Menara Perkebunan 2025, 93.
, 148-157 p-ISSN: 0125-9318/ e-ISSN: 1858-3768 http://dx.
org/10.
22302/iribb.
Accreditation Number: 177/E/KPT/2024 Characterization of sn-1,3 extracellular lipases of Aspergillus niger and Rhizopus oryzae for the crude palm oil hydrolysis Ludwinardo PUTRA1.
Eneng NURHASANAH2, .
Eti ROHAETI2, & Irma KRESNAWATY1* .
Indonesia Oil Palm Research Institute Ae Bogor Unit.
Jl.
Taman Kencana No.
Bogor 16128.
Indonesia .
Department of Chemistry.
IPB University Oe Jl.
Tanjung Kampus.
Dramaga.
Bogor 16680.
Indonesia Received 25 Sep 2025/ Revised 11 Nov 2025/ Accepted 13 Nov 2025 Abstract Hydrolysis of triacylglycerol-rich oils is a crucial step in producing structured lipids such as sn-2 palmitate, which requires sn-1,3-specific lipases capable of selectively cleaving ester bonds at the outer positions of the glycerol backbone.
This study characterized extracellular sn-1,3 lipases from Aspergillus niger and Rhizopus oryzae cultivated in crude palm oil (CPO)enriched submerged fermentation (SmF) media.
The resulting culture filtrates exhibited lipase activities of 2.
76 and 2.
10 U mL-1, respectively.
Partial purification through ammonium sulfate fractionation .
Ae40%) yielded lipases with the highest specific activities of 4.
89 U mg-1 (A.
64 U mg-1 (R.
, with estimated molecular masses of 35 and 29 kDa.
Both enzymes showed sn-1,3 positional specificity toward CPO hydrolysis and exhibited optimal activity at pH 6Ae 7 and 35 AC.
Although thermal stability was moderate, both lipases maintained substantial activity in benzene, ethanol, and methanol, indicating tolerance to both non-polar and lowpolarity solvents.
These findings highlight the potential of indigenous fungal lipases as promising biocatalysts for CPO-based structured lipid synthesis under mild and sustainable conditions.
[Keywords: enzymatic hydrolysis, filamentous fungi, regioselectivity, sn-2 palmitate, vegetable oi.
Introduction Crude palm oil (CPO), extracted from the mesocarp of oil palm fruit, is a major source of vegetable oil rich in triacylglycerols (TAG.
composed of 39Ae44% palmitic acid, 39Ae40% oleic acid, 10Ae11% linoleic acid, about 5% stearic acid, and smaller proportions of lauric, myristic, palmitoleic, linolenic, and arachidic acids (Man et al.
, 1999.
Stavila et al.
, 2023.
Bomfima et al.
, 2.
Previous enzymatic study has demonstrated that CPO can be effectively modified through glycerolysis reactions using microbial lipases, particularly those from Rhizopus oryzae, to (MAG) diacylglycerol (DAG) with desirable structural characteristics (Tri-Panji et al.
, 2.
Extending from these findings, the rich palmitic acid content and balanced fatty acid composition of CPO highlight its potential as a cost-efficient and sustainable substrate for producing structured lipids such as sn-2 palmitate, a key component in human milk fat substitutes (HMFS).
Sn-2 palmitate refers to palmitic acid esterified at the sn-2 position of the TAG molecule, commonly found in 1,3dioleoyl-2-palmitoylglycerol (OPO) and 1-oleoyl2-palmitoyl-3-linoleoylglycerol (OPL) (Wei et al.
HMFS are structured lipids designed to mimic the fatty acid composition and positional distribution of human milk fat (Yuan et al.
, 2.
The enzymatic modification of CPO using sn-1,3specific lipases offer a promising biocatalytic route to generate these high-value lipids under mild and environmentally friendly conditions.
Lipase are enzymes capable of catalyzing the hydrolysis of TAGs into free fatty acids (FFA) and glycerol (Kyse et al.
, 2.
This biocatalyst acts on the carboxylate ester bond (Emmanuel et al.
Based on their positional regioselectivity toward the carboxylate ester bond of TAG, lipases can be classified into sn-1,3-specific and nonspecific lipases.
Sn-1,3-specific lipases can direct the cleavage of ester bonds at the sn-1 (R.
and sn3 (R.
positions (Figure .
, while non-specific lipases completely hydrolyze TAG at all three ester bond positions (Girelli & Chiappini,2.
The use of lipases with sn-1,3 regioselectivity can contribute to the processing of CPO as a source of sn-2 palmitate through hydrolysis.
*)Corresponding author: irmakresnawaty83@gmail.
0125-9318/ 1858-3768 A2025 Authors This is an open access article under the CC BY license .
ttps://creativecommons.
org/licenses/by/4.
Menara Perkebunan is DOAJ indexed Journal and accredited as Sinta 2 Journal .
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id/journals/profile/3.
How to Cite: Putra.
Nurhasanah.
Rohaeti.
, & Kresnawaty.
Characterization of sn-1,3, extracellular lipases of Aspergillus niger and Rhizopus oryzae for the crude palm oil hydrolysis.
Menara Perkebunan, 93.
, 148-157.
http://dx.
org/10.
22302/iribb.
Characterization of sn-1,3 extracellular lipases of A.
niger & R.
oryzae for the CPO hydrolysis .
(Putra et al.
sn-1,3 specific lipase H2O Materials and Methods Rn Figure 1.
Enzymatic hydrolysis of TAG with sn-1,3specific lipase Lipases are enzymes capable of catalyzing the hydrolysis of TAGs into free fatty acids (FFA) and glycerol (Kyse et al.
, 2.
This biocatalyst acts on the carboxylate ester bond (Emmanuel et al.
Based on their positional regioselectivity toward the carboxylate ester bond of TAG, lipases can be classified into sn-1,3-specific and nonspecific lipases.
Sn-1,3-specific lipases can direct the cleavage of ester bonds at the sn-1 (R.
and sn-3 (R.
positions (Figure .
, while non-specific lipases completely hydrolyze TAG at all three ester bond positions (Girelli & Chiappini,2.
The use of lipases with sn-1,3 regioselectivity can contribute to the processing of CPO as a source of sn-2 palmitate through hydrolysis.
Commercial production of sn-2 palmitate commonly employs sn-1,3-specific lipases such as Rhizomucor miehei (Lipozyme RM IM).
Thermomyces lanuginosus (Lipozyme TL IM), and Candida antarctica (Novozyme .
These immobilized enzymes are widely available from manufacturers such as Novozymes and SigmaAldrich, typically priced between USD 100-900 per kilograms (Souza-Gonyalves, 2.
While these imported biocatalysts dominate industrial use, their high cost and limited local supply highlight an opportunity for indigenous enzyme production in Indonesia.
Fermentation using native Indonesian filamentous fungi offers a low-cost and sustainable approach to lipase production (Tri-Panji et al.
Extracellular lipases secreted into the fermentation medium by fungi such as Aspergillus niger and Rhizopus oryzae have gained considerable attention for their biotechnological potential in oil and fat modification (Emmanuel et , 2020.
Vidal et al.
, 2.
These fungal lipases are advantageous because they function under mild reaction conditions, remain active in organic solvents and elevated temperatures, and operate across a broad pH range (Javed et al.
, 2.
Understanding their catalytic stability and physicochemical properties is therefore essential for evaluating their suitability in CPO hydrolysis toward sn-2 palmitate formation for human milk fat substitute (HMFS) development.
Therefore, this study aims to isolate and characterize the extracellular lipases from both fungi to evaluate their functional suitability for CPO hydrolysis in HMFS production.
Materials Aspergillus niger was obtained from the isolate collection of Indonesian Oil Palm Research Institute (IOPRI).
Bogor.
Rhizopus oryzae was isolated from tempeh purchased from a local market in Bogor.
All other materials and reagents were supplied by IOPRI.
Culture filtrate production Production was carried out in a jar containing 100 mL of media with a composition of 2 g CPO, 5 g peptone, 1.
5 g yeast extract, 0.
MgSO4.
7H2O, and an adjusted pH of 5 (Helal et al.
, 2.
The medium was sterilized in an autoclave at 121 AC for 20 minutes and cooled to room temperature.
Spores of A.
niger and oryzae were inoculated into the fermentation medium at a concentration of 1 y 10A spores mL-1 using 5 % .
inoculum and incubated at 25 AC under static conditions for 5 days.
The culture broth was filtered through Whatman No.
1 filter paper .
AAm pore siz.
to remove mycelial debris, and the resulting clear filtrate was used as the crude enzyme source.
Lipase activity assay Lipase activity was measured by the titrimetric method (Tri-Panji et al.
, 2.
The hydrolysis reaction mixture .
otal volume 45 mL) consisted of 1 mL of enzyme isolate, 3 g of CPO, 1 g of poly.
inyl alcoho.
(PVA), and 40 mL of 0.
phosphate buffer .
H .
The mixture was homogenized using a laboratory blender at 10,000 rpm for 3 minutes to obtain a fine emulsion and then gently stirred for 5 minutes to stabilize the dispersion before incubation at 37 AC for 30 minutes.
The reaction was stopped with 10 mL of an acetone : ethanol mixture to completely denature the lipase .
:1 v/.
1% of phenolphtalein was added into the mixture, then titrated with 1N NaOH until a persistent faint pink color A blank sample underwent the same procedure without the addition of the enzyme Lipase activity was calculated using the following equation:
Lipase activity = (A Oe B) y [NaOH] y 1000 v yt : Sample titration volume .
L) : Blank titration volume .
L) [NaOH] : NaOH concentration (N) : Incubation time .
: Total reaction volume .
L) Menara Perkebunan 2025, 93.
, 148-157 Protein concentration determination Protein concentration was determined using the method of Goldring .
using Bovine Serum Albumin (BSA) as a standard.
Through the mathematical equation of the protein standard curve, the total dissolved protein contained in the crude fraction and fractions I and II was obtained.
Partial purification of culture filtrate The crude enzyme fraction was partially purified through (NH.
2SO4 fractionation with two saturation fraction variations: fraction I .
-40%) and fraction II .
-80%).
The (NH.
2SO4 was added gradually at 4 AC using a magnetic stirrer.
Then, the fractions were centrifuged at 8000 y g at 4 AC for 30 minutes.
The precipitate was suspended 1 M phosphate buffer, pH 7, and dialyzed using a semipermeable membrane .
ut-off 10-14 kD.
against the same buffer overnight at 4AC (Barathi et al.
, 2.
The extracellular lipase fractions from A.
niger and R.
oryzae in this study consisted of three types: a crude fraction and two partially purified fractions (Fractions I and II).
All fractions were analyzed for total protein content and lipase activity to determine which had the highest specific activity.
Determination of lipase molecular mass profile The molecular masses of both lipases were measured using SDS-PAGE electrophoresis (Goldring, 2.
Non-induced Escherichia coli BL21(DE.
was used as a negative control for the non-lipase protein.
A 10% polyacrylamide gel was used as a porous membrane for protein migration .
kDa-250 kD.
The gel was stained with Coomassie Brilliant Blue R-250 and visualized under white light using BIO-RAD GeldocGo Imaging System.
Lipase specificity assay The positional specificity of lipase was assessed following the method of Tri-Panji et al.
The reaction mixture consisted of 3 g CPO and 0.
1 mL of crude lipase suspended 1 M phosphate buffer .
H .
and was incubated at 35 AC for 60 minutes.
The hydrolysis products were extracted using n-hexane, and the organic phase was analyzed on a Thin Layer Chromatography (TLC) Silica Gel 60 F254 plate developed in n-hexane : diethyl ether : acetic acid .
: 5 : .
Visualization was performed by exposing the developed plate to iodine vapor in a closed chamber until brown lipid spots appeared The Rf value of each spot was calculated as the ratio between the migration distance of the compound and the solvent front.
The apparent sn-1,3 activity was inferred from the appearance of 2-palmitoylglycerol (MAG standar.
together with the glyceryl tripalmitate (TAG standar.
The optimum pH and temperature, as well as the stability of each enzyme against pH, temperature, and organic solvents, were evaluated using the lipase activity assay described above.
Lipase characterization The optimum pH and temperature, as well as the stability of each enzyme towards pH, temperature, and organic solvents were evaluated using the lipase activity assay described above.
Each treatment was performed in triplicate, and the results are presented as the mean with error bars representing standard deviation.
The optimum pH was evaluated using 0.
05 M phosphate buffer at pH levels of 4, 5, 6, 7, and 8, with incubation at 35 AC for 30 minutes.
The effect of temperature was evaluated at 25, 35, 45, 55, and 65 AC under the previously determined optimal pH condition for 30 minutes.
Lipase activity at both optimum pH and temperature was expressed as relative activity, calculated by comparing the activity under each test condition to that of the enzyme under optimal assay conditions, which was defined as 100%.
The pH stability was evaluated by preincubating the enzyme at pH 4, 5, 6, 7, and 8 in 05 M phosphate buffer at 35 AC for 60 minutes.
Temperature stability was evaluated by preincubating the enzyme at 25, 35, 45, 55 and 65 AC under the previously determined optimum pH stability for 60 minutes.
After incubation, each sample was subjected to the lipase assay at the determined optimum pH and temperature.
Lipase activity of both temperature and pH stability was expressed as residual activity, defined as the percentage of enzymatic activity retained after preincubation compared to the initial activity measured under optimal assay conditions.
The effect of organic solvents was evaluated by incubating the enzyme at room 25AC for 60 minutes in the presence of the 10% of the following solvents: acetone, ethanol, methanol, isopropanol, and benzene.
As a control, the enzyme was also incubated in 0.
05 M phosphate Lipase activity of organic solvent stability was expressed as residual activity defined as the percentage of lipase activity of each sample compared to the control measured under optimal assay conditions.
Results and Discussion Extracellular lipase activity of culture filtrate The culture filtrates of A.
niger and R.
in submerged fermentation (SmF) showed lipase activities of 2.
76 and 2.
10 U mL-1, respectively (Table .
Several studies have previously reported the enzyme activity of both isolates, ranging from 6 to 63 U mL-1 (Mukhtar et al.
, 2015.
Hermansyah et al.
, 2019.
Yassein et al.
, 2.
This suggests potential improvements to the current method, including optimizing culture conditions and nutrient availability.
Innoculation of A.
Characterization of sn-1,3 extracellular lipases of A.
niger & R.
oryzae for the CPO hydrolysis .
(Putra et al.
Table 1.
Activity and total protein of the extracellular lipase in ammonium sulfate fraction Lipase activity (U mL-.
Total protein g mL-.
Specific activities (U mg-.
Recovery of enzyme activity (%) Purification fold Crude fraction Fraction I .
-40%) Fraction II
-40%)
Crude fraction Fraction I .
-40%) Fraction II
-40%)
Test sample Aspergillus niger Rhizopus oryzae and R.
oryzae spores into SmF medium aimed to establish a fungal metabolic system that secretes Several bioprocess components are utilized by fungal species for lipase secretion, including CPO as an inducer and carbon source, peptone and yeast extract as nitrogen sources, and MgSO4.
7H2O as a source of metal ion compounds (Al-Dahlan et al.
, 2.
Both fungal species are responsive to the environmental conditions of the media containing these components, enabling them to secrete lipase into the culture filtrate.
Extracellular lipase fraction The maximum specific activity was obtained in Fraction I .
Ae40% ammonium sulfate saturatio.
, 89 U mg-1 for A.
niger and 9.
64 U mg-1 for R.
The corresponding enzyme activity recoveries were 71.
50% and 67.
57%, with purification folds of 2.
19 and 3.
99, respectively (Table .
These results indicate that the initial fraction retained most of the lipase activity during the partial purification process.
Similar findings were reported by Putra et al.
, who observed that recombinant lipase fractions with low salt saturation .
Ae30%) exhibited higher specific activity than advanced fractions.
Likewise.
Fathi et al.
found that initial low-salt fractions from Lactobacillus fermentum retained superior lipase activity compared to later fractions.
Based on these results.
Fraction I was selected as the enzyme source for subsequent characterization.
The precipitate of fractions I and II is formed due to the salting-out process during fractionation, by changing the solubility of the protein (Nargotra et al.
, 2.
(NH.
2SO4 can increase the saturation of the fraction, so that components with low solubility will be precipitated (Du et al.
, 2.
Protein precipitates of fractions I and II from niger and R.
oryzae were suspended in 1 M phosphate buffer, pH 7, and dialyzed in a semipermeable membrane.
Small molecules in the sample move through the membrane into the dialysis buffer, while large macromolecules .
ncluding lipas.
are retained within the membrane (Thomas et al.
, 2.
Lipase molecular mass profile SDS-PAGE electrophoresis of 10% protein fraction I of A.
niger and R.
oryzae produced bands with estimated molecular masses of 35 and 29 kDa, respectively (Figure .
The determination of the molecular masses of lipase from these two species is relatively consistent with previous studies, with each of A.
niger and R.
oryzae being found to be 5 - 35 kDa and 29-31 kDa, respectively (Chattopadhyaya, 2000.
Lypez-Fernyndez et al.
Xing et al.
, 2020.
Namboodiri & Vidal et al.
Lipase specificity The hydrolysis products of crude palm oil (CPO) by lipases from A.
niger and R.
produced three visible spots on TLC (Figure .
Two of these spots matched those of identified as glyceryl tripalmitate and 2-palmitoylglycerol standards (Figure .
, which suggests that the lipases preferentially hydrolyze ester bonds at the sn-1 and sn-3 positions.
However, this interpretation should be considered indicative rather than conclusive, as TLC alone cannot distinguish between positional isomers.
Additional analyses, such as co-chromatography with authentic 1-MAG standards or instrumental techniques like gas chromatography (GC) or highperformance liquid chromatography (HPLC), would be required to confirm the regioisomeric Menara Perkebunan 2025, 93.
, 148-157 Effect of pH and temperature The A.
niger lipase was optimal at pH 6 with a relative activity of 92.
59 A 6.
42%, while the oryzae lipase was optimal at pH 7 with a relative activity of 88.
89 A 11.
11 % (Figure 4A).
Another sn-1,3 lipase.
Rhizomucor miehei, was also optimal at pH 7 (Tako et al.
, 2.
At optimum conditions, the enzyme conformation is in a highly electrostatically stable form, so the binding of substrate-enzyme is effective (Yao et al.
, 2.
Exposure of the enzyme to unfavorable pH causes changes in the ionization state of the enzyme's amino acid groups, leading to changes in the characteristic electrostatic interaction patterns in the enzyme structure (Nargotra et al.
, 2.
The lipase from fraction I of both fungal species showed optimal activity at 35 AC (Figure 5B).
The sn-1,3 extracellular lipase from niger in the study by Xing et al.
also had an optimum temperature of 35 AC.
Similarly, the non-specific lipase from R.
oryzae in the study by Helal et al.
showed an optimum temperature of 37 AC.
Lipase activity at the optimum temperature indicates that the affinity of substrate for lipase is at optimum conditions for hydrolysis (Remonatto et al.
, 2.
The lowest activity of both fungal species was achieved when the hydrolysis reaction temperature exceeded 45 AC.
However, it should be noted that enzymes are proteins subjected to protein denaturation at high temperatures, preventing them from Figure 2.
maintaining their structure (Alabdalall et al.
, 2.
pH and temperature Stability The stability range of A.
niger lipase was broader .
H 5Ae.
than that of R.
oryzae lipase .
H 6Ae.
(Figure .
However, the overall stability of both free lipases was lower compared to the Nuraliyah et al.
and Yang et al.
This result indicates that free lipases lose more activity after exposure to extreme pH and temperature for 60 minutes, compared to immobilized ones.
According to Kumar et al.
, enzyme immobilization enhances catalytic stability, and further improvements can also be achieved through lipase engineering (Ali et al.
, 2.
Effects of organic solvents Both A.
niger and R.
oryzae lipases followed a similar stability trend, retaining the highest residual activity in benzene, followed by methanol, ethanol, and isopropanol, while losing most activity in acetone (Figure .
This pattern suggests a preference for non-polar environments that preserve the enzyme hydrophobic core, as found in solvents like benzene.
Although industrial studies often preferred n-hexane and toluene as non-polar media, benzeneAos comparable polarity helps indicate the enzyme compatibility within such hydrophobic systems.
SDS-PAGE profile .
% resolving ge.
of crude lipase fractions visualized using Coomassie Brilliant Blue R250 staining.
L: pre-stained Bio-Rad Precision Plus ProteinE Dual Color Standards .
Da indicate.
NTC:
negative control .
on-induced E.
coli BL21(DE.
AN: Aspergillus niger lipase fraction.
RO: Rhizopus oryzae lipase fraction.
Light blue boxes indicate the putative lipase protein bands.
Characterization of sn-1,3 extracellular lipases of A.
niger & R.
oryzae for the CPO hydrolysis .
(Putra et al.
MAG
TAG
Spot Rf TAG*) Spot Rf MAG**) .
*)Retention factor triacylglycerol, **)retention factor 2-monoacylglycerol Figure 3.
TLC in the eluent system n-hexane: diethyl ether: acetic acid .
:5:.
Glyceryl tripalmitate standard, .
2-palmytoylglycerol standard, .
CPO hydrolysis product by lipase fractions I A.
niger, .
CPO hydrolysis product by lipase fractions I R.
The reaction was carried out at pH 7 and a temperature of 35 EE Relative activity (%) Relative activity (%) Aspergillus niger Temperature (EE) Aspergillus niger Rhizopus oryzae Rhizopus oryzae Figure 4.
Characteristics of optimal conditions for lipase in CPO hydrolysis.
(A) At pH 4-8 and 35 EE, (B) at temperatures 25-65 EE and optimal pH Residual activity (%) Residual activity (%) Rhizopus oryzae Temperature (EE) Aspergillus niger Aspergillus niger Rhizopus oryzae Figure 5.
Characteristics of stability conditions for 60-minute lipase in CPO hydrolysis.
(A) At pH 4-8 and 35 EE, (B) at temperatures 25-65 EE and optimal pH Menara Perkebunan 2025, 93.
, 148-157 Aspergillus niger Rhizopus oryzae Residual activity (%) Control Aceton Ethanol Methanol Isopropanol Benzene Organic solvents Figure 6.
Effect of organic solvents on lipase stability for 60 minutes Both lipases also showed good activity in waterAemethanol and waterAeethanol systems, confirming their tolerance toward low-polarity Similar findings were reported by Ayinla et al.
and Xing et al.
, who demonstrated strong methanol compatibility in extracellular lipases.
According to Tian et al.
, enhancing methanol tolerance and catalytic activity in water-soluble lipases is critical for improving biodiesel conversion efficiency.
High lipase activity at the waterAepolar solvent interface is attributed to interfacial activation (Albayati et al.
, 2.
, though a minor decline in hydrolytic activity was noted.
The lower FFA levels detected by NaOH titration likely resulted from a shift in reaction equilibrium toward ester formation through esterification (Dasetty et al.
Despite moderate stability in polar alcohols .
ethanol, ethanol, isopropano.
, the sharp decline in acetone activity indicates that moderately polar solvents are the most disruptiveAibeing polar enough to disturb enzymeAewater interactions but not hydrophobic enough to stabilize the enzymeAos structure (Wang et al.
, 2.
Conclusion The submerged fermentation of Aspergillus niger and Rhizopus oryzae in CPO-enriched media successfully produced extracellular lipases with specific activities of 4.
89 and 9.
64 U mg-1.
The highest enzymatic performance was obtained from the 0Ae40% ammonium sulfate fraction, corresponding to molecular masses of approximately 35 kDa (A.
and 29 kDa (R.
Both enzymes exhibited sn-1,3 positional specificity during CPO hydrolysis, indicating potential application in structured lipid synthesis, although further analysis is needed to confirm regioisomer distribution.
The optimal catalytic conditions were pH 6Ae7 and 35 AC, with notable stability in benzene, ethanol, and methanol, suggesting good compatibility in both non-polar and low-polarity environments.
Acknowledgements We thank the Indonesian Oil Palm Research Institute - Bogor Unit Ae PT Research Perkebunan Nusantara for the research funding.
We also thank
the Bogor Agricultural Institute for supporting this References