1
) than VWH medium. Highest
chlorophyll content of 7.2 mg L-1
was observed in 8:2 ratio of FWH whereas it was 4.3 mg L-1
in
VWH at the same concentration. Carotenoid content was highest in VWH than FWH media with a
maximum content of 0.52 and 0.42 mg L-1
respectively. Fruit waste hydrolysates significantly
increased the total lipid content than the vegetable waste hydrolysate medium. Highest lipid
content of 6.63 mg L-1
was recorded in 8:2 ratio of FWH. This work demonstrates the feasibility of
fruit waste hydrolysate as a nutrient source for algal cultivation and a cost reduction of growth
medium in algal biomass and lipid production.
Keywords: Microalgae, fruit wastes, vegetable wastes, hydrolysates, biomass, lipid
INTRODUCTION
Today’s world is highly dependent on the energy derived
from fossil fuels (Doll and Pachauri, 2010) and
development of alternative energy sources from plants and
microbial origin is necessary (Berg and Boland, 2014;
Gaurav et al., 2017). Oleaginous microorganisms
accumulate lipid in their cell more than 20% of a dry
biomass and the lipid can be used as a potential feedstock
for biodiesel production due to their chemical composition.
Oil accumulated in most microalgae is mainly triglyceride
that can be applied to form biodiesel and glycerol through
transesterification. Microalgae are well-adapted to survive
under a large spectrum of environmental stresses
(Tandeau-de-Marsac et al., 1993). Algal biomass can be
fractionated into both bio-energy and food products
(Wijffels et al., 2010). The price of algal biofuel ultimately
depends on the substrate cost, lipid yield, and the quality
of the products formed by the downstream process (Yang
et al., 2006). Commercialization of biofuels using
microalgae is hindered by the fact that it is more expensive
due to the higher costs of heterotrophic growth nutrients
(Hong et al., 2012). In algal cultivation, the cost of carbon
source represents 50% of the cost of growth medium
(Cheng et al., 2009; Li et al., 2007).
*Corresponding author: Sibi Ganapathi, Department of
Biotechnology, Indian Academy Degree College-
Autonomous, Bengaluru, India. Email address:
gsibii@gmail.com
Journal of Environment and Waste Management
Vol. 4(2), pp. 204-210, July, 2017. © www.premierpublishers.org. ISSN: XXXX-XXXX
Research Article

Fruit and Vegetable Waste Hydrolysates as Growth Medium for Higher Biomass and Lipid Production in Chlorella vulgaris
Pratap et al. 205
Recent studies have found that the biomass and lipid
content of algae can be increased through changing
cultivation conditions especially nutrient content of growth
medium (Chiu et al., 2009; Converti et al., 2009).
Microalgae are being cultivated in various culture media
(Pleissner et al., 2013; Wu et al., 2014; Sibi, 2015) and use
of natural wastes as source of growth medium will not only
fulfil the nutrient requirements of microalgae but also
reduce the cost of growth medium.
Interest in the recovery of waste or by-products has been
increasing for both economic and ecological reasons.
Waste generation through fruits and vegetables use higher
due to increase in world population. Wastes emanating
from fruits and vegetables include peels, pulp and seeds
that constitute about 40% of the total mass. The majority
of these waste materials are often improperly disposed,
hence constitute huge environmental problems (Essien et
al., 2005; Lim et al., 2010). Cultivation of microalgae that
utilizes fruit and vegetable wastes to produce value added
products such as biomass and lipids is a unique approach.
Media formulation and optimization are key considerations
in development of bioprocesses that can produce
affordable by products. The study of growth medium
components affecting significantly growth rate, biomass
production, pigments content, biochemical composition of
microorganisms is a step required to advance in the design
of a low-cost culture medium for the efficient production of
value added products. This study aimed primarily at
utilization of fruit and vegetable waste hydrolysates as the
growth medium for the cultivation of microalgae and
secondly to determine the effect of hydrolysates growth
and biochemical composition of microalgae thereby
optimizing the suitable hydrolysate concentration for
higher biomass and lipid production in microalgae.
MATERIALS AND METHODS
Collection of Fruit and Vegetable Wastes
The fruit and vegetable wastes were collected from juice
shops, supermarkets, local markets located in Bangalore
urban areas. Fruit wastes are mixture of banana peels-
13.4%; sapota-8.3%, sweet lime-11.2%, orange-6.4%;
apple-10.3%; mango-14.7%, pomegranate-8.6%,
watermelon-9%, musk melon-5.8% and papaya-12.3%.
Vegetable wastes are foliage-26.8%, corn-4.3%, pumpkin-
4.9%, carrots and beans-11%, tomatoes-19.2%, potatoes-
9.3%, cucumber-8.4%, capsicum-4.9%, cauliflower-8.5%
and cabbage-3.7%.
Pre-treatment of fruit and vegetable wastes
Size-reduced and homogenised FVW was mixed with
deionised water to achieve 250 g solids L-1 and then
refrigerated at 4°C for 24 h. This process allows for passive
leaching of soluble sugars and essential nutrients, whilst
minimising microbial mediated leaching processes. After
24 h incubation, the fruit and vegetable waste slurry was
pretreated by thermal hydrolysis. Hydrolysis via thermal
hydrolysis employed autoclaving of the fruit and vegetable
waste slurry under standardised heat and pressure
conditions of 121°C for 15 min.
Growth medium
Fruit and vegetable waste hydrolysates prepared as
described earlier was diluted into various concentrations
(7:3, 8:2; 9:1, 10:0) with sterile distilled water and used as
growth medium for the cultivation of C. vulgaris.
Growth rate and Biomass concentration
Specific growth rate (μ) of the microalgae was calculated
according to the following formula (Levasseur et al., 1993).
µ=
ln (Nt
/N0)
Tt-T0
Where, Nt and N0 are the dry cell weight concentration (g
L-1) at the end (Tt) and start (T0) of log phase respectively.
Biomass (g L-1) of microalgae grown in the fruit and
vegetable waste hydrolysate medium was determined by
measuring the optical density of samples at 600 nm
(OD600) using UV-Vis spectrophotometer. Biomass
concentration was then calculated by multiplying OD600
values with 0.6, a predetermined conversion factor
obtained by plotting OD600 versus dry cell weight (DCW).
DCW was determined gravimetrically by centrifuging the
algal cells (3,000×g, 10 min) and drying.
Biomass concentration = OD600 × 0.6 ……….. Eq. (1)
Chlorophyll Estimation
Algal cells were centrifuged and extracted with acetone
overnight. The extract was centrifuged at 3000 x g for 5
mins and the chlorophyll content in the supernatant were
determined by measuring the optical densities at 645 and
663 nm in a spectrophotometer (Becker, 1994) and then
calculated using the Eq. (2).
Chl (mg/L) = 8.02 × OD663 + 20.21 × OD645 ……….. Eq. (2)
Carotenoids Estimation
Algal cells were centrifuged and treated with centrifuging
the algal cells and treated with KOH (60% w/w). The
mixture was homogenized and warmed to 40°C for 40
mins and extracted using ethyl ether. The solvent was
evaporated followed by resuspending in acetone and the
optical density was measured at 444 nm (Whyte, 1987).
Total carotenoids were calculated using the Eq. (3).
Ct (mg/L) = 4.32 × OD444 − 0.0439………….. Eq. (3)

Fruit and Vegetable Waste Hydrolysates as Growth Medium for Higher Biomass and Lipid Production in Chlorella vulgaris
J. Environ. Waste Manag. 206
Fig-1: Specific growth rate of Chlorella species grown in fruit waste hydrolysate
Protein Assay
The extraction of proteins from microalgae was performed
using alkali method. Aliquots of algal sample were
centrifuged and 0.5 N NaOH was added to the pellet
followed by extraction at 80˚C for 10 mins. The mixture
was centrifuged and protein content of the supernatant
was estimated using Bovine Serum Albumin (BSA) as
standard (Lowry et al., 1951).
Carbohydrate Assay
Cellular carbohydrates were estimated using the anthrone
method described by Gerhardt et al., (1994) after hot
alkaline extraction (Levya et al., 2008). Briefly, microalgal
pellets were resuspended in distilled water and then
heated in 40% (w/v) KOH at 90˚C for 1 h. After cooling
down, ice cold ethanol was added and stored at -20˚C
overnight followed by centrifugation. The pellet was
resuspended in distilled water and then reacted with
anthrone reagent. D-glucose was used as standard and
the colour development was read at 578 nm in a
spectrophotometer.
Determination of Total Lipids
Algae cells were harvested by centrifugation and then
dried for the analysis of lipid content. The lipids were
extracted using a one step extraction method (Folch et al.,
1956). Dried algal cells added with distilled water were
ultrasonicated and mixed with chloroform: methanol (2:1).
The mixture was left for 30 mins in a water bath (30˚C) and
filtered through a Whatman No.1 filter paper. The filtrate
was transferred to another screw cap tube containing NaCl
solution (0.9%) and the purified chloroform layer was
evaporated to a constant weight in a fuming hood under
vacuum at 60˚C. The total lipid content of dry weight was
calculated using the following Equation (4).
Lipid content (%) = (m2-m0)/m1 × 100………….. Eq. (4)
where m1 is the weight of the dried algal cells, m0 is the
weight of the empty new screw cap tube and m2 is the
weight of the new screw cap tube with the dried lipids. Lipid
productivity (g·L−1·d−1) was determined using the following
Equation (5).
Lipid productivity = Biomass productivity × Lipid content….
Eq. (5)
Statistical Analysis
All the experiments were carried out in triplicates and the
results were expressed as mean values and the standard
deviation (SD). The statistical differences were obtained
through one-way analysis of variance (ANOVA) (p < 0.05).
RESULTS AND DISCUSSION
Specific growth rate of C. vulgaris grown in fruit waste
hydrolysate (FWH) and vegetable waste hydrolysate
(VWH) media was determined in alternate days for a
period of 14 days. Control flasks contained Bristol media
alone to compare the effect of hydrolysate on microalgal
growth. It was found that fruit waste hydrolysate in the ratio
of 8:2 has produced maximum specific growth rate until the
end of cultivation period (Fig-1) which was significantly
higher than the control. This was followed by 9:1 and 7:3
which recorded 1.68 and 1.52 µ d-1. Lower specific growth
rate was observed when the ratio was 10:0.
0
0.5
1
1.5
2
2.5
2 4 6 8 10 12 14
Specificgrowthrate(µ)d-1
Growth period (days)
7:3 8:2 9:1 10:0 Control

Fruit and Vegetable Waste Hydrolysates as Growth Medium for Higher Biomass and Lipid Production in Chlorella vulgaris
Pratap et al. 207
Fig-2: Specific growth rate of Chlorella species grown in vegetable waste hydrolysate
The effect of vegetable waste hydrolysate on the specific
growth rate of C. vulgaris is represented in Fig-2.
Hydrolysates in the ratio of 9:1 and 8:2 recorded highest
growth rate of 1.82 and 1.42 µ d-1. It was found that
vegetable waste hydrolysate in the ratio of 9:1 has
produced maximum specific growth rate at the end of
cultivation period (Fig-2). Lower specific growth rate was
observed when the ratio was 10:0. The lowest growth rate
of 1.08 µ d-1 was observed in control group at the end of
14 days cultivation period.
The biomass concentration of C. vulgaris grown on FWH
and VWH were represented in Fig-3 and 4. In general,
higher biomass was recorded in growth medium
supplemented with fruit waste hydrolysate (0.16 mg L-1).
The biomass was increased with increasing cultivation
period however the concentration started declining after 12
days. However, increase in biomass concentration was
observed with 10:0 ratio of hydrolysate and control
experiments. The hydrolysates compositions, such as the
carbon, nitrogen, pigments, polyphenols, polypeptide,
minerals, volatile substances, and even the pH of the
hydrolysates, might be the factors that influence the
biomass growth of Chlorella vulgaris.
Both chlorophyll and carotenoid contents of C. vulgaris
grown on hydrolysates medium was shown in Fig-5 and
Fig 6. Highest chlorophyll content of 7.2 mg L-1 was
observed in 8:2 ratio of FWH whereas it was 4.3 mg L-1 in
VWH at the same concentration. Carotenoid content was
highest in VWH than FWH media with a maximum content
of 0.52 and 0.42 mg L-1 respectively.
Both protein and carbohydrate contents were determined
photometrically and higher levels of protein were found in
cells grown in FWH. In the case of carbohydrates, highest
content of 18.5 mg L-1 was found in VWH at the ratio of
10:0.
Gas chromatography analysis revealed that the lipid
extract from C. vulgaris after 14 days of cultivation in fruit
waste hydrolysate medium composed of three main types
of fatty acids that could be present in a triglyceride:
saturated (Cn:0), monounsaturated (Cn:1) and
polyunsaturated with two to four double bonds (Cn:2,3,4)
(Table-1). The fatty acid composition of the microalgal lipid
is similar to that of vegetable oils, and so the lipid with this
fatty acid composition is a promising feedstock for
biodiesel production through transesterification
(Christophe et al., 2012). Considering the amount of
monounsaturated and polyunsaturated fatty acids present
in the sample, the degree of unsaturation (DU) was
calculated in accordance with the empirical equation
[DU=(monounsaturated Cn: 1; wt:%)+2 (polyunsaturated
Cn: 2;3;4; wt:%)] described by Ramos et al. (2009). The
DU, which was calculated as 61.39, might have influenced
the cetane number of the biodiesel to be synthesized with
the extracted lipid. Higher the cetane number, the better
the ignition properties of the biodiesel (Meher et al., 2006).
The higher cetane number is associated with lower DU and
vice versa (Knothe et al., 2003), and higher DU than 137
makes lipids unsuitable to meet the European Standard for
the cetane number. Again, the abundance of lignoceric
acid, the longer saturated fatty acid including other
saturated fatty acids might be associated with higher
cetane number. Furthermore, the lower DU indicated the
higher oxidation stability of the biodiesel to be synthesized
with the microbial lipid produced from fruit waste
hydrolysate by C. vulgaris. However, further research is
necessary to synthesize biodiesel with this microalgal lipid
and to study its physico-chemical properties.
0
0.5
1
1.5
2
2.5
2 4 6 8 10 12 14
Specificgrowthrate(µ)d-1
Growth period (days)
7:3 8:2 9:1 10:0 Control

Fruit and Vegetable Waste Hydrolysates as Growth Medium for Higher Biomass and Lipid Production in Chlorella vulgaris
J. Environ. Waste Manag. 208
Table-1: Fatty acid profiles of C. vulgaris grown on FWH media
Length Fatty acid Percentage
C13:1 Tridecanoic acid 0.92
C16:0 Palmitic acid 3.57
C18:0 Stearic acid 2.74
C18:1 Oleic acid 21.68
C18:3 Linolenic acid 6.59
C20:4 Arachidonic acid 4.37
C24:0 Lignoceric acid 24.36
Fig-3: Biomass concentration of Chlorella species grown in fruit waste Fig-4: Biomass concentration of Chlorella species grown in
hydrolysate vegetable waste hydrolysate
Fig-5: Chlorophyll content of C. vulgaris on FWH and VWH media Fig-6: Carotenoid content of C. vulgaris on FWH and VWH media
Production of lipids by algal species is very much
importance in terms of their use in biodiesel, as lipid is the
essential source of biofuel production. The production and
content of lipids from microalgae can be manipulated by
introducing the algae with various carbon sources.
Enzymatic hydrolysates of carbohydrates from various
agroindustrial wastes were used for heterotrophic
cultivation of microalgae as alternative carbon feed stocks
(Xu et al., 2006; Gao et al., 2010; Lu et al., 2010; Wei et
al., 2009; Li et al., 2011; Cheng et al., 2009; Yan et al.,
2011; Liu et al., 2012).
In this study, fruit waste hydrolysates produced highest
specific growth rate and biomass productivity than
vegetable waste hydrolysates. Similar results were
obtained in cellular pigment levels, carbohydrate and
protein content. The hydrolysates compositions, such as
the pH, carbon, nitrogen, pigments, polyphenols,
polypeptide, minerals, volatile substances might be the
factors that influence the biomass growth of Chlorella
vulgaris. This work demonstrates the feasibility of fruit
waste hydrolysate as a nutrient source for algal cultivation
and a cost reduction of growth medium in algal lipid
production may be expected.

Fruit and Vegetable Waste Hydrolysates as Growth Medium for Higher Biomass and Lipid Production in Chlorella vulgaris
Pratap et al. 209
Fig-7: Protein content of C. vulgaris on FWH and VWH media Fig-8: Carbohydrate content of C. vulgaris on FWH and VWH media
Fig-9: Lipid content of C. vulgaris on FWH and VWH media
REFERENCES
Becker EW (1994). Microalgae: Biotechnology and
Microbiology. Cambridge University Press, New York.
Berg P, Boland A (2014). Analysis of ultimate fossil fuel
reserves and associated CO2 emissions in IPCC
scenarios. Nat. Resour. Res. 23(1): 141-158.
Cheng Y, Zhou W, Gao C, Lan K, Gao Y, Wu Q. (2009).
Biodiesel production from Jerusalem artichoke
(Helianthus Tuberosus L.) tuber by heterotrophic
microalgae Chlorella protothecoides, J. Chem. Technol.
Biotechnol. 84: 777-781.
Chiu SY, Kao CY, Tsai MT, Ong SC, Chen CH, Lin CS.
(2009). Lipid accumulation and CO2 utilization of
Nannochloropsis oculata in response to CO2 aeration,
Bioresour. Technol. 100: 833-838.
Christophe G, Kumar V, Nouaille R, Gaudet G, Fontanille
P, et al. (2012). Recent developments in microbial oils
production: A possible alternative to vegetable oils for
biodiesel without competition with human food. Braz.
Arch. Biol. Technol. 55: 29-46.
Converti A, Casazza AA, Ortiz EY, Perego P, Del Borghi
M. (2009). Effect of temperature and nitrogen
concentration on the growth and lipid content of
Nannochloropsis oculata and Chlorella vulgaris for
biodiesel production, Chem. Eng. Proc. 48: 1146-1151.
Doll CN, Pachauri S (2010). Estimating rural populations
without access to electricity in developing countries
through night-time light satellite imagery. Energy Policy.
38(10): 5661-5670.
Essien J, Akpan E, Essien E. (2005). Studies on mould
growth and biomass production using waste banana
peel. Bioresour. Technol. 96 (13): 1451–1456.
Folch J, Lees M, Stanley GHS. (1956). A simple method
for the isolation and purification of total lipids from animal
tissues. J. Biol. Chem. 226: 497-509.
0
1
2
3
4
5
6
7
8
7:3 8:2 9:1 10:0 Control
Lipidproductivity(mgL-1)
Hydrolysate concentration ratio
FWH
VWH

Fruit and Vegetable Waste Hydrolysates as Growth Medium for Higher Biomass and Lipid Production in Chlorella vulgaris
J. Environ. Waste Manag. 210
Gao C, Zhai Y, Ding Y, Wu Q. (2010). Application of sweet
sorghum for biodiesel production by heterotrophic
microalga Chlorella protothecoides, Appl. Energy. 87:
756-761.
Gaurav N, Sivasankari S, Kiran GS, Ninawe A, Selvin J.
(2017). Utilization of biomass for sustainable biofuels: A
Review. Renewable and Sustainable Energy Reviews.
73: 205-214.
Gerhardt P, Murray RGE, Wood WA, Krieg NR. (1994).
Methods for General and Molecular Bacteriology. ASM,
Washington DC.
Knothe G, Matheaus AC, Ryan TW. (2003). Cetane
numbers of branched and straight chain fatty esters
determined in an ignition quality tester. Fuel .82: 971-
975.
Leyva A, Quintana A, Sanchez M, Rodriguez EN, Cremata
J, Sanchez JC. (2008). Rapid and sensitive anthrone-
sulfuric acid assay in microplate format to quantify
carbohydrate in biopharmaceutical products: Method
development and validation. Biologicals .36: 134-141.
Levasseur M, Thompson PA, Harrison PJ.
(1993). Physiological acclimation of marine
phytoplankton to different nitrogen sources. J.
Phycol. 29. 587-595.
Li P, Miao X, Li R, Zhong J. (2011). In situ biodiesel
production from fast-growing and high oil content
Chlorella pyrenoidosa in rice straw hydrolysate, J.
Biomed. Biotechnol. 141207.
Li X, Xu H, Wu Q. (2007). Large-scale biodiesel production
from microalga Chlorella protothecoides through
heterotrophic cultivation in bioreactors, Biotechnol.
Bioeng. 98: 764-771.
Lim JY, Yoon HS, Kim KY, Kim KS, Noh JG, Song IG.
(2010). Optimum conditions for the enzymatic hydrolysis
of citron waste juice using response surface
methodology (RSM). Food Sci. Biotechnol. 19(5): 1135-
1142.
Liu J, Huang J, Jiang Y, Chen F. (2012). Molasses-based
growth and production of oil and astaxanthin by Chlorella
zofingiensis, Bioresour. Technol. 107: 393-398.
Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951).
Protein Measurement with the Folin Phenol Reagent. J.
Biol. Chem. 193: 265-275.
Lu Y, Zhai Y, Liu M, Wu Q. (2010) Biodiesel production
from algal oil using cassava (Manihot esculenta Crantz)
as feedstock, J. Appl. Phycol. 22: 573-578.
Meher LC, Sagar DV, Naik SN. (2006). Technical aspects
of biodiesel production by transesterification-a review.
Renew. Sust. Energ Rev. 10: 248-268.
Pleissner D, Lam WC, Sun Z, Lin CSK. (2013). Food waste
as nutrient source in heterotrophic microalgae
cultivation. Bioresour. Technol. 137: 139-146.
Ramos MJ, Fernandez CM, Casas A, Rodriguez L, Perez
A. (2009). Influence of fatty acid composition of raw
materials on biodiesel properties. Bioresour. Technol.
100: 261-268.
Sibi G (2015). Low cost carbon and nitrogen sources for
higher microalgal biomass and lipid production using
agricultural wastes. Journal of Environmental Science
and Technology. 8(3): 113-121.
Tandeau-de-Marsac N, Houmard J (1995). Adaptation of
cyanobacteria to environmental stimuli: New steps
towards molecular mechanisms. FEMS Microb. Rev.
104: 119-190.
Wei A, Zhang X, Wei D, Chen G, Wu Q, Yang ST. (2009).
Effects of cassava starch hydrolysate on cell growth and
lipid accumulation of the heterotrophic microalgae
Chlorella protothecoides, J. Ind. Microbiol. Biotechnol.
36: 1383-1389.
Whyte JC. (1987). Biochemical composition and energy
content of six species of phytoplankton used in
mariculture of bivalves. Aquaculture. 60: 231-241.
Wijffels RH, Barbosa MJ, Eppink MHM. (2010). Microalgae
for the production of bulk chemicals and biofuels.
Biofuels Bioprod. Biorefining-Biofpr. 4: 287-295.
Hong WK, Kim CH, Rairakhwada D, Kim S, Hur BK, Kondo
A, Seo JW. (2012). Growth of the oleaginous microalgae
Aurantiochytrium sp. KRS101 on cellulosic biomass and
the production of lipids containing high levels of
docosahexaenoic acid. Bioprocess. Biosyst. Eng. 35:
129-133.
Wu S, XR Xu, KF Sun, HB Li. (2014). Effects of fruit waste
hydrolysates on biomass and chlorophyll a fluorescence
parameters of Chlorella pyrenoidosa. Int. J. Environ.
Bioener. 9(2): 105-121.
Xu H, Miao X, Wu Q. (2006). High quality biodiesel
production from a microalga Chlorella protothecoides by
heterotrophic growth in fermenters, J. Biotechnol. 126:
499-507.
Yan D, Lu Y, Chen YF, Wu Q. (2011). Waste molasses
alone displaces glucose-based medium for microalgal
fermentation towards cost-saving biodiesel production,
Bioresour. Technol. 102: 6487-6493.
Yang JS, Huang X, Ni JR. (2006). Mathematical modelling
of batch fermentation of Zoogloea sp. GY3 used for
synthesizing polyhydroxyalkanoates, J. Chem. Technol.
Biotechnol. 81: 789-793.
Accepted 21 July, 2017
Citation: Pratap A, Kumar M, Sibi G (2017) Fruit and
Vegetable Waste Hydrolysates as Growth Medium for
Higher Biomass and Lipid Production in Chlorella
vulgaris. Journal of Environment and Waste Management
4(2): 204-210.
Copyright: © 2017 Pratap et al. This is an open-access
article distributed under the terms of the Creative
Commons Attribution License, which permits unrestricted
use, distribution, and reproduction in any medium,
provided the original author and source are cited.