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Production and characterization of rhamnolipid using
palm oil agricultural refinery waste
Authors
Mohd Nazren Radzuan1,3, Ibrahim Banat2 and James Winterburn*1
1School of Chemical Engineering and Analytical Science, The University of Manchester,
Manchester, United Kingdom
2 School of Biomedical Sciences, Faculty of Life and Health Sciences, University of Ulster,
Northern Ireland, United Kingdom
3 Department of Biological and Agricultural Engineering, Faculty of Engineering, Universiti
Putra Malaysia, Malaysia.
*Corresponding author: [email protected]
Tel: +44(0)161 306 4891
Abstract
In this research we assess the feasibility of using palm oil agricultural refinery waste as a
carbon source for the production of rhamnolipid biosurfactant through fermentation. The
production and characterization of rhamnolipid produced by Pseudomonas aeruginosa PAO1
grown on Palm Fatty Acid Distillate (PFAD) under batch fermentation were investigated.
Results show that P. aeruginosa PAO1 can grow and produce 0.43 g L-1 of rhamnolipid using
PFAD as the sole carbon source. Identification of the biosurfactant product using mass
spectrometry confirmed the presence of monorhamnolipid and dirhamnolipid. The
rhamnolipid produced from PFAD were able to reduce surface tension to 29 mN m-1 with a
critical micelle concentration (CMC) 420 mg L-1 and emulsify kerosene and sunflower oil,
with an emulsion index up to 30%. Results demonstrate that PFAD could be used as a low-
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cost substrate for rhamnolipid production, utilizing and transforming it into a value added
biosurfactant product.
Keywords
Biosurfactant, fermentation, rhamnolipid, palm fatty acid distillate, Pseudomonas aeruginosa
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1. Introduction
Biosurfactants are surface-active compounds produced by microorganisms (Abdel-
Mawgoud et al., 2010). Rhamnolipid are a type of glycolipid biosurfactant produced mainly
by Pseudomonas aeruginosa (Henkel et al., 2012). The type of rhamnolipid produced
depends on the culture conditions, carbon source used and the microbial strain (Silva et al.,
2010). There are two main types of rhamnolipid; monorhamnolipid and dirhamnolipid,
named depending on the numbers of rhamnose molecules present, each having a fatty acid
chain (Müller et al., 2010). The research attention given to biosurfactants has grown
significantly in the last few years because they have many potential commercial applications
in the food processing industry, biomedical, pharmaceutical and agricultural sectors where
the advantages that biosurfactants have over synthetic surfactants, such as biodegradability,
lower toxicity and effectiveness at an extended range of pH and temperature values, can be
exploited (Banat et al., 2010).
At present the main obstacle to more extensive utilization of biosurfactants is the
disadvantageous economics of their production (Nitschke et al., 2005). The market cost of
biosurfactants is high in comparison to petrochemical surfactants, with biosurfactants high
production cost being influenced by three main factors; the raw materials used in
biosurfactant production which account for 10–30% of the total production cost, the
availability of a suitable, economic production and recovery method, and also the yield and
productivity of the producer microorganism (Wadekar et al., 2012).
The economic issues regarding the production of biosurfactants could be greatly
improved with the use of commonly available, renewable, low-cost nutrients, such as
agricultural waste, as a fermentation feedstock (Campos et al., 2014). Adding value to waste
streams will additionally improve the environmental and economic sustainability of
biosurfactant production processes. The use of wastes from the food and agricultural sectors
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that contain a significant amount of free fatty acids have potential to be used as the main
substrate for rhamnolipid production because these wastes are abundant, renewable and low-
cost. Wastes that contain free fatty acids such as waste frying oil, by-products from biodiesel
production, olive oil mill waste and sunflower oil refining waste have been demonstrated to
be suitable for use as substrates for rhamnolipid production by P. aeruginosa strains in
several studies (Moya Ramírez et al., 2015; Pereira et al., 2013; Wadekar et al., 2012;
Benincasa and Accorsini, 2008).
The palm oil industry is one of the major contributors to the Malaysia economy in
terms of foreign exchange earnings employment and development. In 2015, crude palm oil
(CPO) production in Malaysia was 29 × 106 tons with 0.95 × 106 tons of palm fatty acid
distillate (PFAD) as a by-product. The annual year on year increase in CPO production
causes waste disposal and by-product management problems, especially in the case of PFAD.
To produce edible CPO, there are three refining steps; degumming, bleaching and
deodorization. The degumming process removes the gum, bleaching reduces the colour and
deodorization removes volatile compounds from the CPO. After all these refining processes
have been undertaken, a produce low value by-product known as PFAD remains. At room
temperature PFAD is solid, with a light brown colour which changes to a dark brown liquid
upon heating above its melting point of approximately 40○C (Chabukswar et al., 2013).
PFAD contains mainly free fatty acids and currently there is research interest in converting
PFAD into biodiesel and, more broadly, PFAD can also be used as a source of fatty acids for
the animal food, soap and oleochemical production (Hosseini et al., 2015). There is still need
to identify routes to higher added value products, e.g. biosurfactants, using PFAD. As such
there is potential for PFAD to be used as a low-cost substrate for rhamnolipid production. We
are not aware of any literature report using PFAD as the main carbon source for rhamnolipid
production. The novelty of this paper is the demonstration of a new route for the valorisation
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of PFAD and we give new information on the production and characterization of
rhamnolipids produced using palm oil agricultural refinery waste, contributing to expanding
knowledge of biosurfactant production.
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2. Materials and Methods
2.1 Chemicals and Standards
The Malaysian palm oil agricultural refinery waste known as Palm Fatty Acid
Distillate (PFAD) was provided by Sime Darby Refinery-Jamolina and transported to the
United Kingdom by The Cucurbit Company Sdn. Bhd. D-Glucose Monohydrate, Potassium
Chloride, Trizma-Hydrochloride, Peptone, Kerosene were purchased from Sigma-Aldrich Co
Limited. N-Hexane was purchased from VWR International Ltd and Magnesium Sulphate
Heptahydrate, Ammonium Chloride, Ethyl Acetate were purchased from Fisher Scientific
UK Ltd.
2.2 Determination of PFAD composition
PFAD composition was determined using gas chromatography-mass spectrometry
(GC-MS). The PFAD was first converted into fatty acid methyl esters (FAME) through an
esterification reaction. Firstly, the PFAD was melted and dried in an oven at 70°C for 1 day
to remove all the moisture prior to the esterification process. Methanol was then mixed with
PFAD while stirring at 10:1 ratio with sulfuric acid (2.5% weight of PFAD) as a catalyst in a
flat bottom flask. The flask was connected to a reflux condenser and the mixture was heated
to 100°C using a heating mantle for 1 hour. After the esterification process, the product was
allowed to cool, and transferred to a separating funnel and washed with hot distilled water.
Two distinct layers were formed, the top layer being FAME and bottom layer of excess
water, methanol and catalyst. The washing process was stopped when the water becomes
clear and the pH changed to 7. The produced FAME were analysed by gas chromatography-
mass spectrometry (GC-MS), 0.2 mL FAME were added to 1 mL of N-Hexane in 2 mL
micro-vial and a highly polar capillary column BPX 70 from SGE company (length: 60 m,
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ID: 0.22 mm and film thickness: 0.25 µm) was used for separation of FAME compounds.
1µL of the sample was injected for analysis into the GC-MS injector port. The GC injection
port temperature was 220°C and the detector temperature was 250°C. However, the GC oven
was programmed with an increasing starting temperature from 155°C up to 220°C with
2°C/min to 180°C and continue to 4 °C/min to 220°C.
2.3 Biosurfactant production
2.3.1 Microorganism
The microorganism used in this work was Pseudomonas aeruginosa PAO1 which was
supplied by the University of Ulster from their collection.
2.3.2 Media and Culture Condition
P. aeruginosa PAO1 was maintained on nutrient agar plates at 5°C. The culture
medium in seed culture 1 and 2 contained Proteose Peptone Glucose Ammonium Salt
(PPGas) medium that consisted of Ammonium Chloride (NH4Cl) 1 g L-1, Potassium Chloride
(KCl) 1.5 g L-1, Trizma-Hydrochloride (Tris-HCL) 0.12M 19 g L-1, Peptone 10 g L-1,
Magnesium Sulphate Heptahydrate (MgSO4.7H2O) 0.5 g L-1 and carbon sources in distilled
water (Gunther IV et al., 2005). In order to prepare seed culture 1, a loop of cells taken from
a nutrient agar plate was added to a 250 mL flask containing 50 mL culture medium. Seed
culture 1 was used to inoculate seed culture 2 into 400 mL in a 2L flask. The seed culture 1
and 2 were grown in an incubator for 24 hours, set at 37°C and 200 rpm. The concentration
of glucose for seed culture 1 and 2 were 0.5 and 4 g L-1. For the shake flask fermentation, 100
mL of seed culture 2 was used to inoculate 1 L PPGas culture medium in a 5L flask with
carbon source concentrations 20, 50 and 100 g L-1. Two series of shake flask fermentations
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were done in triplicate with different carbon sources were performed using glucose
(Fermentation 1) and PFAD (Fermentation 2).
2.3.3 Growth Measurement
Dry cell weight and optical density were used to measure cell growth. For dry cell
weight, 1 mL samples from both fermentations with addition 0.5 mL of N-Hexane for
Fermentation 2, were centrifuged using Minispin Centrifuge (Eppendorf) at 13,000 g for
10 min. The supernatant was removed and then 1 mL of distilled water was added into the
eppendorf to resuspend and wash cells. The liquid was then decanted and cells with
Eppendorf tube were dried at 70 °C until constant weight was reached. Optical density was
determined by resuspending cell biomass in physiological saline to measure absorption at
600 nm using a Spectrophotometer UVmini-1240 (Shimadzu, USA). The optical density was
read against the physiological saline as a blank. Both measurements were done in triplicate.
2.3.4 Biosurfactant Extraction and Quantification
The rhamnolipid extraction technique was used in combination with acid precipitation
for more efficient yields. The supernatant was mixed with n-hexane in a 1:1 ratio to first
remove the PFAD from the supernatant. The mixture was then centrifuged at 13000 g for
10 min and the resulting supernatant taken and acidified with concentrated hydrochloric acid
to pH 3. The supernatant was then transferred to a separating funnel and mixed with an equal
volume of ethyl acetate and shaken vigorously. A bottom aqueous layer and top ethyl acetate
layer were then allowed to form. The ethyl acetate layer, containing the biosurfactants was
transferred to a separate flask. This extraction was carried out 3 times until no further colour
persisted in the ethyl acetate layer. Finally, 0.5 g of magnesium sulphate per 100 ml were
added into ethyl acetate portion, to remove the traces of water present and sample filtered and
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evaporated using rotary evaporate to yield a brown biosurfactant extract. This extract was
then weighed and the rhamnolipid concentration determined gravimetrically.
2.3.5 Biosurfactant Purification
Solid phase extraction (SPE) was used in normal phase mode, using silica as the
sorbent in the syringe-like tube of 12 ml volume, with 55 µm pores being selected. The SPE
tubes were loaded at the vacuum manifold and connected to the pump to suck solvent from
the tubes. Chloroform was used to prepare the sorbents before extracting the sample. The
rinsing process was stopped when the middle section of the sorbent tube became clear. The
crude rhamnolipid was then diluted with chloroform and loaded onto the sorbent. Chloroform
was then used to elute the rhamnolipid and the process run until there were no colour changes
observed at the output of the tube. The eluted purified rhamnolipid product was dried in an
oven at 70 °C for 1 day and purified biosurfactant was collected.
2.4 Biosurfactant Identification
Biosurfactant identification was carried out using an Agilent 6510 Q-TOF LC/MS
equipped with Agilent 1200 Liquid Chromatography (LC). Extracted rhamnolipid were
diluted in methanol and 5uL of sample was injected into 50% ACN and 0.1% formic acid
using electrospray ionization (ESI) in negative mode.
2.5 Biosurfactant Characterisation
2.5.1 Surface tension and critical micelle concentration (CMC)
Surface tension and CMC were determined with a Krüss K11 Tensiometer equipped
with a De Nöuy ring, measured in triplicate. The CMC was determined by measuring the
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surface tension of dilutions of crude biosurfactant extract from 1000 mg L-1 in 0.1M Tris-HCl
pH 8.0 solutions until a constant value of surface tension was attained. The CMC value (mg
L-1) was obtained from the plot of surface tension against crude biosurfactant concentration.
2.5.2 Emulsion Index
A mixture of 3 ml of crude biosurfactant extract at 1000 mg L-1 in 0.1M Tris-HCl pH
8.0 mixed with 3ml of kerosene and sunflower oil was vigorously shaken for 1 minute to
obtain maximum emulsification. Emulsion index measurement was carried out in triplicate
and calculated as the percent of the height of the emulsified layer relative to the total height
of the liquid column, determined at 24 h time.
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3. Results and Discussion
3.1 PFAD composition
PFAD is a by-product of the crude palm oil refining process, carried out in refinery
mills. The GC-MS analysis and composition of free fatty acid components of PFAD are
shown in Table 1, PFAD is a complex and variable mixture of fatty acids from C12 to C30 of a
range of different saturated and unsaturated fatty acids; mainly oleic acid 41.86 %,
pentadecanoic acid 17.98 %, tridecylic acid 13.98 % and palmitic acid 13.62 % along with
other components, giving a similar composition to those reported by other researcher (Md
Top, 2010). However, different percentages of free fatty acids were observed when
comparing between this study and Md Top (2010), most likely because PFAD is a complex
waste and the fatty acid content depends on the crude palm oil and the refining process itself.
It was expected that PFAD would be suitable substrate for biosurfactant production based on
previous studies shows that free fatty acids are among the best substrates for biosurfactant
production by P. aeruginosa strains (Moya Ramírez et al., 2016; Singh et al., 2013).
3.2 Fermentation kinetics
Rhamnolipid production using glucose (Fermentation 1) and PFAD (Fermentation 2)
was carried out using P. aeruginosa PAO1 under batch fermentation conditions. Shake flask
fermentation using different initial concentrations of carbon sources (20, 50 and 100 g L-1)
were carried out for 96 hours. Growth kinetics and rhamnolipid production were the main
parameters determined. The changing colour of PPGas medium to green once growth has
occurred with both carbon sources, which is mainly due to the presence of pyocyanin
pigments produced during the growth phase (Kalia et al., 2015). The main difference between
both carbon sources was foam production which was quite prominent in the flasks containing
glucose. This was caused by the production of surface active components, potentially
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rhamnolipid in the growth media at the start of fermentation process and the continuity of the
foaming are likely caused by protein from peptone and biosurfactant produced by the cells
(Banat, 1997). Foaming was not observed in Fermentation 2 because the presence of free
fatty acids in PFAD can effectively act as antifoaming agent (Kim et al., 2006).
Generally, the growth of P. aeruginosa PAO1 using glucose and PFAD shows similar
trends, with a period of accelerated growth for 0-24 h fermentation time. This is expected as
initially the substrate and nutrient concentrations are high, allowing for rapid growth.
Fermentation 1 used 20, 50 and 100 g L-1glucose as the carbon source during the fermentation
process. Figure 1shows the fermentation kinetics that for all glucose concentrations with
Figure 1 (a) and (b) showing rapid growth occurred between 0 to 24 h and continued
throughout the fermentation except for 100 g L-1 when cells entered a death phase at 72 h,
Figure 1 (c). It has been suggested that the cells with 100 g L-1 glucose enter death phase
earlier compared with lower concentrations because excess nutrients may lead to toxic
materials accumulation and limited availability of dissolved oxygen which may inhibit
growth (Clark and Blanch, 1997; Lan et al., 2015). The growth pattern observed in Figure 1
(a) and (b) during this experiment when 20 and 50 g L-1 were used showed a similar trend to
those reported for Pseudomonas putida BD2 reported by Janek et al. (2013). The production
of rhamnolipid steadily increased with the highest production detected at the end of
fermentation for 20 and 50 g L-1 of glucose, at a concentration about 0.35 g L-1 as shown in
Figure 1 (a) and (b). For 100 g L-1 glucose concentration in Figure 1 (c), the maximum
rhamnolipid production was at 60 h about 0.36 g L-1. Different concentrations of glucose do
affect the growth and rhamnolipid production in the fermentation. This result was in
accordance with other findings which show the same trend, the higher the concentration of
glucose, the higher the growth and rhamnolipid production because of the increasing
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availability of carbon to be used by P. aeruginosa for growth and rhamnolipid production
(Goswami et al., 2015).
In Fermentation 2, P. aeruginosa PAO1 had the ability to produce rhamnolipid using
different concentration of PFAD as carbon sources. Figure 2 (a), (b) and (c) shows a fast
growth between 0 to 24 h with a steady increase in biomass until the end of fermentation at
all PFAD concentration used. The rhamnolipid production increased during the growth phase
and reached maximum rhamnolipid production of 0.38, 0.39 and 0.43 g L-1 for 20, 50 and 100
g L-1 PFAD concentrations respectively, as shown in Figure 2 (a), (b) and (c). Similar growth
and rhamnolipid production trends were reported by other researchers using waste oil and oil
mill waste (Moya Ramírez et al., 2015). The comparison of the growth and rhamnolipid
production for different concentration of PFAD showed that there were no significant
differences between them. This is mainly due to the availability of PFAD in the culture
media, which is not directly related to the total initial concentration of the PFAD, due to the
low solubility and heterogeneous system, which suggests that roughly equal amounts of
PFAD were dissolved even for different initial concentrations. Therefore the growth and
production of rhamnolipid were similar across the PFAD fermentations.
The data of product yield on substrate consumed (YP/S, g g-1), product yield on initial
substrate (*YP/S, g g-1), product yield on biomass produced (YP/X, g g-1) and productivity (PRL, g
L-1 h-1) are presented along with the comparable literature values in Table 2. The *YP/S for
glucose and PFAD dropped significantly from 0.017 and 0.019 g g-1 at 20 g L-1 to 0.0036 and
0.0043 g g-1 at 100 g L-1 for both initial substrates. This result also shows that the *YP/S values
for PFAD was 10-20% higher when compared to glucose for each initial concentration. This
indicates that the type and concentration of substrate used have an effect on rhamnolipid
production. Higher concentrations of both substrates were not effective in increasing the
rhamnolipid production. Different patterns were observed for YP/X which for glucose show
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that the YP/X value fluctuate from 0.20 g g-1 to 0.15 g g-1 and then back up to 0.17 g g-1.
Meanwhile for PFAD the YP/X was roughly constant at 0.15 g g-1 as the initial concentration
increased. The fluctuation of YP/X for glucose was due to the higher biomass produced at 50 g
L-1 of glucose compared to the other concentrations tested. On the other hand, an increase in
the initial PFAD concentration did not give any increase in rhamnolipid production. The PRL
were calculated based on the time taken to achieve maximum rhamnolipid concentration. PRL
using glucose was around 0.0036 to 0.0037 g L-1 h-1 with 20 and 50 g L-1 of initial glucose
concentration at 96 h, with a large increase in productivity to 0.0060 g L-1 h-1 when100 g L-1
of glucose at 60 h were initially present. This was due to maximum rhamnolipid production
occurring 36 h earlier compare with others glucose initial concentration. Meanwhile the PRL
for rhamnolipid production on PFAD increased slowly from 0.0040 to 0.0045 g L-1 h-1 at 96 h.
The *YP/S, YP/X and PRL values reported in this study were lower compared with other studies
with different types of fermentation, initial concentration, microbial strain and substrates as
shown in Table 2. When the variation of microbial strain is removed, it was noted that P.
aeruginosa PAO1 strain used in this study has higher *YP/S on PFAD compared to olive mill
waste (Moya Ramírez et al., 2016).
P. aeruginosa PAO1 is able to utilize both water-soluble carbon sources, i.e. glucose
and water-immiscible substrate, i.e. PFAD. There are reports that show biosurfactant
production from water immiscible substrates was generally higher compared with water-
soluble substrate by others researchers. For instance, a study using soybean oil by P.
aeruginosa LB1 gave 11.72 g L-1 of rhamnolipid, whereas to give a reduced biosurfactant
concentration of 4.20 g L-1 (Nitschke et al., 2005). This is because the water-immiscible
substrate has longer carbon chains compared to glucose that supplies carbon sources to the
cells (Singh et al., 2013). It is generally accepted that rhamnolipid production by
microorganisms, as well as biosurfactant production more broadly are natural process aimed
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at increasing the availability of water-immiscible substrates for the further metabolic process
through emulsification (Chrzanowski et al., 2012). However, P. aeruginosa EM1 produce
biosurfactant from glucose shows a higher production of biosurfactant about 7.50 g L-1
compare to olive oil and soybean oil about 3.70 and 2.63 g L-1 (Wu et al., 2008). The
variation of the result suggests that types of P. aeruginosa strain and substrate play an
important role in growth and concentration of biosurfactant production (Abdel-Mawgoud et
al., 2011).
3.3 Biosurfactant Identification
Structural characterization of the crude rhamnolipid material produced was carried out
using Electrospray Ionization Mass Spectrometry (ESI-MS). Rhamnolipid produced when P.
aeruginosa PAO1 was fed with glucose were both monorhamnolipid at 503 m/z and
dirhamnolipid at 649 m/z congeners. When PFAD was the carbon substrate, the
monorhamnolipid peak at 503 m/z was not observed and dirhamnolipid at 649 m/z appear as
the main congeners produced. Many other peaks were detected, most likely fatty acids
components from PFAD, in the range 0-300 m/z value (Ham and MaHam, 2015). Further
purification of crude rhamnolipid produced using PFAD by SPE to remove the free fatty acid
molecules confirms that P. aeruginosa PAO1 can produce monorhamnolipid and
dirhamnolipid from PFAD at 503 and 649 m/z congeners. This result was consistent with
other scholars that found monorhamnolipid at 503 m/z value and dirhamnolipid at 649 m/z
(George and Jayachandran, 2013). This resulting mixture gives different properties of
biosurfactant that have several potential environmental and industrial application such as
biological control agent, surface coating, production of chemicals and environmental
remediation. Specifically, Chrzanowski et al. (2012) have conducted a review regarding the
usefulness of rhamnolipids in various application areas. For example, crude rhamnolipid can
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be used as an anti-microbial agent, dirhamnolipid can act as insecticidal agent and
rhamnolipid can increase biodegradation efficiency in general and itself degrades after a
certain time. This demonstrates the utility of rhamnolipids whether crude, pure or a specific
molecule in many fields such as medical, agricultural and bioremediation.
3.4 Biosurfactant Characterization
There were no significant differences in term of reduction in surface tension when
testing crude rhamnolipid produced using the two different carbon sources. All the
rhamnolipid produce from different carbon sources has an ability to reduce the surface
tension to 28-33 mN/m. Figure 3 shows that the surface tension decreases linearly with
regards to rhamnolipid concentration to 120 mg L-1 for glucose and 420 mg L-1 for PFAD as
carbon sources. The greatest reduction of surface tension was obtained when testing the crude
rhamnolipid produced from PFAD, with a reduction in the surface tension to 29.88 mN m-1.
Thus the estimation of the CMC of rhamnolipid produced from glucose was estimated about
120 mg L-1 which was lower than CMC of rhamnolipid produced from PFAD that was about
420 mg L-1. This means that small amount of rhamnolipid concentration needed to reach
CMC by rhamnolipid produced from glucose compare with PFAD. This is most likely
because the crude rhamnolipid mixture contains components from PFAD, such as fatty acids
that associate with the rhamnolipid molecules during the extraction process thus affecting the
CMC and surface tension value.
Rhamnolipid production using different carbon sources can affect the value of
emulsion index with kerosene and sunflower oil. A white emulsion layer was observed for
crude rhamnolipid produced from both carbon sources used. Figure 4 shows that rhamnolipid
produce from glucose has a higher emulsion index of 44% compared to PFAD of 27% when
using 1000 mg L-1 of rhamnolipid. Meanwhile for emulsion with sunflower oil, crude
rhamnolipid produced from PFAD shows two times higher compared to glucose. This
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suggests that the different interactions between these crude rhamnolipid with kerosene and
sunflower oil, as well as with others components in the rhamnolipid extracted can reduce the
ability of rhamnolipid to emulsify (Sakalle and Rajkumar, 2010).
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4. Conclusions
The results show that P.aeruginosa PAO1 can produce rhamnolipid using PFAD as
the sole carbon source. Mass spectrometry results confirm that for each carbon source a
mixture of monorhamnolipid and dirhamnolipid were produced. The rhamnolipid produced
from PFAD were in the range 0.39-0.43 g L-1 and can lower surface tension to 29 mN m-1.
The CMC value was 420 mg L-1 and emulsification index with kerosene and sunflower oil is
20-30%. This finding is significant as it demonstrates the feasibility of producing
rhamnolipid using PFAD and the great potential of PFAD as a low-cost substrate for
rhamnolipid production.
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5. Acknowledgement
This study is financially supported by Ministry of Education, Malaysia and
Department of Biological and Agricultural Engineering, Universiti Putra Malaysia under
“Hadiah Skim Latihan IPTA (SLAI)”. We also like to thank Sime Darby Refinery-Jamolina
and The Cucurbit Company Sdn. Bhd. for providing the PFAD for this research.
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List of Tables
Table 1 Composition of free fatty acids from PFAD detected by GC-MS
Table 2 Comparison of rhamnolipid producing bioprocess metrics for a variety of
carbon sources and microbial strains.
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List of Figures
Figure 1 Time course profiles of P. aeruginosa PAO1 cell growth and rhamnolipid
production at 200 rpm and 37 °C by using (a) 20, (b) 50 and (c) 100 g L-1 of
glucose as carbon sources. (■) Optical density, (▲) Dry cell weight and (●)
Rhamnolipid production.
Figure 2 Time course profiles of P. aeruginosa PAO1 cell growth and rhamnolipid
production at 200 rpm and 37 °C by using (a) 20, (b) 50 and (c) 100 g L-1 of
PFAD as carbon sources. (■) Optical density, (▲) Dry cell weight and (●)
Rhamnolipid production.
Figure 3 Critical micelle concentration (CMC) of rhamnolipid produced from glucose
and PFAD. (■) Glucose and (●) PFAD.
Figure 4 Emulsion index of rhamnolipid with kerosene and sunflower oil
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Table 1. Composition of free fatty acids from PFAD detected by GC-MS
Name Formula Percentage (%)Oleic acid C18H34O2 41.86Pentadecanoic acid C15H30O2 17.98Tridecylic acid CH3(CH2)11COOH 13.98Palmitic acid C16H32O2 13.62Stearic acid C18H36O2 5.45Squalene C30H50 2.29Myristic acid C15H30O2 1.11Undecylic acid C11H22O2 1.11Eicosanoic acid C20H36O2 0.79Margaric acid C17H34O2 0.75Palmitoleic Acid C16H30O2 0.58Lauric Acid C12H24O2 0.48
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Table 2. Comparison of rhamnolipid producing bioprocess metrics for a variety of carbon sources and microbial strain.
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29
Type of
fermentationMicroorganism
Type of
substrate
Initial
concentration
( g L-1 )
RLmax
(g L-1)
YP/S (g
g-1)
*YP/S
(g g-1)
YP/X (g
g-1)
PRL
(g L-1 h-1)Reference
5L flaskP. aeruginosa
PAO1
Glucose
20 0.350 - 0.0173 0.2005 0.0036
This study
50 0.360 - 0.0071 0.1510 0.0037
100 0.360 - 0.0036 0.1705 0.0060
PFAD
20 0.390 - 0.0193 0.1539 0.0040
50 0.390 - 0.0078 0.1558 0.0040
100 0.430 - 0.0043 0.1563 0.0044
1L flaskP. aeruginosa
PAO1
Olive mill
waste
20 0.012 0.0130 0.0006 N/P N/P(Moya Ramírez et
al., 2016)50 0.030 0.0180 0.0006 N/P N/P
100 0.200 0.0580 0.0020 N/P N/P
1L flaskP. aeruginosa
PA1Glycerol 30 6.75 - 0.23 4.9 0.056
(Pacheco et al.,
2012)
0.5L flaskP. aeruginosa
LB1
Sunflower
refinery
waste
30 4 0.3 0.13 N/P 0.1(Benincasa and
Accorsini, 2008)
0.5L flaskP. aeruginosa
EBN-8
Blackstra
p
molasses
20 1.45 0.392 0.725 0.869 0.015 (Raza et al., 2007)
0.25L flaskP. aeruginosa
EM1
Soybean
Oil6 4.3 - 0.716 N/P N/P
(Rahman et al.,
2002)
547
*N/P - not provided in the reference
*YP/S (g g-1) is the product yield on initial substrate fed, calculated only for this study
30
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549
Figure 1. Time course profiles of P. aeruginosa PAO1 cell growth and rhamnolipid
production at 200 rpm and 37 °C by using (a) 20, (b) 50 and (c) 100 g L-1 of
glucose as carbon sources. (■) Optical density, (▲) Dry cell weight and (●)
Rhamnolipid production.
31
(a)
(b)
(c)
550
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570
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574
Figure 2 Time course profiles of P. aeruginosa PAO1 cell growth and rhamnolipid
production at 200 rpm and 37 °C by using (a) 20, (b) 50 and (c) 100 g L-1 of
PFAD as carbon sources. (■) Optical density, (▲) Dry cell weight and (●)
Rhamnolipid production.
32
(a)
(b)
(c)
575
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579
580
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583
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586
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598
599
Figure 3 Critical micelle concentration (CMC) of rhamnolipid produced from glucose
and PFAD. (■) Glucose and (●) PFAD.
33
CMCCMC
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
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616
617
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619
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623
Figure 4 Emulsion index of rhamnolipid with kerosene and sunflower oil using 1000 mg
L-1 of crude rhamnolipid concentration.
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