Kobe University Repository : Thesis · cellulolytic thermophilic bacteria (e.g., Clostridium...
Transcript of Kobe University Repository : Thesis · cellulolytic thermophilic bacteria (e.g., Clostridium...
Kobe University Repository : Thesis
学位論文題目Tit le
Engineering of high-cellulolyt ic Saccharomyces cerevisiae using cell-surface display technique: Towards consolidated bioprocessing(統合型バイオエタノール生産プロセスに資する高効率セルロース分解酵母の開発)
氏名Author Liu, Zhuo
専攻分野Degree 博士(工学)
学位授与の日付Date of Degree 2016-09-25
公開日Date of Publicat ion 2017-09-01
資源タイプResource Type Thesis or Dissertat ion / 学位論文
報告番号Report Number 甲第6745号
権利Rights
JaLCDOI
URL http://www.lib.kobe-u.ac.jp/handle_kernel/D1006745※当コンテンツは神戸大学の学術成果です。無断複製・不正使用等を禁じます。著作権法で認められている範囲内で、適切にご利用ください。
PDF issue: 2021-07-27
DOCTORAL THESIS
Engineering of high-cellulolytic Saccharomyces
cerevisiae using cell-surface display technique:
Towards consolidated bioprocessing
統合型バイオエタノール生産プロセスに資する
高効率セルロース分解酵母の開発
Department of Chemical Science and Engineering
Graduate School of Engineering
KOBE UNIVERSITY
LIU ZHUO July 2016
CONTENTS
Page
Introduction 1
Synopsis 13
Chapter I. 16
Combined cell-surface display- and secretion-based strategies for production of
cellulosic ethanol with Saccharomyces cerevisiae
Chapter II. 43
Engineering of a novel cellulose-adherent cellulolytic Saccharomyces
cerevisiae for cellulosic biofuel production
Chapter III. 72
Efficient ethanol production from crystalline cellulose using high-cellulolytic
Saccharomyces cerevisiae with optimized cellulase ratios on the cell surface
General conclusion 87
References 91
Acknowledgements 104
Publication lists 106
1
INTRODUCTION
Cellulosic biofuels
With the increasing worldwide industrialization, a steep rise for the demand of
petroleum-based fuels appeared. Currently, fossil fuels take up 80% of the primary
energy consumed in the world1, but the sources of these fossil fuels are becoming
exhausted and generating serious environmental issues, such as greenhouse gas
emissions and acid rains. Increasing energy demand also leads to an increase in crude
oil price, directly affected to global economic activity. Thus, many researchers are
focusing on the renewable, environment-friendly, cost-effective energy alternatives
for fossil fuels. In the past decades, biofuels (e.g., ethanol, methanol, biodiesel)
emerge as the most environment-friendly alternative fuels due to their renewability,
security of supply and generating acceptable quality exhaust gases2.
The feedstock for 1st generation biofuel production is starch/ sucrose-rich food
crops, such as corn, wheat, barley, and sugarcane3. Although the 1st generation
biofuels production is commercial in many countries (e.g., corn ethanol in US,
sugarcane ethanol in Brazil), the major obstacle toward its global commercialization
is the concern of “food-versus-fuel” debate4, that whether to use these starch/
sucrose-rich crops as fuel feedstock or as human food. On the contrary,
lignocellulosic biomass (e.g., sugarcane bagasse, newspaper, rice straw) is of great
interest as its low-price, large-scale availability, and no competition with food,
referring as the 2nd generation feedstock for biofuel production. Cellulose is
composed of β-1,4-linked glucose monomers, representing the most abundant
polysaccharide (up to 30-50%) in lignocellulosic biomass. After physical or
thermochemical pretreatments, cellulosic materials are enzymatically decomposed
into monosaccharides (glucose), and subsequently converted to desirable biofuels via
microbial fermentation. However, the extensive hydrogen linkages among glucose
2
molecules lead to a crystalline and strong matrix structure of cellulose 5. As a result, it
requires large amount of costly cellulases to break the recalcitrance of cellulose,
which makes it challenging for cellulose-based biofuels to be economically feasible 6.
Thus, the decrease of the need for enzyme addition may lead to a feasible path
towards the cost-competitive production of cellulosic ethanol.
Consolidated bioprocessing
Conventional technology for lignocellulosic biofuel production includes a
physical/chemical pretreatment step during which polymeric sugar fractions more
accessible to enzyme. Subsequent bioconversion process generally consists of four
steps: the enzyme production, the hydrolysis of cellulose and hemicellulose into
monosaccharides, and the fermentation of sugars into biofuels (Figure 1). Combining
two or more of these steps into one integrated process will greatly decrease the
processing cost. Consolidated bioprocessing (CBP), which allows the four biological
steps occurring in one reactor using a CBP microorganism7, contributes to reducing
the costs in capital investment, substance and utilities associated with microbial
enzyme production. It has been widely considered as the most promising way to
achieve the commercialization of cellulosic ethanol production7.
Figure 1 Schemes of conventional process and CBP process for lignocellulosic biofuel
production
3
The key to CBP is the engineering of a microorganism capable of both producing
saccharifying enzymes to degrade lignocellulosic biomass and efficiently fermenting
sugars into biofuels. Microorganisms used for CBP can be categorized into two
groups: cellulase producers and ethanol producers. Cellulase producers include
cellulolytic thermophilic bacteria (e.g., Clostridium thermocellum), cellulolytic fungal
(e.g., Trichoderma reesei) and some anaerobic filamentous fungi8. These
microorganisms contain high cellulolytic activities, however, their ethanol production
capacities and tolerances to ethanol are relatively low. On the contrary, ethanol
producer group (e.g., Saccharomyces cerevisiae, Zymomonas mobilis, Kluyveromyces
marxianus) are natively possesses superior fermenting capacity and tolerance to
ethanol, but lacks essential cellulolytic activities to degrade cellulose into glucose. To
resolve this problem, researches are working on expressing heterologous cellulases
derived from filamentous fungi or Clostridium species in yeast strains9, 10. Currently,
the production of cellulases in yeast strain follows two major strategies: secreting
enzymes into the fermentation broth or immobilzing enzymes on the cell exterior
using cell-surface display engineering.
Cell-surface display engineering
Cell-surface display engineering is a promising technique that uses microbial
functional components to locate enzymes or peptides on the cell exterior of
microorganisms. This technique endows the engineered strain with novel functions,
such as whole-cell biocatalysts, bio-adsorbents, biosensor, vaccine-delivery vehicles
and screening platforms 11. As shown in Figure 2, cell-surface display engineering has
been applied to the biorefinery of various types of waste materials. In the biorefinery
of sugar-/protein-rich waste to produce biofuels and biochemicals, hydrolytic
enzymes (e.g. cellulase, hemicellulose, and amylase) are immobilized on the surface
of yeast to perform biodegradation and bioconversion of sugars simultaneously. With
4
regard to waste oils, lipase-displaying yeast cells are employed as whole-cell
biocatalysts, which catalyze the transesterification of triglycerides with short-chain
alcohols into biodiesels. Moreover, yeast cells displaying metal-binding proteins on
the cell surface can also be used as bioadsorbents to retrieve heavy metals from
wastewater.
Cell-surface display systems have been successfully developed in various
microorganisms, such as Escherichia coli and Streptococcus gordonii12. Yeast is one
of the most suitable host strains for cell-surface display, because of its rigid cell walls
(around 110-200 nm wild), as well as useful platform for protein production, since
yeast allows the folding and glycosylation of expressed heterologous eukaryotic
proteins. The cell wall of yeast strain S. cerevisiae mainly constitutes cross-linked β-1,
3/1,6-glucans, mannoproteins, and chitin. Although the β-1,6-glucan takes up
quantitatively a minor component of the cell wall, it plays a central role in anchoring
cell-wall protein. Glycosylphosphatidylinositol (GPI) proteins are kind of
glucanase-extractable cell-wall proteins that are linked with β-1,6-glucan via the GPI
anchor, and play critical roles in the structural rigidity of cell walls (e.g., Cwp2p and
Sed1p) and cell functions (e.g., Flo1p for cell flocculation)13. During the secretory of
a GPI protein, GPI anchor is bound onto GPI-attachment signal on the membrane of
endoplasmic reticulum; while secretion signal directs GPI protein transported to cell
surface. On reaching plasma membrane, GPI protein is cleaved at GPI-anchor site by
a phosphatidylinositol-specific phospholipase C and subsequently released into cell
wall. GPI protein is then covalently bound with β-1,6-glucan via remnant
GPI-anchor13. Yeast cell-surface display technique is to fuse the targeted enzyme or
peptide to a GPI protein/ GPI anchoring domain, and this is anchored covalently on
the cell surface via the GPI anchor14. The characteristics and applications of GPI
5
anchoring domains in yeast strains have been summarized in Table 1.
Anchoring of enzymes or peptides on the cell surface to perform cellulose
degradation has the following advantages: (1) Proteins are produced and
auto-immobilized on the cell surface via easy cellpropagation, which can reduce the
expenditure and facilities needed for protein generation and enrichment37. (2) The
proteins that are anchored on solid surfaces (such as cell surface) are more stable than
free-form proteins under extreme conditions (e.g., high temperature and organic
solvents)38. Improved enzyme stability will facilitate long-term storage and recycling
in industrial processes. (3) Tethering of multiple synergetic enzymes on a single cell
significantly shortens the enzyme-to-enzyme distance16, preventing long-distance
mass transfer of substrates, especially in high-solid fermentation. (4) Monomers (e.g.,
glucose, xylose, and amino acid) that are liberated via enzymatic hydrolysis can be
immediately utilized by cells because of the proximity of the enzyme to cell surface.
As a result, the monomers in the extracellular environment are maintained at low
concentrations, which significantly reduce the risk of contamination (especially when
glucose is released from cellulose) and diminishes the repression effects of substrates
on microorganism. (5) The proteins that are immobilized on the cell surface can be
easily recollected through centrifugation or filtration, implying high potentials of
recycling.
6
7
Was
te S
ugar�
Was
te O
il�
Cel
lulo
se�
Cel
lula
se�
Prot
ease�
Lip
ase�
Met
al-b
indi
ng p
rote
in�
Anc
hor
prot
ein�
Pept
ide�
Yeas
t cel
l�
Glu
cose�
2Pyr
uvat
e�
2Ace
tald
ehyd
e�E
than
ol�
Was
te P
rote
in�
Was
te W
ater�
Met
al io
n�
Abs
orpt
ion� R
ecov
ery�
Cel
l mas
s�
Fatt
y ac
id a
lkyl
est
ers�
Methanol�
Tria
cylg
lyce
ride� Glycerol�
Am
ino
acid�
Glu
tath
ione
Figu
re 2
Sch
emes
for v
ario
us fo
rms o
f was
te b
iore
finer
y us
ing
cell-
surf
ace
disp
lay
engi
neer
ing.
8
Applications of cell-surface display technique in cellulosic ethanol production
Currently, cell-surface display technique has been widely applied in the ethanol
production from cellulosic materials. In order to degrade cellulose into glucose, at
least three kinds of synergistic cellulases are required, including β-glucosidase (BGL),
endoglucanase (EG), and cellobiohydrolase (CBH)8. Display of heterologous
synergistic cellulases derived from filamentous fungi or Clostridium species on the
surface of yeast enables it to directly generate biofuels from cellulosic substrates. As
shown in Table 2, Fujita et al. (2002) and Yanase et al. (2010) showed that BGL- and
EG-displaying yeast strains could efficiently convert β-glucan (linear, water-soluble
polysaccharides composed of six or seven β-1, 4-linked glucose residues) into ethanol,
with a high yield of more than 90% of the theoretical level. In comparison, phosphoric
acid swollen cellulose (PASC) is an insoluble cellulosic material mainly composed of
amorphous cellulose, and more resistant to enzymatic hydrolysis than β-glucan.
Although it has been demonstrated that cellulase-displaying yeasts are able to directly
produce ethanol from PASC, most of their ethanol yields were less than 60% (Table
2). This is because the cellulolytic activities on the cell surface are insufficient to
support the complete degradation of PASC.
In addition, as shown in Table 2, the current cellulase-displaying yeast strains
seem incapable of direct degrading lignocellulosic substrates in the absence of
exogenous cellulases, owing to the recalcitrance of lignocellulose. A recombinant S.
cerevisiae co-displaying BGL, EG and CBHII achieved 89% of the theoretical ethanol
yield from rice straw in the presence of 10 FPU/g-biomass commercial cellulase,
while it obtained an ethanol yield of 65% under 5 FPU/g-biomass cellulase39,
indicating that the cellulosic ethanol yield is strongly related to the support of
additional cellulase. As for natural lignocellulosic materials, crystalline cellulose
serves as the major form of cellulose. The highly rigid structures of crystalline
9
cellulose are resistant to enzymatic depolymerization, making it more difficult to be
degraded than the fluffy cellulosic materials (e.g., β-glucan and PASC). Thus,
improvement of the degradation ability towards crystalline cellulose is of great
importance in achieving CBP using cell-surface display engineering.
To solve these problems, much effort has been devoted to increasing the
enzyme-display efficiency or improving cellulase production. In order to alleviate the
conflict between cell-wall protein and displayed-protein, Kotaka et al. disrupted the
gene encoding cell-wall protein SED1, obtaining a 1.6-fold increase of enzyme
activity on the cell surface 40. In addition, the genes involved in the protein secretory
pathway of S. cerevisiae were over-expressed, achieving a 71% increase in CBH
production 41. However, these improvements only result in a limited increase of
cellulose degradation ability, which are still insufficient for the hydrolysis of
crystalline cellulose. Therefore, in the present study, three novel strategies have been
employed to improve the cellulose degradation ability of cellulase-displaying S.
cerevisiae: study of the most suitable strategy for cellulase production (Chapter I),
improvement of interactions between cell and cellulose (Chapter II), and optimization
of enzyme ratios that displayed on the cell surface (Chapter III).
Firstly, we investigated the suitable strategy for cellulase production from the
view of steric hindrance. The location of cellulase (immobilized on the cell surface or
secreted into medium) is of great importance in enzyme synergism and the mass
transfer of intermediate products. It has been reported that BGL is preferable to be
displayed on the cell surface rather than secreted because of the improved stability 38.
However, "display" systems may suffer from inefficiency of processive enzymes (e.g.,
CBH1 and CBH2), or cause steric restriction in the collision with cellulose. To date,
the best locations (or production strategies) for EG and CBH remained obscure. Thus,
10
in this study, EG and CBH1 were produced heterologously in a BGL-displaying S.
cerevisiae strain using cell-surface display, secretion, or a combined strategy. The
most suitable strategy for producing EG and CBH1 for cellulose degradation was
evaluated. (described in Chapter I)
Secondly, we investigated the interactions between cellulase-displaying cells and
cellulose during cellulose degradation process. Currently, large amounts of researches
have been pursued to improve the cellulolytic activities on yeast cell surface through
delicate design of surface-displayed scaffoldin42, increasing enzyme diversities43, or
increasing the expression level of cellulase genes. However, very limited reports
could provide the information of cellulose degradation mechanism via
surface-displayed cellulase, which correlates with the interactions of
enzyme-to-cellulose and cell-to-cellulose. In this study, we focus on the
cellulose-adherent characteristics of cellulase-displaying cells that could be used to
improve the cellulose hydrolysis efficiency, such as enhancement of cell-to-cellulose
interactions via altering the surface properties of cellulose. (described in Chapter II)
Thirdly, we engineered a recombinant yeast strain with high degradation ability
towards crystalline cellulose through optimizing of the enzyme ratio on the cell
surface. It is postulated that improvement of the crystalline-cellulose degradation
ability in yeast cell will enable it direct utilization of natural cellulosic materials. Most
current reports of cellulase-displaying yeast strain focus on enhancing the degradation
potential of amorphous cellulose (e.g., PASC) (Table 1). However, the enzymatic
hydrolysis pattern of crystalline cellulose, especially the optimal ratios of synergisitic
cellulases, is completely different from that of amorphous cellulose. It has been
reported that engineering of EG as the main proportion in total cellulase mixture is
preferable for amorphous cellulose degradation, while using CBHs as the major
11
component is to be assumed more effective in the degradation of crystalline
cellulose44. In the present study, we integrated multiple-copy of cellulase genes (EG,
CBH1, CBH2) into the genome of BGL-displaying yeast strain using cocktail
δ-integration method44. The transformants were screened based on their degradation
abilities toward Avicel. Moreover, the enzyme ratios that displayed on the cell surface
were determined via nano LC-MS/MS. (described in Chapter III)
12
13
SYNOPSIS Chapter I
Combined cell-surface display- and secretion-based strategies for production of
cellulosic ethanol with Saccharomyces cerevisiae
The production of cellulase is usually pursued by one of the two strategies:
displaying enzyme on the cell surface or secreting enzyme into the medium. However,
to our knowledge, the combination of the two strategies in a yeast strain has not been
employed. In this chapter, heterologous endoglucanase (EG) and cellobiohydrolase 1
(CBH1) were produced in a β-glucosidase (BGL) -displaying S. cerevisiae strain
using cell-surface display, secretion, or a combined strategy. Strains EG-D-CBH1-D
and EG-S-CBH1-S (with both enzymes displayed on the cell surface or with both
enzymes secreted to the surrounding medium) showed higher ethanol production (2.9
g/L and 2.6 g/L from 10 g/L phosphoric acid swollen cellulose, respectively) than
strains EG-D-CBH1-S and EG-S-CBH1-D (with EG displayed on cell surface and
CBH1 secreted, or vice versa). After 3-cycle repeated-batch fermentation, the
cellulose degradation ability of strain EG-D-CBH1-D remained 60% of the 1st batch,
at a level that was 1.7-fold higher than that of strain EG-S-CBH1-S. This work
demonstrated that placing EG and CBH1 in the same space (on the cell surface or in
the medium) was favorable for amorphous cellulose-based ethanol fermentation. In
addition, the cellulolytic yeast strain that produced enzymes by the cell-surface
display strategy performed better in cell-recycle batch fermentation compared to
strains producing enzymes via the secretion strategy.
14
Chapter II
Engineering of a novel cellulose-adherent cellulolytic Saccharomyces cerevisiae
for cellulosic biofuel production
The main obstacle toward the economic feasibility of cellulosic bioethanol
production is the recalcitrance of lignocellulose requiring large amount of costly
enzyme to break. To reduce the need for enzyme addition, we successfully engineered
a high-efficiency cellulolytic Saccharomyces cerevisiae by displaying four synergistic
cellulases (BGL, EG, CBH1, and CBH2) on the cell surface. The cellulase-displaying
yeast strain exhibited clear cell-to-cellulose adhesion and a “tearing” cellulose
degradation pattern; the adhesion ability correlated with enhanced surface area and
roughness of the target cellulose fibers, resulting in higher hydrolysis efficiency. The
engineered yeast could directly produced ethanol from rice straw despite a more than
40% decrease in the required enzyme dosage for high-density fermentation. Thus,
improved cell-to-cellulose interactions provided a novel strategy for increasing
cellulose hydrolysis, suggesting a mechanism for promoting the feasibility of
cellulosic biofuel production.
15
Chapter III
Efficient ethanol production from crystalline cellulose using high-cellulolytic
Saccharomyces cerevisiae with optimized cellulase ratios on the cell surface
Crystalline cellulose is the most rigid, degradation-resistant structure in natural
cellulosic materials, and is widely considered as the major obstacle to achieve the
efficient utilization of lignocellulose. To improve the degradation ability towards
crystalline cellulose in cellulolytic yeast strain, the optimal ratio of synergisitic
cellulases (BGL, EG, CBH1, and CBH2) displayed on the cell surface was
investigated in this chapter. We constructed a pool of cellulase-displaying yeast strain
with various enzyme ratios through cocktail δ-integration method. Strain A26, which
achieved the highest cellulosic ethanol yield (57%, corresponding to 1.46-fold of that
in control strain (cellulase ratio is 1:1:1:1)) without any metabolic burden on cell
growth was screened from the yeast pool. The cellulase ratios on the cell surface were
analyzed using nano LC-MS/MS. To our knowledge, this is the first work on relative
quantitation of four kinds of cellulase (BGL, EG, CBH1, and CBH2) anchored on
single yeast cell, and the first to optimize enzyme ratio towards crystalline cellulose
degradation.
16
Chapter I
Combined cell-surface display- and secretion-based strategies for
production of cellulosic ethanol with Saccharomyces cerevisiae
17
Introduction
Due to the limitations in fossil fuel supplies and environmental issues, bioethanol
derived from lignocellulosic materials has recently gained increased attention1, 5.
Saccharomyces cerevisiae is the most commonly used microorganism for ethanol
production, but lacks essential cellulolytic enzyme activities to degrade cellulose into
glucose52. To resolve this problem, the construction of recombinant yeast strains
capable of producing heterologous cellulases, including β-glucosidase (BGL),
endoglucanase (EG), and cellobiohydrolase (CBH), has been pursued over the last
two decades10, 39.
Currently, the production of cellulases follows two major strategies: displaying
enzymes on the cell surface or secreting enzymes into the fermentation broth. The
glycosylphosphatidylinositol (GPI) anchoring system enables the display of various
kinds of enzymes on the cell surface14. The cell-surface display strategy increases the
effective concentration of enzymes, and promotes a greater degree of synergy16. In
addition, glucose liberated from cellulose in proximity to the cell surface is
immediately taken up, thereby minimizing the risk of contamination or product
inhibition53. Furthermore, immobilizing enzymes on the cell surface enables the
re-use of enzymes and cells in multi-batch fermentations, which reduces the cost of
yeast propagation and that of supplementation with extraneous enzymes54, 55. In
contrast, secreting enzymes into the medium recreates the “free enzyme system”,
which is similar to the cellulase system of filamentous fungi. The quantity of secreted
enzymes is limited only by the production capacity of cells, not by physical
restrictions, such as the incorporation capacity of yeast cell wall associated with
cell-surface display56. Moreover, free cellulases can penetrate into the secondary cell
walls of plant biomass57, increasing the accessibility of cellulose, which was reported
as the critical factor in enzymatic hydrolysis58.
18
Thus, each strategy has both advantages and disadvantages. The selection of an
optimal strategy for enzyme production should be based on the characteristics of a
given enzyme and its reaction mechanism. It has been reported that
cell-surface-displayed BGL exhibited higher efficiency in cellobiose usage than
secreted BGL because of the improved stability caused by immobilization on the cell
wall38. However, “display” systems may suffer from inefficiency of processive
enzymes (e.g., CBH), leading to decreased hydrolysis efficiency compared to free
enzyme systems56. Thus the combination of cell-surface display and secretion
strategies into one recombinant yeast strain was expected to achieve improved
hydrolysis of cellulose compared to either single strategy of displaying or secreting
cellulases. Such a combined strategy may allow the various kinds of enzymes to be
produced in their most appropriate location, assembling the advantages of the two
strategies into one system of enzyme production.
Cellobiohydrolase I (CBH1) is the major component (~ 60%) of the total
cellulolytic protein of the cellulase system of Trichoderma reesei7. CBH1 acts by
hydrolyzing from the reducing end of crystalline cellulose fibers in a progressive
manner. Recently, CBH1 was reported as the main contributor to overall cellulose
degradation, with other enzymes synergistically enhancing its hydrolytic efficiency59.
Although CBH1 has been heterologously expressed and secreted in S. cerevisiae, the
relatively low titer60, 61 and low specific activity62 of secreted CBH1 has limited the
study of co-expression of CBH1 with other cellulolytic enzymes. Recently, CBH1
originating from Talaromyces emersonii fused with the T. reesei C-terminal
carbohydrate-binding module (CBM) was efficiently expressed in S. cerevisiae with a
yield of 100-200 mg/L, which is approximately 20-fold higher than the expression
levels of T. emersonii CBH1 reported elsewhere63. In separate work, immobilization
19
of enzyme on the cell surface was reported to improve the stability of the enzyme38.
However, to our knowledge, there has been no previous report of displaying CBH1 on
the cell surface of yeast strain.
In the present study, EG and CBH1 were produced heterologously in a
BGL-displaying S. cerevisiae strain using cell-surface display, secretion, or a
combined strategy. The most suitable strategy for producing EG and CBH1 for
cellulose degradation was evaluated. Direct conversion of cellulose into ethanol was
conducted by cellulolytic yeast strains and then applied in cell-recycle batch
fermentation for further evaluation. To our knowledge, the work reported here is the
first study on displaying CBH1 on the yeast cell surface and the first study on the
feasibility of combining the cell-surface display and secretion strategies in one yeast
strain for heterologous cellulase production. We believe that this work will
significantly increase our knowledge of how to engineer optimal yeast strains for
biofuel production from cellulosic biomass.
Materials and methods
Microbial strains and media
The relevant features and sources of the yeast strains used in this study are listed
in Table I-1. Strain Escherichia coli NovaBlue (Novagen, Inc., Madison, WI, USA)
was used for the propagation of the plasmids. Bacterial cells were grown at 37 °C in
Luria-Bertani broth (10 g/L tryptone, 5 g/L yeast extract, and 5 g/L sodium chloride)
containing 100 mg/L ampicillin. Haploid yeast S. cerevisiae BY4741 (Life
Technologies, Carlsbad, CA, USA) was used for cellulase production. Yeast
strainswere screened and pre-cultivated in synthetic dextrose (SD) medium (6.7 g/L of
yeast nitrogen base without amino acids (Difco Laboratories, Detroit, MI, USA) and
20 g/L of glucose) supplemented with the appropriate amino acids in a shaker
20
21
incubator (150 rpm) at 30 °C, and then aerobically cultivated at 30 °C in YPD
medium (20 g/L peptone (Bacto-PeptoneTM, Difco Laboratories), 10 g/L yeast extract
and 20 g/L glucose). Ethanol fermentation was performed in YP medium (10 g/L
yeast extract and 20 g/L peptone) containing either 10 g/L PASC or 10 g/L β-glucan
from barley (Megazyme, Bray, Ireland). PASC was prepared from Avicel PH-101
(Fluka Chemie GmbH, Buchs, Switzerland), as previously described 9.
Plasmid and strain construction
The plasmids and primers used in this study are summarized in Table I-2 and
Table I-3, respectively. To construct the plasmid pRDH225, the gene T. reesei EG2
was cloned as a 1277 bp PacI/AscI fragment from pRDH14764 into pBZD265 to
form pRDH225. To construct pRDH226, the T. emersonii CBH1 encoding gene
containing a domain encoding a carboxy-terminal CBM originating from the T. reesei
CBH1 was amplified using Phusion hi-fidelity polymerase (Thermo Scientific) as
directed by the manufacturer from pMI529 63 as template with primers TeCBH1-L
and TeCBH1-R. The resulting 1567 bp fragment was cloned as a PacI/AscI fragment
into pBZD2 to form pRDH226.
The integrative plasmids for cell-surface display with the SED1 anchor were
constructed as follows: the DNA fragment encoding EG2 from T. reesei was
amplified from plasmid pRDH225 by PCR using the primers TrEG2-F and TrEG2-R.
The cell-surface display cassette, which includes the I2 region (the 3’ non-coding
region between gene YFL021W and YFL020C, used for integration), LEU2, SED1
promoter, SED1 anchoring region, and SAG1 terminator was amplified from plasmid
pIL2GA-SS using primers P-EG2 and EG2-A. Two DNA fragments were connected
by the isothermal assembly method66, generating the plasmid pIL2-EGD. To construct
the secretion expression cassette without the SED1 anchoring region, amplification
22
was performed using the plasmid pIL2GA-SS as a template with primers P-EG2 and
EG2-T. Primers TrEG2-F and TrEG2-R2 were used for amplifying the fragment
encoding EG. The resulting plasmid from the combination of two DNA fragments
was named pIL2-EGS.
Table I-2 Characteristics of the integrative plasmids used in this study
Plasmid Relevant features References
pRDH225 KanMX, expression of T. reesei EG2 gene This study
pRDH226 ZeoR, expression of T. emersonii CBH1 gene This study
pIL2GA-SS LEU2, display of R. oryzae glucoamylase [67]
pIU5GA-SS URA3, display of R. oryzae glucoamylase [67]
pIBG-SS HIS3, display of A. aculeatus BGL1 [25]
pIL2-EGD LEU2, display of T. reesei EG2 This study
pIL2-EGS LEU2, secretion of T. reesei EG2 This study
pIU5-CBH1D URA3, display of T. emersonii CBH1 This study
pIU5-CBH1S URA3, secretion of T. emersonii CBH1 This study
R. oryzae, Rhizopus oryzae; A. aculeatus, Aspergillus aculeatus; T. reesei, Trichoderma
reesei; T. emersonii, Talaromyces emersonii; BGL1, β-glucosidase 1; EG2, endoglucanase 2;
CBH1, cellobiohydrolase 1
The construction of CBH1 integrative plasmids was performed by a process
similar to the above description. For the cell-surface display plasmid, the DNA
fragment of CBH1 from T. emersonii was amplified from plasmid pRDH226 using
primers TeCBH1-F and TeCBH1-R, and fused with the PCR product amplified from
plasmid pIU5GA-SS by primers P-CBH1 and CBH1-A. The resulting plasmid, which
was named pIU5-CBH1D, can integrate into I5 region (the 3’ non-coding region of
gene YLL055W and YLL054C). Plasmid pIU5-CBH1S is the integrative plasmid with
the secretion expression cassette, connected by the segment of CBH1-encoding gene
(primes TeCBH1-F and TeCBH1-R2) and the PCR products amplified from plasmid
23
pIU5GA-SS (primers P-CBH1 and CBH1-T for secretion expression cassette).
Table I-3 The primers used in this study
Plasmids were transformed into S. cerevisiae BY4741 using lithium acetate as
described68. The transformants were identified using colony PCR to check the
integration of the cellulase gene expression cassettes (primers I2-F and I2-R for I2
insertion of EG-encoding cassette, and primers I5-F and I5-R for I5 insertion of
CBH1-encoding cassette). Transformants with one copy of the cassette were selected
for subsequent experiments.
Primers Sequence (5’-3’)
TeCBH1-L GACTTTAATTAAAATGCTAAGAAGAGCTTTACTATTG
TeCBH1-R GACTGGCGCGCCTTACAAACATTGAGAGTAGTATGGG
TrEG2-F AATACGTTCGCTCTATTAAGATGAACAAGTCTGTTGCTCCATTG
TrEG2-R GTTGATAATTTACTCGAGCCTAACTTTCTAGCCAAACATGAAGAAACC
TrEG2-R2 CTCAATGTACTAACTGTACATTATAACTTTCTAGCCAAACATGAAGAAAC
TeCBH1-F AATACGTTCGCTCTATTAAGATGCTAAAGAAGAGCTTTACTATTGAGC
TeCBH1-R GTTGATAATTTACTCGAGCCCAAACATTGAGAGTAGTATGGGTTT
TeCBH1-R2 CTCAATGTACTAACTGTACACTACAAACATTGAGAGTAGTATGGGTTT
P-EG2 GGAGCAACAGACTTGTTCATCTTAATAGAGCGAACGTATTTT
EG2-A CATGTTTGGCTAGAAAGTTAGGCTCGAGTAAATTATCAACTGTCC
EG2-T GTTTGGCTAGAAAGTTATAATGTACAGTTAGTACATTGAGTCTAAATA
P-CBH1 AGTAAAGCTCTTCTTAGCATCTTAATAGAGCGAACGTATTTT
CBH1-A CATACTACTCTCAATGTTTGGGCTCGAGTAAATTATCAACTGTCC
CBH1-T ACTACTCTCAATGTTTGTAGTGTACAGTTAGTACATTGAGTCTAAATA
I2-F GAAGCCGCGAGTACGAACAATGATG
I2-R TGGTATTTTCGTGAGCAAACCCAAC
I5-F CATTGAAGAAGGGAAAGTGGTAACC
I5-R TCCCTCTCTAATCTGGGTGAGAC
rt-ACT1-F TGGATTCCGGTGATGGTGTT
rt-ACT1-R TCAAAATGGCGTGAGGTAGAGA
rt-EG-F GGTTGTTTGTCTTTGGGTGCTTAC
rt-EG-R AATTGAGCATTTGTTGGACCACCTT
rt-CBH1-F CAACTTACTGTCCAGACGACGAAAC
rt-CBH1-R AAGGAAGAACCAGAGGAGGTAACAC
24
Quantification of the transcription level of cellulase-encoding genes by real-time
PCR
The transcription levels of the cellulase- encoding genes were quantified by
real-time PCR as described previously69. The PCR primers BGL 761F and BGL
858R44 were used to determine the transcription level of gene BGL1. Primers rt-EG-F
and rt-EG-R were used for the EG2 gene and primers rt-CBH1-F and rt-CBH1-R
were used for the CBH1 gene. Transcription levels of the target genes were
normalized to the housekeeping gene ACT1 (primers rt-ACT1-R and rt-ACT1-F).
Yeast cell growth assay
To measure cell growth, the parent strain and the engineered strains were
cultivated individually in SD medium at 150 rpm for 24 h at 30 °C. The pre-cultured
medium was inoculated into 5 mL YPD medium in a L-shaped vitreous tube at the
initial OD660 of 0.05 and cultivated at 30 °C. The value of the OD660 was measured
once hourly using a TVS062CA Bio-photorecorder (Advantec Toyo, Tokyo, Japan).
The value of the OD660 was taken as an indicator of cell growth.
Ethanolic fermentation
Recombinant yeast strains were pre-cultivated in SD medium for 24 h, then
inoculated into YPD medium and aerobically cultured in YPD medium at 30 °C for
72 h. Cells were harvested by centrifugation at 3,000 × g for 10 min at 4 °C, and then
washed twice with sterile distilled water. The wet cell pellet was weighed and then
resuspended in 20 mL YP medium containing 10 g/L PASC or β-glucan from barley
at an initial cell concentration of 150 g wet cells/L (PASC) or 50 g wet cells/L
(β-glucan). Ethanol fermentation was performed at 37 °C for 96 h with 200 rpm
agitation in 100 mL closed bottles, each equipped with a siliconized tube and check
valve (Sanplatec Corp., Osaka, Japan) as a CO2 outlet under the oxygen-limited
25
conditions. The ethanol concentration in the fermentation medium was determined
using a gas chromatograph (model GC-2010; Shimadzu, Kyoto, Japan), as described
previously48.
In the cell recycle batch fermentation, after the 96-h batch fermentation
described above, cells were collected by centrifugation at 8,000 × g for 10 min at 4 °C.
The pelleted cells were inoculated into fresh YP medium supplemented with 10 g/L
PASC. The fermentation was repeated three times sequentially under the
oxygen-limited conditions.
To measure PASC amount in fermentation, the fermentation broth (including the
cells and residual PASC) was sterilized at 121 °C, 20 min (to terminate glucose
consumption by yeast cells) and then cooled to room temperature. Sterilized medium
was incubated with 3 FPU/g-biomass PASC commercial cellulase (Cellic CTec2;
Novozymes Inc., Bagsvaerd, Denmark) for 2 h at 50 °C. After the hydrolysis reaction,
the supernatant was obtained by centrifugation at 8,000 × g, 10 min, 4 °C. Glucose
concentration in the supernatant was measured by the Glucose CII kit (Wako Pure
Chemical Industries, Ltd., Osaka, Japan) and taken as the amount of PASC remnant.
Enzyme assay
At the 0-h and 96-h time points of ethanol fermentation, fermentation medium
was assayed for PASCase, and individual cellulase activities. PASCase activity
represents the PASC degradation ability of all enzymes present. Fermentation broth
was added into a final concentration of 5 g/L PASC in 50 mM sodium citrate buffer
(pH 5.0) and 100 mM methyl glyoxal (Nacalai Tesque, Inc., Kyoto, Japan); the
methyl glyoxal prevents the assimilation of glucose by yeast cells70. The reaction was
performed at 50 °C for 4 h using a heat block (Thermo Block Rotator SN- 06BN;
26
Nissin, Tokyo, Japan) with shaking at 35 rpm, and the supernatant was collected by
centrifugation for 10 min at 8,000 × g at 4 °C to remove cells and debris. The amount
of glucose in the supernatant was measured by the Glucose CII kit. One unit of
PASCase activity (U/mL) was defined as the amount of enzyme needed to produce 1
µmoL of glucose per minute at 50 °C, pH 5.0.
The medium of PASC fermentation was used for the BGL, EG, and CBH1
activity assays. The BGL and EG activities were determined as previously described25.
One unit of the BGL activity was defined as the enzyme amount required for
production of 1 µmoL p-nitrophenol (pNP) in 1 min at 30 °C (U/mL). One unit of EG
activity was defined as the absorption at 590 nm of released blue dye in 1 h at 38 °C
(U/mL). p-nitrophenyl-β-lactopyranoside (pNPL, Sigma Co. Ltd, St. Louis, MO, USA)
was used for the measurement of CBH1 activity as previously described71. One unit of
CBH1 activity (U/mL) was defined as the enzyme amount required for production of
1 µmoL pNP in 1 min at 50 °C.
Results
Construction of yeast strains
In this study, the haploid yeast strain S. cerevisiae BY4741 was used as the host
strain for the heterologous expression of cellulase genes. The plasmids containing
gene expression cassettes are listed in Table I-2. All the gene expression cassettes
included the SED1 promoter and the SAG1 terminator. The secretion signal peptide of
BGL1 was derived from Rhizopus oryzae glucoamylase, while EG2 and CBH1 were
produced with their native secretion signals. The GPI-anchoring region used for
cell-surface display was constructed using the full length S. cerevisiae SED1 gene to
display cellulases on the cell surface. It has been reported that the combination of
SED1 promoter and SED1 anchoring region in a gene cassette enables highly efficient
27
immobilization of enzyme into the cell wall25. Each cellulase gene was integrated into
a separate internal open reading frame (ORF) region (I2 region for EG2 gene and I5
region for CBH1 gene) and confirmed by polymerase chain reaction (PCR). Figure I-1
shows the cellulase production scheme of the recombinant yeast strains constructed in
this study. Strain BY-BG-SS, which displayed Aspergillus aculeatus BGL1 on the
cell surface, was reported previously25. The T. reesei EG2 gene expression cassettes,
with and without the SED1 anchoring region, were integrated into the genome of
strain BY-BG-SS to yield strains EG-D and EG-S, respectively. Next, the expression
cassettes of the T. emersonii CBH1 gene, with and without the SED1 anchoring region,
were integrated into the genome of strain EG-D to yield EG-D-CBH1-D and
EG-D-CBH1-S or into the genome of strain EG-S to yield EG-S-CBH1-D and
EG-S-CBH1-S. The engineered yeast strains in this study are listed in Table I-1.
Figure I-1 Schematic description of the recombinant yeasts strains constructed in this study.
In addition, the transcription levels of cellulase genes were determined after 72 h
of cultivation. The transcription levels of the BGL1, EG2, and CBH1 genes, each of
28
which were under the control of a SED1 promoter, were similar among all
transformants (Figure I-2).
Figure I-2 Relative transcription levels of cellulolytic enzyme-encoding genes in
recombinant yeast strains. Gene ACT1 was used as the internal standard. The relative
transcription levels were shown normalized to the level observed in strain EG-D-CBH1-D,
whose relative transcription level was defined as 1. For each strain, data are presented as the
mean ± SD from three independent experiments.
Effect of multiple gene expression on cell growth
Cell growth was profiled to determine the metabolic burden caused by the
expression of heterologous cellulase genes. Each of the engineered strains was
inoculated into liquid YPD media and cultivated aerobically for 72 h at 30 °C. The
host strain S. cerevisiae BY4741 was used as a reference strain. As shown in Figure
I-3, no apparent difference in cell growth was observed between the host strain and
any of the recombinant yeast strains.
29
Figure I-3 Time-course profiles of cell growth using host strain BY4741 and recombinant
yeast strains in YPD medium. Each strain was inoculated in YPD medium to an initial OD660
of 0.05 and then cultured aerobically at 30 °C, 150 rpm for 72 h. For each strain, data are
presented as the mean ± SD from three independent experiments.
Direct ethanol production from cellulosic materials
Barley β-glucan and phosphoric acid swollen cellulose (PASC) were utilized as
fermentation substrates. β-glucan is a linear, water-soluble polysaccharide composed
of 6 or 7 β-1,4-linked glucose residues49. PASC, which is derived from phosphoric
acid treatment of Avicel PH-101, is an insoluble cellulosic material with more
amorphous regions and a lower degree of crystallinity compared to Avicel72, 73.
As depicted in Figure I-4A, ethanol production from 10 g/L β-glucan was
performed using strains EG-D and EG-S. Yeast strains were cultivated in YPD
medium for 72 h; cells then were collected by centrifugation and inoculated into
fermentation medium at an initial cell concentration of 50 g wet cells/L. The ethanol
fermentation was conducted under oxygen-limited conditions at 37 °C for 24 h.
30
Ethanol production by strain EG-D initiated immediately after the start of
fermentation and reached a maximum of 4.1 g/L after 6 h of fermentation. In contrast,
strain EG-S exhibited a long lag phase before the start of ethanol production; no
ethanol was detected until 9 h of fermentation.
Figure I-4 Time course of direct ethanol production from cellulosic materials by recombinant
strains EG-D and EG-S. (A) Ethanol production from β-glucan. (B) Ethanol production from
PASC. For each strain and time point, data are presented as the mean ± SD from three
independent experiments.
The fermentation abilities of EG-D and EG-S also were evaluated by performing
direct ethanol production from 10 g/L PASC (Figure I-4B). The fermentation was
conducted under oxygen-limited conditions at 37 °C for 96 h with an initial cell
concentration of 150 g wet cells/L. Ethanol production by strain EG-D peaked at 1.5
g/L ethanol at 72 h, while the production by strain EG-S peaked at 0.6 g/L at 48 h.
These results revealed that locating EG on the cell surface improved the ethanol
production from both soluble and insoluble cellulosic materials.
31
Figure I-5 Time course of direct ethanol production from PASC by recombinant strains
EG-D-CBH1-D, EG-D-CBH1-S, EG-S-CBH1-D, and EG-S-CBH1-S. For each strain and
time point, data are presented as the mean ± SD from three independent experiments.
To investigate the most suitable strategy for EG and CBH1 production, direct
ethanol production from 10 g/L PASC was evaluated using recombinant strains
EG-D-CBH1-D, EG-D-CBH1-S, EG-S-CBH1-D, and EG-S-CBH1-S. As shown in
Figure I-5, after 96 h of fermentation, ethanol production by strains EG-D-CBH1-D
and EG-S-CBH1-S peaked at 2.9 g/L and 2.6 g/L, respectively. Ethanol production by
strain EG-D-CBH1-S peaked at 2.3 g/L at 96 h while strain EG-S-CBH1-D peaked at
1.2 g/L at 24 h. To further characterize the fermentation capacity of our constructs,
the strains were compared by evaluating the PASCase and individual cellulase
enzyme activities in PASC fermentation, and by testing the strains in cell-recycle
batch fermentation.
Enzyme activity in direct ethanol production from PASC
The cellulose degradation ability of cellulolytic strains is considered as one of
the critical factors in the conversion of cellulose into ethanol. In this study, PASCase
32
Figure I-6 PASCase activities and cellulase activities of the cellulolytic S. cerevisiae strains
in PASC fermentation. (A) PASCase activity at 0 h of the fermentation. (B) PASCase activity
at 96 h of the fermentation. (C) Cellulase activity at 0 h of the fermentation. (D) Cellulase
activity at 96 h of the fermentation. For each strain and time point, data are presented as the
mean ± SD from three independent experiments.
33
activity represents the PASC-degradation capability of the cellulolytic yeast strains.
The PASCase activity at 0 h and 96 h of ethanol production from 10 g/L PASC was
investigated. As shown in Figures I-6A and I-6B, PASCase activity was highest
(among the four recombinant yeast strains) in EG-D-CBH1-D at both 0 h (50.9
mU/mL) and 96 h (53.6 mU/mL). The PASCase activity of strain EG-S-CBH1-S
increased from 26.9 mU/mL at 0 h to 47.8 mU/mL at 96 h. By contrast, the PASCase
activity of strain EG-S-CBH1-D decreased after 96 h of fermentation, exhibiting the
lowest activity (34.1 mU/mL) compared with other recombinant yeast strains. The
activity of BGL, EG, and CBH1 from yeast strains at 0 h and 96 h of PASC
fermentation also was determined, as illustrated in Figures I-6C and I-6D. BGL
activity was similar among the four recombinant strains and remained similar (within
a given strain and among different strains) during the fermentation. EG activity at 96
h was highest in strain EG-D-CBH1-D (24.2 U/mL) when compared with that in the
other cellulolytic yeast strains. Initial (0 h) CBH1 activity was highest in strain
EG-D-CBH1-D (4.7 U/mL), and lowest in strain EG-S-CBH1-S. After 96 h of
fermentation, CBH1 levels appeared to rise similarly in the four strains, achieving
activities of ~ 7-10 U/mL.
Cell-recycle batch fermentation
To investigate the efficiency of strains EG-D-CBH1-D and EG-S-CBH1-S in a
continuous process, cell-recycle fermentation was conducted under anaerobic
conditions (Figure I-7). Recombinant yeast cells were collected for recycling and 10
g/L PASC was added into the fermentation medium at the beginning of each run. In
the first batch, 69.3% and 55.9% of PASC (corresponding to 6.9 g/L and 5.6 g/L
PASC) was converted into 2.9 g/L and 2.5 g/L ethanol by strain EG-D-CBH1-D and
strain EG-S-CBH1-S, respectively. In the 3rd batch, 41.8% of PASC was consumed
34
0 10
20
30
40
50
60
70
80
90
100
0
0.5 1
1.5 2
2.5 3
3.5
0 24
48
72
96
12
0 14
4 16
8 19
2 21
6 24
0 26
4 28
8
PASC (%)
Ethanol (g/L)
Tim
e (h
)
EG-D
-CB
HI-
D e
than
ol
EG-S
-CB
HI-
S et
hano
l EG
-D-C
BH
I-D
PA
SC
EG-S
-CB
HI-
S PA
SC
Cyc
le 1�
Cyc
le 2�
Cyc
le 3�
Figu
re I
-7 T
hree
cyc
les
of C
RB
F us
ing
reco
mbi
nant
S. c
erev
isia
e st
rain
s E
G-D
-CB
H1-
D a
nd E
G-S
-CB
H1-
S. P
ASC
(%)
indi
cate
s th
e pe
rcen
tage
of
resi
dual
PA
SC n
orm
aliz
ed t
o th
e in
itial
con
cent
ratio
n in
the
res
pect
ive
cycl
e. T
he i
nitia
l
amou
nt o
f PA
SC in
eac
h cy
cle
was
def
ined
as
100%
. For
eac
h st
rain
and
tim
e po
int,
data
are
pre
sent
ed a
s th
e m
ean
± SD
from
thre
e in
depe
nden
t exp
erim
ents
.
35
by EG-D-CBH1-D, corresponding to ~60% of the consumption in the 1st batch.
In contrast, the consumption of PASC in strain EG-S-CBH1-S decreased from
55.9% to 19.4% after 3-cycle repeated fermentation. Correspondingly, the final
ethanol titer generated by strain EG-D-CBH1-D was 1.7-fold higher than that
generated by strain EG-S-CBH1-S in the 3rd batch. These results suggested that
associating EG and CBH1 with cells facilitated retention of cellulolytic activity even
after three cell-recycles.
Discussion
In this study, we integrated heterologous EG and CBH1 genes into the genome of
a BGL-displaying S. cerevisiae, permitting the production of EG and CBH1 via
cell-surface display, secretion, or a combined strategy. The recombinant strains that
produced EG and CBH1 in the same space (on the cell surface or in the medium)
showed superior performance in the production of cellulosic ethanol. To our
knowledge, this is the first report on combining cell-surface display and secretion
strategies in a single yeast strain for heterologous cellulase production.
The benefits of attaching BGL to the yeast cell wall have been reported
previously38. However, suitable strategies for EG and CBH production by cells
remained obscure. In the present study, direct ethanol production from β-glucan and
PASC was performed using cellulolytic yeast strains. Specifically, the production of
ethanol was compared to investigate the suitable strategy for producing various kinds
of cellulases. When β-glucan was used as the fermentation substrate, the ethanol
production rate of strain EG-D was apparently higher than that of EG-S. Notably,
strain EG-D was able to convert β-glucan into ethanol immediately after the start of
fermentation, indicating that the displayed BGL and EG were successfully transferred
36
into the fermentation medium with the cell inoculum. In contrast, strain EG-S
converted β-glucan into ethanol after a 9-h lag, consistent with the need for this strain
to accumulate (via secretion) EG in the medium following inoculation. This
phenomenon demonstrated the advantage of an EG-display system to the cellulose
fermentation process, since display permitted early onset of the production of ethanol.
Compared with β-glucan, PASC contains a higher degree of polymerization and
crystallization, rendering this substrate more difficult to degrade by BGL and EG.
Indeed, fermentation with PASC yielded apparently lower rates of ethanol production
than those seen upon fermentation with β-glucan. The decline of ethanol observed in
PASC fermentation is probably due to the consumption via yeast cells when cellulases
could not hydrolyze sufficient glucose from PASC. Notably, the rate of ethanol
production from PASC by EG-D appeared higher than that by EG-S.
Subsequently, the CBH1 gene was successfully expressed along with BGL and
EG genes in S. cerevisiae. The T. emersonii CBH1 used in this study was fused with
the CBM of T. reesei CBH163; use of the CBM has been shown to enhance the
adsorption of enzyme to its substrate and modify substrate surfaces to facilitate
enzymatic hydrolysis74. We observed that co-production of CBH1 with displayed EG
enhanced ethanol production by up to 2-fold, demonstrating that the heterologous
CBH1 that was present assisted in the degradation of PASC. In the display strategy,
the CBH1 gene was expressed via the SED1 expression cassette and anchored on the
cell surface of S. cerevisiae using the SED1 anchoring domain25. It has been reported
that the expression level of the SED1 gene was highly induced in the stationary phase
by various environmental stresses, such as ethanol75. In the present work, expression
of both CBH1 and EG gene by cell-surface expression cassettes (in strain
EG-D-CBH1-D) permitted a doubling of CBH1 activity at 96 h vs. 0 h of
fermentation, confirming the utility of the stress-induced SED1 expression cassette.
37
However, we note that the SED1 anchoring domain was fused to the C-terminus of
the CBH1-CBM chimera protein; N- and C-terminal of CBM were fused with CBH1
and SED1 anchoring domain respectively, which may hinder the function of the CBM.
Alternatively, an N-terminal anchoring domain, such as the N-terminus flocculation
functional domain of Flo1p, may be more suitable for displaying the chimeric CBH1
containing a C-terminal CBM14. Nonetheless, to our knowledge, the present work
represents the first report on displaying a CBH1 on the cell surface of yeast strain.
In a previous report, a yeast strain displaying BGL, EG, and cellobiohydrolase 2
(CBH2) on the cell surface yielded higher ethanol production than a strain secreting
the corresponding enzymes in free form76. Immobilized BGL on the cell wall is
considered more appropriate for cellulosic ethanol production compared to free BGL
38; our use of cell-surface-displayed BGL is presumably one of the reasons for the
elevated ethanol yields in the present study. Ethanol production by strain
EG-S-CBH1-S was similar to that of strain EG-D-CBH1-D. To understand this
interesting result, the mechanism of the enzymatic hydrolysis of cellulose should be
taken into account. EG can randomly cleave the amorphous regions of cellulose to
produce oligosaccharides and provide free chain ends for CBH activity. Then CBH1
can initialize cleavage from the free chain ends, degrading crystalline cellulose into
cellobioses in a processive manner 77. The binding of EG and CBH1 onto the
cellulose surface, along with the processive movement of CBH1, are considered
essential steps in the degradation of cellulose 77. Besides enzymes, the presence of a
living microorganism is also important to the hydrolysis of cellulose. Generally, the
radius of a spherical yeast cell is around 2 µm 78, which is nearly 400-fold larger than
the hydrodynamic radius of a cellulase protein (≥ 5 nm) 79. The immobilization of
enzymes onto the cell wall may block the movement of processive enzymes (e.g.,
38
CBH1) or cause steric restriction in the collision with cellulose. As shown in Figure
I-8A, in the case of strain EG-D-CBH1-D, three different cellulases were displayed on
the cell-surface in a relatively close proximity (e.g., CBH1-BGL distance) compared
to the strains constructed by the other strategies. Such co-localization is expected to
increase the occurrence of synergistic interactions among cellulases16 and to facilitate
the transportation of glucose into cells. However, due to the size of the cell-enzyme
complex, the penetration of EG and CBH1 into the internal space of cellulose is
expected to be limited; such enzyme penetration has been reported as an important
determinant of hydrolytic rate58 80. Additionally, the processive movement of CBH1
may be retarded due to its immobilization on the cell. In previous pre-steady-state
analyses of CBH1 activity on cellulose, stalling of the processive movement of CBH1
was reported to lead to lower specific activity 81, 82. By contrast, EG and CBH1 in
strain EG-S-CBH1-S were produced as free forms (Figure I-8D), a strategy that was
expected to decrease steric hindrance and to increase the chance of collision with the
substrate. Although the cellulases appeared to accumulate in fermentation with
EG-S-CBH1-S (rising from 27 mU/mL at 0 h to 48 mU/mL at 96 h), the cellulolytic
enzyme activity was still lower than that of EG-D-CBH1-D. Additionally, the
diffusion efficiency of enzymes might affect the ability to degrade cellulose,
especially in a substrate with higher viscosity (e.g., PASC). Thus, co-locating EG and
CBH1 in the same space (on the cell-surface or in the medium) is favorable for
amorphous cellulose-based ethanol fermentation.
By contrast, ethanol production levels were apparently lower in strains
EG-D-CBH1-S and EG-S-CBH1-D. We infer that yeast cells attach to the surface of
cellulose to facilitate the collision between immobilized cellulases and cellulose, but
39
that this proximity may block the access of secreted EG or CBH1 to the surface of the
substrate (Figure I-8B, I-8C). For instance, the cellulase activities of strain
EG-D-CBH1-S at 96 h were higher than those of EG-S-CBH1-S, while the PASCase
Figure I-8 The effect of different locations of EG and CBH1 on the conversion of
cellulose into ethanol. A EG-D-CBH1-D. B EG-D-CBH1-S. C EG-S-CBH1-D. D
EG-S-CBH1-S. The diagrams suggest the multiple factors involved in the degradation of
cellulose, such as the distance between synergistic enzymes (CBH1-BGL distance), the effect
of cell-surface display on the processive movement of CBH1 (Retarded/ Un-retarded CBH1
movement), and the steric restriction during the binding of cellulases to cellulose surface
(Enzyme binding path)
activity was lower than that of strain EG-S-CBH1-S, effects that may be due to the
steric hindrance mentioned above. Additionally, the hydrolysis efficiency of strain
EG-S-CBH1-D appeared to be further decreased, presumably by the retarded
40
movement of immobilized CBH1 on the cell surface, as evidenced by the lower
PASCase activity and ethanol production obtained in strain EG-S-CBH1-D.
Interestingly, the individual enzyme activities in strain EG-D-CBH1-D were raised
while PASCase activity kept constant. We assume that in the case of cell-surface
displayed cell, only the enzymes located in the contacting region between yeast cell
and cellulose can involve in cellulose degradation. Although the individual enzyme
activities in overall cell were raised, the improvement of cellulolytic activity in
“attaching region” is limiting. In contrast, most of free cellulases can bind to cellulose
and participate in hydrolysis; the increase of individual enzyme activities can be fully
reflected on improvement of cellulolytic activity. This may also explain why the
overall enzyme activity of strain EG-S-CBH1-S (secretion system) observably rose
(77%) along with the increase of individual activities. Nonetheless, our results do
indicate that the involvement of the microorganism in the synergism of cellulases may
affect their hydrolysis efficiency towards cellulose. Moreover, the cellulose
degradation process perhaps involves more complex interactions, such as between
microbial cells and cellulose. Francisco et al. displayed Cex CBH on the surface of E.
coli and found that engineered E. coli exhibited specific adhesion capacity toward
cellulose 83. Such cell-to-cellulose adhesion pattern may lead to a distinct mechanism
from the “ablative mechanism” 57 of free cellulases to break down cellulose.
From an industrial point of view, the reuse of yeast cells through various rounds
of fermentation may be important in the ethanol production process 84. Cell-recycle
batch fermentation (CRBF) is a semi-continuous operation strategy using high
densities of recycled cells to produce ethanol continuously. In previous reports, the
CRBF of lignocelluloses was performed by the addition of large amount of
commercial cellulases, a step that represents one of the main bottlenecks for
commercialization85, 86. The application of a cellulolytic yeast strains to CRBF is
41
expected to decrease the need for the addition of costly commercial enzymes 39. Thus,
in this study, the direct ethanol production from PASC using recycled cellulolytic
cells was conducted without the addition of commercial cellulases. After a 3-batch
recycle fermentation, our strain EG-D-CBH1-D retained 60% of the PASC
degradation ability that the strain had in the 1st batch. This retention of activity was
1.7-fold higher than that seen in strain EG-S-CBH1-S. The apparently lower
degradation ability in strain EG-S-CBH1-S likely can be attributed to the loss of
cellulases in each cycle, as most secreted enzymes were separated from cells during
the cell collection step at the beginning of each batch. Although the amount of
enzyme produced was sufficient for saccharification in the first batch, the ability to
generate cellulases in each new cycle apparently declined in subsequent batches. In
contrast, the enzymes immobilized on strain EG-D-CBH1-D showed more consistent
activity than the activities in strain EG-S-CBH1-S during the recycling. Khaw et al.
reported that the ethanol production rate of a yeast strain producing displayed
α-amylase was maintained during a number of repetitions87. Consequently, the
construction of cellulolytic yeast by cell-surface display strategy is more applicable to
CRBF compared with the secretion strategy. Matano et al. even reported that with the
addition of extraneous cellulases (10 FPU/g-biomass), the fermentation ability of a
cellulase-displaying yeast strain remained constant after 5-cycle repeated-batch
fermentation 46. However, the required amount of additional cellulase was still too
high, precluding economic feasibility for an industrial process. In future work, we
propose to further improve the cellulolytic activity of yeast strains and apply the
cell-surface display strategy to ethanol production from lignocellulosic biomass (e.g.,
rice straw) with addition of trace amounts of exogenous cellulases.
Conclusion
In this study, we investigated suitable strategies for heterologous production of
42
EG and CBH1 production in the engineering of cellulolytic S. cerevisiae strains. We
demonstrated that cell-surface display system enhanced the production rate of ethanol.
Placement of EG and CBH1 in the same space (both on the cell surface or both
secreted into the medium) was favorable for ethanol production from amorphous
cellulose. In addition, a cellulolytic yeast strain producing cellulases via cell-surface
display was more effective in the cell-recycle batch fermentation than a strain
producing secreted cellulases.
43
Chapter II
Engineering of a novel cellulose-adherent cellulolytic Saccharomyces
cerevisiae for cellulosic biofuel production
44
Introduction
As the demand for fossil fuels increases and atmospheric CO2 levels continue to
rise, biofuels have attracted increasing attention as sustainable and renewable
alternative energy sources88. Lignocellulose is a potential resource for biofuel
generation because of its low cost and large-scale availability89. However, the need
for high-dosages of costly commercial cellulases in the saccharification process
makes it challenging for cellulose-based biofuels to be economically feasible6.
Although two leading enzyme companies (Genencor and Novozymes) have
significantly reduced cellulase prices (to 15-20 cents per gallon of ethanol produced),
these prices are still 5- to 10-fold higher than those of the amylases used for
starch-based biofuel production90. Thus, development of a microorganism that is
capable of both producing cellulases and fermenting resultant sugars into biofuels is a
promising approach to alleviate the economic burden imposed by the need for
commercial enzymes7.
Saccharomyces cerevisiae has been reported to have a superior capacity for
converting glucose into ethanol91. However, yeast lacks the cellulolytic enzymes
needed to degrade cellulose into glucose, such as β-glucosidase (BGL),
endoglucanase (EG), and cellobiohydrolase (CBH). Several engineered S. cerevisiae
strains capable of producing heterologous cellulases have been reported9, 10. However,
these engineered strains are unable to degrade crystalline cellulose effectively
(achieving only 8-23% hydrolysis efficiency)92, 93. This is due to insufficient
production (or secretion) of CBH63, as CBHs are responsible for the degradation of
crystalline cellulose and have been considered key forces in disrupting the recalcitrant
structure of cellulose59.
Up to now, cellulolytic S. cerevisiae strains have been constructed primarily via
45
either secretion or cell-surface display56. Secretion system, which releases enzymes
into extracellular environment, is the most common route of cellulase production in
recombinant strains. Although free enzymes can easily penetrate into the secondary
cell wall of plant cells57, they are incapable of being recycled usage during an
industrial process56. In comparison, a cell-surface display system permits
immobilization of enzymes on the cell surface through glycosylphosphatidylinositol
(GPI) anchoring of proteins14. Immobilization of numerous cellulases on a given
microbial cell provides an effective increase of local enzyme concentration; facilitates
synergistic interactions among enzymes; and, most importantly, enables re-utilization
of enzymes in repeated fermentation, improving the economic feasibility of the
process37. The cellulose degradation mechanisms of free-form cellulases and
cellulosomes (complexed cellulase system) have been studied intensively in the last
few decades57, 94, however, few investigations have focused on the mechanisms
employed by cell-surface-displayed enzymes (non-complexed cellulase system).
Cellulosic ethanol production using cellulase-displaying cells is gaining increased
attention37, but information on how these cells digest cellulose to fermentable sugars
is still unclear.
To achieve commercial scale, high bioethanol productivity will need to be
obtained from lignocellulosic waste (e.g., rice straw). However, most studies select
only pure/model cellulose (e.g., phosphoric acid swollen cellulose (PASC) or Avicel)
as an experimental substrate37, a model that is significantly different to real word
ethanol fermentation from lignocellulosic feedstock. Here, we report the successful
engineering of a cellulose-adherent S. cerevisiae strain producing heterologous BGL,
EG, CBH1, and CBH2 via cell-surface display, permitting direct ethanol production
from a realistic lignocellulosic substrate (rice straw). The interactions between
engineered yeast cells and cellulose were imaged by scanning electron microscopy
46
(SEM) to demonstrate clearly the dynamic mechanisms of cell-to-cellulose adhesion.
This work provided valuable information on how to increase cellulose hydrolysis
efficiency by enhancing cell-to-cellulose interactions for various types of substrate,
and therefore may lead to a feasible path towards the cost-competitive production of
cellulosic ethanol.
Materials and methods
Media and materials
Escherichia coli NovaBlue was grown in Luria-Bertani broth at 37 °C, and 100
mg/L ampicillin was added to the medium when required. Yeast strains were screened
on synthetic dextrose (SD) agar plates (6.7 g/L of yeast nitrogen base without amino
acids and 20 g/L of glucose) supplemented with appropriate amino acids. Yeast
strains were pre-cultured at 30 °C for 72 h in yeast extract-peptone (YP) medium (10
g/L yeast extract and 20 g/L peptone) containing 20 g/L glucose (YPD). Cellulosic
ethanol production was carried out at 37 °C in YP media containing different
cellulosic materials depending on the purpose of different fermentation experiments.
PASC was prepared from Avicel PH-101 (Fluka Chemie GmbH, Buchs, Switzerland)
as described previously9. Rice straw was pretreated with a liquid hot water method
(130-300 °C under a pressure of less than 10 MPa) and then subjected to 4 cycles of
milling using a CMJ01 nano-mech reactor (Techno Eye, Tokyo, Japan)95; the
resultant biomass was designated MC6. The composition of MC6 was 43% (w/w)
glucan, 2% (w/w) xylan, 42.3% (w/w) ash and lignin, and 12.7% (w/w) other
materials39.
47
48
Plasmid and strain constructions
The plasmids and primers used in this study are summarized in Table II-1 and
Table II-2, respectively. The plasmid pRDH227 was constructed by removing the C.
lucknowense open reading frame as a 1468 bp PacI/AscI fragment from the plasmid
pMU78463 and cloning it into the corresponding sites of pBHD197. Other plasmids
were constructed by connecting DNA fragments using the isothermal assembly
method66. For plasmid pDI9-CBH1D, four PCR products were connected. The four
PCR products included the following: I9 region fragments (the 3’ non-coding region
between gene YOR191W and YOR192C, amplified from the genome of S. cerevisiae
BY4741 using primers pairs I9a-M-F + I9a-O-R and I9b-O-F + I9b-C1-R; ori-ampR
fragment (from plasmid pIU-CBH1D) using primers O-I9a-F + O-I9b-R; MET15
fragment (from plasmid pRS401) using primers M-I9a-F + M-C1-R; and T. emersonii
CBH1-encoding surface-display cassette (from plasmid pI5-CBH1D) using primers
C1-M-F + C1-I9b-R. To construct plasmid pDI9-CBH2D, a PCR-amplified C.
lucknowense CBH2-encoding gene (from plasmid pRDH227, using primers C2-F +
C2-R) and the vector backbone containing the enzyme-display cassette (from plasmid
pDI9-CBH1D, using primers D-C2-F + P-C2-R) were linked together. Similarly, the
CBH2 gene (primers C2-F and C2-R2) was ligated with the vector backbone
containing the enzyme-secretion-cassette (from plasmid pDI9-CBH1D, using primers
D-C2-F2 + P-C2-R), to yield plasmid pDI9-CBH2S. In addition, the CBH2 gene,
enzyme-display cassette, and I5 region (the 3’ non-coding region of gene YLL055W
and YLL054C, amplified from plasmid pI5-CBH1-D using primers D-C2-F + P-C2-R)
were connected to yield plasmid pIU5-CBH2D.
Plasmids were transformed into S. cerevisiae BY4741 using lithium acetate as
described68, and integrated into either the I5 or I9 region by homologous
recombination. The transformants were identified using colony PCR to check for the
49
integration of cellulase genes (using screening with primers I9-F + I9-R for I9 region
integration or primers I5-F + I5-R for I5 region integration).
Table II-2 PCR primers used in this study
Primers Sequence (5’-3’)
I9a-M-F ATTAATGAATCGGCCAACGCTGGATATGACTGTGTTGTTGCTGATA
I9a-O-R GGGGGCGGAGCCTATGGAAAAACGCCAGCAACGCGG
O-I9a-F AAGGCCGCGTTGCTGGCGTTTTTCCATAGGCTCCGCCCCC
O-I9b-R GCACTTTTCGGGGAAATGTGCGCGGAACCCCTATTTGTTTATTTTTC
I9b-O-F AAACAAATAGGGGTTCCGCGCACATTTCCCCGAAAAGTGCCACC
I9b-C1-R TTTTCACCGTCATCACCGAAGGGCCCATGGCTAGGTGT
M-I9a-F ACACACCTAGCCATGGGCCCTTCGGTGATGACGGTGAAAA
M-C1-R TTTCACACCGCATAGATCCGACTTGTGAGAGAAAGTAGGTTTAT
C1-M-F ACCTACTTTCTCTCACAAGTCGGATCTATGCGGTGTGAAATAC
C1-I9b-R CAACAACACAGTCATATCCAGCGTTGGCCGATTCATTA
C2-F AATACGTTCGCTCTATTAAGATGGCCAAGAAGTTGTTCATTACC
C2-R GTTGATAATTTACTCGAGCCGAATGGTGGATTTGCGTTCGTTAAC
C2-R2 CTCAATGTACTAACTGTACATTAGAATGGTGGATTTGCGTTCG
D-C2-F CGAACGCAAATCCACCATTCGGCTCGAGTAAATTATCAACTGTCC
P-C2-R ATGAACAACTTCTTGGCCATCTTAATAGAGCGAACGTATTTT
D-C2-F2 ACGCAAATCCACCATTCTAATGTACAGTTAGTACATTGAGTCTAAATA
I9-F AAGAAGAAATCCGTGCTTACACATT
I9-R GCTATCCCATGCAAAGATTGTCAACG
I5-F CATTGAAGAAGGGAAAGTGGTAACC
I5-R TCCCTCTCTAATCTGGGTGAGAC
rt-CBH1-F CAACTTACTGTCCAGACGACGAAAC
rt-CBH1-R AAGGAAGAACCAGAGGAGGTAACAC
rt-CBH2-F AGAAGTCCCTAGTTTCCAATGGCTT
rt-CBH2-R CGGCCTTATTCAAAGCTCTAACCTG
rt-ACT1-F TGGATTCCGGTGATGGTGTT
rt-ACT1-R TCAAAATGGCGTGAGGTAGAGA
50
Cell growth assay
To measure the growth of yeast cells, parent strain S. cerevisiae BY4741 and
recombinant strains were cultivated individually in SD medium at 30 °C with shaking
at 150 rpm for 24 h. The resulting pre-cultures were inoculated into 5 mL YPD
medium at an initial optical density (OD660) of 0.05 and cultivated at 30 °C with
shaking at 70 rpm. The value of the OD660 was measured once hourly using a
TVS062CA Bio-photorecorder (Advantec Toyo, Tokyo, Japan). The value of the
OD660 was taken as an indicator of cell growth.
Quantitative real-time PCR
The transcription levels of the cellulase-encoding genes were quantified as
described previously 69. Primers rt-CBH1-F and rt-CBH1-R were used to determine
the transcription level of the CBH1 gene; primers rt-CBH2-F and rt-CBH2-R were
used to determine the transcription level of the CBH2 gene. Transcription levels of the
target genes were normalized to those of the housekeeping gene ACT1 (tested using
primers rt-ACT1-R and rt-ACT1-F).
Scanning electronic microscopy and optical microscopy
Yeast cells were resuspended in phosphate buffer (pH 5.0) at a concentration of
30 g wet cells/L. Subsequently, 1% (w/v) PASC or Avicel was added to the cell
suspension, which then was incubated at 37 °C for 2 h. Cellulose fibers were fixed
with 4% paraformaldehyde and 4% glutaraldehyde (GA) in 0.1 M cacodylate buffer
(pH 7.4) at 4 °C. Thereafter, fibers were fixed with 2% GA in cacodylate buffer
overnight. The samples were additionally fixed with 1% tannic acid at 4 °C for 2 h.
After the fixation the fibers were washed with cacodylate buffer 4 times, followed by
post fixation with 2% osmium tetroxide in cacodylate buffer for 4 h. The samples next
were dehydrated in a graded series of ethanol solutions (50, 70, 90, and 100%), and
51
then were substituted into tert-butyl alcohol and dried by vacuum freeze drying. After
drying, the samples were coated using an osmium plasma coater (NL-OPC80NS,
Nippon Laser & Electronics Laboratory, Nagoya, Japan). The samples were
visualized using a scanning electron microscope (JSM-6340F; JEOL Ltd., Tokyo,
Japan) at an acceleration voltage of 5 kV. For observation using optical microscopy,
post-incubation samples were directly applied onto microscope slides and imaged.
Ethanol production from cellulosic materials
Fermentations were performed at 37 °C under oxygen limited conditions at an
agitation speed of 200 rpm in 100-mL closed bottles equipped with a bubbling
CO2 outlet and a stir bar. Pre-cultivated cells in YPD medium were centrifuged and
washed twice by sterilized water. The collected cells were inoculated into 20 mL YP
medium containing 20 g/L PASC, 10 g/L Avicel, or 100 g/L MC6. Initial cell
densities were adjusted to approximately 150 g wet cells/L. Yeast cell wet weight was
determined by weighing a cell pellet that was harvested by centrifugation at 1,000 × g
for 5 min. To evaluate cellulase dosages in the SSF process, commercial enzyme
(Novozymes Cellic CTec2; Novozymes Inc., Bagsvaerd, Denmark) was added into
medium at 0-h of fermentation at enzyme concentrations of 0, 0.2, 0.6, 1.0, 1.4, 1.8,
2.2 FPU/g-biomass. Measurement of the filter paper cellulase units (FPU) of CTec2
was based on the standard NREL analytical procedure 98 performed at 37 °C. The
ethanol concentrations in the fermentation medium were determined using a gas
chromatograph, as described previously 48.
Cellulolytic activity assay
At the 0-h and 96-h time points of ethanol production from 100 g/L MC6,
fermentation medium was assayed for cellulolytic activities. Cellulolytic activity
represents the degradation ability of all enzymes present in the fermentation broth.
52
Fermentation broth was added into 50 mM sodium citrate buffer (pH 5.0) containing a
final concentration of 1% (w/v) MC6 and 100 mM methyl glyoxal (Nacalai Tesque,
Inc., Kyoto, Japan); the methyl glyoxal prevented the assimilation of glucose by yeast
cells 70. The reaction was performed at 37 °C using a heat block (Thermo Block
Rotator SN- 06BN; Nissin, Tokyo, Japan) with shaking at 35 rpm, and the supernatant
was collected by centrifugation. The amount of glucose in the supernatant was
determined by the Glucose CII kit (Wako Pure Chemical, Osaka, Japan). One unit of
cellulolytic activity (expressed as U/L) was defined as the amount of enzyme needed
to produce 1 µmoL of glucose per minute at 37 °C, pH 5.0.
Results
Heterologous expression of cellulases in S. cerevisiae
The relevant features of the recombinant yeast strains used in this study are listed
in Table II-1. Four codon-optimized genes that encode Aspergillus aculeatus BGL1,
Trichoderma reesei EG2, Talaromyces emersonii CBH1, and Chrysosporium
lucknowense CBH2 were expressed and used for assembly of enzyme cocktails in S.
cerevisiae BY4741. Cellulase enzyme encoding genes were all expressed under the
control of the SED1 promoter, which is highly induced during the stationary-phase
growth 75. The secretion signal peptide of BGL1 was derived from Rhizopus oryzae
glucoamylase, while EG2, CBH1, and CBH2 were produced with their native
secretion signals. The schemes of engineered cellulolytic yeast strains producing
enzymes via cell-surface display or secretion are illustrated in Figure II-1. In the strain
expressing cell-surface-displayed proteins, the enzymes were encoded fused to the
N-terminus of Sed1, a S. cerevisiae cell wall protein rich in threonine/serine residues
that contains a putative GPI attachment signal. Sed1p becomes the most abundant cell
wall protein during the stationary phase of growth75. The GPI-anchor is attached to
the Sed1-enzyme chimeric proteins in the endoplasmic reticulum and subsequently
53
Cel
l-sur
face
dis
play
syst
em�
CB
H1
SS
ASE
D1
T SA
G1
P SE
D1
CB
H2
SS
ASE
D1
T SA
G1
P SE
D1
EG
SS
A
SED
1 T S
AG
1 P S
ED
1
BG
L SS
A
SED
1 T S
AG
1 P S
ED
1
Pyru
vate�
Ace
tald
ehyd
e�
CO
2�
ATP�
AD
P�
Eth
anol�
NA
D+�
NA
DH�
Glu
cose�
CB
H1�
EG�
CB
H2�
BG
L�
Glu
cose�
Cel
lulo
se�
Sacc
haro
myc
es c
erev
isia
e ce
ll�
SED
1 an
chor�
Secr
etio
n sy
stem�
Pyru
vate�
Ace
tald
ehyd
e�
CO
2�
ATP�
AD
P�
Eth
anol�
NA
D+�
NA
DH�
Glu
cose�
CB
H1
SS
T SA
G1
P SE
D1
CB
H2
SS
T SA
G1
P SE
D1
EG
SS
T S
AG
1 P S
ED
1
BG
L SS
P S
ED
1 A
SED
1 T S
AG
1
Figu
re I
I-1
Engi
neer
ing
cellu
loly
tic S
. cer
evis
iae
for
cellu
losi
c et
hano
l pro
duct
ion
via
eith
er c
ell-s
urfa
ce d
ispl
ay o
r
secr
etio
n of
enz
ymes
. P S
ED1,
SED
1 pr
omot
er;
SS,
secr
etio
n si
gnal
; A S
ED1,
SED
1 an
chor
ing
regi
on;
T SAG
1, SA
G1
54
transferred to the cell surface through an S. cerevisiae secretory pathway. Upon
reaching outer layer of the cell wall, chimeric proteins are covalently bound to
β-1,6-glucan via the GPI anchor99. In contrast, a chimeric protein lacking the Sed1
domain passes through cell wall and is released into the extracellular medium
(secretion system). Metabolic burden often occurs due to the expression of
heterologous protein genes, resulting in inhibition of cell growth100. However, in the
present study, the growth rates of the recombinant strains were similar to those of the
parent strain (Figure II-2), suggesting that production of the heterologous enzymes did
not impose an obvious metabolic burden.
Figure II-2 Time-course profiles of cell growth using parent strain BY4741 and recombinant
yeast strains in YPD medium. Each strain was inoculated in YPD medium to an initial OD660
of 0.05 and then cultured aerobically at 30 °C, 70 rpm, for 72 h. For each strain, data are
presented as the mean ± SD from three independent experiments.
Assembly of high-efficiency enzyme cocktail
To develop a high-efficiency cellulase system, various cellulase combinations of
some or all of our candidate proteins were engineered, including BGL, EG, CBH1,
0.01
0.1
1
10
100
0 24 48 72
Cel
l gro
wth
(OD
660
)�
Time (h)�
BY4741
EG-D-CBH1-D-CBH2-D
EG-S-CBH1-S-CBH2-S
EG-D-CBH1-D-CBH1-D
EG-D-CBH2-D
EG-D-CBH1-D
55
and CBH2. All enzyme gene combinations were expressed in S. cerevisiae and
produced through cell-surface display technique. Cellulosic ethanol production using
different recombinant strains was investigated. Pure celluloses consisting of PASC
(amorphous type) or Avicel (crystalline type) were used as substrates. Two
recombinant strains designated EG-D-CBH1-D and EG-D-CBH2-D (i.e., containing
combinations of BGL+EG+CBH1 and BGL+EG+CBH2, respectively) were designed
as “minimum enzyme cocktails”. As shown in Figure II-3, a maximal ethanol
production of 2.3 g/L (equivalent to 23% of the theoretical value) was achieved in
both engineered strains using PASC as the substrate (Figure II-3a). In contrast, strain
EG-D-CBH1-D showed a higher ethanol production rate than strain EG-D-CBH2-D
when fermenting Avicel (Figure II-3b), suggesting that the T. emersonii CBH1 is
more effective than C. lucknowense CBH2 in the degradation of crystalline cellulose.
Figure II-3 Time course of direct ethanol production from cellulosic materials. Ethanol was
produced from 20 g/L PASC (a) and from 10 g/L Avicel (b). Data are presented as the means
and standard deviations of triplicate measurements.
Subsequently, an additional copy of the CBH1-encoding gene was added to
BGL+EG+CBH1 cocktail, yielding strain EG-D-CBH1-D-CBH1-D. The transcription
level of CBH1 in strain EG-D-CBH1-D-CBH1-D was approximately 1.8-fold that of
0
1
2
3
4
5
6
7
8
0 24 48 72 96
Eth
anol
(g/L
)�
Time (h)�
a�
EG-D-CBH1-D EG-D-CBH2-D EG-D-CBH1-D-CBH1-D EG-D-CBH1-D-CBH2-D
0
0.2
0.4
0.6
0.8
1
1.2
1.4
1.6
0 24 48 72 96
Eth
anol
(g/L
)�
Time (h)�
b�
56
strain EG-D-CBH1-D (Figure II-4). In PASC and Avicel fermentations, strain
EG-D-CBH1-D-CBH1-D produced ethanol at 3.1 g/L (30% of the theoretical value)
and 0.56 g/L (11% of the theoretical value), respectively, corresponding to 1.3- and
1.9-fold the levels seen in strain EG-D-CBH1-D (Figure II-3). To further increase the
cellulose degradation efficiency, a strain (designated EG-D-CBH1-D-CBH2-D) that
contained genes encoding BGL, EG, CBH1, and CBH2 was designed; this strain
generated an ethanol titer of 6.7 g/L from PASC (66% of the theoretical value) and
1.4 g/L from Avicel (27% of the theoretical value), which are 2.9-fold and 4.5-fold
higher than strain EG-D-CBH1-D. These results suggested that introduction of a
second copy of the CBH1-encoding gene only slightly improved the cellulolytic
activities, while the combination of both CBH1- and CBH2-encoding loci yielded a
greater enhancement in the efficiency of cellulose hydrolysis.
Figure II-4 Relative transcription levels of CBHI and CBH2 genes in recombinant yeast
strains. Gene ACT1 was used as the internal standard. The relative transcription levels are
shown normalized to the level observed in strain EG-D-CBH1-D-CBH2-D, whose relative
transcription level was defined as 1. For each strain, data are presented as the mean ± SD
from three independent experiments.
0
0.5
1
1.5
2
2.5
CBH1 CBH2
Rel
ativ
e ex
pres
sion
leve
l (fo
ld)��
EG-D-CBH1-D-CBH2-D
EG-D-CBH1-D
EG-D-CBH2-D
EG-D-CBH1-D-CBH1-D
EG-S-CBH1-S-CBH2-S
57
Performance of enzyme-secreting and -displaying cells in cellulose degradation
We next compared the cellulose fermenting efficacy of S. cerevisiae strains
expressing the EG, CBH1, and CBH2 cellulases via either cell-surface display or
secretion. As depicted in Figure II-5, the cellulase-displaying cells
(EG-D-CHB1-D-CBH2-D) showed better ethanol yield (63% of the theoretical yield)
from PASC than was seen from cellulase-secreting cells (EG-S-CBH1-S-CBH2-S, 54%
of the theoretical yield), which is in good agreement with previous studies 101.
Figure II-5 Time course of direct ethanol production using recombinant strains
EG-D-CBH1-D-CBH2-D and EG-S-CBH1-S-CBH2-S. Ethanol was produced from 20 g/L
PASC (a) and from 10 g/L Avicel (b). Data are presented as the means and standard
deviations of triplicate measurements.
Observation of enzyme-secreting and -displaying cells in cellulose degradation
Based on the SEM observation shown in Figure II-6a, PASC appeared to form a
more sponge-like surface material than Avicel. After incubation of yeast cells with
cellulose, numerous cellulase-displaying cells (EG-D-CBH1-D-CBH2-D) remained
attached to the PASC surface, while very few cellulase-secreting cells
0
1
2
3
4
5
6
7
8
0 24 48 72 96
Eth
anol
(g/L
)�
Time (h)�
a�
EG-D-CBH1-D-CBH2-D EG-S-CBH1-S-CBH2-S
0
0.4
0.8
1.2
1.6
2
2.4
0 24 48 72 96
Eth
anol
(g/L
)�
Time (h)�
b�
58
(EG-S-CBH1-S-CBH2-S) were retained on the cellulose surface (Figure II-6b).
Figure II-6 SEM micrographs of the interactions between cellulolytic S. cerevisiae cells and
cellulosic materials. Interactions with PASC (a, b) and with Avicel (c, d). Cellulolytic cells
(30 g/L) were incubated with 1% cellulosic materials (PASC or Avicel) for 2 h, and the
cellulosic materials were used for SEM imaging. Scale bars are 10 µm.
EG-D-CBH1-D-CBH2-D� EG-S-CBH1-S-CBH2-S�
A PASC�
c Avicel �
b PASC�
d Avicel�
10 µm� 10 µm�
10 µm� 10 µm�
59
Notably, the adhesion between displayed cells and sponge-like cellulose (PASC)
was clearly observed in Figure II-7, showing that the displayed cells are tightly bound
to cellulose filaments. Unlike the enzyme-secreting strain, which digests the cellulose
via free-form enzymes (such that yeast cell need have no direct interaction with
cellulose), the enzyme-displaying strain apparently mediates cellulose degradation via
attachment to the cellulose surface, which presumably significantly shortens the
distance between microbe and substrate. However, interestingly, no obvious
difference in ethanol titer between enzyme-displaying and -secreting cells was
observed in Avicel fermentation (around 32% Avicel conversion, Figure II-5b), which
may reflect lower binding efficiency of displayed cells to Avicel than to PASC
(Figure II-6). We hypothesize that rough, sponge-like structure of PASC provides
greater surface area and fine structures, thereby facilitating adhesion of displaying
cells and promoting degradation of the substrate. To our knowledge, this is the first
report on observation of the adhesion between cellulolytic yeast cells and cellulose.
To better understand the dynamic degradation mechanisms employed by
displaying or secreting cells, the morphological changes of celluloses after 24 h
degradation were monitored (Figure II-8). Cellulose fibers (both PASC and Avicel)
were observed to be disrupted into small, irregular pieces. In contrast, the cellulose
fibers digested by secreting cells appear to be evenly ablated from the outer layer of
microfibers, which is consistent with a previous report 57.
60
1 µm�
1 µm�
Figu
re I
I-7
SEM
mic
rogr
aphs
of
the
inte
ract
ions
bet
wee
n PA
SC a
nd E
G-D
-CB
H1-
D-C
BH
2-D
cel
ls.
The
obse
rvat
ion
cond
ition
s wer
e th
e sa
me
as th
ose
used
in F
igur
e II
-6. S
cale
bar
s ar
e 1
µm.
61
Figure II-8 Micrographs of PASC (a, b) and Avicel (c, d) degradation process by cellulolytic
S. cerevisiae. Cellulolytic cells (30 g/L) were incubated with 1% cellulosic materials for 24 h
at 37 °C. Cellulosic materials were sampled and directly applied to microscope slides for
observation. Scale bars are 10 µm. The arrows indicate the different cellulose degradation
tendencies of cellulolytic yeasts: strain EG-D-CBH1-D-CBH2-D tended to conduct
degradation via tearing cellulose fibers into smaller pieces, while EG-S-CBH1-S-CBH2-S
equably eroded cellulose from the outer surface.
a PASC�
cAvicel�
b PASC�
d Avicel�
EG-D-CBH1-D-CBH2-D� EG-S-CBH1-S-CBH2-S�
62
Targeted pre-treatment of rice straw
To verify our hypothesis, that higher ethanol yield can be obtained using rough,
sponge-like cellulose as a substrate that provides more surface area and finer structure
for adhesion by enzyme-displaying yeast cells, the surface properties of cellulose
obtained from agricultural waste (specifically, from rice straw) were altered. Liquid
hot water (LHW) pretreatment has been reported to disrupt recalcitrant microstructure
and remove 35–60% of the lignin and all of the hemicellulose from lignocellulose
materials102; separately, milling has been reported to reduce particle size and
crystallinity of lignocellulose, resulting in increased surface area for enzymatic attack 103. In the present study, rice straw was treated with LHW plus 4 cycles of milling,
and the resultant biomass was designated MC6. The morphological characteristics of
rice straw and MC6 were observed by SEM (Figure II-9aI, II). Unprocessed rice straw
exhibited a flat, rigid, and compact surface, while the surface of MC6 was rough and
sponge-like. The average diameters of MC6 particles were about 20- to 50-fold
smaller than those of rice straw. These results suggested that LHW and milling
pretreatment significantly altered the surface characteristics of rice straw into a more
sponge-like structure with increased surface area.
It was also observed that more cellulolytic cells adhere to MC6 fibers than to
unprocessed rice straw (Figure II-9aI’, II’), a result that correlated with the obtained
ethanol concentrations (Figure II-9b). These results suggested that rough, sponge-like
cellulose particles facilitated cell-to-cellulose adhesion, thereby improving cellulose
degradation.
63
Figure II-9 Comparison of cellulosic feedstock rice straw with MC6 for cellulosic ethanol
production. (a) SEM micrographs of the surface structures on cellulosic materials (I, II);
observation of adhesion between strain EG-D-CBH1-D-CBH2 and cellulosic materials using
optical microscopy (I’, II’). (b) Ethanol yields at 96 h of fermentation from 25 g/L rice straw
and MC6, respectively. Data are presented as the means and standard deviations of triplicate
measurements.
I’�
I�
a�Rice straw�
III�
II’�
MC6�
I� II�
0 0.2 0.4 0.6 0.8
1 1.2 1.4 1.6 1.8
Rice straw MC6
Eth
anol
(g/L
)
b
64
Evaluating the feasibility of high-density cellulosic ethanol production
To test the feasibility of high-density cellulosic ethanol production using strain
EG-D-CBH1-D-CBH2-D, fermentation was conducted using 100 g/L MC6 as the
substrate. Only 1.3 g/L ethanol (7% of the theoretical yield) was produced by
enzyme-displaying cells after 96 h fermentation, while no ethanol was obtained by
wild type strain (BY4741). Considering the low ethanol yields in both two strains
(Figure II-9b), the inclusions of small amounts of exogenous commercial enzyme
(C-Tec2) were still required to support hydrolysis of MC6. To address the economic
feasibility of fermentation of MC6 using the enzyme-displaying strain, the potential
diminution of C-Tec2 supplementation by using displaying cells was assessed. As
shown in Figure II-10a, the ethanol yield from EG-D-CBH1-D-CBH2-D fermentation
of MC6 was 7-fold increased by addition of trace amount of exogenous C-Tec2 (0.2
FPU/g-biomass). Specifically, the inclusion of 1.0 FPU/g-biomass C-Tec2 together
with displaying cells provided a high cellulosic ethanol yield of 18 g/L, equivalent to
80% of the theoretical value. The result represents an approximately 44% decrease of
the required enzyme dosage compared to that required for M6 fermentation by the
wild-type strain. Additionally, although approximately half of the C-Tec2 activity was
lost after 96 h of fermentation, the cellulolytic activities from displaying cells
(indicated as the activity difference between EG-D-CBH1-D-CBH2-D and BY4741
fermentations) persisted (Figure II-10b), demonstrating the feasibility of recycling of
cellulase-displaying yeast cells.
65
0 2 4 6 8 10
12
14
16
18
20
22 0
0.2
0.4
0.6
0.8
1 1.
2 1.
4 1.
6 1.
8 2
2.2
2.4
Ethanol (g/L)�
Cel
lula
se a
dditi
on (F
PU/g
bio
mas
s)�
EG-D
-CB
H1-
D-C
BH
2-D
B
Y47
41
0 10
20
30
40
50
60
70
80 0
0.2
0.4
0.6
0.8
1 1.
2 1.
4 1.
6 1.
8 2
2.2
2.4
Cellulolytic activity (mU/mL)�
Cel
lula
se a
dditi
on (F
PU/g
bio
mas
s)��
EG-D
-CB
H1-
D-C
BH
2-D
0 h
B
Y47
41 0
h
EG-D
-CB
H1-
D-C
BH
2-D
96
h B
Y47
41 9
6 h
a�b�
Figu
re I
I-10
Eva
luat
ions
of c
omm
erci
al c
ellu
lase
add
ition
in e
than
ol p
rodu
ctio
n fr
om 1
00 g
/L M
C6.
(a)
Eth
anol
prod
uctio
n at
96
h of
ferm
enta
tion
with
the
addi
tion
of 0
, 0.2
, 0.6
, 1.0
, 1.4
, 1.8
, and
2.2
FPU
/g-b
iom
ass
cellu
lase
(C-T
ec2)
. (b)
Cel
lulo
lytic
act
iviti
es in
the
ferm
enta
tion
med
ia a
t 0 a
nd 9
6 h
of f
erm
enta
tion
with
the
resp
ectiv
e
stra
ins.
Dat
a ar
e pr
esen
ted
as th
e m
eans
and
sta
ndar
d de
viat
ions
of t
riplic
ate
mea
sure
men
ts.
66
Discussion
One of the main bottlenecks impeding the widespread production of cellulosic
bioethanol is the large amount of costly commercial enzymes required for hydrolysis
of cellulose7. As described in this work, a cellulose-adherent S. cerevisiae that
displayed BGL, EG, CBH1, and CBH2 on the cell surface was constructed through a
series of rational designs. It was demonstrated that the cellulase-displayed yeast strain
employed a cell-to-cellulose adhesion and a “tearing” pattern as parts of its
cellulose-degradation mechanism, which differed from those of strains producing
free-form enzyme. Appropriate pretreatment of lignocellulose materials aiming to
enhancing cell-to-cellulose interactions dramatically enhanced cellulosic ethanol yield.
The resultant cellulose-adherent S. cerevisiae may significantly reduce the need for
exogenous enzyme, potentially alleviating the bottleneck in commercial production of
cellulosic bioethanol.
A crucial point in assembly of the cellulase cocktail is to maximize the
synergistic actions among enzymes. Both CBH1 and CBH2 are known to act as
exoglucanases, initiating cleavage of cellulose chains from reducing and non-reducing
ends, respectively8. In this study, T. emersonii CBH1 appeared more important than C.
lucknowense CBH2 in the synergism with BGL and EG (especially in Avicel
fermentation), but we found that providing both CBH1 and CBH2 in enzyme cocktail
was more effective on improving cellulolytic activities than increasing the proportion
of CBH1 (here, by doubling the gene dose of the CBH1 encoding locus). This is
because CBH2 can synergistically enhance the hydrolysis efficiency of CBH1, that it
diminished the bumpy surface on cellulose, apparently preventing CBH1 from getting
stuck during processive movement104. To date, a maximum of three kinds of
cellulases (non-complexed cellulase system) have been simultaneously displayed on
one single cell, because of the difficulty in displaying stable and functional
67
heterologous proteins on the microbial surface. In the present work, we provided the
first report (to our knowledge) of tethering four types of heterologous cellulases to the
surface of a single yeast cell. The resulting strain exhibited significantly higher
ethanol yield from both amorphous cellulose and crystalline cellulose compared to the
cellulolytic yeast strains previously described in relevant studies.
Most reports have assessed enzyme production strategies using amorphous
cellulose as substrate, and the cellulosic ethanol yield obtained with cell-surface
display systems was generally higher than that obtained with secretion systems 42, 105.
However, we proved that the performance of each system is actually
substrate-dependent. The higher ethanol yields in enzyme-displaying systems are
obtained only using amorphous cellulose as substrate, indicating that there is likely
unique interactions between displaying cells and substrate, differing from those that
occur in secreting cell. Cellulase-displaying cells degrade cellulose via tight
attachment onto cellulose filaments, while cellulase-secreting cells did not exhibit
obvious interactions with substrate. One of the reasons for the cell-to-cellulose
adhesion is the high affinity of the carbohydrate binding domain (CBD) in cellulase
towards cellulose106, and anchoring cellulase on cell surface results in a higher affinity
of cells towards cellulose. Increased adhesion of Escherichia coli to cotton fibers has
been demonstrated via anchoring the CBD from CBH onto the cell surface 83. We also
found that the adhesion between displaying cells and cellulose correlates with the
hydrolysis efficiency of cellulose. Electron microscopy (Figure II-7) clearly illustrated
that the swollen, rough microfibers of PASC can encircle the cellulase-displaying
cells, thereby enhancing the adhesion between cells and cellulose, which may explain
the higher ethanol yield obtained from PASC using displaying cells. The
cellulose-adherent characteristic of enzyme-displaying cells is expected to shorten the
distance between cells and substrates, thereby facilitating the mass transfer of
68
hydrolysis products, particularly in high-density fermentation.
Notably, two distinct cellulose-degradation mechanisms are proposed based on
the morphological changes of cellulose caused by both displayed and secreted cells
(Figure II-8). The results indicated that the cell-surface display system tends to
increase the accessible surface area for cell-adhesion by tearing microfibers from
cellulose particles. In contrast, free (secreted) enzymes equably erode cellulose
surface and diminish particle size, consistent with a previous report that free enzymes
employ an ablative, fibril-sharpening mechanism during cellulose degradation107. A
potential explanation for the different mechanisms is the distinct enzyme-to-substrate
interactions represented by the two systems. It has been known that free enzymes
repeatedly associate and dissociate with cellulose to avoid getting stuck during
processive movement81. In contrast, owing to cell-to-cellulose adhesion, the
processive movements of surface-displayed enzymes are slowed down; enzymes are
entrapped on cellulose and digestion proceeds without repeated cycles of association
and dissociation with the substrate. As a result, the displayed cellulases are more
likely to carry out deeper directed digestion of cellulose particles than are free
enzymes, resulting in the splayed morphology observed in Figure II-8.
Natural lignocellulose (e.g., rice straw) is usually rigid and lacks the rough,
intricate surface structures presented in PASC. Thus, to increase the adhesion between
natural lignocellulose and cellulase-displaying yeast, the surface structures of rice
straw were broken into more “sponge-like” materials via pretreatment by LHW and
milling. In our work, cellulase-displaying yeast cells preferred to attach to MC6 and
exhibited dramatically higher hydrolysis rates than those seen with untreated rice
straw, presumably because the MC6 has more surface area with favorable sites for
cell-to-cellulose adhesion. Thus, appropriate pretreatment of lignocellulose, which
69
adopts the cell-to-cellulose adhesion pattern in combination with cellulase-displaying
yeast, is expected to effectively promote the degradation efficiency. Although the
optimal biomass pretreatment combined with free-cellulase-mediated saccharification
has been extensively studied103, the pretreatment process suitable for
surface-displayed cellulases still remains obscure. Pretreatment by steam explosion
has been reported to be capable of producing pores with 3-nm to 1-µm diameters on
the surface of lignocellulose108, facilitating the access of free enzymes (approximately
5.1 nm in diameter) to the interior of cellulose particles109. However, the
enzyme-displaying cells (around 3 µm) would not be able to enter pores of this size;
such cells would instead be expected to digest the substrate primarily by “shaving”
cellulose fibers from the external surface of substrate particles. As a result, such a
“pore-punching” pretreatment process may be not ideal for cell-surface display
hydrolysis systems. On the contrary, the method of ammonia fiber expansion (AFEX)
pretreatment can effectively increase the surface roughness of lignocellulose materials 110, which likely will be of greater use in combination with cell-surface display
systems.
Natural lignocellulose is commonly considered as the most rigid substrate
resistant to digestion, and impossible to degrade by cellulolytic yeast strains in the
absence of exogenous enzymes. Even with addition of exogenous enzymes, natural
lignocelluloses rarely provide high ethanol yields (over 80% of theoretical value)
when used as the substrate for simultaneous saccharification and fermentation (SSF) 111. In the reported SSF for bioethanol production mediated by S. cerevisiae, cellulase
loadings of 15-30 FPU/g-biomass are generally used, depending on the specific
substrate 112, 113. Recently, Matano et al. reported the use of cellulase-displayed yeast
strain to diminish cellulase dosage to 10 FPU/g-biomass in ethanol production from
LHW-pretreated rice straw 46. Surprisingly, the engineered cellulose-adherent S.
70
cerevisiae in the present study is capable of converting pretreated rice straw into
ethanol without commercial enzyme addition. To our knowledge, this is the first
report of direct ethanol production from nature lignocellulose using a cellulolytic
yeast strain, presenting a new potential towards making efficient use of lignocellulose.
Moreover, nearly 80% of ethanol theoretical yield was achieved in the presence of
only 1 FPU/g-biomass C-Tec2, a quantity that represents a 44% decrease of enzyme
loading compared with that required in non-cellulolytic S. cerevisiae and a much
lower enzyme dosage compared with other reports. Meanwhile, the enzymes
immobilized on the cell surface appeared higher stability during the fermentation than
did commercial enzymes. This observation is in agreement with the previous report
that tethering of enzymes to solid supports can increase the protein stability under
non-optimal reaction conditions such as low temperature and organic solvent
composition 114. The retention of activities in displayed enzymes also demonstrates the
feasibility of using cell-surface display in cell-recycling processes. By constructing a
novel cellulose-adherent S. cerevisiae with surface-displayed cellulases, we
successfully demonstrated the concept of enhancing cell-to-cellulose interactions to
effectively alleviate the requirement for high doses of supplemental enzymes in
cellulosic ethanol production. Further improvement of cell-to-cellulose adhesion (e.g.,
increased display of carbohydrate binding domains) combined with the increased
cellulolytic activities on yeast surface is expected to provide a viable model for
implementation of economically feasible cellulosic ethanol production.
Conclusion
In this chapter, we engineered a high-efficient cellulolytic S. cerevisiae with
displaying four different synergistic cellulases (BGL, EG, CBH1, and CBH2) on the
cell surface simultaneously. The engineered yeast cells exhibited clear
cell-to-cellulose adhesion and a “cellulose-tearing” pattern during cellulose
71
degradation. Notably, this strain could directly produce ethanol from rice straw; it
requires only 1.8 FPU/g-biomass commercial cellulases to achieve ethanol yield of 80%
from 100 g/L rice straw, cutting down more than 40% of the required enzymes under
high-dense fermentation. Our study also provides a novel strategy that improves
cellulose hydrolysis via enhancement of cell-to-cellulose interactions.
72
Chapter III
Efficient ethanol production from crystalline cellulose using
high-cellulolytic Saccharomyces cerevisiae with optimized cellulase
ratios on the cell surface
73
Introduction
Production of biofuels to replace the exhausting fossil fuels is gaining increasing
prominence worldwide. Cellulosic biomass presents the most renewable feedstock for
sustainable production of biofuels because of its abundance, low-price, and favorable
environmental properties115. Cellulose is composed of long-chain of glucose
monomers via β-1,4-glycosidic bonds. The extensive hydrogen linkages among chains
lead to a rigid structure of mostly well-ordered crystalline domains and small part of
disordered amorphous domains80. The crystallinities in natural cellulosic materials are
90-100% in plant-based fibers and 60-70% in wood-based fibers116. The crystalline
cellulose is resistant to the access of reactant and enzyme, whereas amorphous
cellulose is readily digestible6. It has been postulated that the efficient hydrolysis of
crystalline cellulose will enable the commercialization of cellulosic biofuels
production.
Efficient degradation of cellulose requires a synergistic reaction of the
cellulolytic enzymes including β-glucosidase (BGL), endoglucanase (EG), and
cellobiohydrolase (CBH). EG randomly degrades the non-crystalline region of the
cellulose; CBH attacks the exposed end of cellulose chain, degrading crystalline
cellulose into cellobioses; BGLdegrades cellooligosaccharide into glucose117. To date,
a variety of cellulolytic biofuel-producing microorganisms have been constructed via
expressing heterologous cellulases in a strategy of either cell-surface display or
secretion9, 50, such as displaying cellulases on the surface of Saccharomyces
cerevisiae. Although these microbes are capable of well degrading amorphous
cellulose (e.g., phosphoric acid swollen cellulose (PASC)), few of them can
efficiently hydrolyze crystalline cellulose (usually 10-30% hydrolysis of cellulose)63,
92. Liu et al. obtained a recombinant S. cerevisiae with BGL, EG, CBH1 and CBH2
activities, which achieved an ethanol yield of 66% from PASC, but only a yield of 27%
74
from Avicel (microcrystalline cellulose)118. This is because most of current researches
are focusing on overexpression of cellulase genes25 or improving the secretory level
of cellulases41, while few of them working on the optimization of enzyme ratios in
recombinant cellulolytic strains. The optimal ratio of synergisitic cellulase is one of
the most critical determinants in the hydrolysis of crystalline cellulose, and it differs
depending on the types of cellulose (e.g., resource, proportion of crystalline and
amorphous domains). Yamada et al. constructed a variety of cellulolytic yeast strains
containing different cellulase ratios (BGL, EG, CBH2). They found that the strains
with high degradation abilities towards amorphous cellulose generally used EG as the
principal proportion in total cellulase mixture44. Nonetheless, due to the preferences
of cellulase to substrate (EG prefers amorphous domain, while CBH targets on
crystalline domain)115, CBH activity should be more important than EG activity
during the degradation of crystalline cellulose. Thus, engineering of a recombinant
strain with the optimal cellulase ratio (perhaps taken CBH as the major proportion) is
assumed to achieve the efficient utilization of cellulosic biomass. Currently, the
optimal cellulase ratio that immobilized on cell surface for crystalline cellulose
degradation remains obscure.
In this chapter, to engineer the cellulase-displaying yeast pools with various
enzyme ratios, we employed cocktail δ-integration method44 to incorporate
multi-copy of cellulase genes (EG, CBH1, and CBH2) into the genome of a
BGL-displaying S. cerevisiae simultaneously. The cocktail-strains exhibiting higher
Avicel degradation abilities than strain EG-D-CBH1-D-CBH2-D (the strain
containing single-copy of each cellulase gene, constructed in Chapter II) were
selected to analyze the enzyme ratios on the cell surface using nano LC-MS/MS. To
our knowledge, this is the first work on relative quantitation of four kinds of cellulase
(BGL, EG, CBH1, and CBH2) anchored on one yeast cell, and the first to investigate
75
the optimal enzyme ratio for crystalline cellulose degradation, which therefore may
lead to a feasible path towards the efficient utilization of natural cellulosic materials.
Materials and methods
Strains and media
All yeast strains used in this study (Table III-1) are derived from strain S.
cerevisiae BY4741 (MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0). The Escherichia coli
strain NovaBlue (Novagen, Inc., Madison, WI, USA) was used for maintenance and
amplification of plasmid DNA. Bacterial cells were grown at 37 °C in Luria-Bertani
broth (10 g/L tryptone, 5 g/L yeast extract, and 5 g/L sodium chloride) containing 100
mg/L ampicillin. Yeast strains were pre-cultivated in synthetic dextrose (SD) medium
(6.7 g/L of yeast nitrogen base without amino acids (Difco Laboratories, Detroit, MI,
USA) and 20 g/L of glucose) supplemented with appropriate amino acids in a shaker
incubator (150 rpm) at 30 °C, and then aerobically cultivated at 30 °C in YP medium
(20 g/L peptone, 10 g/L yeast extract) and 20 g/L glucose. Ethanol fermentation was
performed in YP medium containing either 10 g/L Avicel PH-101 (Fluka Chemie
GmbH, Buchs, Switzerland) or 100 g/L MC6 as carbon sources. MC6 was prepared
from rice straw, as previously described95.
Plasmid construction and cocktail δ-integration
The vectors and primers used in this study are listed in Table III-1 and Table
III-2, respectively. The δ-integrative plasmid pδEG-D was constructed as followed:
the δ DNA fragment was amplified from plasmid pδW by PCR using primers δ-F and
δEG-R. The cell-surface display cassette, which includes SED1 promoter, SED1
anchoring region, and SAG1 terminator was amplified from plasmid pIL2-EGD using
primers SEG-F and S-R. Two DNA fragments were linked together by the isothermal
assembly method66 to yield the plasmid pδEG-D. The constructions of plasmids
76
77
pδCBH1-D and pδCBH2-D were performed by a process similar to the above
description. The δ DNA fragment was amplified from plasmid pδW using primers δ-F
and δC1-R for the construction of pδCBH1-D (primers δ-F and δC2-R for
pδCBH2-D), and fused with the PCR product amplified from plasmid pIU5-CBH1D
via primers SC1-F and S-R to construct pδCBH1-D (amplified from pDI9-CBH2D via
primers SC2-F and S-R to construct pδCBH2-D).
Table III-2 PCR primers used in this study
Primers Sequence (5’-3’)
δ-F TCACTGGATCTGCATTGAGAAATGGGTGAATGTTGAG
δEG-R CCACAGTTGATGTGGTGTTGGAATAGAAATCAACTATCATC
δC1-R GCTGTGGCAGGTTGTTGTTGGAATAGAAATCAACTATCATC
δC2-R TCTCTTTTATAGGAATGTTGGAATAGAAATCAACTATCATC
SEG-F ATTTCTATTCCAACACCACATCAACTGTGGGAATAC
SC1-F ATTTCTATTCCAACAACAACCTGCCACAGCTTTTCAA
SC2-F ATTTCTATTCCAACATTCCTATAAAAGAGAAGCGTATAAAAC
S-R TTCACCCATTTCTCAATGCAGATCCAGTGAGCG
Plasmids were transformed into S. cerevisiae BY4741 using lithium acetate as
described68. In cocktail δ-integration method, identical amounts of three δ-integrative
plasmids (over 20 µg of each plasmid), pδEG-D, pδCBH1-D, and pδCBH2-D, were
mixed and transformed simultaneously. The transformants with the highest
cellulolytic activities were selected based on their degradation abilities towards
crystalline cellulose. Cell growth of parent strain S. cerevisiae BY4741 and
recombinant strains were measured as described by Liu et al96.
Avicelase activity assay
The Avicelase activity of yeast cells was determined to indicate their degradation
ability towards microcrystalline cellulose. Recombinant colonies were picked up and
78
cultivated at 30 °C for 24 h in SD medium. The culture was then inoculated into YPD
medium, and grew at 30 °C, 300 rpm for 72 h. Yeast cells were subsequently
collected via centrifuge and washed twice by sterilized water. The Avicelase activities
of transformants were performed in a 48-well microplate. Each well involves 1%
Avicel solution, 100 mM methylglyoxal (Nacalai Tesque, Inc., Kyoto, Japan) and 100
g wet cells/L. The enzymatic reaction was carried out at 50 ºC, 300 rpm for 24 h. The
Avicelase activity of strain EG-D-CBH1-D-CBH2-D was used as control.
Methylglyoxal has been reported to inhibit the transportation of glucose into cells119.
The yeast cell surface glucose sensors Rgt2 and Snf3 act as glucose receptors that
detect extracellular glucose and generated an inductive signal for gene expression of
glucose transporters. Methylglyoxal can inactivate these glucose sensors, blocking the
production of glucose transporter119. As a result, the glucose generated from Avicel
degradation is remained in supernatant, rather than being assimilated by yeast cells.
After the reaction, the supernatant was obtained by centrifugation at 8,000 × g, 10 min,
4 °C. Glucose concentration in the supernatant was measured by the Glucose CII kit
(Wako Pure Chemical Industries, Ltd., Osaka, Japan). The principle of Glucose CII
kit is followed: the α-D-glucose existing in the sample is converted rapidly to
β-D-glucose under the action of mutarotase. β-D-glucose is oxidized by glucose
oxidase (GOD) to produce hydrogen peroxide, which is subsequently converted into
red pigment using peroxidase (POD). The absorbance of pigment is measured at 505
nm. One unit of Avicelase activity (U/mL) was defined as the amount of enzyme
needed to produce 1 µmoL of glucose per 24 h at 50 °C, pH 5.0.
Ethanol production from cellulosic materials
Fermentations were performed at 37 °C at an agitation speed of 200 rpm in 30
mL glass serum bottles, which are sealed using rubber stoppers with a needle for the
removal of CO2 produced during fermentation. Pre-cultivated cells in YPD medium
79
were centrifuged and washed twice by sterilized water. The collected cells were
inoculated into 10 mL YP medium containing 10 g/L Avicel, or 100 g/L MC6.
Pre-cultures of yeast strains were inoculated with an average cell concentration of
150 g wet cells/L. The ethanol concentrations in the fermentation medium were
determined using a gas chromatograph, as described by Yamada et al48.
Results
Screening of transformants with increased cellulose degradation abilities
Three cellulase genes, EG, CBH1, and CBH2 were introduced into a
BGL-displaying yeast (strain BY-BG-SS, described in Chapter I) simultaneously
using different amino acid markers (Figure III-1). As a result, a pool of recombinant
yeasts with various enzyme ratios was constructed. Then the strains with optimized
enzyme ratios were screened from over 500 numbers of transformants based on their
degradation abilities toward Avicel (namely, Avicelase activity). Twenty-nine
transformants exhibiting higher Avicelase activities than that of control strain
EG-D-CBH1-D-CBH2-D were illustrated in Figure III-2. The Avicelase activities in
δ-integrated strains were mostly over 20% higher than that in single-integrated strain
(Control). Strain B6 showed the highest Avicelase activity (97 U/mL), corresponding
to 1.5-fold the level seen in control strain. Alternatively, the Avicelase activity of
strain A26 (89 U/mL) was appropriately 43% higher than that of control strain. These
results demonstrate that it is efficient to reach high cellulolytic activities in yeast cells
through expression of multiple-copy cellulase gene. The enzyme ratios on the cell
surface are still under analysis.
80
….�
Avi
cel�
Mea
sure
cel
l gro
wth
and
A
vice
l fer
men
tatio
n�
Coc
ktai
l of E
G, C
BH
1,
CB
H2
expr
essi
on p
lasm
ids�
Qua
ntita
tion
of su
rfac
e-di
spla
yed
cellu
lase�
Pick
up
posi
tive
colo
nies�
Scre
enin
g�
Nan
o L
C-M
S/M
S�
Pool
of c
ellu
loly
tic
yeas
t str
ains�
BG
L-d
ispl
ayin
g y
east
stra
in�
Met
hylg
lyox
al�
Glu
cose�
BG
L�
EG�
CB
H1�
CB
H2�
Yeas
t cel
l�
Figu
re I
II-1
Sch
emat
ic d
escr
iptio
n of
the
con
stru
ctio
n an
d sc
reen
ing
of h
igh-
cellu
loly
tic y
east
stra
ins
in t
his
stud
y. B
GL,
β-gl
ucos
idas
e ; E
G, E
ndog
luca
nase
; CB
H1,
Cel
lobi
ohyd
rola
se 1
; CB
H2,
Cel
lobi
ohyd
rola
se 2
.
81
50
60
70
80
90
100
110
Contro
l A1 A15
A16
A18
A23
A24
A26
A27
B6 B12
B15
B19
B20
B22
B23
B24
B26
B30
B37
C1 C4 C5 C6 C9 C11
C18
C20
C21
C25
Avicelase activity �U/mL)�
Coc
ktai
l δ-in
tegr
ated
str
ains
Figu
re I
II-2
The
Avi
cela
se a
ctiv
ities
of c
ockt
ail δ
-inte
grat
ion
trans
form
ants
. The
Avi
cela
se a
ctiv
ity o
f stra
in E
G-D
-CB
H1-
D-C
BH
2-D
(sin
gle-
copy
int
egra
ted
stra
in)
was
tak
en a
s co
ntro
l. Th
e A
vice
lase
act
ivity
of
yeas
t ce
lls i
ndic
ates
the
ir de
grad
atio
n ab
ility
tow
ards
mic
rocr
ysta
lline
cel
lulo
se. O
ne u
nit o
f Avi
cela
se a
ctiv
ity w
as d
efin
ed a
s the
am
ount
of e
nzym
e ne
eded
to p
rodu
ce 1
µm
ol o
f glu
cose
per
24 h
at 5
0 °C
, pH
5.0
(U/m
L). V
alue
s ar
e pr
esen
ted
as th
e m
ean
± SD
from
thre
e in
depe
nden
t exp
erim
ents
.
82
Cell growth and cellulosic ethanol production
Generally, overloaded expression of heterologous proteins in yeast cell leads to
decreases in cell growth and ethanol fermentation capacity, referred as the metabolic
burden. Thus, the growth deficiency associated with metabolic burden should be
considered in the development of cellulolytic yeast strains. A cell growth comparison
of cocktail δ-integrated strains and control strain is presented in Table III-3. Aerobic
conditions were used to fully understand the underlying metabolic effects associated
with recombinant protein production. Although most of the δ-integrated strains
showed limited metabolic burden on cell growth compared with reference strain, six
transformants exhibited significant decrease in cell growth. The maximum specific
growth rate (µmax) of strains B12, B26 (0.35 h-1) are significantly lower than that of
control (0.44 h-1); the periods of lag phase in strains B24 (3 h), B26 (4 h), and B37 (4
h) are longer than in the other strains (2 h). Notably, although strain A24 showed
similar values of µmax and lag phase to those of control strain, it still yielded a
relatively low biomass concentration after 12 h cultivation, because its increase speed
of specific growth rate (µ) was slower than the control in exponential phase.
Subsequently, to obtain a high-cellulolytic yeast strain without growth deficiency
associated with metabolic burdens, the cellulosic ethanol yields of cocktail
δ-integrated strains were investigated. As shown in Figure III-3, most of
transformants gained higher ethanol yields from Avicel than the single-integrated
control strain (39%). Strain A26 achieved the highest cellulosic ethanol yield (57%,
corresponding to 1.46-fold of control strain) without any metabolic burden on cell
growth. On the contrary, some growth-deficient strains appeared much lower ethanol
yield than the control, indicating that metabolic burdens probably influence the
ethanol fermentation capacities in yeast cells. Interestingly, growth-deficient strain
A24 (with an ethanol yield of 55%) showed 1.41-fold of reference strain, suggesting
83
Table III-3 Growth of cocktail δ-integrated strains. Cell growth was measured under aerobic
condition in YPD medium. Mean values from triplicate experiments with standard deviations
are shown.
Strains Lag phase
(h)
Specific growth rate
(µmax/ h)
12 h biomass concentration
(OD660)
Control 2 0.44±0.02 1.83±0.07
A1 2 0.40±0.01 1.62 ±0.19
A15 2 0.41±0.02 1.65±0.05
A16 2 0.38±0.01 1.31±0.17
A18 2 0.44±0.02 1.82±0.18
A23 2 0.42±0.02 1.60±0.03
A24 2 0.40±0.02 0.92±0.05
A26 2 0.44±0.01 1.73±0.10
A27 2 0.41±0.02 1.65±0.09
B6 2 0.47±0.02 2.05±0.13
B12 2 0.35±0.02 1.11±0.09
B15 2 0.49±0.04 2.08±0.25
B19 2 0.44±0.02 2.00±0.33
B20 2 0.43±0.01 1.83±0.07
B22 2 0.41±0.01 1.61±0.12
B23 2 0.44±0.02 1.89±0.22
B24 3 0.45±0.02 1.00±0.16
B26 4 0.35±0.01 0.40±0.38
B30 2 0.43±0.02 1.91±0.04
B37 4 0.40±0.02 0.93±0.12
C1 2 0.41±0.02 1.98±0.13
C4 2 0.46±0.01 2.03±0.05
C5 3 0.42±0.03 1.51±0.07
C6 2 0.45±0.04 2.05±0.06
C9 2 0.45±0.01 2.00±0.17
C11 2 0.42±0.02 1.97±0.19
C18 2 0.46±0.02 2.00±0.24
C20 2 0.45±0.01 1.94±0.03
C21 2 0.44±0.02 2.30±0.15
C25 2 0.41±0.02 1.50±0.09
84
that its cellulolytic activities on the cell surface is possibly high enough to make up
the deficiency of metabolic burden. The amount of cellulase displayed on yeast
surface will be evaluated via nano LC-MS/MS in the further experiment. Considering
of heterologous protein expression and metabolic burdens, strain A26 emerges as the
best one among the cocktail δ-integration strains.
Figure III-3 Cell growth and cellulosic ethanol yields of cocktail δ-integrated strains.
Single-integrated strain EG-D-CBH1-D-CBH2-D was used as control. Ethanol was produced
from 10 g/L Avicel. The condition of cell growth measurement is the same with that in Table
III-3. Data are presented as the means and standard deviations of triplicate measurements.
Control
A18�
A23�A24�
A26
B6
A16
B24�B26�
B37�
B12
C5
C25�
15
20
25
30
35
40
45
50
55
60
65
0.2 0.7 1.2 1.7 2.2
Cel
lulo
sic
etha
nol y
ield
(%)�
Cell growth (12 h biomass concentration)�
85
Discussion
In this study, a series of recombinant yeast strains with high degradation abilities
towards crystalline cellulose were developed via optimizing the ratio of three types of
cellulase displayed on the cell surface. Notably, we obtained a strain A26 producing a
cellulosic ethanol titer corresponding to 57% of the theoretical yield. To our
knowledge, this is the highest ethanol yield from crystalline cellulose using
cellulolytic yeast among the current reports84, 92.
Currently, much effort has been done to engineer yeast strains to directly convert
cellulose, especially the crystalline cellulose into bioethanol, but the cellulolytic
activities in these strains are still insufficient. Fan et al. designed a delicate
miniscaffoldin on the surface of S. cerevisiae, achieving an ethanol yield of 27% from
10 g/L Avicel84. In addition, a yeast strain expressing a cocktail of cellulase genes was
constructed92; but this strain could only generate an ethanol yield of 12% from Avicel.
In contrast, using cocktail δ-integration method, several kinds of cellulase genes are
integrated into yeast chromosomes simultaneously in one step, and strains with high
cellulolytic activity (that is, expressing the optimum ratio of cellulases) can be easily
obtained. In Figure III-2, around one-second of the twenty-nine selected transformants
produced ethanol yields of over 45%. Especially, strain A26 exhibited an ethanol
yield of 57%, demonstrating the effectiveness of this method.
However, the production of large amounts of heterologous protein leads to
obvious growth deficiency in recombinant strains. We found that the lag phases of
some transformants were extended, indicating that the adaptive capacity of these
strains to cultivation environment declined; on the other hand, the µmax of some
transformants were decreased. A possible explanation for the lower µmax value could
reside in the metabolic burden from the overloaded heterologous protein expression.
Generally, the metabolic burden in yeast cells generates a decrease in the maximum
86
specific growth rate, biomass yield and respiratory capacity as well as an increase in
maintenance requirement120. The generation of metabolic burden has been attributed
to a broad variety of causes, such as the cellular stress associated with protein folding
and degradation of misfolded protein in the endoplasmic reticulum (ER)121, and
competition for resources to synthesize native protein and heterologous enzyme122.
These stresses would lead to a drain on the energy metabolism, which have negatively
impacted on the µmax of recombinant strain. The relationships between cell growth
(Table III-3) and Avicelase activities (Figure III-2) also support this hypothesis, that
the strains with higher cellulolytic activities (e.g., strain A16, A24, and B26) also
possessed defects in cell growth. Notably, growth-deficient strain A24 showed higher
cellulosic ethanol yield than control, indicating that its intracellular burden probably
emerged as the cell-growth-related stress in ER, rather than competing energy
resource (e.g., NADPH) with glycolysis pathway. On the contrary, strain B6 exhibited
the highest cellulolytic activities among transformants and similar cell growth to
control, but its ethanol yield is relatively low. It is postulated that the increased carbon
flux (more available glucose caused by improved cellulose degradation ability) in B6
did not flow into the synthesis of ethanol owing to the effect of metabolic burden.
Another potential reason for the growth deficiency is the random, vast
integration of genes into the δ-regions. δ-Integration simultaneously occurs on
multiple chromosomes due to the presence of around 425 δ-regions dispersed
throughout the yeast genome123. The random integration of genes into δ-regions
possibly up-/ down-regulates the expression level of some critical genes involved in
cell propagation. Currently, no obvious growth deficiency caused by δ-integration has
been observed in other reports, except that the stability of the integrated gene varied
with the integration sites on the genome after 50 generations of subculture124.
87
Many researchers have attempted to reduce the metabolic burden derived from
protein expression in yeast strains, such as employing of a non-growth related
expression system for heterologous protein gene expression125, or releasing of ER
stress through over-expression of chaperone and ubiquitin126. Canonaco et al. reported
the functional expression of arginine kinase in S. cerevisiae, setting up an intracellular
pool of phosphoarginine to maintain constant intracellular ATP levels upon
fluctuating energy demands; the engineered strain conferred a clear reduce of lag
phase period in growth127. However, owing to the disproportionate complexity of
intracellular activities, more detail information including metabolic flux balance
between amino acid biosynthesis pathway and glycolysis pathway, and the genes
against cellular stress sensitivity should be clarified to break the limitation of
metabolic burdens.
Conclusion
In this chapter, we successfully constructed a pool of cellulolytic S. cerevisiae
with diverse enzyme ratios on the cell surface using cocktail δ-integration method.
Strain A26 achieved the highest cellulosic ethanol yield from crystalline cellulose
(57%, corresponding to 1.46-fold of that in control strain (cellulase ratio is 1:1:1:1))
without any metabolic burden on cell growth. This study provided the insight that
optimization of cellulase ratios displayed on the yeast surface would significantly
increase its degradation ability towards crystalline cellulose, and ultimately result in
the overall viability of low-cost, sustainable cellulosic ethanol production.
88
GENERAL CONCLUSION
In this thesis, to achieve low-cost, environment-friendly degradation of cellulose
for bioethanol production, novel cellulolytic yeast strains were developed using
cell-surface display technique. Investigation of the suitable strategy for cellulase
production, optimization of enzyme ratio on the cell surface, and improvement of
cell-to-cellulose interactions were used to improve the cellulose degradation ability in
yeast strains. A summary of the results from each chapter is listed in Table IV. The
following can be concluded:
1. It was demonstrated that placement of EG and CBH1 in the same space (both
on the cell surface or both secreted into the medium) was favorable for ethanol
production from amorphous cellulose.
2. The cellulolytic yeast constructed via cell-surface display strategy was able to
accelerate the production rate of ethanol from cellulosic materials; it appeared
more effective in the cell-recycle batch fermentation than the strain
constructed via secretion strategy.
3. Increase of the enzyme variety (co-display of BGL, EG, CBH1, and CBH2) on
the cell surface could efficiently enhance the synergism between cellulases, as
a result, significantly improving the efficiency of cellulose hydrolysis.
4. Through cocktail δ-integration method, the enzyme ratio on the cell surface
was optimized, yielding a 1.46-fold higher cellulosic ethanol yield than the
unoptimized strain (enzyme ratio of 1:1:1:1).
5. The cellulase-displaying yeast strain exhibited clear cell-to-cellulose adhesion
and a “tearing” cellulose degradation pattern; the adhesion ability correlated
89
with enhanced surface area and roughness of the target cellulose fibers. Our
engineered strain could directly produce ethanol from rice straw, as well as
cutting down more than 40% of the required enzymes under high-dense
fermentation
6. This thesis provides a novel strategy to improve the efficiency of cellulose
hydrolysis: enhancement of the interactions between displayed-cellulase and
cellulosic substrate based on the mechanism of cell-to-cellulose adhesion.
This thesis presents some new understanding of the cellulose degradation
mechanism via cellulase-displaying yeast strain. The following is recommended for
future work.
1. Carbohydrate binding domain (CBD) has been report to possess high affinity
towards cellulose. Therefore, display of CBD on yeast surface is expected to
enhance the interactions of cell-to-cellulose as well as the cellulose
degradation ability.
2. Although the optimal biomass pretreatment method combined with
free-cellulase-mediated saccharification has been extensively studied, the
pretreatment method suitable for surface-displayed cellulases still remains
obscure, which will undoubtly improve the efficiency of cellulose hydrolysis.
Thus, the corresponding suitable pretreatment approach needs to be studied in
the future.
3. The combination of cell-surface display technique with metabolic engineering
aimed at redesigning microbial metabolic pathway will enable the production
of advanced biofuels and biochemicals from cellulose, presenting a new
opportunity in the biorefinery of cellulosic biomass.
90
91
REFERENCE
1. Nigam PS, Singh A. Production of liquid biofuels from renewable resources. Prog
Energy Combust Sci 37, 52-68 (2011).
2. Lynd LR, Van Zyl WH, McBride JE, Laser M. Consolidated bioprocessing of
cellulosic biomass: an update. Curr Opin Biotechnol 16, 577-583 (2005).
3. Ajanovic A. Biofuels versus food production: Does biofuels production increase food
prices? Energy 36, 2070-2076 (2011).
4. Naik SN, Goud VV, Rout PK, Dalai AK. Production of first and second generation
biofuels: A comprehensive review. Renew Sust Energ Rev 14, 578-597 (2010).
5. Limayem A, Ricke SC. Lignocellulosic biomass for bioethanol production: current
perspectives, potential issues and future prospects. Prog Energy Combust Sci 38,
449-467 (2012).
6. Himmel ME, et al. Biomass recalcitrance: engineering plants and enzymes for
biofuels production. Science 315, 804-807 (2007).
7. van Zyl WH, Lynd LR, den Haan R, McBride JE. Consolidated bioprocessing for
bioethanol production using Saccharomyces cerevisiae. Adv Biochem Eng Biotechnol
108, 205-235 (2007).
8. Hasunuma T, Okazaki F, Okai N, Hara KY, Ishii J, Kondo A. A review of enzymes
and microbes for lignocellulosic biorefinery and the possibility of their application to
consolidated bioprocessing technology. Bioresour Technol 135, 513-522 (2013).
9. Den Haan R, Rose SH, Lynd LR, van Zyl WH. Hydrolysis and fermentation of
amorphous cellulose by recombinant Saccharomyces cerevisiae Metab Eng 9, 87-94
(2007).
10. Yamada R, Taniguchi N, Tanaka T, Ogino C, Fukuda H, Kondo A. Direct ethanol
production from cellulosic materials using a diploid strain of Saccharomyces
cerevisiae with optimized cellulase expression. Biotechnol Biofuels 4, 8 (2011).
92
11. Wu CH, Mulchandani A, Chen W. Versatile microbial surface-display for
environmental remediation and biofuels production. Trends Microbiol 16, 181-188
(2008).
12. Lee SY, Choi JH, Xu Z. Microbial cell-surface display. Trends Biotechnol 21, 45-52
(2003).
13. Orlean P. Architecture and biosynthesis of the Saccharomyces cerevisiae cell wall.
Genetics 192, 775-818 (2012).
14. Kondo A, Ueda M. Yeast cell-surface display-applications of molecular display. Appl
Microbiol Biotechnol 64, 28-40 (2004).
15. Hara KY, et al. Development of a glutathione production process from proteinaceous
biomass resources using protease-displaying Saccharomyces cerevisiae. Appl
Microbiol Biotechnol 93, 1495-1502 (2012).
16. Bae J, Kuroda K, Ueda M. Proximity effect among cellulose-degrading enzymes
displayed on the Saccharomyces cerevisiae cell surface. Appl Environ Microb 81,
59-66 (2015).
17. Kuroda K, Ueda M. Bioadsorption of cadmium ion by cell surface-engineered yeasts
displaying metallothionein and hexa-His. Appl Microbiol Biotechnol 63, 182-186
(2003).
18. Liu W, Zhao H, Jia B, Xu L, Yan Y. Surface display of active lipase in
Saccharomyces cerevisiae using Cwp2 as an anchor protein. Biotechnol Lett 32,
255-260 (2010).
19. Kambe-Honjoh H, Ohsumi K, Shimoi H, Nakajima H, Kitamoto K. Molecular
breeding of yeast with higher metal-adsorption capacity by expression of
histidine-repeat insertion in the protein anchored to the cell wall. J Gen Appl
Microbiol 46, 113-117 (2000).
20. Mormeneo M, Andrés I, Bofill C, Díaz P, Zueco J. Efficient secretion of Bacillus
subtilis lipase A in Saccharomyces cerevisiae by translational fusion to the Pir4 cell
wall protein. Appl Microbiol Biotechnol 80, 437-445 (2008).
93
21. Matsumoto T, Fukuda H, Ueda M, Tanaka A, Kondo A. Construction of yeast strains
with high cell surface lipase activity by using novel display systems based on the
Flo1p flocculation functional domain. Appl Environ Microbiol 68, 4517-4522 (2002).
22. Sato N, Matsumoto T, Ueda M, Tanaka A, Fukuda H, Kondo A. Long anchor using
Flo1 protein enhances reactivity of cell surface-displayed glucoamylase to polymer
substrates. Appl Microbiol Biotechnol 60, 469-474 (2002).
23. Xu L, et al. Preparation of a promising whole cell biocatalyst of Geotrichum sp.
lipase and its properties. J Chem Technol Biotechnol 87, 498-504 (2012).
24. Nam KT, Lee YJ, Krauland EM, Kottmann ST, Belcher AM. Peptide-mediated
reduction of silver ions on engineered biological scaffolds. ACS Nano 2, 1480-1486
(2008).
25. Inokuma K, Hasunuma T, Kondo A. Efficient yeast cell-surface display of exo-and
endo-cellulase using the SED1 anchoring region and its original promoter. Biotechnol
Biofuels 7, (2014).
26. Ni X, Yue L, Chi Z, Li J, Wang X, Madzak C. Alkaline protease gene cloning from
the marine yeast Aureobasidium pullulans HN2-3 and the protease surface display on
Yarrowia lipolytica for bioactive peptide production. Mar Biotechnol (NY) 11, 81-89
(2009).
27. Song H, et al. Construction of a whole-cell catalyst displaying a fungal lipase for
effective treatment of oily wastewaters. J Mol Catal B: Enzym 71, 166-170 (2011).
28. Pan X-X, et al. Efficient display of active Geotrichum sp. lipase on Pichia pastoris
cell wall and its application as a whole-cell biocatalyst to enrich EPA and DHA in
fish oil. J Agric Food Chem 60, 9673-9679 (2012).
29. Han Z-l, Han S-y, Zheng S-p, Lin Y. Enhancing thermostability of a Rhizomucor
miehei lipase by engineering a disulfide bond and displaying on the yeast cell surface.
Appl Microbiol Biotechnol 85, 117-126 (2009).
30. Liang X-x, et al. Quantitative evaluation of Candia antarctica lipase B displayed on
the cell surface of a Pichia pastoris based on an FS anchor system. Biotechnol Lett 35,
367-374 (2013).
94
31. Yan YJ, Xu L, Dai M. A synergetic whole-cell biocatalyst for biodiesel production.
RSC Adv 2, 6170-6173 (2012).
32. Jin Z, Han SY, Zhang L, Zheng SP, Wang Y, Lin Y. Combined utilization of
lipase-displaying Pichia pastoris whole-cell biocatalysts to improve biodiesel
production in co-solvent media. Bioresour Technol 130, 102-109 (2013).
33. Moura MVH, da Silva GP, de Oliveira Machado AC, Torres FAG, Freire DMG,
Almeida RV. Displaying lipase B from Candida antarctica in Pichia pastoris using
the yeast surface display approach: Prospection of a new anchor and characterization
of the whole cell biocatalyst. PloS One 10, e0141454 (2015).
34. Wang Q, Li L, Chen M, Qi Q, Wang PG. Construction of a novel Pichia pastoris
cell-surface display system based on the cell wall protein Pir1. Curr Microbiol 56,
352-357 (2008).
35. Yanase S, et al. Direct ethanol production from cellulosic materials at high
temperature using the thermotolerant yeast Kluyveromyces marxianus displaying
cellulolytic enzymes. Appl Microbiol Biotechnol 88, 381-388 (2010).
36. Kim SY, Sohn J-H, Pyun Y-R, Yang IS, Kim K-H, Choi E-S. In vitro evolution of
lipase B from Candida antarctica using surface display in hansenula polymorpha. J
Microbiol Biotechnol 17, 1308-1315 (2007).
37. Hasunuma T, Kondo A. Development of yeast cell factories for consolidated
bioprocessing of lignocellulose to bioethanol through cell surface engineering.
Biotechnol Adv 30, 1207-1218 (2012).
38. Pack SP, Park K, Yoo YJ. Enhancement of β-glucosidase stability and
cellobiose-usage using surface-engineered recombinant Saccharomyces cerevisiae in
ethanol production. Biotechnol Lett 24, 1919-1925 (2002).
39. Matano Y, Hasunuma T, Kondo A. Display of cellulases on the cell surface of
Saccharomyces cerevisiae for high yield ethanol production from high-solid
lignocellulosic biomass. Bioresour Technol 108, 128-133 (2012).
40. Kotaka A, Sahara H, Kuroda K, Kondo A, Ueda M, Hata Y. Enhancement of
95
β-glucosidase activity on the cell-surface of sake yeast by disruption of SED1 J
Biosci Bioeng 109, 442-446 (2010).
41. Van Zyl JHD, Den Haan R, Van Zyl WH. Over-expression of native Saccharomyces
cerevisiae exocytic SNARE genes increased heterologous cellulase secretion. Appl
Microbiol Biotechnol 98, 5567-5578 (2014).
42. Tsai SL, DaSilva NA, Chen W. Functional display of complex cellulosomes on the
yeast surface via adaptive assembly. ACS Synth Biol 2, 14-21 (2013).
43. Nakatani Y, Yamada R, Ogino C, Kondo A. Synergetic effect of yeast cell-surface
expression of cellulase and expansin-like protein on direct ethanol production from
cellulose. Microb Cell Fact 12, 66 (2013).
44. Yamada R, Taniguchi N, Tanaka T, Ogino C, Fukuda H, Kondo A. Cocktail
δ-integration: a novel method to construct cellulolytic enzyme expression
ratio-optimized yeast strains. Microb Cell Fact 9, 32 (2010).
45. Nakanishi A, Kuroda K, Ueda M. Direct fermentation of newspaper after
laccase-treatment using yeast codisplaying endoglucanase, cellobiohydrolase, and
β-glucosidase. Renew Energ 44, 199-205 (2012).
46. Matano Y, Hasunuma T, Kondo A. Cell recycle batch fermentation of high-solid
lignocellulose using a recombinant cellulase-displaying yeast strain for high yield
ethanol production in consolidated bioprocessing. Bioresour Technol 135, 403-409
(2013).
47. Nakanishi A, et al. Effect of pretreatment of hydrothermally processed rice straw
with laccase-displaying yeast on ethanol fermentation. Appl Microbiol Biotechnol 94,
939-948 (2012).
48. Yamada R, Nakatani Y, Ogino C, Kondo A. Efficient direct ethanol production from
cellulose by cellulase- and cellodextrin transporter-co-expressing Saccharomyces
cerevisiae. AMB Express doi: 10.1186/2191-0855-3-34. , (2013).
49. Fujita Y, Ito J, Ueda M, Fukuda H, Kondo A. Synergistic saccharification, and direct
fermentation to ethanol, of amorphous cellulose by use of an engineered yeast strain
codisplaying three types of cellulolytic enzyme. Appl Environ Microb 70, 1207-1212
96
(2004).
50. Baek S-H, Kim S, Lee K, Lee J-K, Hahn J-S. Cellulosic ethanol production by
combination of cellulase-displaying yeast cells. Enzyme Microb Technol 51, 366-372
(2012).
51. Fujita Y, et al. Direct and efficient production of ethanol from cellulosic material
with a yeast strain displaying cellulolytic enzymes. Appl Environ Microb 68,
5136-5141 (2002).
52. Garvey M, Klose H, Fischer R, Lambertz C, Commandeur U. Cellulases for biomass
degradation: comparing recombinant cellulase expression platforms. Trends
Biotechnol 31, 581-593 (2013).
53. Hasunuma T, Kondo A. Consolidated bioprocessing and simultaneous
saccharification and fermentation of lignocellulose to ethanol with thermotolerant
yeast strains. Process Biochem 47, 1287-1294 (2012).
54. Sanda T, Hasunuma T, Matsuda F, Kondo A. Repeated-batch fermentation of
lignocellulosic hydrolysate to ethanol using a hybrid Saccharomyces cerevisiae strain
metabolically engineered for tolerance to acetic and formic acids. Bioresour Technol
102, 7917-7924 (2011).
55. Kondo A, et al. High-level ethanol production from starch by a flocculent
Saccharomyces cerevisiae strain displaying cell-surface glucoamylase. Appl
Microbiol Biotechnol 58, 291-296 (2002).
56. den Haan R, van Rensburg E, Rose SH, Görgens JF, van Zyl WH. Progress and
challenges in the engineering of non-cellulolytic microorganisms for consolidated
bioprocessing. Curr Opin Biotechnol 33, 32-38 (2015).
57. Resch MG, et al. Fungal cellulases and complexed cellulosomal enzymes exhibit
synergistic mechanisms in cellulose deconstruction. Energy Environ Sci 6, 1858-1867
(2013).
58. Arantes V, Saddler JN. Access to cellulose limits the efficiency of enzymatic
hydrolysis: the role of amorphogenesis. Biotechnol Biofuels 3, (2010).
97
59. Hu J, Gourlay K, Arantes V, Van Dyk JS, Pribowo A, Saddler JN. The accessible
cellulose surface influences cellulase synergism during the hydrolysis of
lignocellulosic substrates. ChemSusChem 8, 901-907 (2015).
60. Den Haan R, Mcbride JE, Grange DlCL, Lynd LR, Van Zyl WH. Functional
expression of cellobiohydrolases in Saccharomyces cerevisiae towards one-step
conversion of cellulose to ethanol. Enzyme Microb Technol 40, 1291-1299 (2007).
61. Takada G, Kawaguchi T, Sumitani J, Arai M. Expression of Aspergillus aculeatus No.
F-50 cellobiohydrolase I (cbhI) and beta-glucosidase 1 (bgl1) genes by
Saccharomyces cerevisiae. Biosci Biotechnol Biochem 62, 1615-1618 (1998).
62. den Haan R, Kroukamp H, van Zyl J-HD, van Zyl WH. Cellobiohydrolase secretion
by yeast: current state and prospects for improvement. Process Biochem 48, 1-12
(2013).
63. Ilmén M, et al. High level secretion of cellobiohydrolases by Saccharomyces
cerevisiae. Biotechnol Biofuels 4, 30 (2011).
64. Brevnova E MJ, et al. Yeast expressing saccharolytic enzymes for consolidated
bioprocessing using starch and cellulose. PCT/US2011/039192(WO/2011/153516)
12-8-2011 3-6-0110.
65. McBride JE BE, et al. Yeast expressing cellulases for simultaneous saccharification
and fermentation using cellulose. PCT/US2009/065571 5-27-2010.
66. Gibson DG, Young L, Chuang R-Y, Venter JC, Hutchison CA, Smith HO. Enzymatic
assembly of DNA molecules up to several hundred kilobases. Nat Methods 6,
343-345 (2009).
67. Inokuma K, Yoshida T, Ishii J, Hasunuma T, Kondo A. Efficient co-displaying and
artificial ratio control of α-amylase and glucoamylase on the yeast cell surface by
using combinations of different anchoring domains. Appl Microbiol Biotechnol 99,
1655-1663 (2014).
68. Chen D-C, Yang B-C, Kuo T-T. One-step transformation of yeast in stationary phase.
Curr Genet 21, 83-84 (1992).
98
69. Ismail KSK, Sakamoto T, Hatanaka H, Hasunuma T, Kondo A. Gene expression
cross-profiling in genetically modified industrial Saccharomyces cerevisiae strains
during high-temperature ethanol production from xylose. J Biotechnol 163, 50-60
(2013).
70. Kalapos MP. Methylglyoxal in living organisms: chemistry, biochemistry, toxicology
and biological implications. Toxicol Lett 110, 145-175 (1999).
71. Yamada R, et al. Effective saccharification of kraft pulp by using a cellulase cocktail
prepared from genetically engineered Aspergillus oryzae. Biosci Biotechnol Biochem,
1-4 (2015).
72. Bunterngsook B, Eurwilaichitr L, Thamchaipenet A, Champreda V. Binding
characteristics and synergistic effects of bacterial expansins on cellulosic and
hemicellulosic substrates. Bioresour Technol 176, 129-135 (2015).
73. Andersen N, Johansen KS, Michelsen M, Stenby EH, Krogh KBRM, Olsson L.
Hydrolysis of cellulose using mono-component enzymes shows synergy during
hydrolysis of phosphoric acid swollen cellulose (PASC), but competition on Avicel.
Enzyme Microb Technol 42, 362-370 (2008).
74. Reyes-Ortiz V, et al. Addition of a carbohydrate-binding module enhances cellulase
penetration into cellulose substrates. Biotechnol Biofuels 6, (2013).
75. Shimoi H, Kitagaki H, Ohmori H, Iimura Y, Ito K. Sed1p is a major cell wall protein
of Saccharomyces cerevisiae in the stationary phase and is involved in lytic enzyme
resistance. J Bacteriol 180, 3381-3387 (1998).
76. Yanase S, et al. Ethanol production from cellulosic materials using cellulase‐
expressing yeast. Biotechnol J 5, 449-455 (2010).
77. Griggs AJ, Stickel JJ, Lischeske JJ. A mechanistic model for enzymatic
saccharification of cellulose using continuous distribution kinetics I:
depolymerization by EGI and CBHI. Biotechnol Bioeng 109, 665-675 (2012).
78. Peckys DB, Mazur P, Gould KL, de Jonge N. Fully hydrated yeast cells imaged with
electron microscopy. Biophys J 100, 2522-2529 (2011).
99
79. Ju X, Grego C, Zhang X. Specific effects of fiber size and fiber swelling on biomass
substrate surface area and enzymatic digestibility. Bioresour Technol 144, 232-239
(2013).
80. Lynd LR, Weimer PJ, Van Zyl WH, Pretorius IS. Microbial cellulose utilization:
fundamentals and biotechnology. Microbiol Mol Biol Rev 66, 506-577 (2002).
81. Cruys-Bagger N, et al. Pre-steady-state kinetics for hydrolysis of insoluble cellulose
by cellobiohydrolase Cel7A. J Biol Chem 287, 18451-18458 (2012).
82. Kurasin M, Valjamae P. Processivity of cellobiohydrolases is limited by the substrate.
J Biol Chem 286, 169-177 (2011).
83. Francisco JA, Stathopoulos C, Warren RA, Kilburn DG, Georgiou G. Specific
adhesion and hydrolysis of cellulose by intact Escherichia coli expressing surface
anchored cellulase or cellulose binding domains. Biotechnology (N Y) 11, 491-495
(1993).
84. Fan L-H, Zhang Z-J, Yu X-Y, Xue Y-X, Tan T-W. Self-surface assembly of
cellulosomes with two miniscaffoldins on Saccharomyces cerevisiae for cellulosic
ethanol production. PNAS 109, 13260-13265 (2012).
85. Jin M, Gunawan C, Uppugundla N, Balan V, Dale BE. A novel integrated biological
process for cellulosic ethanol production featuring high ethanol productivity, enzyme
recycling and yeast cells reuse. Energy Environ Sci 5, 7168-7175 (2012).
86. Watanabe I, Miyata N, Ando A, Shiroma R, Tokuyasu K, Nakamura T. Ethanol
production by repeated-batch simultaneous saccharification and fermentation (SSF)
of alkali-treated rice straw using immobilized Saccharomyces cerevisiae cells.
Bioresour Technol 123, 695-698 (2012).
87. Khaw TS, Katakura Y, Koh J, Kondo A, Ueda M, Shioya S. Evaluation of
performance of different surface-engineered yeast strains for direct ethanol
production from raw starch. Appl Microbiol Biotechnol 70, 573-579 (2006).
88. Sheehan J. Engineering direct conversion of CO2 to biofuel. Nat Biotechnol 27,
1128-1129 (2009).
100
89. Lynd LR, Wyman CE, Gerngross TU. Biocommodity Engineering. Biotechnol Prog
15, 777-793 (1999).
90. Schubert C. Can biofuels finally take center stage? Nat Biotechnol 24, 777-784
(2006).
91. Piskur J, Rozpedowska E, Polakova S, Merico A, Compagno C. How did
Saccharomyces evolve to become a good brewer? Trends Genet 22, 183-186 (2006).
92. Chang J-J, et al. Assembling a cellulase cocktail and a cellodextrin transporter into a
yeast host for CBP ethanol production. Biotechnol Biofuels 6, 19-31 (2013).
93. Wen F, Sun J, Zhao H. Yeast surface display of trifunctional minicellulosomes for
simultaneous saccharification and fermentation of cellulose to ethanol. Appl Environ
Microbiol 76, 1251-1260 (2010).
94. Morais S, Shterzer N, Lamed R, Bayer EA, Mizrahi I. A combined cell-consortium
approach for lignocellulose degradation by specialized Lactobacillus plantarum cells.
Biotechnol Biofuels 7, 112 (2014).
95. Sasaki K, et al. Mechanical milling and membrane separation for increased ethanol
production during simultaneous saccharification and co-fermentation of rice straw by
xylose-fermenting Saccharomyces cerevisiae. Bioresour Technol 185, 263-268
(2015).
96. Liu Z, et al. Combined cell-surface display- and secretion-based strategies for
production of cellulosic ethanol with Saccharomyces cerevisiae. Biotechnol Biofuels
8, 162 (2015).
97. Kroukamp H, den Haan R, van Wyk N, van Zyl WH. Overexpression of native PSE1
and SOD1 in Saccharomyces cerevisiae improved heterologous cellulase secretion.
Appl Energy 102, 150-156 (2013).
98. Adney B, Baker J. Measurement of cellulase activities. Laboratory analytical
procedure 6, 1996 (1996).
99. Moukadiri I, Armero J, Abad A, Sentandreu R, Zueco J. Identification of a
mannoprotein present in the inner layer of the cell wall of Saccharomyces cerevisiae.
101
J Bacteriol 179, 2154-2162 (1997).
100. Glick BR. Metabolic load and heterologous gene expression. Biotechnol Adv 13,
247-261 (1995).
101. Goyal G, Tsai S-L, Madan B, DaSilva NA, Chen W. Simultaneous cell growth and
ethanol production from cellulose by an engineered yeast consortium displaying a
functional mini-cellulosome. Microb Cell Fact 10, 89 (2011).
102. Li XZ, Lu J, Zhao J, Qu YB. Characteristics of corn stover pretreated with liquid hot
water and fed-batch semi-simultaneous saccharification and fermentation for
bioethanol production. Plos One 9, (2014).
103. Alvira P, Tomas-Pejo E, Ballesteros M, Negro MJ. Pretreatment technologies for an
efficient bioethanol production process based on enzymatic hydrolysis: A review.
Bioresour Technol 101, 4851-4861 (2010).
104. Valjamae P, Sild V, Pettersson G, Johansson G. The initial kinetics of hydrolysis by
cellobiohydrolases I and II is consistent with a cellulose surface-erosion model. Eur J
Biochem 253, 469-475 (1998).
105. Tsai SL, Oh J, Singh S, Chen R, Chen W. Functional assembly of minicellulosomes
on the Saccharomyces cerevisiae cell surface for cellulose hydrolysis and ethanol
production. Appl Environ Microbiol 75, 6087-6093 (2009).
106. Wang AA, Mulchandani A, Chen W. Specific adhesion to cellulose and hydrolysis of
organophosphate nerve agents by a genetically engineered Escherichia coli strain
with a surface-expressed cellulose-binding domain and organophosphorus hydrolase.
Appl Environ Microbiol 68, 1684-1689 (2002).
107. Brunecky R, et al. Revealing nature's cellulase diversity: the digestion mechanism of
Caldicellulosiruptor bescii CelA. Science 342, 1513-1516 (2013).
108. Meng XZ, Foston M, Leisen J, DeMartini J, Wyman CE, Ragauskas AJ.
Determination of porosity of lignocellulosic biomass before and after pretreatment by
using Simons' stain and NMR techniques. Bioresour Technol 144, 467-476 (2013).
109. Wang QQ, et al. Evaluations of cellulose accessibilities of lignocelluloses by solute
102
exclusion and protein adsorption techniques. Biotechnol Bioeng 109, 381-389 (2012).
110. Li C, et al. Influence of physico-chemical changes on enzymatic digestibility of ionic
liquid and AFEX pretreated corn stover. Bioresour Technol 102, 6928-6936 (2011).
111. Paulova L, Patakova P, Branska B, Rychtera M, Melzoch K. Lignocellulosic ethanol:
Technology design and its impact on process efficiency. Biotechnol Adv 33,
1091-1107 (2014).
112. Lopez-Linares JC, Romero I, Cara C, Ruiz E, Moya M, Castro E. Bioethanol
production from rapeseed straw at high solids loading with different process
configurations. Fuel 122, 112-118 (2014).
113. Ko JK, et al. Ethanol production from rice straw using optimized aqueous-ammonia
soaking pretreatment and simultaneous saccharification and fermentation processes.
Bioresour Technol 100, 4374-4380 (2009).
114. Lupoi JS, Smith EA. Evaluation of nanoparticle-immobilized cellulase for improved
ethanol yield in simultaneous saccharification and fermentation reactions. Biotechnol
Bioeng 108, 2835-2843 (2011).
115. La Grange DC, Den Haan R, Van Zyl WH. Engineering cellulolytic ability into
bioprocessing organisms. Appl Microbiol Biotechnol 87, 1195-1208 (2010).
116. Thygesen A, Oddershede J, Lilholt H, Thomsen AB, Stahl K. On the determination of
crystallinity and cellulose content in plant fibres. Cellulose 12, 563-576 (2005).
117. Yamada R, Hasunuma T, Kondo A. Endowing non-cellulolytic microorganisms with
cellulolytic activity aiming for consolidated bioprocessing. Biotechnol Adv 31,
754-763 (2013).
118. Liu Z, et al. Engineering of a novel cellulose-adherent cellulolytic Saccharomyces
cerevisiae for cellulosic biofuel production. Sci Rep 6, 24550 (2016).
119. Roy A, Hashmi S, Li Z, Dement AD, Cho KH, Kim J-H. The glucose metabolite
methylglyoxal inhibits expression of the glucose transport genes by inactivating the
cell surface glucose sensors Rgt2 and Snf3 in yeast. Mol Biol Cell, mbc. E15-11-0789
(2016).
103
120. van Rensburg E, den Haan R, Smith J, van Zyl WH, Görgens JF. The metabolic
burden of cellulase expression by recombinant Saccharomyces cerevisiae Y294 in
aerobic batch culture. Appl Microbiol Biotechnol 96, 197-209 (2012).
121. Travers KJ, Patil CK, Wodicka L, Lockhart DJ, Weissman JS, Walter P. Functional
and genomic analyses reveal an essential coordination between the unfolded protein
response and ER-associated degradation. Cell 101, 249-258 (2000).
122. Snoep JL, Yomano LP, Westerhoff HV, Ingram LO. Protein burden in Zymomonas
mobilis: negative flux and growth control due to overproduction of glycolytic
enzymes. Microbiol 141, 2329-2337 (1995).
123. Dujon B. The yeast genome project: what did we learn? Trends Genet 12, 263-270
(1996).
124. Lee FW, Da Silva NA. Sequential delta-integration for the regulated insertion of
cloned genes in Saccharomyces cerevisiae. Biotechnol Prog 13, 368-373 (1997).
125. Gorgens JF, van Zyl WH, Knoetze JH, Hahn-Hagerdal B. The metabolic burden of
the PGK1 and ADH2 promoter systems for heterologous xylanase production by
Saccharomyces cerevisiae in defined medium. Biotechnol Bioeng 73, 238-245
(2001).
126. Kim MD, Han KC, Kang HA, Rhee SK, Seo JH. Coexpression of BiP increased
antithrombotic hirudin production in recombinant Saccharomyces cerevisiae. J
Biotechnol 101, 81-87 (2003).
127. Canonaco F, Schlattner U, Pruett PS, Wallimann T, Sauer U. Functional expression
of phosphagen kinase systems confers resistance to transient stresses in
Saccharomyces cerevisiae by buffering the ATP pool. J Biol Chem 277, 31303-31309
(2002).
104
ACKNOWLEDGEMENT
I would like to express my sincere gratitude and appreciation to the followings:
l My supervisors, Professors Akihiko Kondo and Tomohisa Hasunuma, for
their assistance, guidance, advice and ideas. They gave me a wonderful
opportunity to study in this top-level laboratory. They indicated me new
perspectives and inspired me to challenge more attractive goals. I will
always remember their encouragement and enthusiasm.
l Professor Shih-Hsin Ho and Associate Professor Chiaki Ogino for their
invaluable suggestions and kind support during the preparation of this
thesis.
l Assistant Professor Kentaro Inokuma for his warm guidance on
experimental manipulations.
l Associate Professor Kengo Sasaki, for his kind supply of MC6 for the
experimental usage.
l Professor Willem H. van Zyl and Riaan den Haan, for their constructive
comments and revisions on my manuscripts.
l Professor Xinqing Zhao, Fengwu Bai, and Jo-Shu Chang, for their sincere
concerns on my thesis and helpful guidance for my future career.
l Mr. Takahiro Bamba, Musashi Takenaka, Kenta Morita, Ms. Mami
Matsuda, and all the members of Professor Kondo’s laboratory for their
technical assistance and encouragement.
l My friends, Iwamoto Ryo, Xiao Xue and Shan Wang, for their
encouragements and supports to complete my Ph.D. thesis.
105
l My parents, Juan Zhai and Dawei Liu, for their understanding, financial
support and deeply care on me. Without my family’s support, I would not
have been able to devote myself to the research and complete my thesis.
The present work was supported by Special Coordination Funds for Promoting
Science and Technology, Creation of Innovative Centers for Advanced
Interdisciplinary Research Areas (Innovative Bioproduction Kobe), MEXT, Japan, the
National Research Foundation (NRF) of South African’s Research Chair (SARChi)
for Biofuels, and through an international collaborative project supported by Japan
Society for the Promotion of Science (JSPS) and NRF of South Africa.
Liu Zhuo
Department of Chemical Science and Engineering
Graduate School of Engineering
Kobe University
106
PUBLICATION LISTS
Introduction
Liu Z, Ho SH, Hasunuma T, Chang JS, Ren NQ, Kondo A. (2016) Recent
advances in yeast cell-surface display technologies for waste biorefinery
(Review), Bioresourse Technology, S0960-8524 (16) 30435-7
Chapter I.
Liu Z, Inokuma K, Ho SH, den Haan R, Hasunuma T, van Zyl WH, Kondo A.
(2015) Combined cell-surface display- and secretion-based strategies for
production of cellulosic ethanol with Saccharomyces cerevisiae, Biotechnology
for Biofuels, 8: 162
Chapter II.
Liu Z, Ho SH, Sasaki, K, den Haan R, Inokuma K, Ogino C, van Zyl WH,
Hasunuma T, Kondo A. (2016) Engineering of a novel cellulose-adherent
cellulolytic Saccharomyces cerevisiae for cellulosic biofuel production,
Scientific Reports, 6: 24550
Chapter III.
Liu Z, Ho SH, Inokuma K, den Haan R, van Zyl WH, Hasunuma T, Kondo A.
Efficient ethanol production from crystalline cellulose by engineering of a
high-cellulolytic Saccharomyces cerevisiae: Towards consolidated bioprocessing.
under preparation.
107
Doctor Thesis, Kobe University
“Engineering of high-cellulolytic Saccharomyces cerevisiae using cell-surface display
technique: Towards consolidated bioprocessing”, 107 pages
Submitted on August 29th, 2016
The date of publication is printed in cover of repository version published in Kobe
University Repository Kernel.
© LIU ZHUO
All Right Reserved, 2016