Robust production of pigment-free pullulan from lignocellulosic hydrolysate by a new fungus co-utilizing glucose and xylose

Carbohydrate Polymers 241 (2020) 116400

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Carbohydrate Polymers
journal homepage: www.elsevier.com/locate/carbpol

Robust production of pigment-free pullulan from lignocellulosic hydrolysate
by a new fungus co-utilizing glucose and xylose

T

Guanglei Liua,c,1, Xiaoxue Zhaoa,1, Chao Chenb,d,e, Zhe Chia,c, Yuedong Zhangb,d,e, Qiu Cuib,d,e,
Zhenming Chia,c, Ya-Jun Liub,d,e,*
a

College of Marine Life Science, Ocean University of China, China
CAS Key Laboratory of Biofuels, Shandong Provincial Key Laboratory of Synthetic Biology, Qingdao Institute of Bioenergy and Bioprocess Technology, Chinese Academy
of Sciences, China
c
Laboratory for Marine Biology and Biotechnology, Pilot National Laboratory for Marine Science and Technology, China
d
Dalian National Laboratory for Clean Energy, China
e
University of Chinese Academy of Sciences, Chinese Academy of Sciences, China
b

A R T I C LE I N FO

A B S T R A C T

Keywords:
Aureobasidium
Consolidated biosaccharification
Co-utilization
Lignocellulosic hydrolysate
Pigment-free pullulan

Cost-efficient production of pullulan is of great importance but remains challenging due largely to the high-cost
carbon sources. Lignocellulosic biomass is considered an alternative carbon source for industrial pullulan production, while new fungus producers that co-utilizing lignocellulose-derived glucose and xylose are required. In
this study, a new fungus Aureobasidium melanogenum TN2-1-2 showed simultaneously assimilation of glucose and
xylose and could produce pigment-free pullulan due to its deficiency in melanin synthesis. The ability of TN2-1-2
producing pullulan was remarkably robust in the presence of varying glucose to xylose ratios and ionic salt
concentrations. Furthermore, condensed lignocellulosic hydrolysates obtained by consolidated biosaccharification was used as the pullulan production medium without supplying any nutrients, and pigment-free pullulan
was produced by TN2-1-2 with the titer and yield of 55.1 g/L and 0.5 gPullulan/gCBS hydrolysate, respectively.
Hence, this work provides a potential industrial pullulan producer TN2-1-2 and new insight into the lignocellulose bioconversion to pullulan.

1. Introduction
Pullulan is an imperative natural polymer produced extracellularly
by yeast-like fungus Aureobasidium spp. (Li et al., 2015). Structurally,
pullulan is primarily composed of maltotriose repeating units crosslinked by α-(1→6) glycosidic bonds, and the glucose units of maltotriose are attached by α-(1→4) linkages (Sugumaran & Ponnusami,
2017). The unique linkage pattern endows pullulan distinctive physical
traits, adhesive properties, and capability to form fibers, compression
moldings, and strong films that are impervious to oxygen. Besides,
pullulan can be derivatized by substituting its hydroxyl groups with
desired chemical moieties to extend its biomedical applications, including targeted drug delivery, DNA carrier, tissue engineering, vaccination, molecular chaperons, and medical imaging (Singh, Kaur, &
Kennedy, 2015; Singh, Kaur, Rana, & Kennedy, 2017).
Owing to the important properties of pullulan, the bioprocess for

pullulan production has been widely studied to enhance the production
and yield. So far, sucrose is used as the main substrate for the commercial production of pullulan (Jiang et al., 2018; Sugumaran &

Ponnusami, 2017). However, the relative shortage of sucrose resource
and its high cost limit the industrial production of pullulan. Moreover,
the by-product accumulation of fructooligosaccharides caused by the
pullulan production from sucrose should also be concerned (Liu et al.,
2017). To reduce the carbon source cost for pullulan production, various substrates, including glucose, molasses, hydrolyzed potato starch
waste, and inulin, have been used to substitute sucrose (Goksungur,
Uzunogullari, & Dagbagli, 2011; Jiang et al., 2018; Ma, Liu, Chi, Liu, &
Chi, 2015; Srikanth et al., 2014), but it remains challenging to develop
processes for cost-efficient pullulan production.
Lignocellulosic biomass is the most abundant sustainable carbon
source on earth, thus has the potential to be used as an alternative
carbon source for industrial fermentation. Because of the complex and

⁎

Corresponding author at: CAS Key Laboratory of Biofuels, Shandong Provincial Key Laboratory of Synthetic Biology, Qingdao Institute of Bioenergy and
Bioprocess Technology, Chinese Academy of Sciences, China.
E-mail address: (Y.-J. Liu).
1
The authors contribute equally to this work.
/>Received 25 January 2020; Received in revised form 26 April 2020; Accepted 28 April 2020
Available online 03 May 2020
0144-8617/ © 2020 Elsevier Ltd. All rights reserved.


Carbohydrate Polymers 241 (2020) 116400

G. Liu, et al.

previously described (Jiang et al., 2018). Fungal fermentation and

carbon source assimilation analyses were performed according to a
published standard method (Kurtzman, Fell, & Boekhout, 2011). The
genomic DNA was extracted using a TIANamp Yeast DNA Kit
(TIANGEN, Beijing, China). Amplification and sequencing of the internal transcribed spacer region (ITS) of the rRNA gene cluster were
performed using the common primers ITS1 and ITS4 according to the
methods described by (Ma et al., 2014). The ITS1 sequence obtained
was aligned using BLAST analysis ( />cgi). The sequence which shared over 98% similarity with the currently
available sequence was considered to be the same species. The phylogenetic tree was constructed and visualized using Mega 7 software
(Kumar, Stecher, & Tamura, 2016).

recalcitrant structure of lignocellulose, one of the premises to produce
pullulan from lignocellulosic substrates is the efficient bioconversion of
the insoluble substrate into fermentable sugars. So far, various strategies have been developed for lignocellulose bioconversion, including
separate enzymatic hydrolysis and fermentation (SHF), simultaneous
saccharification and fermentation (SSF), consolidated bioprocessing
(CBP), and consolidated bio-saccharification (CBS) (Liu, Liu, Feng, Li, &
Cui, 2019; Parisutham, Kim, & Lee, 2014). SHF and SSF are off-site
saccharification strategies depending on fungal cellulases, and the enzyme cost severely limits their commercial applications (Lynd et al.,
2017; Taha et al., 2016). CBP integrates enzyme production, cellulose
hydrolysis, and fermentation in one step for lignocellulose bioconversion to greatly reduce the enzyme cost (Lynd, van Zyl, McBride, & Laser,
2005) and is mainly used for lignocellulosic biofuel production (Xu,
Singh, & Himmel, 2009). CBS is a newly proposed strategy for lignocellulose bioconversion by separating fermentation from the integrated CBP process (Liu, Li, Feng, & Cui, 2020). CBS employs cellulosome-producing strains as the whole-cell biocatalyst for lignocellulose
deconstruction and determines fermentable sugars as the target products. The produced sugar-rich CBS hydrolysates can be potentially
used as the carbon sources for various downstream fermentations (Liu
et al., 2019, 2020), including pullulan production.
It should be noted that CBS hydrolysates derived from complex
lignocellulosic biomass are usually with low sugar purity and concentration. For example, the CBS end-products from pretreated wheat
straw contained 22.9 g/L glucose and 7.0 g/L xylose (Liu et al., 2019).
Thus, to construct a complete bioprocess from lignocellulose to pullulan, the pullulan production process should be compatible with the
CBS process, and the Aureobasidium strains should co-ferment C6/C5

sugars under various sugar ratios and tolerate high salt conditions. To
be specific, the pullulan producers should be able to efficiently co-ferment glucose and xylose to achieve high yield. Robust pullulan production under various sugar ratios is also critical for industrial applications. Additionally, because CBS hydrolysate will be concentrated to
improve the reducing sugar concentrations, the high osmotic pressure
caused by increased salinity in the CBS system should be tolerated by
the pullulan producers as well. Therefore, in the present study, a new
pullulan producing yeast-like fungal was screened and characterized,
and a CBS-compatible fermentation process was developed for the robust production of pullulan from lignocellulosic biomass.

2.3. Preparation of CBS hydrolysates
The CBS process was performed as previously described (Liu et al.,
2019). In brief, the strain C. thermocellum ΔpyrF::KBm was initially
cultivated with 5 g/L Avicel as the sole carbon source to the exponential
stage. 5% (v/v) of the cells were then inoculated into the GS-2 medium
to initiate the saccharification process with 40 g/L sulfite pretreated
wheat straw (SPS) as a cellulosic substrate. The sulfite pretreatment was
performed in a cooking reactor (VRD-42SD-A China Pulp and Paper
Research Institute, Beijing, China) at 160 °C for 60 min with 20% (w/w,
based on dry substrates) dosage of ammonium sulfite and a liquid to
solid ratio of 6. Afterwards, the pretreated wheat straw samples were
washed with tap water before the saccharification process. The anaerobic bottles were horizontally shaken in a 55 °C incubator at 200 rpm
for 8 days. 1.5-mL cultures were sampled every 2 days to determine
sugar production. The obtained lignocellulosic hydrolysates were then
treated with 3% (w/v) activated carbon in a water bath at 80 °C shaking
for 3 hours. The mixtures were concentrated at 10,000 g for 10 min to
remove carbon powder. The supernatants were placed in a 50 °C drying
container for moisture evaporation until the reducing sugars were
concentrated to the determined concentration. The protein concentrations of the hydrolysates were determined as previously described
(Bradford, 1976).
2.4. Fungal fermentation for pullulan production
The fungal strains were aerobically grown in YPD medium at 28 °C

for 24 h, and then 5-ml cultures were inoculated into the 250-mL flasks
containing 30.0 ml of pullulan production medium with xylose, glucose, or a mixture of xylose and glucose as the carbon source. The
concentration of supplemented sugar varied from 100.0 to 140.0 g/L.
When a mixture of xylose and glucose was used as the carbon source,
the total sugar concentration was 110.0 g/L with different glucose to
xylose ratios (100%:0, 75%:25%, 50%:50%, 25%:75%, 0:100%). After
treatment with activated carbon and concentration, CBS hydrolysates
were used as the medium for pullulan production without adding relative nutrients. If required, different concentrations of NaCl (0.0, 10.0,
20.0, 30.0, 40.0, 50.0, and 60.0 g/L) was supplemented at the beginning of fermentation. The electrical strength of the cultures was measured using an AquaPro Water Quality Tester (HM Digital, US) at 20 °C.
All cultivations were performed aerobically at 28 °C, 180 rpm for 120 h.

2. Materials and methods
2.1. Bacterial and fungal strains and cultivation
The yeast-like fungal strains TN12-1, TN12-2, TN5-3, TN1-2, TN3-3
and TN2-1-2 used in this study were isolated from natural honeycomb
(Jiang et al., 2018). A. melanogenum strain P16 was isolated from a
mangrove ecosystem (Ma, Fu, Liu, Wang, & Chi, 2014). The yeast-like
fungal strains were maintained in yeast-polypeptone-dextrose (YPD)
medium and on potato dextrose agar (PDA) at 28 °C. The pullulan
production medium was composed of 110.0 g/L carbon source (xylose
and glucose), 2.0 g/L yeast extract, 0.2 g/L (NH4)2SO4, 5.0 g/L
Na2HPO4·12H2O, and 0.15 g/L MgSO4·7H2O, pH 6.5. Clostridium thermocellum strain ΔpyrF::KBm (Liu et al., 2019) was cultivated anaerobically at 55 °C in GS-2 medium (1.5 g/L KH2PO4, 3.8 g/L
K2HPO4·3H2O, 2.1 g/L Urea, 1.0 g/L MgCl2·6H2O, 150 mg/L
CaCl2·2H2O, 1.25 mg/L FeSO4·6H2O, 1.0 g/L cysteine-HCl, 10 g/L
MOPS-Na, 6.0 g/L yeast extract, 3.0 g/L trisodium citrate·2H2O, 0.1
mg/L resazurin, pH 7.4) (Johnson, Madia, & Demain, 1981) with 5 g/L
Avicel (PH-101, Sigma-Aldrich LLC.) as the carbon source.

2.5. Pullulan purification and quantification
The pullulan purification and quantitative determination were

performed according to a previously reported method (Ma et al., 2014).
The fermentation broth was first heated in a boiling water bath for 15
min, cooled to room temperature, and centrifuged at 14,000 g and 4 °C
for 10 min to remove cells. Two volumes of ice-cold ethanol were added
in the supernatant to precipitate polysaccharides at 4 °C for 12 h. The
precipitate was then dissolved in deionized water at 80 °C and the
ethanol precipitation step was repeated. The obtained precipitate was

2.2. Phenotypic, biochemical and molecular analyses of the fungal strain
The colonies formed on the PDA plates were photographed, and the
phenotypic analysis of the cells in the cultures was performed as
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G. Liu, et al.

SunFire C18 (4.6 mm × 250 mm) chromatographic column at 35 °C
with the mobile phase of ethanol/water (volume ratio of 1:4) and a flow
rate of 1 mL/min. Soluble lignin was estimated by UV spectrophotometry at 280 nm (Zhang et al., 2014) and calculated according to
a previous method (Mussatto & Roberto, 2006). The concentration of
reducing sugar in CBS hydrolysate was determined by the 3,5-dinitrosalicylic acid (DNS) method. The carbon to nitrogen (C/N) ratio was
determined by analyzing the carbon and nitrogen elements on an elemental analyzer (Vario EL cube, Elementar Co., Germany) through
burning with oxygen at the temperature of 1200 °C for 70 s.
The statistical analysis was performed based on three separate experiments using the GraphPad Prism 6.01 (GraphPad Software Inc.,
USA). Mean values were compared and analyzed using either t-test or
one-way analysis of variance (ANOVA) with Tukey HSD post hoc multiple comparison test. A probability value of p < 0.05 was considered
significant.

lyophilized and weighed.
2.6. Pullulan characterization
For thin-layer chromatography (TLC), the purified pullulan and the
pullulan standard were dissolved in deionized water to reach a concentration of 10 mg/mL and were hydrolyzed by a commercial pullulanase (400 U/ml, Sigma-Aldrich LLC.) at 60 °C for 15 min. The TLC
analysis was carried out with a solvent system of N-butanol-pyridinewater (6:4:3) and a detection reagent comprising 20.0 g/L diphenylamine in acetone, 20.0 g/L aniline in acetone, and 850.0 g/L phosphoric
acid (5:5:1, v/v/v) (Silica gel 60, MERCK, Germany) (Jiang et al.,
2018). For determining the pullulan purity, 10 mg/mL of either the
purified pullulan or the pullulan standard was completely hydrolyzed
by the commercial pullulanase at 37 °C. The released maltotriose was
determined by high-performance liquid chromatography (HPLC) using
Agilent 1260 equipped with a Venusil HILIC column (4.6 × 250 mm, 5
μm) and a RID detector at 35 °C. The mobile phase was ultra-pure water
at a flow rate of 0.5 mL/min. The purity was calculated with the following equation (Eq1):

3. Results and discussion
3.1. The honey-derived strain TN2-1-2 converted both glucose and xylose to
pullulan efficiently

Purity(%) =
The amount of the releasedmaltotriosefromthe purifiedpullulan
×100%
The amount of the releasedmaltotriosefrom thepullulanstandard

Efficient co-utilization of glucose and xylose is essential for the
economically feasible production of pullulan from lignocellulosic hydrolysates because lignocellulosic hydrolysates usually contain both C6
and C5 sugars. However, most microorganisms prefer glucose over
other monomeric sugars (Gancedo, 1992). For pullulan production,
numerous reported Aureobasidium strains can efficiently synthesize
pullulan from glucose (Sugumaran & Ponnusami, 2017) but the pullulan production from xylose is relatively less investigated. So far, only
A. pullulans AY82 and A. pullulans ATCC 42023 were reported to produce pullulan from xylose with titers of 17.63 and 11.2 g/L, respectively, under their optimized conditions (Chen et al., 2014; Kennedy &

West, 2018).
To increase pullulan titer, high sugar concentrations are generally
required but may cause high osmotic stress to pullulan producers
(Choudhury, Saluja, & Prasad, 2011). Thus, the robust osmotolerant
ability is regarded as a desirable criterion for commercial pullulanproducing strains. Honey is a saturated or supersaturated solution of
sugars with extremely low water activity and is considered a natural
environment for the isolation of osmophilic yeasts (Cadez, Fulop,
Dlauchy, & Peter, 2015). In this study, six yeast-like fungal strains
isolated from natural honey samples in our previous study (Jiang et al.,
2018) were used for pullulan production with a high concentration of
xylose as the carbon source. As shown in Fig. 1, the strain TN2-1-2
showed the highest pullulan yield of 49.5 and 58.3 g/L from 110 g/L
xylose and glucose, respectively (Fig. 1), which were significantly
higher than those of the other tested strains, including a mangrove

(1)
For NMR analysis, 20 mg of the purified sample was dissolved in 0.5
mL of deuterated water. One-dimensional 13C NMR and 1H NMR experiments were performed on a Bruker Avance III 600 MHz NMR
spectrometer equipped with a z-gradient triple resonance cryoprobe
using the internal DSS as previously described (Lazaridou, Roukas,
Biliaderis, & Vaikousi, 2002).
The molecular weight of pullulan was determined using a Waters™
1515 Gel Permeation Chromatography (GPC) system with a Multi-angle
laser light scattering detector (MALLS) as described by (Jiang et al.,
2018).
2.7. Melanin extraction and determination
The strain TN2-1-2 and the type strain CBS105.22T were cultured in
the pullulan production medium for 120 h to determine the production
and accumulation of melanin after the fungal fermentation for pullulan
production according to previously described methods with modifications (Kumar, Mongolla, Pombala, Kamle, & Joseph, 2011). In detail,

the fungal cells grown in the pullulan production medium for 120 h
were separated by centrifugation at 8000 g for 10 min and suspended in
1 mol/L NaOH, followed by autoclaving at 120 °C for 20 min. The
supernatant of the autoclaved solution was further acidified to pH 2.0
with 1 N HCl to precipitate the pigment. The precipitate was recovered
by centrifugation at 5,000 g for 10 min and washed with deionized
water for three times. The purified melanin was lyophilized and
weighted to determine melanin production.
Melanin in the purified pullulan was determined based on the absorbance from 500 to 600 nm using a Multiskan Sky Microplate
Spectrophotometer (Thermo Fisher Scientific, USA) according to previously reported methods (Li et al., 2009).
2.8. Analysis methods
The cell biomass was determined by monitoring the cell dry weight
according to the methods described by (Chen et al., 2019). The CBS
hydrolysates were analyzed by HPLC. Glucose, xylose, arabinose and
cellobiose concentrations were determined using a refractive index
detector equipped with a Bio-Rad HPX-87H column as previously described (Zhang, Liu, Cui, & Cui, 2015). Furfural and 5-hydroxymethyl
furfural (HMF) was detected using a UV detector (284 nm) and a

Fig. 1. Pullulan producing ability of different yeast-like fungal strains isolated
from natural honey. 110 g/L of glucose or xylose were used as the carbon
source. Values were means of three independent determinations. ABCD Data in
the xylose group with different superscripts differ (p < 0.05). abcd Data in the
glucose group with different superscripts differ (p < 0.05).
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G. Liu, et al.

Fig. 2. Phenotypic and molecular analyses of the fungal strain TN2-1-2. A,
The colonies of the strain TN2-1-2 and
the type strain A. melanogenum
CBS105.22T on the PDA plate after 3, 6,
and 8-day cultivation. B, The cell morphology of the strain TN2-1-2. C, The
phylogenetic tree of TN2-1-2 with other
yeast relatives based on neighborjoining analysis of D1/D2 26S rDNA
sequences. Bootstrap values at the notes
are from 1000 replicates.

A. melanogenum NG swollen cells producing melanin-free pullulan because of its pH-based regulation of cell morphogenesis and melanin
biosynthesis (Li et al., 2009). The melanin biosynthesis after the pullulan production process was also analyzed. As shown in Fig. S1, after
cultivation in the pullulan production medium for 5 days, the type
strain CBS105.22T produced 0.026 ± 0.003 g/gcell dry weight melanin
while almost no melanin was detected in the culture of TN2-1-2. Thus,
the strain TN2-1-2 showed natural deficiency in melanin biosynthesis
thereby is considered a promising candidate for the production of
pigment-free pullulan.
For molecular identification, the ITS sequence of the strain TN2-1-2
(Accession number MN752213) exhibited 99 % similarity to that of the
type strain A. melanogenum CBS105.22T. Thus, as the topology of the
phylograms in Fig. 2D confirmed, the strain TN2-1-2 belonged to the
species A. melanogenum. So far, three A. melanogenum strains, including
TN2-1-2, P16, and TN1-2 have been proved to produce high titer of
pullulan (Jiang et al., 2018; Ma et al., 2014). This suggested that, although A. pullulans strains are generally regarded as the important
pullulan producer (Sugumaran & Ponnusami, 2017), strains of A. melanogenum also have great potential in the commercial production of
pullulan.

strain P16 (8.7 and 45.1 g/L) (Ma et al., 2014) and another honeyderived strain TN1-2 (15.2 and 48.7 g/L) (Jiang et al., 2018). It was also
noteworthy that the strain TN2-1-2 had similar pullulan productivity

with either xylose or glucose as the sole carbon source compared to
most of the other tested strains (Fig. 1), suggesting that TN2-1-2 could
efficiently assimilate both glucose and xylose to produce pullulan.

3.2. The strain A. melanogenum TN2-1-2 was naturally deficient in melanin
biosynthesis
The TN2-1-2 colonies grown on the PDA plate showed weak pink
and sticky and were surrounded by extracellular polysaccharides
(Fig. 2A). All the yeast-like fungal cells were ellipsoidal and oval and
were budding to generate the secondary conidia without the formation
of chlamydospores and arthroconidia (Fig. 2C), which were similar to
reported strains of Aureobasidium spp. (Li et al., 2015). Carbon source
assimilation experiments were performed and the results also showed
that TN2-1-2 had characteristics closely related to the type strain A.
melanogenum CBS105.22 T 584.75 (Table S1). Interestingly, unlike most
of known Aureobasidium spp. strains that usually synthesize melanin
thereby being named as “black yeast” (Li et al., 2015), TN2-1-2 showed
significantly reduced ability to synthesize melanin because the blackish
color was undetectable in the colonies on the PDA plate after 6 days’
cultivation, and was barely observed after 8 days’ cultivation (Fig. 2A).
In contrast, the type strain A. melanogenum CBS105.22T produced a
large amount of melanin within 6 days under the same condition
(Fig. 2B).
The melanin production is well-known as an obstacle to pullulan
industrial production by increasing the cost of pullulan purification
(Singh, Saini, & Kennedy, 2009). The pullulan fermentation process
commonly goes on for 5 days (Sugumaran & Ponnusami, 2017) and
such long-term cultivation usually causes severe accumulation of melanin pigment. Many studies have been carried out to obtain strains
deficient in pigment formation by mutagenesis and genetic engineering
(Chen et al., 2019; Yu, Wang, Wei, & Dong, 2012). For instance, Chen

et al. construct a mutant strain of A. melanogenum producing no melanin
by inactivating two copies of the PKS1 and PKS2 genes involved in the
DHN-melanin biosynthesis (Chen et al., 2019). Additionally, pH control
during fermentation was considered effective and convenient to harvest

3.3. Effect of glucose and xylose concentrations on pullulan production
It has been well documented that a high initial carbon/nitrogen
ratio (nitrogen starvation) is required to boost pullulan biosynthesis (Li
et al., 2015). Therefore, the effects of different concentrations of glucose and xylose on pullulan production and cell growth were investigated. The results indicated that as the glucose concentration increased from 100 to 140 g/L, the pullulan titer was gradually improved.
The glucose utilization maintained at a level of above 99% and the
production of cell biomass also showed no significant change (Fig. 3A).
When 110.0 g/L of glucose was used as the carbon source, the highest
pullulan yield of 0.53 gPullulan/gGlucose was obtained with the pullulan
titer of 58.3 g/L, which were higher than previously reported A. pullulans strains. For example, A. pullulans CCTCCM2012259 produced 39.8
g/L from glucose under nitrogen-limiting conditions in a 5 L fermenter
(Wang, Chen, Wei, Jiang, & Dong, 2015). Yu et al. reported an A.
pullulans SZU 1001 mutant which produced 25.65 g/L pullulan with a
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G. Liu, et al.

Fig. 3. Effects of glucose (A) and xylose
(B) concentrations on pullulan production and yield, cell dry weight, and
sugar utilization. Values were means of
three independent determinations. ABC
Data in the yield group with different
superscripts differ (p < 0.05). abc Data

in the pullulan production group with
different superscripts differ (p < 0.05).

yield of 0.51 gPullulan/gGlucose by fermenting in flasks (Yu et al., 2012).
As shown in Fig. 3B, the xylose utilization ratio of TN2-1-2 was
above 95% except for the setup containing 140.0 g/L xylose. When the
xylose concentration in the pullulan production medium ranged from
100 to 120 g/L, the pullulan yield and titer were all at the highest level
of 50.2 g/L and 0.46 gPullulan/gGlucose, respectively. According to a
previous report, A. pullulans AY82 could produce pullulan from xylose
with a maximal pullulan titer of 17.63 g/L under the optimized conditions (Chen et al., 2014). This suggested the excellent capability of the
strain TN2-1-2 to produce pullulan from not only glucose but also xylose. In addition, when the xylose concentration increased, the pullulan
production by TN2-1-2 maintained at a similar level under our condition but the xylose utilization decreased significantly especially when
the xylose concentration increased from 120 g/L to 140 g/L. This indicated the effect of xylose concentration on the pullulan production by
TN2-1-2 was more pronounced than that of glucose, and 110.0 g/L was
determined as the optimum concentration for both glucose and xylose
in terms of the pullulan production and yield. For further experiments,
the concentration of supplemented reducing sugars was adjusted to
110.0 g/L for fermentations based on either pure sugar or lignocellulosic hydrolysates.

was used as the sole carbon source (100% xylose). This result indicated
that the strain TN2-1-2 was capable of robust production of pullulan
with different glucose to xylose ratios ranging from 100%:0 to
50%:50%. Thus, TN2-1-2 could be potentially used as a robust industrial strain for pullulan production from various lignocellulosic hydrolysates.
Carbon catabolite repression is known as a wide-spread cellular
regulation that cells utilize one of two or more carbon sources preferentially when multiple carbon sources are provided (Deutscher,
2008). The presence of glucose may inhibit the utilization of xylose and
thus cause decreased sugar utilization and product yield (Kwak & Jin,
2017). Thus, besides the ability to utilize mixed glucose and xylose to
produce pullulan, whether the supplemented xylose and glucose were

utilized simultaneously by the strain should be concerned as well. The
co-assimilation ability of glucose and xylose by TN2-1-2 was subsequently determined by fermentation using a mixed sugar with an equal
amount of glucose and xylose as the carbon source (Fig. 4B). The result
showed that the amount of glucose and xylose decreased simultaneously along with the cultivation, indicating the C6/C5 co-fermenting
capability of TN2-1-2. It took 72 hours and 120 hours to exhaust 55 g/L
of glucose and xylose under this condition, respectively, indicating a
higher assimilation rate of glucose than that of xylose. Although TN2-12 could assimilate glucose and xylose simultaneously, it still showed
preference to glucose as the carbon source as other reported yeast
strains (Agbogbo, Coward-Kelly, Torry-Smith, & Wenger, 2006).

3.4. Robust pullulan production by the strain TN2-1-2 with mixed sugars
The C6 and C5 sugar compositions in lignocellulosic hydrolysates
vary depending on the type of substrate, pretreatment method, and
cellulolytic enzymes (Singh, Shukla, Tiwari, & Srivastava, 2014). Thus,
the industrial pullulan producers should have the robustness in the
efficient utilization of mixed sugars with various glucose to xylose ratios. To address this, the effects of glucose to xylose ratios on the cell
growth, substrate utilization, and pullulan production of the strain TN21-2 were tested. As shown in Fig. 4A, the glucose to xylose ratio varied
from 100%:0 to 0:100% and the total sugar concentration was 110 g/L.
The utilization rates of the total sugar maintained at a level of above
99% with various sugar ratios, and high pullulan titers and yields were
detected without significant difference (p > 0.05) except when xylose

3.5. Halotolerance of the strain TN2-1-2 in pullulan production
According to a previous study, 40 g/L SPS was considered the optimal substrate load for the current CBS process (Liu et al., 2019), and
approximately 30 g/L of reducing sugar would be produced. Because
110 g/L sugar was required for pullulan production, further concentration of the CBS hydrolysates would be required. Since the GS-2
medium used for CBS contained phosphate and various metal ion elements, the residual medium components in the CBS hydrolysate might
be concentrated along with the reducing sugar, resulting in increased
Fig. 4. The co-utilization of glucose and
xylose by the strain TN2-1-2 to produce

pullulan. A, Effects of glucose (G) to
xylose (X) ratios on pullulan production, yield, cell dry weight, and sugar
utilization. Values were means of three
independent determinations. ABC Data
in the yield group with different superscripts differ (p < 0.05). abc Data in
the pullulan production group with
different superscripts differ (p < 0.05).
B, The time course of residual glucose
during the fermentation with 50% glucose and 50% xylose as the carbon
source.
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G. Liu, et al.

substrate following the previously reported CBS process (Liu et al.,
2019). As determined by the DNS and HPLC methods, 32.84 g/L reducing sugar including 25.1 g/L glucose and 7.0 g/L xylose concentration was produced, and a trace amount of cellobiose was also
detected (Fig. S2).
Lignocellulosic hydrolysates may contain various lignin-derived
compounds that have an inhibitory or toxic effect on fermenting organisms (Kont, Kurasin, Teugjas, & Valjamae, 2013). But for fermentation with CBS hydrolysate as the carbon source, this may not be a big
issue as CBS itself is basically a biological process involving alive cells
and the pretreatment is generally performed under the mild conditions
and produce a low amount of toxins (Liu et al., 2020). According to the
HPLC analysis, furfural and 5-hydroxymethyl furfural (HMF) was not
detectable in the CBS hydrolysate, indicating the slight inhibitory effect
on downstream fungal fermentation and pullulan production. Although
lignin can be partially removed by pretreatment, there is still lignin left
in pretreated substrates which can be gradually released during the

saccharification process (Li, Liu, Yu, Zhang, & Mu, 2017; Tian, Zhao, &
Chen, 2018). Both soluble lignin and extracts contain chromophores
and auxophores, resulting in the deep color of the hydrolysates
(Korntner et al., 2015; Sixta, 2006). Furthermore, the hydrolysates
should be concentrated to reach the optimal sugar concentration of 110
g/L for pullulan production by TN2-1-2, and the color would become
even deeper as the sugar solution is concentrated. Because the dark
color of the cultivation medium must be avoided for pullulan production, activated carbon was used to discolorize the lignocellulosic hydrolysates before using as the carbon source. As shown in Fig. 6, the
hydrolysate color changed from dark red to light yellow. The protein,
sugar, and soluble lignin concentrations were determined before and
after discolorization. The results showed that the reducing sugar concentration maintained at a similar level, only slightly reduced by 3.3%
(from 32.84 g/L to 31.74 g/L) but the protein and soluble lignin concentrations decreased significantly from 0.16 g/L and 1.05 g/L to 0.01
g/L and 0.20 g/L, respectively. The C/N ratio of the decolorized CBS
hydrolysate was determined as 13.07 ± 0.63 based on the element
analysis. The decolorized hydrolysates were then concentrated until the
reducing sugar concentration reached 110 g/L and were used for
pullulan production. It was notable that the final electrical conductivity
of the condensed hydrolysates was 3.35 S/m, which was in the electrical conductivity tolerance range of the strain TN2-1-2 (0-3.86 S/m)
(Fig. 5). This suggested that the salinity of the prepared CBS hydrolysates would have little effect on the cell growth and pullulan

Fig. 5. Effects of NaCl concentrations on pullulan production, yield, cell dry
weight, and sugar utilization. The corresponding electrical conductivity values
were also given for comparison. Values were means of three independent determinations. ABC Data in the cell dry weight with different superscripts differ
(p < 0.05). abc Data in the pullulan production group with different superscripts
differ (p < 0.05).

salinity. To confirm whether the pullulan producer TN2-1-2 could tolerate high salt concentration, different amount of NaCl was supplied
into the pullulan production medium, and the total salinity was estimated by monitoring the corresponding electrical conductivities. As
shown in Fig. 5, the cell growth was not significantly influenced when
the NaCl concentration increased from 0 to 50 g/L (electrical conductivity increased from 0.46 to 6.37 S/m). When the NaCl concentration increased to 60 g/L and resulted in electrical conductivity of

7.60 S/m, the cell dry weight only declined by 13.9%, indicating high
halotolerance of the strain TN2-1-2. In terms of pullulan production, the
strain TN2-1-2 maintained a relatively high pullulan production level
when the NaCl concentration increased to 30 g/L (electrical conductivity increased to 3.86 S/m). It is known that the electrical conductivity of standard seawater with a salinity of 3.5% is about 3 S/m
(20 °C), implying that the strain TN2-1-2 was able to tolerate a high
level of salinity comparable to seawater for pullulan production.

3.6. Preparation of CBS hydrolysates for pullulan fermentation
The genetically engineered C. thermocellum strain ΔpyrF::KBm was
previously constructed as a whole-cell biocatalyst to produce lignocellulose-derived sugars via CBS process (Liu et al., 2019). After precultivation with Avicel as the sole carbon source, the ΔpyrF::KBm cells
were inoculated into the saccharification system with 40 g/L SPS as the

Fig. 6. Schematic representation of the
whole process for pullulan production
from lignocellulosic biomass. The
whole process contains two main steps,
consolidated biosaccharification (CBS)
and fungal fermentation. In the CBS
processs, the engineered C. thermocellum strain ΔpyrF::KBm was used as
the whole-cell biocatalyst to solubilize
sulfite pretreated wheat straw (SPS) to
sugar-rich CBS hydrolysates. Cellulose
and hemicellulose of the lignocellulosic
biomass were converted to glucose and
pentose (mainly xylose), respectively.
The obtained CBS hydrolysate was decolorized using activated carbon and
concentrated to reach a reducing sugar
concentration of 110 g/L. Afterwards,
the CBS hydrolysate was directly used
for fungal fermentation to produce

pullulan using a newly isolated fungus
strain A. melanogenum TN2-1-2.
Because TN2-1-2 is deficient in melanin
biosynthesis, the produced pullulan is
pigment-free without blackish color.
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G. Liu, et al.

pullulan production from glucose is usually with relatively low efficiency (Sugumaran & Ponnusami, 2017). Recently, Chen et al. developed an engineered strain TN3-1 that can produce 103.5 g pullulan
from 140 g glucose under the experimental conditions (Chen et al.,
2019). Nevertheless, glucose is mainly produced from corn starch. Due
to the big concern of the worldwide food shortage, especially in developing countries, the use of grains for non-food production should be
avoided.
Agro-wastes are promising non-food substrates for pullulan production (Mishra, Zamare, & Manikanta, 2018). For example, proteinrich corn steep liquor and de-oiled seed cake were used to grow A.
pullulans strains, and over 70 g/L pullulan were produced (Choudhury,
Sharma, & Prasad, 2012; Sharma, Prasad, & Choudhury, 2013). Although high pullulan yields were obtained, these agro-wastes were
usually supplemented as a nitrogen nutrient and starch-derived glucose
was still considered the main carbon source. Among all the carbon
sources of agro-wastes, lignocellulose is known as the most abundant
alternative carbon source for industrial fermentation but difficult to
utilize due to its recalcitrant structure. Chen et al. reported that the
maximal production of pullulan by A. pullulans AY82 from sugarcane
bagasse hydrolysate was 12.65 g/L with a yield of 0.25 g/g after 7-day
cultivation (Chen et al., 2014). While the hemicellulose hydrolysate
was obtained by stream explosion and sulfuric acid hydrolysis rather
than biosaccharification and various medium nutrients were supplemented (Chen et al., 2014; Prakash, Varma, Prabhune, Shouche, & Rao,

2011). The fungus A. pullulans ATCC 42023 was used to produce pullulan from prairie cordgrass hydrolysate obtained by cellulase hydrolysis. A high yield of 0.79 g/g was obtained but the pullulan titer was
only 11.2 g/L after cultivation for 168 h with supplementation of yeast
extract (Kennedy & West, 2018). Wang et al. isolated an adapted A.
pullulans mutant to produce pullulan from the hydrolysate of untreated
rice hull and obtained a maximal yield of 22.2 g/L (Wang, Ju, Zhou, &
Wei, 2014). Thus, although the pullulan production from lignocellulosic hydrolysates has been reported previously, we obtained
higher pullulan yield and titer by coupling CBS and fermentation of the
strain TN2-1-2.
The competitiveness of CBS sugars compared to starch sugar would
affect the feasibility of pullulan production from lignocellulose to a
great extent. As we have calculated previously, the cost of CBS sugar
should be competitive to starch sugar (∼500 US$ per ton currently)
(Liu et al., 2020). As a newly developed technology, CBS is considered
promising because it enjoys a major advantage in reducing enzyme
costs but further improvements, including process optimization and
development of new biocatalysts, are still required to make breakthroughs in terms of practical cost-effectiveness (Liu et al., 2020). It is
worth noting that the cost of sugar purification using activated carbon
accounted for over 50% of the total cost when the sugar yield of CBS
was 30 g/L in this study. Although further optimization on the purification process should be carried out, if the sugar yield of CBS could
increase to the optimal sugar concentration required for pullulan production (110 g/L), the cost of both cell cultivation and sugar purification would be greatly reduced. Additionally, pH control and optimization would also enhance pullulan production performance and reduce
the cost (Xia, Wu, & Pan, 2011).

production by the strain TN2-1-2.
3.7. Pullulan production by the strain TN2-1-2 from lignocellulosic
hydrolysates
Because the GS-2 medium used for the cultivation of wholt-cell
biocatalyst contained all metal ions and macronutrients that pullulan
production medium required, together with the remained YPD medium
in the inoculum, the residual nutrients in the CBS hydrolysate might
support the pullulan production by TN2-1-2. Besides, the pH value of

the CBS hydrolysate which was detected as 6.2 was close to that of the
pullulan production medium (pH 6.5). Thus, the concentrated CBS
hydrolysates containing 110 g/L reducing sugar was directly used as the
medium for pullulan production without supplementation of any nutrients, and the strain TN2-1-2 could produce 55.1 ± 2.1 g/L pullulan
with a yield of 0.50 gPullulan/gCBS hydrolysate. We also carried out the
fermentation using CBS hydrolysates supplemented with nutrients of
the pullulan production medium, and obtained the pullulan titer and
yield of 55.7 ± 2.3 g/L and 0.50 gPullulan/gCBS hydrolysate, respectively. As
shown in Fig. S3, the time courses of the cell growth, pullulan production, and sugar consumption were all similar, indicating that no
further supplementation of nutrients was required using the CBS hydrolysate for pullulan production. The pH value decreased to 3.6 after
120-h fermentation which was similar to the previous study (Chen
et al., 2014).
In this study, 5 ml of cells grown in the YPD medium were inoculated in 30-ml fermentation systems for pullulan production and the
remained yeast extract and peptone in the inoculum might contain
sufficient nitrogen sources for pullulan production. The C/N ratio of the
pullulan production media containing 110 g/L glucose was calculated
as 42.56 based on the content of nitrogen sources in the pullulan production medium and YPD medium, the inoculum size, and the nitrogen
contents according to a previously reported method (Guerfali et al.,
2019). The C/N ratio of the decolorized CBS hydrolysate was determined as 13.07 based on the element analysis. Taking account with
the nitrients derived from the inoculum, the C/N ratio was calculate to
be 10.96. It is known that a high C/N ratio usually plays an important
role in pullulan production (Li et al., 2015) because high concentration
of ammonium and glutamine mediates severe nitrogen metabolite repression in fungi (Tudzynski, 2014). Interestingly, although the CBS
hydrolysate had a lower C/N ratio than the pullulan production
medium, similar pullulan titers of 58.3 g/L and 55.1 g/L were obtained
(Fig. 3A and S3A). This result might be explained by the various nitrogenous metabolites such as protein-derived amino acids, lipid-derived phosphocholine and colamine, and nucleotides produced by C.
thermocellum in the CBS hydrolysate that could be detected by element
analysis but showed slight nitrogen repression effects on fungal fermentation, and also implied that the strain had the robustness on C/N
ratios for pullulan production and the tolerance to the metabolites in
reused media.

The glucose to xylose ratio in CBS hydrolysate was about 78%:22%,
thus the pullulan production was compared to that with mixed sugars
(glucose to xylose ratio of 75%:25%) as the carbon source, which was
58.9 g/L (Fig. 4A). The result suggested the pullulan titer and yield of
TN2-1-2 using the concentrated CBS hydrolysate were comparable to
those with pure sugars as the carbon source even without further supplementation of medium components
Because the fungal fermentation and pullulan purification processes
are relatively mature in industry, the cost-effectiveness of the pullulan
production process may greatly depend on the cost of carbon sources,
i.e., sugars. Sucrose is mainly used as the substrate for the industrial
pullulan production and the pullulan titer and yield could reach 67.4 g/
L and 0.56 gPullulan/gsucrose, respectively (Ma et al., 2014). However, the
high cost of carbon source (over 900 US$ per ton sucrose) still limits the
industrial production of pullulan. Glucose is a universal carbon source
with lower price (∼500 US$ per ton) compared to sucrose, but the

3.8. Characterization of pullulan produced from lignocellulosic hydrolysate
Pullulan is connected by α-1,6-D-glucosidic and α-1,4-D-glucosidic
linkages (Fig. 6), and pullulanase could selectively hydrolyze α-1,6-Dglucosidic linkages of pullulan to release maltotriose (Sugumaran &
Ponnusami, 2017). Indeed, as indicated by the TLC analysis (Fig. 7), the
pullulan produced from CBS hydrolysate by the strain TN2-1-2 was
hydrolyzed by a commercial pullulanase to maltotriose. Additionally,
we performed pullulanase hydrolysis of equal amount of the produced
pullulan sample and the pullulan standard and analyzed the hydrolysates using HPLC (Fig. S4). By comparing the amounts of released
7


Carbohydrate Polymers 241 (2020) 116400

G. Liu, et al.

standard, and 102. 8753 and 102.4281 ppm for the produced pullulan).
All these results suggested that the pullulan produced from CBS hydrolysates by the strain TN2-1-2 is with identical structure with the
commercial pullulan standard.
The product quality of the CBS hydrolysate-derived pullulan was
determined by GPC chromatogram analysis. As showed in Fig. S5, the
weight-average (Mw) and number-average molecular weight (Mn) of
the pullulan produced from CBS hydrolysates were 1.862 × 105 and
1.377 × 105 g/mol, respectively, which were higher than the values of
the pullulan produced from pure glucose (1.537 × 105 and 0.77 × 105
g/mol for Mw and Mn, respectively, suggesting the feasibility of utilizing CBS hydrolysate for pullulan production by TN2-1-2. The produced pullulan was pure white observed by eyes (Fig. 6) and the melanin in the purified pullulan was rarely detected (Fig. S6), suggesting
that the pullulan produced by TN2-1-2 was pigment-free. Thus, the
pullulan-producing strain that deficient in melanin biosynthesis would
have a wide range of application prospects in the food and medicine
industry.
4. Conclusion

Fig. 7. TLC analysis of the pullulan hydrolyzed by commercial pullulanase.
Lane 1, glucose; Lane 2, maltotriose; Lane 3 and 4, TN2-1-2 produced pullulan
hydrolyzed by activated and inactivated pullulanase, respectively; Lane 5,
pullulan produced by TN2-1-2 without treatment.

To adapt the conditions of the lignocellulosic hydrolysate containing mixed C5/C6 sugars and high salt concentration, the pullulan
production by the strain TN2-1-2 was investigated with various concentrations and ratios of glucose and xylose and different salinities. The
results suggested robust pullulan production by TN2-1-2 utilizing glucose and xylose simultaneously. As shown in Fig. 6, based on the robust
and melanin-free pullulan producer TN2-1-2 and the previously developed CBS biocatalyst, we constructed a complete bioprocess to
produce pigment-free pullulan from lignocellulosic biomass effectively.
Thus, this study provided an insight into the cost-effective pullulan
industrial production.

maltotriose, the purity of the produced pullulan was calculated to be
93.7%.
Furthermore, the produced pullulan was verified by 1H-NMR and
13
C-NMR structural analyses (Fig. 8). As indicated in the unidimensional 1H-NMR optical spectrum (Fig. 8A), the proton peak displacements of both the pullulan standard and purified pullulan were distributed between W3.3 and W5.4. Moreover, the anomeric proton at the
site of α-(1→6) linkages was detected based on the chemical shifts at
4.9483 ppm for the pullulan standard and 4.9681 ppm for the produced
pullulan, and the signal distribution at 5.3717 and 5.4090 ppm (produced pullulan) could be attributed to α-(1→4) linkages while the
analogs of pullulan standard were 5.3562 and 5.3928, respectively.
Furthermore, as shown in the 13C-NMR results (Fig. 8B), the signal
distribution of anomeric carbon region of the pullulan standard and
produced pullulan appeared to be consistent, especially for the chemical shifts corresponding to the α-(1→6) linkages (100.5473 ppm for
the pullulan standard and 100.5434 ppm for the produced pullulan)
and α-(1→4) linkages (102.8521 and 102.3890 ppm for the pullulan

Fig. 8. 1H-NMR (A) and

13

Declaration of Competing Interest
The authors declare that they have no competing interests.
CRediT authorship contribution statement
Guanglei Liu: Conceptualization, Data curation, Writing - original
draft, Funding acquisition. Xiaoxue Zhao: Investigation, Data curation.
Chao Chen: Investigation, Data curation. Zhe Chi: Visualization,
Validation, Writing - review & editing. Yuedong Zhang: Visualization,

C-NMR (B) spectra of pullulan standard and the pullulan produced by TN2-1-2.
8

Carbohydrate Polymers 241 (2020) 116400

G. Liu, et al.

Validation. Qiu Cui: Writing - review & editing, Resources. Zhenming
Chi: Writing - review & editing, Resources. Ya-Jun Liu:
Conceptualization, Visualization, Supervision, Writing - review &
editing, Funding acquisition.

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Acknowledgments
This research was supported by the National Natural Science
Foundation of China [grant number 31970069], QIBEBT and Dalian

National Laboratory For Clean Energy (DNL), CAS (Grant number
QIBEBT I201905), Key Laboratory of Biofuels, Qingdao Institute of
Bioenergy and Bioprocess Technology, Chinese Academy of Sciences
(Grant number CASKLB201803), the “Transformational Technologies
for Clean Energy and Demonstration”, Strategic Priority Research
Program of the Chinese Academy of Sciences (Grant Number XDA
21060201), and the Major Program of Shandong Provincial Natural
Science Foundation (Grant Number ZR2018ZB0208).
Appendix A. Supplementary data
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