Accepted Manuscript

Title: Identification of Lipolytic Enzymes Isolated from

Bacteria Indigenous to Eucalyptus Wood Species for
Application in the Pulping Industry

Authors: L. Ramnath, B. Sithole, R. Govinden

PII: S2215-017X(17)30110-8
DOI: http://dx.doi.org/doi:10.1016/j.btre.2017.07.004
Reference: BTRE 212

To appear in:

Received date: 24-4-2017

Revised date: 21-6-2017
Accepted date: 11-7-2017

Please cite this article as: L.Ramnath, B.Sithole, R.Govinden, Identification

of Lipolytic Enzymes Isolated from Bacteria Indigenous to Eucalyptus
Wood Species for Application in the Pulping Industry, Biotechnology
Reportshttp://dx.doi.org/10.1016/j.btre.2017.07.004

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Identification of Lipolytic Enzymes Isolated from Bacteria Indigenous to
Eucalyptus Wood Species for Application in the Pulping Industry

Short title: Substrate specificity of lipolytic enzymes isolated from bacteria in Eucalyptus

wood

*L. Ramnath1, B. Sithole2, 3, and R. Govinden1

1
Discipline of Microbiology, School of Life Sciences, College of Agriculture, Engineering
and Science, University of KwaZulu-Natal, Westville Campus, P/Bag X54001, Durban 4000,
South Africa.
2
Forestry and Forest Products Research Centre, Council for Scientific and Industrial
Research, Durban 4000, South Africa
3
Discipline of Chemical Engineering, University of KwaZulu-Natal, Durban 4000, South
Africa.

*Corresponding author. E-mail: lucretiaramnath@gmail.com. Tel: +2731 260 7401. Fax:

+2731 260 7809.

Abstract

This study highlights the importance determining substrate specificity at variable

experimental conditions. Lipases and esterases were isolated from microorganisms cultivated

from Eucalyptus wood species and then concentrated (cellulases removed) and characterized.

Phenol red agar plates supplemented with 1% olive oil or tributyrin was ascertained to be the

most favourable method of screening for lipolytic activity. Lipolytic activity of the various

enzymes were highest at 45-61 U/ml at the optimum temperature and pH of between at 30-

35°C and pH 4-5, respectively. Change in pH influenced the substrate specificity of the

enzymes tested. The majority of enzymes tested displayed a propensity for longer aliphatic

acyl chains such as dodecanoate (C12), myristate (C14), palmitate (C16) and stearate (C18)

indicating that they could be characterised as potential lipases. Prospective esterases were

also detected with specificity towards acetate (C2), butyrate (C4) and valerate (C5). Enzymes

1
maintained up to 95% activity at the optimal pH and temperature for 2-3 hours. It is essential

to test substrates at various pH and temperature when determining optimum activity of

lipolytic enzymes, a method rarely employed. The stability of the enzymes at acidic pH and

moderate temperatures makes them excellent candidates for application in the treatment of

pitch during acid bi-sulphite pulping, which would greatly benefit the pulp and paper

industry.

Keywords: Lipase, esterase, substrate specificity, pitch, pulp, paper

Introduction

Lipase and esterase are two major classes of hydrolase enzymes [1]. Lipases (triacylglycerol

acylhydrolases, EC 3.1.1.3) catalyse the hydrolysis of long chain triacylglycerol substrates

(>C8), whereas esterases (EC 3.1.1.x) catalyse the hydrolysis of glycerolesters with short acyl

chains (<C8) [2]. The three-dimensional (3D) structures of both enzymes exhibit the

characteristic α/β-hydrolase fold [3] a definite order of α-helices and β-sheets. The catalytic

triad is comprised of Ser-Asp-His (Glu instead of Asp for some lipases) and typically also a

consensus sequence (Gly-x-Ser-x-Gly) is found around the active site serine [4]. These

lipolytic enzymes have been isolated from plants, animals, and microorganisms [1,5],

however, microbial lipolytic enzymes are reported to be more robust in nature than plant or

animal enzymes [6,7]. They are also appealing due to their low cost of production and they

are simple to manipulate [1]. Some microbial species reported to produce these enzymes

include Bacillus sp., Pseudomonas sp., Burkholderia sp., Candida rugose, Candida

Antarctica, Galactomyces geotricum, Saccharomyces cerevisiae, Yarrowia lipolytica,

Trichosporon fermantans, Cryptococcus albidus, Aspergillus flavus, Thermomyces

lanuginosus and Rhizopus oryzae [8-13]. Due to the versatility of lipases and esterases, they

2
have various applications in industries such as detergents, starch and fuels, food, baking, pulp

and paper, fats and oils, organic synthesis, leather and environmental application [14,15].

In the pulp and paper industry, the presence of wood extractives plays a vital role. During

pulping, pitch particles (composed of extractives such as triglycerides, fatty acid esters,

glycosides, free and conjugated sterols) [16] tend to coalesce to form black pitch deposits in

the pulp and on machinery which has a negative impact on the process and quality of pulp

[17,18]. Sulphite pulps (acidic) in particular retain greater amounts of extractives in relation

to kraft pulps (alkaline), as the alkaline method disbands and dissolves the wood resin [19].

The production of dissolving pulp, which is a high grade cellulose pulp, is generated using

the acid bi-sulphite method.

Traditional methods for the control of pitch include seasoning and the addition of

chemicals [20]. The biotechnological approach of using enzymes for pitch control is an

alternative choice, especially for removal of glycerides. The treatment of pulp with lipases

has been effective in reducing triglycerides (TG), however, steryl esters (SE) are frequently at

the source of pitch formation [17]. Nonylphenol ethoxylates (NPEs) are the best chemicals

for removing pitch components in chemical pulping. Unfortunately, their use is frowned

upon due to their estrogen mimicking effects. Indeed, their use has been banned in North

American and European chemical pulp mills as pulp handlers in European markets are

reluctant to handle pulps treated with NPEs [21,22]. Also, the residual NPE in sulphite pulps

are undesirable since the pulps are commonly used in pharmaceutical and food applications.

Based on a mill study conducted by Sithole et al. (2010) it was suggested that the inclusion of

an enzyme to target residual steryl esters could deliver a strategic solution to removing the

extractives present in sulphite pulps [21].

3
Oxidative enzymes such as laccases have also been implemented in the degradation of

various lipophilic extractives such as triglycerides, free and conjugated sterols, fatty acids and

resin acids [23]. Laccases are typical for white-rot fungi and have been described as prime

lignin degraders. Treatment of wood or pulp with these enzymes could offer a dual advantage

in the company of redox mediators [24,25]. Redox mediators facilitate laccase removal of

residual lignin, in conjunction with extensive degradation of extractives [26]. A decrease in

kappa number and improved pulp brightness can also observed [19,26].

The enzymes characterized in this study are for application in the pulp and paper industry, for

reduction or elimination of pitch deposit formation during pulping. Previous studies have

reported the incomplete degradation of pitch by lipases [19,21], hence we are confident that

the inclusion of esterases will assist in targeting the side groups that are theoretically present

once the longer chain acyl chains (triacylglycerides) have been degraded by lipases. Lipases,

esterases and laccases were included as part of this study and were selected based on their

stability and activity at temperatures and pH levels employed during the acid bi-sulphite

pulping of Eucalyptus wood species. To our knowledge, the lipolytic enzymes produced by

microorganisms indigenous to Eucalyptus sp. wood have not been previously investigated.

The results of the present study will provide more information on the characteristics of these

enzymes and their potential for reduction of pitch components in pulps. For this study it was

important to include different types of enzymes that could benefit the pulping process.

Therefore purifications of the enzymes of interest were not necessary, as a cocktail of

enzymes (excluding cellulases) is required and ideal in this study for the removal or

degradation of all unwanted compounds (excluding cellulose). Combinations of

hemicellulases, ligninases and other accessory enzymes are known to be essential for

hydrolysis of plant biomass [27]. It was also important to test the effects of various conditions

4
on substrate specificity as most researchers focus only on the pH and temperature optima of

the enzyme and thereafter test substrate specificity at optimum conditions. Neglecting to

investigate the effects of pH and temperature on substrate specificity of enzymes could have

drastic implications for its efficiency and effectiveness. Therefore, the aim of this study was

to screen indigenous microflora from Eucalyptus species for lipolytic activity and to

determine the effects of pH and temperature on the hydrolysis of different substrates of these

lipolytic enzymes (lipases, esterases and laccases).

Materials and Methods

Isolation and Identification of Bacterial and Fungal Cultures

Five grams of wood chips from a commercial wood chip pile and individual Eucalyptus spp.

were thoroughly washed by vortexing with 5 ml of phosphate buffer (pH 8.0) for 5 min. The

washings were serially diluted and spread onto nutrient agar (NA) and potato dextrose agar

(PDA) (Merck, South Africa) and incubated at 37°C and 40°C for 1 and 5 days, for the

growth of bacteria and fungi, respectively. Colonies were selected based on morphological

features; size, shape, pigmentation, margin, consistency and elevation and sub-cultured till

pure isolates were obtained [28]. DNA was extracted from isolates and 16S rRNA and 18S

rRNA for bacteria and fungi, respectively, were amplified according to Ramnath et al. (2014)

[28]. Following PCR, the amplicons were sequenced (Inqaba Biotech, South Africa), and the

sequences edited and entered in the Basic Alignment Search Tool (BLAST) algorithm [29]

for identification of microorganisms.

Optimization of Plate Screening Assays for Lipolytic Activity

5
There are a number of methods currently available for the screening of lipases and esterases.

However, they vary with sensitivity, cost and ease of preparation. In this study a few methods

were tested and evaluated.

All strains were pre-cultivated in Luria-Bertani (LB) medium and malt extract broth for

bacteria and fungi, respectively. For detection of esterase activity a basal medium containing

0.5% (w/v) peptone, 0.3% (w/v) yeast extract and 2% bacteriological agar (pH 7)

supplemented with 1%, 2% and 5% tributyrin was used. Five millimetre wells were bored

into the agar plates and inoculated with 50 µl of pure bacterial cultures. Plates were incubated

at 37°C for 48 hours. After incubation the isolates were observed for zones of hydrolysis

(clear halos) around the colonies. Lipase activity was screened for on olive oil/rhodamine B

agar plates. Rhodamine B (1 mg/ml; Sigma Chemical Co., Munich, Germany) was dissolved

in distilled water and filter-sterilized. The agar plates contained 8 g nutrient broth, 4 g sodium

chloride, 10 g agar (per litre) (pH 7). After autoclaving the medium was cooled to 60°C,

31.25 ml olive oil and 10 ml of Rhodamine B solution (0.001% [wt/vol]) was added and

stirred vigorously for 1 min. The medium was allowed to stand for 10 min to reduce foaming

before pouring into sterile petri dishes. Lipase production was detected by irradiating plates

with UV light at 350 nm [30]. Due to difficulty encountered with reading the screening plates

using the above mentioned methods, two additional screening methods were tested, viz.,

assay with phenol red and tween agar plate screenings. Phenol red olive oil/tributyrin agar

plates were prepared as follows (g/L); 0.01% (w/v) phenol red, 0.1% (w/v) CaCl2, 1% (v/v)

substrate, 2% (w/v) agar and pH adjusted to 7.3-7.4 with 0.1 N NaOH [31]. Organisms were

inoculated onto the phenol red agar plates supplemented with 1% substrate and incubated at

37°C for 2-4 days. The principle behind this assay is that a slight drop in pH from 7.3 (end

point of the phenol red dye) to a more acidic pH will result in a change of colour from red to

6
orange. The increase in acidity is due to the release of fatty acids following lipolysis [31]. A

precipitation test using Tween 20 and Tween 80 agar plates was carried out to confirm

lipolytic activity. Tween substrate plates were prepared as follows (g/L); 10 g peptone, 5 g

NaCl2, 0.1 g CaCl2.2H2O, 20 g agar and 10 ml (v/v) Tween 20/80 [32]. This method is based

on the principle of calcium salt precipitation. The hydrolysis of tween releases fatty acids

which bind with the calcium in the medium to form insoluble crystals around the point of

inoculation. Tween 80 is used for the detection of lipases as it contains esters of oleic acid,

whilst Tween 20 is used for esterases as it contains esters of lower chain fatty acids [32]. The

organisms were inoculated onto the plates and incubated at 37ºC for 2-4 days. A white

precipitation around the boundary of the colony was indicative of lipase activity [31].

Fungal isolates were screened for laccase activity on PDA plates supplemented with and

0.2% bromophenol blue [33] (Merck, South Africa). Plates were incubated at 40°C for 5

days, and then visually examined to evaluate the decolourizing ability of the fungal enzymes.

To establish cellulase activity, substrates specific for the detection of exoglucanase (1% (w/v)

avicel) and endoglucanase (1% (w/v) carboxymethyl cellulose (CMC)) were used to screen

isolates on NA and PDA agar plates, for bacteria and fungi, respectively. All screening assays

were performed in duplicate.

Enzyme Assays

Lipolytic activity was determined spectrophotometrically by measuring the release of p-

nitrophenol. P-nitrophenyl (p-NP) esters with various lengths of aliphatic acyl chains were

used to determine esterase; p-NP acetate (C2), p-NP butyrate (C4), p-NP valerate (C5) and

lipase; p-NP octanoate (C8), p-NP dodecanoate (C12), p-NP myristate (C14), p-NP palmitate

(C16), and p-NP stearate (C18) activity. The substrate mixture consisted of 0.5 mM p-NP

7
substrate in methanol, 50 mM Tris-HCl buffer (pH 8) and 0.1% Triton X-100. The standard

assay mixture contained 200 μl of substrate mixture and 20 μl of the crude supernatants,

which were incubated at 37°C for 1 hour. The enzyme activity was determined by measuring

the release of p-NP at an absorbance of 405 nm. One unit (U) of enzyme activity was defined

as the amount of enzyme required to release 1 nM of p-NP per min under the assay

conditions. Lipase/esterase and laccase activity was calculated from the formula derived from

the Beer-Lambert Law: enzyme activity (U.ml-1) = ΔA.V/ε.t.v. ΔA is the change in

absorbance over time; V is the total volume of reaction mixture (ml); ε is the molar extinction

coefficient in nM-1.cm-1; t is the incubation time in minutes, and v is the volume of the

enzyme in the assay mixture (ml) [34]. The appropriate extinction coefficient for each

substrate under these assay conditions was used to calculate activity [35].

Laccase activity was determined based on the oxidation of syringaldazine substrate

according to a protocol from Sigma-Aldrich (USA) [36]. The assay mixture (1 ml) contained

733 µL of acetate buffer (100 mM, pH 4/5) and 167 µL of laccase enzyme extract. The

reaction vessels were equilibrated to 37°C and absorbance monitored at 530 nm until

constant. Thereafter 100 µL of 0.216 mM syringaldazine were added to the assay (to begin

the reaction), followed by immediate mixing by inversion. The assays were incubated for 10

min and the increased absorbance was recorded using a UV-1800 Shimadzu UV

Spectrophotometer (Japan). Production of the corresponding quinone was monitored at 530

nm (ε530 = 65 000 M-1.cm-1). One enzyme unit is defined as the amount of enzyme that

will oxidise 1 µmol of syringaldazine per min, under the assay conditions [37].

The dinitrosalicylic acid (DNS) assay was used to determine cellulase activity by

detecting reducing sugars which are liberated by the hydrolytic action of endo and exo-

glucanase on different cellulose substrates (avicel and carboxymethylcellulose) [38].

8
 Effects of Temperature and pH on Lipase/Esterase Activity and Stability

The effect of temperature on enzyme activity was determined by conducting assays at

incubation temperatures ranging from 25–50ºC (with 5°C increments) and various p-NP

esters as substrates [39]. Temperature stability of purified enzyme was determined by

incubating the enzyme at various temperatures (25–50ºC) and estimating residual enzyme

activities after incubation for 30 min, 1, 1.5, 2, 2.5, and 3 hours. The effect of pH on enzyme

activity was determined by assaying enzyme activity over a pH range of 3-12 using p-NP

esters as substrates [39]. Citrate–phosphate buffer (pH 3 to 6), Tris–HCl buffer (pH 7 and 8),

Carbonate–bicarbonate buffer (pH 9 and 10) and sodium-bicarbonate and sodium-phosphite

buffer (pH 11 and 12) were used as buffer systems. Stability of the purified enzyme over a

range of pH was also determined by measuring the residual activity after incubating 200 μl of

the enzyme in 1800 μl of the above mentioned buffer systems (pH 3–12) for 3 hours at the

optimum temperature. Absorbance was read at 405 nm.

Production of Crude Enzyme Extracts

The selected bacterial isolates were grown in basal medium containing 0.5% (w/v) peptone

and 0.3% (w/v) yeast extract supplemented with 1% tributyrin. Flasks were incubated at 37°C

for 24 hours at 180 rpm. Cells were harvested by centrifugation at 10 000 rpm for 10 min.

Cell pellet was then resuspended in lysis buffer (20 mM Tris-HCl, 0.5 M NaCl, pH 8.0) and

disrupted by ultrasonic treatment for 10 min in 10 second intervals. The cell lysate was

centrifuged at 10 000 rpm for 10 min at 4 °C, and the supernatant was recovered to test

intracellular activity. To test extracellular activity the cell free supernatant was collected and

concentrated 10-fold by ultrafiltration with an Amicon system (Millipore, Massachusetts,

USA) using first a 3 kDa cut-off membrane after which a 50 kDa cut-off membrane was used

on the concentrated sample to remove proteins larger than 50 kDa.

9
 Native & SDS-PAGE

Protein sizes were determined by Sodium Dodecyl Sulphate Polyacrylamide Gel

Electrophoresis (SDS-PAGE) as outlined by Judd (1996) [40]. Samples were electrophoresed

by Native-PAGE (no SDS included) and SDS-PAGE in 12 % polyacrylamide gels according

to the method of Laemmli (1970) [41]. Protein concentration was determined using the

Bradford assay (Bradford, 1976) [42].

Native SDS-PAGE was utilized to ensure removal of potential cellulases. To identify

endo and exo-glucanases, 12% native-PAGE gels containing 1% avicel and

carboxymethylcellulose, respectively, (prepared in 50 mM phosphate buffer pH 7) were

prepared. Following electrophoresis at 100 V for approximately 90 min at room temperature,

the gel slab was cut in two halves; one half was stained using 0.5% Coomassie Brilliant Blue

R250 (Sigma-Aldrich, Germany) to determine the size of the proteins and the other portion

was used to detect enzyme activity. The gel for activity staining was washed with 50 mM

phosphate buffer (pH 7) for 5 min, followed by staining in Congo-Red solution (0.1%, [w/v])

for 15 min. The gel was then destained with 1 M NaCl to visualise the clearing zone of

hydrolysis, and then fixed with 0.5% (v/v) acetic acid [43].

Statistical analysis

Results shown here are the means of three independent determinations. Standard deviations

for each of the experimental results were calculated using Microsoft Excel software and

represented as error bars.

Results and Discussions

Identification of isolated bacteria and fungi

10
A total of ten different bacterial strains were isolated using the traditional culture and

identification method using 16S rRNA sequencing: three Bacillus spp., three different

Pseudomonas aeruginosa isolates, Inquilinus sp., Micrococcus sp., Pantoea sp., Klebsiella,

Streptomyces sp. and Cellulosimicrobium sp. (Table 2) (all with a similarity index of more

than 97%). Bacillus spp. were the predominant bacterial species (33%). Some of these genera

such as Bacillus, Pantoea, Klebsiella and Pseudomonas have previously been identified in

other woods [44,45], whilst others such as Inquilinus and Mucilaginibacter have not observed

in woods. The two fungal isolates described in this study were identified as Paecilomyces

formosus (F4) and Phialophora alba (X) using 18S rRNA sequencing. Both these fungal

isolates have not been previously identified in Eucalyptus spp. woods.

Optimization of plate screening assays for lipase and esterase activity

One percent tributyrin (esterase activity) was optimal for bacteria isolated from the mixed

wood sample (Table 2), however, 2% was optimal for bacteria from individual wood species

(Table 3). Slight halos were observed for a few of the bacterial isolates in 5% tributyrin

plates. Plate screening assays for lipase activity revealed minimum lipase activity for isolates

from mixed wood species; however, for bacteria isolated from individual Eucalyptus species,

1% substrate concentration was optimal. Sixty-seven percent, 28% and 28% of the isolates

displayed activity on 1%, 2% and 5% tributyrin plates, respectively. Bacillus firmus was

capable of hydrolysing all three concentrations of tributyrin, but largest halos were observed

at 1% substrate concentration. Micrococcus luteus, Pseudomonas aeruginosa, and

Cellulosimicrobium cellulans were also identified as esterase producers. Eight percent, 63%

and 22% of the isolates displayed activity on 1%, 2% and 5% tributyrin plates, respectively.

Curtobacterium flaccumfaciens, Bacillus thuringiensis, B. cereus, Pantoea agglomerans and

P. vagans produced the greatest zones of hydrolysis indicating esterase activity, with a halo

11
zone of 2-5 mm (Figure 2). Other studies have also had some degree of success with the use

of tributyrin and olive oil/rhodamine B as substrates and methods for screening for lipolytic

activity [32,46,47].

Due to difficulty encountered with visualization and of the clearing zones, additional assays

such as phenol red and tween agar plate screenings were also performed to validate the

results obtained. Both assays confirmed the results, however, the phenol red agar plate assay

was more sensitive than the other assays. Distinct clearings for the phenol red plates and

precipitation zones for the tween plates were observed (Figure 3). The phenol red screening

plates were used to quantify activity (Table 2 and 3).

Lipases and esterases have been identified by screening microorganisms on various types of

agar plates such as phenol red, rhodamine B, tween, Nile blue and so forth [48]. However

varying degrees of success have been reported with the different methods of screening. An

extracellular lipase isolated from a psychrotrophic Pseudomonas strain was discovered by

screening on olive oil agar plates. Some researchers have found success with the rhodamine

B dye method developed by Kouker and Jaeger (1987) [30,32,49,50]. However, others

encountered difficulties in preparing the media, as well as visualizing activity of weaker

lipases [51]. Based on the results from this study, the recommended method of screening for

lipolytic activity would therefore be, phenol red agar plates supplemented with 1% olive oil

or tributyrin.

12
In addition, isolates were also screened for cellulose activity. In the pulp and paper industry,

the presence of cellulases has undesirable effects on the quality of pulp generated,

particularly in the production of dissolving pulp (high grade cellulose pulp, >98% cellulose

content). Potential cellulases would hydrolyze the cellulose fibres resulting in a decrease in

alpha cellulose, thus impacting yield [52]. Consequently, the detection and elimination of

cellulase activity is important. Both the quantitative (screening plates) and qualitative (DNS

assay) revealed negligible cellulase activity except for Curtobacterium flaccumfaciens (Table

4). This was addressed by using spin columns with specific cut-off sizes to eliminate the

larger proteins (>50 kDa) which could be potential cellulases.

Native & SDS-PAGE

Native PAGE gels supplemented with carboxymethylcellulose and avicel were used to ensure

that the minimal endoglucanase and exoglucanase activity observed was eliminated. Samples

concentrated with the 3 kDa spin column were thereafter passed through a 50 kDa spin

column to remove the larger proteins, presumably thought to be cellulases (Figure 8). It is

imperative that the enzyme extracts characterized here, contain no cellulase activity that may

degrade the cellulose fibers. All other accessory enzymes such as xylanases, laccases, and

ligninases that may be present will positively contribute to the production of high quality

cellulose pulp. Bacterial lipases and esterases generally have an expected protein size of

between 15 and 45 kDa [53]. Proteins larger than 50 kDa were regarded as potential

cellulases. Cellulases have a negative impact on the final pulp by reducing cellulose chains.

An esterase as small as 1.57 kDa from Bacillus stearothermophilius has been described by

Simoes et al. (1997) [54]. Bacillus thuringiensis has been reported to produce a 38 kDa

phospholipase [55].

13
 Lipase and esterase activity

Upon evaluation of the preliminary screenings, the following isolates were selected for

further study, DF3 - Curtobacterium flaccumfaciens, DF7 - Bacillus thuringiensis, B9 –

Pantoea sp. and BT - Bacillus thuringiensis. In addition to the bacterial isolates selected, two

fungal isolates F4 – Paecilomyces formosus and X – Phialophora alba were chosen based on

similar preliminary plate screenings (data not shown) as well as previous studies on laccase

activity [56]. The effect of initial pH on the extracellular and intracellular lipase/esterase

activity of the selected isolates was investigated at pH 8 and 37°C with acetate and butyrate

as substrates (generally selected for initial investigations). The results in Table 4 show a

higher enzyme activity in the extracellular fractions of BT, DF7, and DF3, whilst B9

demonstrated higher activity in its intracellular fraction. Therefore, the appropriate fractions

were used for further characterization of these enzymes. Fungi are known to produce

extracellular enzymes to degrade polymers that cannot be absorbed [57], therefore it was not

unexpected that the intracellular fraction yielded no enzyme activity.

Effects of temperature and pH on enzyme activity

Specificity of lipases are directed by a variety of properties such as type of substrate, position

of esters fatty acids, stereospecificity and a combination of all four. These include factors that

alter the binding of the enzyme to the substrate, the molecular properties of the enzyme, and

structure of the substrate [58]. Therefore, in the work reported here, it was vital to institute an

experimental design to test the effects of pH and temperature on a range of substrates. A

majority of reported studies elect to determine pH and temperature optima and then test the

substrate specificity of the optimal expressed enzyme [59,60]; less detailed studies have

14
demonstrated some effect of pH on substrate specificity of lipases and esterases [61]. Ertuğrul

and colleagues found that at pH 6, lipases from a Bacillus strain demonstrated highest activity

towards the long chain triglyceride trimyristin (C14), however, at pH 9, the shorter chain

triglycerides such as tributyrin (C4) and triacetin (C2) provided higher lipase activity

compared to the longer chain triglycerides (C8-C14) [61]. This behaviour has also been

reported for acetyl esterases from Thermomyces lanuginosus where, no activity was observed

against pNP-acetate at pH 9.0, however, activity at pH 4.0 was recorded [62]. This reveals the

varying degrees of lipase and esterase activity depending on the pH of the medium, which

may be attributed to the presence of isoenzymes. Results of our study indicate that substrate

specificity is affected by changes in pH and temperature.

The enzymes in our study showed a preference for acidic conditions which is fairly

uncommon amongst bacterial lipases. The majority of lipases are known to display their

highest activities at a neutral or alkaline pH [63,64,65]. However, there are reports of the

production of acidic lipases from bacteria although with varying amounts of activity. Ramani

et al. (2010) described the production of an acidic lipase by Pseudomonas gessardii which

had a maximum activity of 156 U/ml at a pH of 3.5 [66]. On the lower end of the scale, an

acidic lipase produced by Aeromonas sp. demonstrated optimal activity of 0.7 U/ml at a pH

of 6 [67].

The highest hydrolysis rates were obtained with potential lipases isolated from Bacillus

thuringiensis (BT and DF7) on p-NP-valerate (C5) p-NP-octanoate (C8), p-NP-dodecanoate

(C12), and p-NP-myristate (C14), indicating the enzymes’ propensity for longer acyl chain

lengths (Figure 5). The p-NP esters of palmitic and stearic acids were also good substrates,

however the shorter acyl chain esters such as acetate, butyrate and valerate were hydrolysed

15
at a lower rate but with relatively comparable activity to the longer chain acyl chain

substrates. This suggests that the enzymes from both B. thuringiensis isolates could

potentially produce both lipases and esterases. Lipases from Bacillus species such as Bacillus

stearothermophilus have been reported to hydrolyse synthetic substrates with acyl group

chain lengths between C8 and C12 with optimal activity on C10 p-NP-caprate [68]. On the

other hand, a lipase isolated from B. stearothermophilus had a wide substrate specificity

towards triglycerides with C4 to C18 [69].

Initially, when the enzymes were tested at pH 8, greater activity was observed with p-NP

acetate and p-NP butyrate (data not shown). However, at the optimal pH of 4 and 5, greater

activity towards dodecanoate, myristate and palmitate was noted (Table 6). This suggests that

changes in pH have an influence on the substrate specificity of the enzyme. These findings

may be explained by the phenomenon of induced fit model. This model claims that the

substrate may cause substantial transformation in the three-dimensional link of the amino

acids at the active site and these modifications in protein structure initiated by a substrate will

bring the catalytic groups into a suitable orientation for reaction [70]. Post and Ray (1995)

showed that conformational changes can enhance the specificity of an enzyme with

suboptimal catalytic efficiency [71].

The enzymes isolated from the other microorganisms (DF3, F4, X) showed a preference for

dodecanoate, palmitate, myristate, octanoate and stearate substrates. The enzymes’ specificity

in relation to lipids with fatty acid residues of C8-C18 chain length compellingly suggests that

the enzymes described in this study could be true lipases. Enzymes isolated from Pantoea sp.

(B9) could potentially be classified as esterases due to their specificity towards butyrate and

valerate. The criteria used to differentiate esterases from lipases, is that esterases do not

16
hydrolyse esters containing an acyl chain length of longer than 10 carbon atoms [72]. It is

unusual for isolate B9 to prefer pNP-butyrate over pNP-acetate, such specificity is

uncommon in nature, however, novel esterases from Lactobacillus casei and Escherichia coli

have previously demonstrated such catalytic preference [72,73]. Curtobacterium

flaccumfaciens (DF3) displayed highest activity of 60 U/ml at 30°C with substrate specificity

towards palmitate. Curtobacterium flaccumfaciens is an endophytic bacteria associated with

crops such as rice, potato, yam, tobacco, and cucumber and is capable of producing lipases

[74]. This could be the first report of a characterized lipase from Curtobacterium

flaccumfaciens isolated from Eucalyptus wood.

Low activities were obtained for laccases (Figure 4), and this is expected as extracellular

laccases from basidiomycete fungi are known to be produced in low amounts [75]. It is

recognized that when fungi are grown in a medium of pH 5, laccases will be produced in

excess, however most studies show that a pH range of 3.6 to 5.2 is suitable for enzyme

production [76]. Optimal temperatures for laccase activity can vary significantly amongst

organisms. There are reports of activities in the range of 25 to 80°C, with most enzymes

having an optimum at 50 to 70°C [77]. In this study the optimum temperatures of the lipases

and esterases were 30 and 35°C, respectively. Therefore, laccase activity and stability were

tested at these temperatures as the final application of this study would be to create an

enzyme cocktail to treat pulp for effective removal of lipophilic extractives. Nevertheless,

there was minimal variation in activity from the optimal pH and temperature of isolates F4

and X. Isolate F4 displayed 6.8% and 9.7% more activity at the optimal conditions of 40°C

and pH 5.5, respectively. Isolate X showed 15.3% more activity at 50°C, whilst the optimal

pH remained the same. Our results are comparable to another study where the maximum

17
production of laccase from Trichoderma harzianum was observed at 35°C and pH 5 after 6

days [78].

In addition to demonstrating laccase activity (up to 3.1 U/ml) (Figure 4), Paecilomyces

formosus (F4) and Phialophora alba (X) also demonstrated high substrate specificity towards

dodecanoate at 35 and 30°C, respectively. Limited information has been published on the

enzymes produced by P. alba, however, previous work indicate that xylanases from this

microorganism were characterized with activity of up to 420 IU/ml [79]. The presence of

enzymes from this microorganism could greatly assist in the reduction of pitch formation as

well as the breakdown of xylan which will reduce the amount of chemicals used in the

downstream processing of pulp [80,81]. Laccases also have the ability to degrade both

phenolic and non-phenolic compounds. Plant phenols released by hardwoods during pulping

may have an inhibitory effect on enzyme activity [82], therefore the inclusion of fungal

laccases in this study could mitigate the inhibitory effects of phenolic compounds.

Effects of temperature and pH on stability of enzymes

In the pulp and paper industry, the enzyme pre-treatment of pulp is a tricky affair. When

considering the addition of enzymes to pulp, a number of variables such as dosage,

incubation period, temperature, pH and combination of enzymes needs to be taken into

account. Time is money, so minimal amount of time for enzyme pre-treatment would be

optimal. Therefore, when determining enzyme stability, a shorter range for the incubation

period was selected. Stability was however tested at 18 hours to establish a broader range for

incubation time, however, in industry pre-treatment times of up to 18 hours are not feasible.

18
The enzymes from the various microorganisms appear to be relatively stable over a period of

18 hours at their optimal temperature. Enzymes from DF3, DF7, and X maintained their

lipolytic activity over a period of 3 hours with minimal loss in activity and retained at least

60% activity after 18 hours (Figure 6). Enzymes isolated from BT, X, F4, DF3 and DF7 were

fairly stable up to 2 hours and thereafter a 30-40% decrease in activity was observed. More

than 90% of the original activity was retained after 18 hours for DF3 with dodecanoate and

palmitate as substrates. Enzymes from DF7 and F4 retained more than 75% activity after 18

hours with butyrate and valerate as substrates, respectively. B9 on the other hand, initially

demonstrated high stability after 1 hour of incubation followed by a drop in activity to 70%

after 3 hours of incubation. These results fare well in comparison to other studies under

similar conditions. For example, in a study by Eggert et al. (2001) a variant of an esterase

(LipB, EC 3.1.1.1) from Bacillus subtilis was found to be stable at pH 5 and 45°C for 1 hour

[83].

Specificity of enzymes from DF3, DF7, F4 and X towards both the shorter and longer

aliphatic acyl chains over the 18 hour incubation period indicates the broad range of

substrates these enzymes are able to act upon. The stability of these enzymes is a desirable

characteristic and would offer an advantage in potential industrial applications. However, for

the purpose of this study the addition of these enzymes to pulp as a pre-treatment step would

be optimal up to 2-3 hours. Similar results were reported by Massadeh and Sabra (2011)

where a lipase isolated from Bacillus stearothermophilus remained stable at a pH range of 7

to 9 after incubation for 1 hour at 30°C, with a residual activity remaining above 50% for pH

7, 8 and 9 [84]. However, extremophilic organisms are capable of producing hardier lipases.

A thermostable lipase from Geobacillus thermodenitrificans IBRL-nra was found to have an

optimal temperature of 65°C, at which it retained its initial activity for 3 hours. Its highest

19
lipase activity was reported at pH 7.0 and stable for 16 hours at 65°C [85]. Borkar et al.

(2009) reported a lipase from a Pseudomonas aeruginosa strain which was found to be

completely stable at 55°C after 2 hours at pH 6.9 [86]. A lipase from a psychrotolerant

Pseudomonas fluorescens strain was active at a temperature range of 15-65°C, however, it

exhibited maximum activity at 45°C and pH 8.0. This enzyme demonstrated high stability,

retaining 100% and 70% of its activity after an incubation period of 45 and 100 minutes,

respectively, at 45°C and pH 8.0. This particular lipase also showed a broad substrate

specificity acting on p-nitrophenyl esters with C8-C18 acyl groups as substrates [60].

Many researchers elect to clone genes coding for enzymes of interest in order to increase

activity and improve production [87,88,89]. However, in industry this may not be a practical

approach as screening of clone libraries involves conventional agar plate-based methods,

which would require approximately 10,000 petri plates, each containing 10,000 clones. This

is time-consuming and would greatly increase expenditure [90]. The enzyme activities

observed in this study are comparable to, if not higher, than those of lipases and esterases

which have not been modified or cloned (Table 7). The activities recorded in this study (up to

60 U/ml) could be invaluable in the reduction of pitch formation in the pulp and paper

industry. In addition, the enzymes described here are indigenous to Eucalyptus wood species

and have not been modified in any way, thus making them feasible and ideal for industrial

applications. This is particularly the case for the acid-bisulphite pulping process used to

20
produce dissolving pulp, as this process involves acidic pH process conditions which would

be suitable for the enzymes described in this study.

Conclusions

In the present work, a cellulose-free cocktail of lipolytic and other enzymes was obtained

from microorganisms indigenous to South African Eucalyptus wood chips. Lipases and

esterases showed optimal activity at moderate temperatures (30 and 35°C) and acidic pH

range (pH 4 and 5). The enzymes’ stability and activity on a broad range of lipophilic

substrates could lead to potential biotechnological applications in the removal of lipophilic

components that cause pitch problem in the manufacture of high purity chemical pulps such

as dissolving wood pulp. The inclusion of laccases have the potential to assist in further

degradation of these problematic lipophilic compounds. Future work will focus on applying

these enzymes directly to the pulped wood chips and evaluating their potential to reduce the

agglomeration of lipophilic compounds that cause pitch formation during pulping. The

application of enzymes produced by indigenous microflora will aid in reducing cost and is a

greener alternative to chemical treatments.

Conflict of Interest.

Acknowledgements

This work was supported by the National Research Foundation (NRF) and the Forestry and

Forest Products (FFP) division at the Council for Scientific and Industrial Research (CSIR),

Durban, South Africa.

21
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30
 A B

Figure 2: A- 1% tributyrin plate screening assays for the detection of esterase activity of pure
bacterial isolates from Eucalyptus wood species, B- 1% olive oil/rhodamine B plate screening
assays for the detection of lipase activity of pure bacterial isolates from Eucalyptus wood
species.

Figure 3: 1% Tween 80 agar plates (left) and 1% tributyrin phenol red agar plates (right) for
the detection of lipase and esterase activity, respectively, of pure bacterial isolates from
Eucalyptus wood species.

31
 A B

Figure 4: Activity and stability of laccases from fungal isolates F4 and X. A: activity at
30°C, 35°C, pH 4 and pH 5; B: enzyme stability at 35°C and pH 4 for F4 and 30°C and pH 5

32
 for X.
A 25 degrees B 30 degrees
Acetate Acetate
60 60

Enzyme Activity (U/ml)

Enzyme Activity (U/ml)

ure Butyrate Butyrate

50 50
Valerate Valerate
40 40
Octanoate Octanoate
30 30
Dedecanoate Dedecanoate
20 20
Myristate Myristate
10 10
Palmitate Palmitate
0 0
Stearate Stearate
DF3 DF7 B9 F4 BT X DF3 DF7 B9 F4 BT X

C 35 degrees D 40 degrees
Acetate Acetate
60 Butyrate 60 Butyrate
Enzyme Activity (U/ml)

Enzyme Activity (U/ml)

50 Valerate 50 Valerate
40 Octanoate 40 Octanoate
30 Dedecanoate 30 Dedecanoate
20 20
Myristate Myristate
10 10
Palmitate Palmitate
0 0
DF3 DF7 B9 F4 BT X Stearate DF3 DF7 B9 F4 BT X Stearate

E 45 degrees F 50 degrees
Acetate
Acetate
60 Butyrate 60 Butyrate

Enzyme Activity (U/ml)

Enzyme Activity (U/ml)

50 Valerate 50 Valerate
40 Octanoate 40 Octanoate
30 Dedecanoate 30 Dedecanoate
20 Myristate 20 Myristate
10 10
Palmitate Palmitate
0 0
DF3 DF7 B9 F4 BT X Stearate Stearate
DF3 DF7 B9 F4 BT X

33
Figure 5: Effect of temperature (at optimum pH 4 or 5) on the activity of esterases/lipases from isolates DF3 (pH 4), DF7 (pH 4), B9 (pH 4), F4
(pH 4), BT (pH 5) and X (pH 5) on p-NP esters (C2-C18).

34
 A B DF3, 30 degrees
100

Residual Activity (%)

Octanoate
80
Dedecanoate
60
Myristate
40 Palmitate
20 Stearate
0
0h 1h 2h 3h 18 h

C D

E F

35
Figure 6: Stability of esterases/lipases from DF3 (A); DF7 (B); B9 (C); F4 (D); BT (E) and X (F) at optimum temperature and pH.

36
 1 2 3 1 2 3 1 2 3
A B C
180 kDa

72

55

34

Figure 8: Native PAGE gels supplemented with carboxylmethylcellulose (CMC) to confirm removal of any potential endoglucanases. A: SDS-
PAGE of crude enzymes, 1- DF7, 2- DF3, 3-BT. B: Native PAGE of crude enzymes, 1- BT, 2-DF3, 3- DF7. C: Native page of crude enzymes
after partial purification, 1- BT, 2- DF3, 3- DF7.

37
 Table 2: Lipase and esterase activity of bacteria isolated from a mixed Eucalyptus wood chip pile
Accession Esterase Esterase Esterase Lipase Lipase
Species
Number 1% Trb 2% Trb 5% Trb 1% Oil 2.5% Oil
B1 Pseudomonas aeruginosa JX945659 + + - - -
B2 Pseudomonas aeruginosa JX945660 ++ - - - -
B4 Bacilllus firmus JX945657 + + - + +
B5 Micrococcus luteus JX945661 + + - + -
B6 Bacillus sp. JX945662 ++ - - + -
B7 Inquilinus limosus JX945663 +++ - + - -
B9 Pantoea sp. JX945664 ++ - - + -
B10 Klebsiella sp. JX945665 + - - - -
B12 Bacillus ginsengihumi JX945658 ++ ++ + + -
B14 Streptomyces costaricanus JX945666 - - - - -
B15 Pseudomonas aeruginosa JX945667 - - + - -
B16 Cellulosimicrobium cellulans JX945668 - - - + +
Key: + = slight halos (1-2 mm), ++ = medium halos (2-5 mm), +++ = large halos (>5 mm), Trb= tributyrin, Oil= olive oil, - = no halos

38
 Table 3: Lipase and esterase activity of bacteria isolated from different Eucalyptus spp.
Lipase
GenBank Esterase Esterase Esterase Lipase
Species 2.5%
Number 1% Trb 2% Trb 5% Trb 1% Oil
Oil
E. dunnii
DF1 Mucilaginibacter sp. JF999998.1 - - + - -
DF2 Unidentified - - ++ + - +
DF3 Curtobacterium flaccumfaciens HE613377.1 - ++ + - -
DF5 Pantoea vagans CP002206.1 - - + + -
DF6 Unidentified - - ++ + + -
DF7 Bacillus thuringiensis FN667913.1 - ++ + + +
DF8 Unidentified - - + + - -
E. grandis
G1 Pantoea agglomerans FJ11844.1 - ++ + - -
G2 Curtobacterium flaccumfaciens JF706511.1 - ++ - - -
G3 Pantoea vagans CP002206.1 - ++ + + -
G4 Unidentified - - - + - -
E. nitens
N1 Bacillus cereus JF758862.1 ++ ++ + - -
N2 Pantoea sp. JN853250.1 - - + - +
N3 Curtobacterium sp. HQ219967.1 - +++ + - -
N4 Bacillus cereus JQ308572.1 - - + - +
N5 Bacillus cereus EU621383.1 - - + - -
N6 Bacillus sp. EU162013.1 - ++ + - +
N7 Bacillus thuringiensis FN667913.1 - ++ + - -

Key: + = slight halos (1-2 mm), ++ = medium halos (2-5 mm), +++ = large halos (>5 mm), Trb= tributyrin, Oil= olive oil, - = no halos

39
 Table 4: Lipase/esterase and cellulase activity (endoglucanase and exoglucanase activity using the DNS assay) and protein concentrations of the
intracellular and extracellular fractions from the different isolates
Protein
Acetate Butyrate Endoglucanase Exoglucanase
Conc.
(U/ml) (U/ml) Activity Activity
(μg/ml)
(U/ml) (U/ml)
Ext. Int. Ext. Int. Ext. Int.
Bacillus
BT 5.55 5.24 9.75 5.78 212.9 1.57 0.057 0.043
thuringiensis
Bacillus
DF7 10.71 5.16 10.98 4.34 414.3 1.84 0.021 0.013
thuringiensis
B9 Pantoea sp. 5.12 6.75 2.82 5.27 1.69 25 0.012 0.015
Curtobacterium
DF3 10.35 4.09 10.70 3.44 62.86 1.88 0.203 0.121
flaccumfaciens
Paecilomyces
F4 7.78 ─ 18.89 ─ 51.43 ─ 0.019 0.029
formosus
Phialophora
X 2.18 ─ 30.11 ─ 98.57 ─ 0.034 0.041
alba

Table 6: Optimized pH, temperature and substrates for lipolytic enzymes from the different isolates
Optimum Optimum
Isolate Substrate Specificity
pH Temperature
BT 5 30°C Dodecanoate, Myristate, Octanoate, Acetate
DF7 4 35°C Dodecanoate, Octanoate, Valerate, Butyrate
B9 4 35°C Valerate, Dodecanoate, Butyrate, Octanoate
DF3 4 30°C Palmitate, Dodecanoate, Myristate, Octanoate
F4 4 35°C Dodecanoate, Palmitate, Octanoate, Myristate
X 5 30°C Dodecanoate, Stearate, Myristate, Octanoate

40
 Table 7: Comparison of optimal temperature and pH of some lipases and esterases isolated from different bacteria
Isolate Enzyme pH Temperature (°C) Enzyme Activity (U/ml) Reference
Bacillus THL027 Lipase 7 70 8.3 Dharmsthiti and Luchai (1999) 91
Bacillus coagulans BTS-3 Lipase 8.5 55 1.16 Kumar et al. (2005) 65
Geobacillus zalihae sp. Lipase 6.5 65 0.15 Rahman et al. (2007) 92
Pseudomonas aeruginosa LP602 Lipase 8 55 3.5 Dharmsthiti and Kuhasuntisuk (1998) 93
Pseudomonas gessardii Lipase 3.5 30 156 Ramani et al. (2010) 66
Burkholderia multivorans Lipase 7 30 58 Gupta et al. (2007) 94
Burkholderia multivorans V2 Lipase 8 37 14 Dandavate et al. (2009) 95
Burkholderia sp. ZYB002 Lipase 8 65 22.8 Shu et al. (2012) 96
Enterococcus durans NCIM5427 Lipase 4.6 30 207.6 Vrinda (2013) 97
Streptomyces exfoliates LP10 Lipase 6 37 6.9 Aly et al. (2012) 98
Salinivibrio sp. strain SA-2 Lipase 7.5 50 5.1 Amoozegar et al. (2008) 64
Anoxybacillus gonensis A4 Esterase 5.5 60-80 0.8 Faiz et al. (2007) 99
Bacillus sp. strain DVL2 Esterase 7 37 5.2 Kumar et al. (2012) 100
Bacillus licheniformis Esterase 8-8.5 45 12 Alvarez-Macarie et al. (1999) 101
Geobacillus sp. DF20 Esterase 7 50 27.9 Özbek et al. (2014) 102
Lactobacillus brevis NJ13 Esterase 8 50 48.12 Kim et al. (2013) 103
Acaligens faecalis Esterase 8 30 0.27 Poornima and Kasthuri (2016) 104
Burkholderia fungorum A216 Esterase 6.5 37 0.014 Jiao et al. (2014) 105
Achromobacter denitrificans strain SP1 Esterase 8 50 89.5 Pradeep et al. (2015) 106
Janthinobacterium lividum Esterase 7 30 0.00568 Park et al. (2001) 107
Pseudomonas sp. KWI-56 Esterase 7.5 22 51.6 Sugihara et al. (1994) 108

41