1.0
INTRODUCTION
Saccharomyces cerevisiae is well known for its superior ethanol fermenting ability
and widely used in industry for producing bioethanol as fuel.
In addition, this yeast is also
considered an efficient expression system for heterologous genes.
It has a eukaryotic
cellular organization similar to that of plants and animals, making it a desirable host for the
production of proteins that require posttranslational modifications for full biological activity.
Starch currently is the major raw material for bioethanol production in North America.
Since wild type yeast cannot directly utilize starch for vegetative growth and fermentation,
it has been suggested that using genetically engineered yeast expressing amylolytic enzymes
could greatly advance bioethanol production (de Moraes et al., 1995; Murai et al., 1997;
Lipke et al., 1998; Murai et al. 1999; Kondo et al., 2002; Matsumoto et al., 2002).
The
traditional
starch
fermentation
process
requires
large
amounts
of
starch-hydrolyzing enzymes and high temperature cooking to break down raw starch, which
result in high operation and energy costs.
While the development of cold starch hydrolysis
could eliminate the high temperature cooking step, it also requires higher amounts of
starch-hydrolyzing enzymes (Hill et al., 1997; Textor et al., 1998; Robertson et al., 2006).
Thus, suitable recombinant yeast need to generate sufficient amounts of starch-hydrolyzing
enzymes to perform simultaneous starch hydrolysis and fermentation.
To date, functional
amylolytic enzymes from various sources have been expressed at high levels in yeast, and
the amylolytic enzymes have been expressed as either cell wall-anchored or secreted forms
(Rothstein et al., 1984; Filho et al., 1986; Toshihiko et al., 1986; de Moraes et al., 1995;
Murai et al., 1997; Kondo et al., 2002; Matsumoto et al., 2002; Wong et al., 2002).
However, most of the strains engineered to convert starch into ethanol have not been
particularly efficient.
Previous findings showed that barley (Hordeum vulgare) -amylase was superior to
bacterial and fungal -amylase for cold hydrolysis of raw wheat starch (Hill et al., 1997;
Textor et al., 1998), suggesting that a yeast strain could be designed to optimize the
amylolytic activity and ethanol production of recombinant yeast in the fermentative process.
Thus, we developed a novel yeast strain that expressed and anchored barley -amylase on
1
the cell surface.
This recombinant strain showed the ability to hydrolyze soluble wheat
starch under fermentation conditions. However, the starch assimilation rate was slow and
no ethanol was generated (Bo Liao, Master’s Thesis, University of Saskatchewan, 2008).
Therefore, the goal of the current study was to improve the strain’s ability to hydrolyze
starch and thus allow for ethanol production on not only soluble starch, but also on raw
starch.
The following literature review section will first discuss the developing bioethanol
industry, industrial production of bioethanol from starch and the key starch-hydrolyzing
enzymes that are employed to break down starch.
Then, the role of S. cerevisiae in ethanol
fermentation and its ability in heterologous protein expression will be reviewed.
Lastly,
the efforts that have been undertaken to generate yeast stains that express amylolytic
enzyme will be discussed.
2
2.0
Literature Review
2.1
Introduction of Bioethanol as a Renewable Energy
2.1.1 World’s Energy Crisis and Sustainable Energy
Crude petroleum is currently the dominant raw material for the energy and chemical
sectors worldwide. Since the world’s economy and fuel supply are so dependent on a
single source, we are now facing a huge crisis with continued increasing demand for energy
worldwide and a gradual depletion of petroleum reserves. According to the 2007 U.S.
Energy Information Administration's (EIA) annual report, by 2030, world crude oil demand
will increase 37% over 2006 levels (Otero et al., 2007). However, reports from the U.S.
Department of Energy and others are showing that the world’s petroleum reserves are
running out (Bothast et al., 2005; Otero et al., 2007).
In June 2008, the price of crude oil
rose above $146/barrel compared to under $25/barrel in September 2003.
This huge
increase is having a significant impact on the global economy (Bothast et al., 2005;
Editorial, Nature/Biotechnology, 2006; Farrel et al., 2006).
Furthermore, petroleum,
mainly consumed as transportation fuel worldwide, is burned in vehicles and generates large
quantities of greenhouse gases, which according to the majority of academic and technical
reports, may lead to dire consequences to the Earth’s climate (Otero et al., 2007).
In
response to the world’s energy crisis and climate changes due to greenhouse gas emissions,
green energy and sustainable energy sources have recently gained increasing popularity and
are considered to be partial solutions to the petroleum-related energy and environmental
issues we are facing.
2.1.2 Bioethanol as an Alternative Fuel
Biofuel generally represents solid, liquid or gas fuel derived from biological material
such as bioalcohols, biodiesel (methyl and ethyl esters), biohydrogen, alkanes and various
other hydrocarbon mixtures (such as wood gas and syngas). Among those, bioethanol is
by far the most available. For example, in 2005 bioethanol constituted 99% of all biofuel
produced in the U.S. (Bothast et al., 2005). Today, bioethanol is mainly produced from the
3
conversion of biomass by microorganism through fermentation.
Biomass used for
bioethanol production ranges from agricultural products such as corn grains to forest waste
such as wood chips.
In the U.S. and Canada, the raw material of choice for bioethanol
production is starch from corn and wheat.
In Brazil, however, the raw material is
sugarcane (Robertson et al., 2006).
Intensive research focusing on the conversion of biomass to bioethanol can be traced
back to the 1970s.
This area has recently gained significant political and scientific
attention mainly due to concerns about dwindling petroleum supplies, g Lobal energy
security and climate change. Compared to fossil fuels, bioethanol produced from the
conversion of biomass has several advantages related to environmental issues and
dependence on the supply of raw materials: 1) the biomass used for bioethanol production is
plant-derived, which is renewable and environmental friendly, and is available in large
amounts as domestic and commercial waste (such as wood chips, corn stalks, and waste
papers) and fast growing plant species (such as willow trees and switchgrass) (Bothast et al.,
2005; Editorial, Nature Biotechnology, 2006; Farrel et al., 2006; Robertson et al., 2006); 2)
carbon dioxide released through fermentation or by biomass burning processes is recycled
from the atmosphere to produce biomass for the next generation of plants and crops, so the
net regeneration of carbon dioxide (the greenhouse gases) is reduced (Farrel et al., 2006;
Fortman et al., 2008; Mukhopadhyay et al., 2008); 3) bioethanol production using
agricultural crops could benefit farmers by creating substantial new markets for grain crops,
and more job opportunities will be created for refinery workers in economically-depressed
rural areas (Bothast et al., 2005; Fortman et al., 2008; Mukhopadhyay et al., 2008); 4)
compared to gasoline, ethanol has a significant reduction in CO2 and NO2 emission, and
ethanol is the favored fuel additive for increasing oxygenation comparing to MTBE
(methyl-tertiary-butyl ether) and ETBE (ethyl-tertiary-butyl ether) (Aristidou et al., 2000).
2.1.3 Does Bioethanol as an Alternative Fuel Make Sense?
A complete and accurate answer to this question is somewhat difficult.
There are
still debates in the scientific literature as to whether bioethanol as an alternative fuel is
feasible.
The debates around bioethanol concern its economic feasibility, sustainability
4
and energy balance.
An editorial (Nature biotechnology, 2006) had critically commented
that “At present, ethanol is not price competitive by any stretch of the imagination - even
with the absurd and decidedly anti-free-market 54-cent per gallon tariff Washington imposes
upon Brazilian ethanol.”
Since starch-based ethanol plants rely totally on cereal grains as
substrate, which is about 70% of the total ethanol cost, this has led to the food versus fuel
debate (Ingledew, 1993).
The use of corn and wheat for bioethanol production has been
contributing to a dramatic rise in food cost over the past 10 years (Bourne, 2007; Robert,
2007).
In addition, studies have estimated that even if all of the crops available in North
America were used for bioethanol production, the supplied amount of starchy materials
would be far from sufficient if bioethanol is going to replace fossil fuel as the main energy
source (Bourne, 2007; Robert, 2007; Wald, 2007).
Further complications arise when determining if ethanol from starch is sustainable.
Use of large quantities of nitrogen fertilizer, pesticides and herbicides that is associated with
excessive cultivation of corn and sugar cane can cause enormous contamination to
surrounding environments and rivers (Editorial, Nature Biotechnology, 2006).
Moreover,
it seems that the reduction in greenhouse gas emission by starch based ethanol is not that
great compared to gasoline. A reduction of 18% in greenhouse gas emission compared to
gasoline was reported by Farrell et al. (2006).
Given that most ethanol plants are still
employing fossil fuels for transporting raw materials and powering the plants, questions
have also been raised about the energy balance of starch-based bioethanol.
However, recent life-cycle analysis, which is a systematic approach for answering
the fundamental question “Is more energy contained in the fuel than is used in the
production of that fuel?” (Otero et al., 2007), has suggested that new strategies, such as
process integration and energy recycling, will contribute to make bioethanol more energy
favorable (Otero et al., 2007).
It is believed that alternative approaches generated by
advances in biotechnology research could contribute to more economical production of
bioethanol (Ingeldew, 1993).
Diversification of end-products, such as valuable
pharmaceutical products or animal foods, rather than just ethanol could improve the
economic gains of bioethanol plants and make the industry competitive on the market
(Ingeldew, 1993; Otero et al., 2007).
Farrell et al. (2006) suggested that use of cellulose
5
as the raw material can also help to achieve the energy and environmental goals of
bioethanol production in the future.
Cellulose is a much cheaper carbon resource and available in abundance.
However,
it has a more complex structure than starch. Cellulosic materials that could be potentially
used for bioethanol production can be obtained from agricultural residues such as leaves,
stalks and husks of corn plants, forest wastes such as wood chips and tree barks, paper pulp
and grasses (Farrell et al., 2006; Bourne, 2007; Robert, 2007; Wald, 2007). Studies have
been suggesting that bioethanol produced from cellulosic materials could reduce greenhouse
gas emission by 88%-90% compared to gasoline (Farrell et al., 2006; Bourne, 2007). Even
though still in its early stage, much effort has gone into the investigation to commercialize
cellulosic fermentation (Ho et al., 1999; Aristidou et al., 2000; Lynd et al., 2005; Himmel et
al., 2007).
To date, there are still no cellulose-based commercial plants for bioethanol
production.
Despite the controversy, the starch-based bioethanol industry keeps growing at a
dramatic rate (Otero et al., 2007). Although sugar cane is reported to be the most widely
used raw material for bioethanol production, in North America, starch is currently the most
economical raw material.
An editorial “Bioethanol needs biotech now” in Nature
Biotechnology (2006) stated that, even though in the long-term bioethanol production from
cellulose is the best way to contribute to our energy and environment goals, currently starch
fermentation is the most economical way for bioethanol production, and biotechnology
should play an important role in advancing starch fermentation into a more economical and
environmental friendly industry.
Furthermore, research discoveries on starch fermentation
could also potentially benefit cellulose fermentation in the future.
Finally, with respect to the future of the bioethanol industry, Otero et al. (2007)
suggested that “bioethanol, biorefineries, and industrial biotechnology will not be successful
and expand to novel areas if we do not focus on engineering public perception and policy.”
2.1.4 Expanding the Bioethanol Industry
The bioethanol industry has become an increasing Ly fast growing industry in the
last 30 years, and shows a great potential to help meet the world’s growing energy demands.
6
Bioethanol was introduced into the commercial transportation fuel supply chain during the
1970s by the Brazilian government.
Brazil has emerged as the second largest producer of
bioethanol and has become energy self-sufficient by supplementing internal petroleum
supplies and refining capacity with bioethanol production (Aristidou et al., 2000; Bothast et
al., 2005). Governments around the world are looking to diversify their energy sources
Bioethanol helps to meet both the world’s
away from the dependence on foreign oil.
immediate and future demands for sustainable energy (Aristidou et al., 2000; Fortman et al.,
2008; Bourne 2007).
In August 2005, the U.S. enacted the Energy Policy Act (EPACT)
into law, creating the national Renewable Fuels Standard (RFS).
The RFS calls for 15.1
billion L of renewable fuels (primarily ethanol but may include alternative fuels such as
biodiesel) to be used by 2006, increasing by 2.6 billion L per year until 2012 when a final
volume of 28.4 billion L will be produced (Himmel et al., 2007; Otero et al., 2007).
Argentina is requiring a 5% ethanol blend over the next 5 years. Thailand requires that all
gasoline sold in Bangkok must be composed of 10% ethanol.
India is requiring 5%
ethanol blends in their gasoline. Canada has provided tax benefits for ethanol producers
and consumers since 1992 (Himmel et al., 2007; Otero et al., 2007).
2.2
A Review of Starch-Based Bioethanol Production
2.2.1 Current Industrial Starch Fermentation Process
Ethanol production from starch is generally initiated using either a dry-grind process
or a wet-mill process.
The dry-grind process focuses on maximizing the ethanol yield
from the starch raw materials.
The wet-mill process tries to extract valuable by-products
from the starch fermentation process, such as distilled-dry grains with soluble (DDGS).
In
the U.S.A, over 70% of bioethanol plants are now using dry-grind based starch fermentation
(Bothast and Schlicher, 2005).
Currently, S. cerevisiae is the primary microorganism used
for industrial bioethanol production.
S. cerevisiae has been used by the brewing industry
for thousands of years for wine and beer production.
Compared to other
ethanol-fermenting yeast, S. cerevisiae is highly ethanol-tolerant, easy to grow in large-scale
7
plants and has the property of GRAS (Generally Regarded As Safe).
Many strains of S.
cerevisiae have already been selected in the brewing industry and characterized as being
able to generate ethanol with high efficiency under stress conditions, which are commonly
encountered in industrial scaled plants.
Although other yeast species and bacterial strains
have been intensively studied and recommended for their potential value in industrial
applications, S. cerevisiae is still the favored microorganism for bioethanol production.
When incubated under anaerobic conditions, S. cerevisiae converts each glucose residue
into two ethanol molecules and two carbon dioxide molecules with the net generation of two
ATPs. This process is generally referred to as fermentation.
Starch exists as an insoluble polymer of glucose residues, with glucose residues
joined together by -1,4 and -1,6 linkages. Starch is generally composed of amylose and
amylopectin.
chain.
In amylose, glucose residues are linked by -1,4 linkages to form a linear
However, amylopectin, which contains both -1,4 and -1,6 linkages, is highly
branched (Fig. 2.1).
In normal corn (Zea Mays) or wheat (Triticum aestivum) starch, 27%
is amylose and 73% is amylopectin (Tester et al., 2004).
A
B
Figure 2.1. Structure of amylose and amylopectin. In amylose, the glucose residues are
linked together by -1,4 linkage (A). In amylopectin, parallel amylose chains are linked
together by two glucose residues at the branch points by -1,6 linkage (B). The diagram is
taken from “Starch in general” by Archer Daniels Midland Company.
8
Since S. cerevisiae does not express starch-hydrolyzing enzymes, it is unable to
directly utilize starch for vegetative growth.
In industry, starch is broken down into single
glucose residues before it can be used by yeast for fermentation.
In order to completely
hydrolyze starch, it is treated with either strong acids or starch-hydrolyzing enzymes which
break the -1,4 and -1,6 linkages in the starch molecules which releases glucose
monomers. Up until the 1970s, acid treatment was commonly used for starch hydrolysis
(Robertson et al., 2006). Starch was mixed with diluted acid and cooked at 120-150 C to
release fermentable sugars.
However, the treatment was reported to have the
disadvantages of being costly, causing corrosion of equipment, forming undesirable
by-products, and having limited yield (Robertson et al., 2006).
After the discovery of
thermostable starch-hydrolyzing enzymes, the process gradually evolved to the
high-temperature cooking and enzyme-mediated hydrolysis of starch.
Generally,
enzyme-catalyzed starch hydrolysis is performed as follows: a 30% (by weight) slurry of
starch is cooked in the presence of thermostable -amylase to 90-165 C (the liquefaction
stage).
The liquefied starch is held at 90 C for 1-3 h, and then cooled further to 60 C
followed by the addition of glucoamylase (the saccharification stage).
Finally, the
completely hydrolyzed starch is cooled to 30 C and then mixed with the yeast in the
fermentor to produce ethanol (Fig. 2.2).
In some cases, the saccharification process is
carried out at the same time as the fermentation process at 30 C, which is referred to as
simultaneous saccharification and fermentation (SSF) (Robertson et al., 2006).
However, an excess energy input of 10-20% of the fuel value of ethanol produced
was estimated for the current high temperature-cooked starch hydrolysis process (Robertson
et al., 2006). This calculation does not include the energy lost due to any plant-specific
thermal conversion efficiencies or heat transfer efficiencies.
Thus, reducing the energy
input during the starch hydrolysis could greatly increase the net energy output of bioethanol.
9
Figure 2.2. Schematic representation of the action of amylases on starch. Raw starch is
mixed with water and then cooked at high temperature to its gelatinized state (more soluble),
and with addition of thermal stable -amylase, the mash is further liquefied into dextrins
(referred to as liquefaction). Next, the mash is cooled and glucoamylase is added to
convert the dextrins into fermentable sugars (referred to as saccharification). Then yeast is
added to the mash to ferment the sugars to ethanol and carbon dioxide (fermentation).
2.2.2 Cold Starch Hydrolysis
In order to avoid the high energy input required for cooking raw starch materials, an
energy-conserving alternative is to lower the starch processing temperature to that of the
fermentation temperature, which is lower than the gelatinization temperature which is 54 C
for wheat, 60 C for potato (Solanum tuberosum), and 65 C for maize (Zea Mays) starches
(Rahman, 1995).
The low-temperature hydrolysis alternative has been described as cold or
raw starch hydrolysis.
Using this strategy, the excess energy input can be either reduced or
altogether eliminated.
This hydrolysis strategy can be applied to raw starch that is
produced, in the case of cereal grains, by wet or dry milling processes.
10
In cold starch
hydrolysis, the degradation of the original starch granule structure primarily relies on the
raw starch-hydrolyzing ability of -amylase, which quickly decreases the average molecular
weight of starch granule and releases oligosaccharides. Upon the addition of glucoamylase,
the starch fragments are further broken down into soluble glucose residues. Yeast are
added before the completion of starch hydrolysis to achieve simultaneous starch hydrolysis
and fermentation.
Few cold hydrolysis, corn-to-ethanol dry-mill plants currently operate on a
commercial scale (Berven, 2005). There are several advantages of cold starch hydrolysis
compared to the conventional high temperature, enzymatic hydrolysis.
First, there is a
significant reduction in energy input, and complete elimination of the cooking step can
significantly increases the energy output of bioethanol.
Second, thermally-solubilized and
liquefied starch has viscosities that can be 20-fold higher than that of the starch slurry and
make it difficult to pump and stir (Robertson et al., 2006).
Low-temperature liquefaction
using cold starch hydrolysis can save additional energy input and increase the equipment
capacity (Kelsall and Lyons, 1999).
Third, the elimination of high-temperature cooking
can increase the ethanol yield by reducing the formation of undesirable reaction by-products
(Galverz, 2005).
As one of the pioneer companies invested in commercializing cold starch
hydrolysis for bioethanol production, Genencor reported simultaneous saccharification and
fermentation of raw starch on a commercial scale at 32C. The final ethanol yield shows a
2.5% increase compared to the conventional cooking-fermentation process (Genencor,
2010).
However, a number of potential concerns have been raised regarding the use of cold
starch hydrolysis for bioethanol production.
Starch-lipid complexes, which cannot be
simply separated by the milling process and are resistant to enzyme digestion, are usually
destroyed by high-temperature cooking.
This may pose a problem for cold starch
hydrolysis, resulting in a large portion of starch not being hydrolyzed.
In addition,
elimination of high-temperature cooking may also cause a greater chance of microbial
contamination, since the raw starch is not sterilized before fermentation. Other issues
include high enzyme requirements because of low enzyme activity, poor enzyme stability,
and enzyme inhibition by glucose and maltose. One published study showed that the
11
amount of malt barley -amylase required for digesting raw wheat starch is about 1000X of
the amount require to digest soluble starch (Textor et al., 1998). This high demand on the
starch-hydrolyzing enzyme and its rapid deactivation during the fermentation process results
in the enzyme being the highest cost after the raw material (about 70% of the raw grain
material cost) (Lang et al., 2001).
Amylolytic enzymes play critical roles in efficient starch hydrolysis, especially for
raw starch hydrolysis.
In nature, efficient starch assimilation has been found in organisms
throughout all three kingdoms.
In the last three decades, by employing recombinant DNA
techniques, researchers have studied the structure-function relationship of a large number of
amylolytic enzymes in detail.
Successful attempts have been made to engineer amylolytic
enzymes with improved industrial characteristics.
2.3
Starch-Hydrolyzing Enzymes
2.3.1 An Overview of Starch-Hydrolyzing Enzymes
Starch-hydrolyzing enzymes are very common in nature, and can be found in a
variety of species throughout animals, plants and microorganisms.
Different
starch-hydrolyzing enzymes show different specificity towards starch and/or different
oligosaccharides, and vary in their ability to perform hydrolysis and/or synthesis of -1,4
and -1,6 linkages in starch and/or oligosaccharides. Many organisms that are known to
efficiently utilize starch produce a combination of starch-hydrolyzing enzymes to optimize
starch assimilation.
It has always been of great interest in the industry to understand the
catalytic functions of different starch-hydrolyzing enzymes so that the “best” ones can be
used for designed tasks under specific industrial conditions.
There are many different ways to categorize amylolytic enzymes, including their
substrate specificity, product types, sequence/domain similarities etc..
One commonly
accepted method is to categorize amylolytic enzymes by their cleavage action towards
starch or oligosaccharides.
Based on this, all amylolytic enzymes can be divided into four
major groups (with a few exceptions (van der Maarel et al., 2002)), including endoamylases,
12
exoamylases, debranching enzymes, and transferases.
Among the four groups,
endoamylase and exoamylase groups are the ones that have been studied the most, and
many species in the groups have been widely used in industrial applications.
Endoamylases are amylolytic enzymes that cleave -1,4 glycosidic linkages in an
endo-acting fashion.
-Amylase (EC 3.2.1.1) is a well-known endoamylase.
-Amylase
activity generates oligosaccharides with varying lengths with an -configuration. Since
-amylase does not cleave -1,6 linkages, its end-products constitute branched
oligosaccharides.
Conversely, exoamylases attack on starch or oligosaccharides in an
exo-acting fashion, which specifically release glucose or maltose from the non-reducing
ends.
Based on the specificity towards the glycosidic linkages, exoamylases can be
divided into two groups. The first group exclusively cleaves -1,4 glycosidic linkage,
such as -amylase (EC 3.2.1.2).
The second group cleaves both -1,4 and -1,6
glycosidic linkages, such as glucoamylase (EC 3.2.1.3).
-1,6 linkages at a much slower rate than -1,4 linkages.
However, glucoamylase cleaves
It is well-known in industrial
starch processing that -amylase and glucoamylase attack starch and oligosaccharides in a
synergistic fashion, thus optimal starch hydrolysis rate is achieved by incorporating both
enzymes.
Debranching enzymes and transferases have been receiving increasing attention,
since under many situations starch hydrolysis cannot go to completion employing only
-amylase and glucoamylase activity. The addition of appropriate debranching enzymes
and transferases can increase the efficiency of starch hydrolysis.
Debranching enzymes are
a group of starch-converting enzymes that exclusively cleave -1,6 glycosidic linkages in
amylopectin and pullulan. Pullulan is a polysaccharide with a repeating unit of maltotriose
that is -1,6 linked.
Based on substrate specificity, there are two major types of starch
debranching enzymes: isoamylase (EC 3.2.1.68) and pullulanase type I (EC 3.2.1.41).
Isoamylase hydrolyzes -1,6 glycosidic linkages exclusively in amylopectin, while
pullulanase hydrolyzes -1,6 linkages in both amylopectin and pullulan.
The end-products
of both isoamylase and pullulanase are long linear polysaccharides.
Despite their
important role in starch hydrolysis, industrial applications of debranching enzymes are still
very limited since the commercially available enzymes lack sufficient thermostability.
13
The last group of amylolytic enzymes are transferases, which cleave an -1,4
linkage of one molecule and then transfers part of the molecule to another one by generating
a new glycosidic linkage.
2.4.1.25)
and
This group of amylolytic enzymes includes amylomaltase (EC
cyclodextrin
glycosyltransferase
(EC
2.4.1.19).
Cyclodextrin
glycosyltransferase is currently used commercially for production of the sweetener
stevioside, which can reduce the bitterness of syrup by increasing its solubility (Pedersen et
al., 1995). Amylomaltase was proposed to be used for the production of thermoreversible
starch gel, which has the property to be repetitively dissolved upon heating (van der Maarel
et al., 2000).
However, neither the enzyme nor the starch gel is commercially available
(van der Maarel et al., 2002).
To date, among all the amylolytic enzymes that have been recorded, -amylase and
glucoamylase are the ones that have been studied the most due to their industrial importance
in starch processing.
The employment of cloning and recombinant DNA techniques made
it possible to study many of those enzymes at the molecular level, and the knowledge has
provided valuable insights into designing amylolytic enzymes with industrial-favored
characteristics.
The following part of the review will devote separate sections for
-amylase and glucoamylase, respectively, with emphasis on their molecular structures and
functions, which have been intensively studied in the last several decades.
-Amylase and
glucoamylase, engineered with improved industrial characteristics, will also be discussed.
2.3.2 -Amylase
2.3.2.1 The Catalytic Function and Industrial Applications of -Amylase
-Amylase (1, 4--D-glucan glucanohydrolase, EC 3.2.1.1) catalyzes the hydrolysis
of -1,4 glycosidic linkages in starch and oligosaccharides with the release of maltose and
glucose.
So far, there are 21 different enzyme specificities recorded in the -amylase
family.
Based on their catalytic functions, they can be categorized into two different
groups.
One group is composed of enzymes that perform starch hydrolysis while the other
group is involved in starch-modification or –transglycosylation.
Members of the
-amylase family are widely used in the food, chemical, pharmaceutical, detergent, textile
14
and bioethanol industries.
Starch-hydrolyzing -amylases with high thermostabilities are
extensively used in starch fermentation.
They are also employed to remove starch in beer,
fruit juices, from clothes and porcelain and pretreat animal feed to improve its digestibility.
High thermostability of many -amylases is the key for their production in large-scale
fermentation and commercialization.
The use of thermostable -amylases has transformed the starch processing industry.
Early starch processing was based on acid hydrolysis.
Incubating starch in acidic solution
at high temperature degrades starch into glucose monomers. However, the use of acid
causes the production of unwanted by-products and is highly corrosive to the equipment.
Following the discovery of thermostable -amylase from Bacillus amyloliquefaciens, the
old method was replaced by the enzyme-mediated hydrolysis of starch.
Recently, the B.
amyloliquefaciens -amylase has been replaced by a more thermostable -amylase from B.
stearothermophilus or B. licheniformis.
Since the -amylases used in industry are
inactivated at a pH lower than 5.9 at the high temperature currently being used, the pH
needs to be adjusted to 6 by the addition of NaOH.
New findings showed that Pyrococcus
furiosus -amylase is active even at 130°C (Jorgensen et al., 1997).
commercialization of this enzyme should advance the industry further.
The
However, instead
of exclusively employing -amylase with high thermostability, the potential of using cold
starch hydrolysis for bioethanol production now encourages researchers to search for
-amylase with raw starch-binding and hydrolytic activity.
A new generation of starch
processing enzymes will likely transform the industry again in the very near future.
2.3.2.2 General Molecular Structure of -Amylase
Structural studies are important to understand enzyme structure and function
relationships, and can provide valuable insight for further protein engineering.
The
-amylase family of enzymes share a structural similarity of having a (/)8 catalytic core
structure. To date, three-dimensional structures of many -amylases are available. The
basic framework for -amylase structure consists of three domains, A, B and C.
Domain A
possesses a catalytic core (/)8 structure with a small domain (domain B) protruding
between the third -strand and the third -helix. Domain A is composed of eight parallel
15
-strands, which are highly symmetrical, folded in a barrel-shaped structure surrounded by
eight -helices (van der Maarel et al., 2002).
Some B domains contain several -strands
and one or two helices, while others, such as the domain in barley -amylase, have no
well-defined secondary-structure. Domain B is generally involved in substrate or Ca2+
binding (van der Maarel et al., 2002).
Domain C is not involved in catalytic activity, but
for many members of -amylase, it plays a role in oligosaccharide binding such as in barley
-amylase (Fig. 2.3).
Figure 2.3. Three dimensional structure of barley -amylase 1. Calcium ions (green dots)
in domain B, the active site in domain A (top), the starch granule binding site in domain A
(left) and the sugar tongs binding site (bottom) in domain C are shown as indicated above.
The three binding maltoheptaose molecules (mimic molecules of short starch chain) are
shown in a gray transparent surface. The diagram is taken from Robert et al. (2005).
16
There are nine other domains that have been identified throughout the members of
-amylase family, and different -amylases have different domains attached to its catalytic
core (domains A and B). The attachment of additional domains and their arrangement
determine the substrate and product specificities of each -amylase.
For example, the
additional domains can give the enzyme the ability to hydrolyze -1,6 linkages (van der
Maarel et al., 2002) or to provide an additional starch binding site, which allows for raw
starch hydrolysis (Dalmia et al., 1995; Knegtel et al., 1995; Penninga et al., 1996;).
However, the functions of many domains are still not identified (van der Maarel et al., 2002).
Further study and understanding on the domain structure and functions are valuable for
engineering -amylase with improved thermostability or raw starch binding ability.
2.3.2.3 Protein Engineering of -Amylase for Industrial Applications
In order to obtain enzymes with improved characteristics for industrial application,
screening for naturally-occurring enzymes with the desired properties is one valid approach.
Since industrial processes are usually performed under extreme conditions, such as high
temperature and extreme pH, screening microorganisms living in extreme environmental
conditions has proven to be effective for obtaining enzymes with favored characteristics.
However, this method is labor-intensive and time-consuming.
By employing DNA cloning
techniques and site-directed mutagenesis, enzymes with improved functions can be
generated through rational design in the laboratory.
For different application conditions,
protein engineering can be used to generate mutants with novel characteristics (Barnett et al.,
1998). Even the product specificity of the enzyme can be modified through site-directed
mutagenesis (Schulz and Candussio, 1995; Cherry et al., 1999; Dijkhuizen et al., 1999).
Since the current industrial starch processing is carried out at high temperature,
much effort has been focused on generating -amylases with high thermostability.
To
increase structural stability, strategies such as introducing prolines into the loop regions of
domain A in -amylase (Matthews et al., 1987; Bisgard-Frantzen et al., 1996) or
introducing disulfide bonds have been attempted (Day, 1999).
Adopting the enzyme
structure of naturally occurring, thermostable -amylases have also proven to be effective.
Thermostable -amylases from B. licheniformis and B. amyloliquefaciens have been
17
intensively studied, and the regions and amino acid residues, which are critical for enzyme
thermostability, are identified (Suzuki et al., 1989; Conrad et al., 1995; Borchert et al.,
1999).
Based on these studies, a large number of mutants were generated and the ones
with improved thermostability were isolated.
Moreover, a mutant of B. licheniformis
-amylase with 23 times higher stability than the wild type enzyme at pH 5.0 and 83°C was
constructed by employing a random mutagenesis approach (Shaw et al., 1999).
Engineered -amylase with improved raw starch hydrolysis has also been reported
(Juge et al., 2006).
More research into raw starch-hydrolyzing enzymes should be
expected due to the concern of employing more energy efficient methods (such as cold
starch hydrolysis) for starch processing in industrial bioethanol production.
In conclusion, by combining protein structure knowledge and DNA recombinant
techniques, significant improvement in -amylase can be achieved to improve its
performance in industrial bioethanol applications.
2.3.3 Glucoamylase
2.3.3.1 General Information and Catalytic Function of Glucoamylase
Glucoamylase (1,4--D-glucan glucohydrolase; EC 3.2.1.3) hydrolyzes -1,4 and
-1,6 glucosidic linkages from starch in an exo-acting fashion and releases glucose.
It is a
key industrial starch processing enzyme and commonly used in conjunction with -amylase
to convert starch into glucose.
Glucoamylase is found throughout many species of animals, plants and
microorganisms.
Industrial production of glucoamylase is carried out in large-scale
bioreactors by submerged, solid state and semi-solid state fermentation.
Among the
glucoamylase-producing microorganisms, filamentous fungi species Aspergillus niger, A.
awamori and Rhizopus oryzae are traditionally used as the source of glucoamylase. These
fungi are able to secrete glucoamylases extracellularly with high efficiency and can be
cultivated into high biomass using simple medium.
Microorganisms with these
characteristics are favorably selected for industrial production of glucoamylase.
In
addition, glucoamylases produced by A. niger, A. awamori and R. oryzae were observed to
18
possess good thermostability and high activity at or near neutral pH values (Frandsen et al.,
1999; Reilly, 1999), which are favored characteristics for industrial starch processing.
Glucoamylase has been extensively studied.
The genes of glucoamylases from
different sources were isolated and cloned (Innis et al., 1985; Meaden et al., 1985;
Yamashita et al., 1985; Ashikari et al., 1986, 1989; Erratt and Nasim, 1986; Pardo et al.,
1986; Pretorius et al., 1986; Cole et al., 1988; Inlow et al., 1988), and fully functional forms
of many of these glucoamylases have been expressed in E. coli, S. cerevisiae and P. pastoris.
Structural analysis and employment of site-directed mutagenesis have provided valuable
insights into the structure-function relationship of glucoamylase, which has made it possible
to generate mutants of glucoamylase with improved characteristics for industrial
applications. The following reviews will focus on the molecular studies performed on
glucoamylases and efforts on the development of related mutants.
2.3.3.2 General Molecular Structure of Glucoamylase
Based on the resolved three-dimensional structures of several fungal glucoamylase
enzymes, generally, glucoamylase is composed of two distinct domains connected by an
O-glycosylated linker region.
The catalytic domain (CD) is located at the N-terminus with
a (/)6-barrel fold, and the C-terminal domain is composed of anti-parallel -sheets with
two binding sites for starch or oligosaccharides (also known as the starch binding domain,
SBD).
The majority of glucoamylases have the SBD positioned at the C-terminal end,
with glucoamylase from R. oryzae, Arxula adeninovorans, and Mucor circinelloides as
exceptions, which have their SBD located at the N-terminus (Kumar and Satyanarayana,
2003).
Although the complete three-dimensional structure of the intact glucoamylase
(including both domains and the linker region) is still not known, the detailed structural
information of each separate domain and the linker region has provided insights into the
structure-function relationships of glucoamylase.
Three-dimensional structures of the glucoamylases from A. awamori var. X100,
Saccharomycopsis fibuligera and Thermoanaerobacterium thermosaccharolyticum have
been determined by different research groups (Aleshin et al., 1992, 1994, 1996; Harris et al.,
1993; Stoffer et al., 1995; Sevcik et al., 1998), with A. awamori var. X100 mostly
19
extensively studied.
Preliminary structural analysis of A. niger glucoamylase showed that
the enzyme shares 94% sequence identity with A. awamori var. X100 glucoamylase (Stoffer
et al., 1995). Both enzymes are among the most commonly used in industrial starch
processing.
Using A. awamori var. X100 as an example, the enzyme is composed of an
N-terminal CD and C-terminal SBD connected by an O-glycosylated linker region.
The
CD of A. awamori contains 13 -helices with 12 of them folding into an (/)6-barrel
structure, in which the funnel-shaped active site is seated.
The SBD contains eight
-strands folded into two anti-parallel -sheets (Jacks et al., 1995; Sorimachi et al., 1996).
The heavily glycosylated linker region was suggested to be a semi-rigid rod shaped structure,
which provides the proper separation between the CD and the SBD (Williamson et al.,
1992).
Figure 2.4.
A predicted structure of Aspergillus niger glucoamylase. Since the
complete three-dimensional structure of an intact glucoamylase is still not known, this
model shows a possible arrangement of the three domains in A. niger glucoamylase. Each
of these domains has been studied in details. The model is constructed by P.M., Coutinho
and P.J., Peilly, Department of Chemical Engineering, Iowa State University, U.S.A.
(http://nte-serveur.univ-lyon1.fr/heyde/www.public.iastate.edu/_pedro/glase/glase_insight.h
tml).
20
2.3.3.3 Structure-Function Relationships of Glucoamylase
The catalytic domains of A. awamori var. X100, A. niger and S. fibuligera share very
similar folds (Sauer et al., 2000).
It was shown that the CD itself is sufficient for
hydrolyzing dextrin (Libby et al., 1994), however, the SBD is required for efficient raw
starch binding and hydrolysis (Svensson et al., 1982; Southall et al., 1999).
Svensson et al.
(1982) showed that the isolated CD of glucoamylase from A. niger had very low activity
towards raw starch.
Furthermore, Southall et al. (1999) showed that the SBD functions by
disrupting the compact structure of the starch granule and thus facilitates the hydrolysis of
the substrate by the CD, and this synergistic effect was structurally confirmed by employing
various heterobifunctional inhibitors (Sigurskjold et al., 1998; Payre et al., 1999).
The linker region is not directly involved in starch binding, but its glycosylation and
length play important roles in structural stability and secretion of many glucoamylases.
The linker regions of most glucoamylases are serine- and threonine-rich that are
O-glycosylated with a high degree of heterogeneity of the amount of attached sugars
(Christensen et al., 1999; Fierobe et al., 1997).
In most cases, glycosylation enhances the
enzyme secretion and thermal stability, and also protects the enzyme from protease cleavage
(Le Gal-Goefer et al., 1995; Cutinho and Reilly, 1997). The length of the linker regions of
different glucoamylases varies greatly (Zalkin et al., 1984).
A minimum length is required
for the secretion of some glucoamylases in their functional forms (Sauer et al., 2000).
Deletion in the linker region can cause no secretion or reduced secretion of glucoamylase or
the enzyme to be secreted in a destabilized form (Yamashita, 1989; Evans et al., 1990;
Coutinho and Reilly, 1994).
An understanding of the structure and function relationship makes engineering
glucoamylase with improved properties possible; some of these examples will be reviewed
in the following section.
2.3.3.4 Engineering of Glucoamylase for Improved Industrial Properties
Current industrial starch processing employs glucoamylase for the saccharification
of dextrin, which is obtained from -amylase-catalyzed starch hydrolysis at high
temperatures. Since glucoamylases from most sources are unstable at high temperatures,
21
the dextrin-rich mash has to be cooled to a much lower temperature before the addition of
glucoamylase.
Industrial saccharification is currently carried out at 60°C.
The
thermostability of glucoamylase at or above this temperature is considered an important
characteristic due to reduced bacterial contamination and viscosity of sugar syrup (Ford,
1999). Use of thermostable glucoamylase could greatly reduce energy consumption during
saccharification and simplify starch processing by achieving simultaneous liquefaction
(-amylase catalyzed starch hydrolysis stage) and saccharification.
Protein engineering has been playing an important role in improving the
thermostability of glucoamylase.
Glucoamylase mutants have been constructed by
site-directed mutagenesis, with showing improved thermostability and/or lower inactivation
rate. Successful ones include mutants generated by replacing glycine 137 with alanine in
the -helix of CD (Chen et al., 1996), replacing serine 30 with proline (Allen et al., 1998),
substituting asparagine 182 to alanine in the Asparagine-Glycine sequence (Chen et al.,
1994), or introducing an extra disulfide bond by replacing alanine 246 to cysteine, which
forms a disulfide bond with cysteine 326 (Fierobe et al., 1996). The recorded successful
mutants showed 1.2 to 4°C increase in the melting temperature without any observed
decrease in the specific activity or secretion rate (Ford, 1999).
In addition to possessing hydrolysis activity towards -1,4 linkages, glucoamylase
has slow hydrolytic activity towards -1,6 linkages. The specific activity (Kcat/Km) of
glucoamylase towards -1,6 linkages was reported to be only 0.2% of that towards -1,4
linkages (Hiromi et al., 1966; Sierks and Svensson, 1994; Frandsen et al., 1995; Fierobe et
al., 1996).
Glucoamylase can also catalyze a reverse reaction to synthesize glucose
residues into -1,6 linked isomaltose and isomaltotriose (Ford, 1999).
In the industrial
saccharification process, these catalytic properties of glucoamylase were reported to limit
the yield of glucose to only 96% (Sauer et al., 2000), which is a considerable loss at the
industrial scale. Thus, glucoamylase with suppressed activity towards -1,6 linkage is
considered of great importance for the industry.
By employing site-directed mutagenesis,
researchers have isolated a large number of mutants with reduced activity towards -1,6
linkage while retaining -1,4 activity (Fang and Ford, 1998; Fang et al., 1998a, 1998b; Liu
et al., 1998; Liu et al., 1999). The most successful one had serine 411 mutated to alanine,
22
resulting in a 2% increase in the final glucose yield (Fang et al., 1998a).
Further studies
are focusing on combining the characteristics of thermostability and suppressed activity
towards the formation of -1,6 linkages in a single mutant (Liu et al., 1998).
Advances in
protein engineering will continue to benefit the industry in the future.
2.3.4 Amylolytic Enzymes with Raw Starch-Hydrolyzing Ability
Only about 10% of amylolytic enzymes have been shown to have raw
starch-hydrolyzing ability, and the presence of a distinct SBD is essential for this ability.
The SBD is always positioned at the C-terminal part of an amylase except for the
glucoamylase from R. oryzae which contains the SBD at its N-terminus.
Starch binding
domains from different amylolytic enzymes share significant sequence identity (Svensson et
al., 1989; Janecek and Sevcik, 1999).
The similarity among the SBDs is more correlated
with the species origin rather than the given amylolytic enzyme family, suggesting that this
independent structure was attached or removed from these enzymes during evolution (Wong
et al., 2002).
The SBD, consisting of several -strand segments forming an open-sided, distorted
-barrel structure, is responsible for the ability of an amylase to bind and digest the native
raw, granular starch.
In -amylases, the SBD may govern the enzyme’s thermostability,
while in glucoamylase the domain does not appear to participate in thermostability.
Although the SBD may behave as an independent module, it was shown to be more stable in
the whole enzyme molecule compared to an isolated domain (Southall et al., 1998).
Proteolytic cleavage of glucoamylase can separate the SBD from the CD, which causes the
CD to be incapable of hydrolyzing raw starch granules. Several strategies have been
shown to effectively alleviate protease production during glucoamylase production,
including nutrient and pH control, bioreactor design and use of protease inhibitors (Pandy
and Radhakrishna, 1992; Pedersen et al., 2000; Xu et al., 2000; Bertolin et al., 2003).
23
2.3.5 Barley -Amylase and its Heterologous Expression
2.3.5.1 Structure and Catalytic Functions of Barley -Amylase
Barley -amylase has been extensively studied over the past 20 years since its raw
starch-hydrolyzing ability has huge potential for industrial applications.
Two -amylase
isozymes, barley -amylase 1 (AMY1) and barley -amylase 2 (AMY2), are produced by
germinating barley seeds.
The two enzymes are encoded by separate genes and their
expressions are regulated by plant hormones (Rogers, 1985).
The two isozymes display 80%
sequence identity, with AMY1 composed of 414 amino acids and AMY2 composed of 403
amino acid residues. They are also distinguished by their pIs with AMY1 known as the
low-pI (pI = 4.9) form, and AMY2 as the high-pI (pI = 5.9) form (Rogers et al., 1983, 1984;
Rogers, 1985; Khursheed et al., 1988).
With such high similarities of their primary sequences, studies have shown that the
two isozymes significantly differ from each other with respect to their catalytic action on
amylose and maltooligosaccharides (Wong et al., 2000, 2002).
higher affinity for Ca2+ ions than AMY2 (Bush et al., 1989).
suitable candidate for industrial starch hydrolysis.
In addition, AMY1 has a
Barley -amylase 1 is a more
Compared to AMY2, AMY1 is more
stable at acidic pH, but less stable at elevated temperatures (Rodenburg et al., 1994).
Barley -amylase 1 was shown to have higher affinity and activity toward starch granules
than AMY2 (MacGregor and Ballance, 1980; Sogaard et al., 1990; MacGregor et al., 1992).
In addition, compared to bacterial and fungal -amylases, the kinetic property and pH
optimum of AMY1 were shown to be more compatible with fermentation conditions (Textor
et al., 1998).
The crystal structure of AMY2 has been determined (Kadziola et al., 1994).
A
much more complete picture of the three-dimensional structure of AMY1 has been recently
solved and the functions of the major domains were described (Kadziola et al., 1998;
Gottschalk et al., 2001; Robert et al., 2003, 2005). Barley -amylase is composed of three
domains, including a major domain consisting of 288 amino acids forming a (/)8-barrel
(domain A), a small loop with 65 amino acids protruding between 3 and 3 of domain A
(domain B) and a C-terminal domain with 61 amino acids organized into a five-stranded
24
anti-parallel -sheet (domain C). Domain A was first characterized and recognized as the
main location of starch binding and catalytic activity.
starch-binding site was found in domain A.
Recently, an additional
Its function is proposed to be involved in
enhancing starch binding by the enzyme. Domain B is responsible for binding of calcium
ions and maintenance of the enzyme’s three-dimensional structure. Domain C has recently
been recognized as the third potential starch-binding site in the enzyme. Robert et al.
(2003) showed that even though AMY1 and AMY2 share high similarity in domain C, this
binding site does not exist in AMY2. It has been suggested that the two extra starch
binding sites located in domains A and C play important roles for orienting the substrates in
the correct position at the enzyme’s catalytic site, and are critical to barley -amylase’s
starch hydrolysis rate (Robert et al., 2003, 2005) (Fig. 2.3).
The work of Wong et al. (2000)
confirmed the presence of the second starch binding site in domain C.
However, domain C
alone is not sufficient for starch binding (Wong et al., 2002). This additional non-catalytic
starch binding site requires a portion of domain A and domain C for function (Wong et al.,
2002). Further study showed that a highly conserved region 401VWEK404 in domain C,
might interact with the remaining structure of AMY1 to create stability, though this region is
not involved in either catalytic activity or substrate binding (Wong et al., 2002).
Interesting Ly, the ten amino acids at the C-terminus of AMY1 are not required either for
enzymatic activity or structural stability (Wong et al., 2002).
2.3.5.2 Heterologous Expression and Secretion of Barley -Amylase 1
Barley -amylase 1 has been successfully expressed and secreted using its own
signal sequences in S. cerevisiae (Sogaard and Svensson, 1990; Sogaard et al., 1993a, 1993b;
Wong et al., 2002), P. pastoris (Juge et al., 1996), A. niger (Gottschalk, 1995), Xenopus
laevis oocytes (Aoyagi et al., 1990).
Tibbot et al. (2000) reported successful expression of
the C-terminal domain of AMY1 in E. coli.
However, the attempt to express intact AMY1
in E. coli was unsuccessful (Sogaard, 1989).
Svenssen et al. (1993a) reported efficient secretion of AMY1 from S. cerevisiae into
the culture medium using its own secretion signal peptide, with no significant level of
AMY1 detected in the intracellular space.
The amount of AMY1 secreted into the medium
25
was about 1% of the total cell protein.
However, the specific activity of purified
recombinant AMY1 was 30-40% lower than that of the corresponding enzyme isolated from
barley malt.
Isoelectric focusing analysis indicated that transformed yeast cells secreted
four major forms of AMY1.
After separating the four recombinant forms of AMY1, they
found that only one species showed similar activity and a pI value as the native enzyme
found in barley malt.
Two of the other three forms had much lower specific activity, and
the other one had an unusually high pI value compared to the malt enzyme.
They
speculated that the variants with either low activity or unusually high pI resulted from
inefficient/incorrect processing.
Sogaard et al. (1993a) examined the post-translational modifications of the
recombinant AMY1, which revealed two modifications that are responsible for the
generation of the four different recombinant forms of AMY1.
Two forms were found to be
processed by Kex1p, a membrane-bound carboxypeptidase located in the Golgi, and
resulted in the removal of the dipeptide Arginine-Serine from the C-terminal end of AMY1.
Furthermore, two forms were modified by glutathione at the cysteine 95 residue, which is
located in strand 3 of domain A (Jensen et al., 2003), resulting in formation of an extra
disulfide bond in the recombinant enzyme. Mature forms of AMY1 secreted from barley
aleurone cells were found to be processed at the C-terminal end by malt carboxypeptidase II,
with the unprocessed forms mainly being retained intracellularly (Jacobsen et al., 1988;
Sogaard et al., 1993a, 1993b). The two recombinant forms with unprocessed C-termini
were speculated to be due to inefficient carboxypeptidase reaction in yeast.
The
unprocessed C-terminus may cause the enzyme to be secreted with less efficiency without
any effects on the enzyme activity.
However, the recombinant forms with the extra
disulfide bond were observed to have much lower specific activity, due to improper pairing
of two SH groups.
Barley -amylase 1 from barley malt is believed to contain one
disulfide bond and two free thiol groups (Sogaard et al., 1992). Sogaard et al. (1993a)
determined that the disulfide bond was formed between cysteine 95 and nearby cysteine 106
or cysteine 125.
disulfide to form.
The redox potential in the yeast secretory pathway may not allow this
Moreover, it has been shown that cells produce elevated levels of
glutathione under stress.
Thus culture conditions may preclude the formation of the
26
rAMY1 disulfide bond in yeast, while an increased level of glutathione that could result
from overexpression of AMY1, might well lead to extensive glutathionylation.
Sogaard et
al. (1993a) demonstrated that genetic removal of the C-terminal Arginine-Serine, along with
mutation of cysteine 95 to alanine, led to the secretion of a single, fully active rAMY1 form.
Wong et al. (2001) also reported constitutive expression and secretion of AMY1
from S. cerevisiae.
The recombinant AMY1 was secreted in three different forms with
slightly different pI values.
In contrast to the findings by Sogaard et al. (1993a), they
found the purified recombinant AMY1 had comparable enzymatic activity to that from the
native seed enzyme.
Recombinant AMY1 secreted from P. pastoris also showed multiple
forms with different pI in the range of pH 4.5-5.1 (Juge et al., 1996).
Six different
recombinant forms were identified, differing with respect to processing at the C-terminus
and post-translational modifications.
The purified recombinant AMY1 was found to be
fully active and contained only a minor fraction that was glutathionylated, which had no
significant effect on activity (Jude et al., 1996).
Moreover, the expression level was 50
fold higher compared to that obtained in S. cerevisiae.
2.4
Saccharomyces cerevisiae for Bioethanol Production
2.4.1 An Overview of Saccharomyces cerevisiae
Saccharomyces cerevisiae, the budding yeast, is an important microorganism not
only in various commercial applications such as in the food and beverage industries, but
also in biomedical science research.
The microorganism is very well known for its superior ethanol producing ability,
which has been harnessed by humans for brewing and wine production since the very early
ages of human civilization.
bread dough quality.
Saccharomyces cerevisiae is also widely used for improving
Compared to other closely related yeast species, S. cerevisiae can be
cultivated in large-scale by using a simple growth medium.
carbon sources that it can utilize.
ethanol-tolerant.
It has a very broad range of
It has fast growth and fermentation rates, and is highly
In addition, as a food organism, S. cerevisiae consists of 48%
27
high-quality proteins and has high public acceptance due to the property of GRAS.
Thus, not only does S. cerevisiae still play vital roles in our daily lives, but it has
also been used as a very important tool to advance biomedical science research due to
several useful properties.
As a eukaryote, S. cerevisiae has a cellular organization similar
to those of plants and animals, but its genetic properties allow it to be manipulated almost as
readily as E. coli.
Saccharomyces cerevisiae is a very efficient expression system for
heterologous genes and a model system for unraveling molecular, biochemical and genetic
details of gene expression and the secretion process.
Lately, with bioethanol considered an important alternative transportation fuel now
and for the future, S. cerevisiae has been intensively studied to examine how it can be
exploited and manipulated to benefit the bioethanol industry.
2.4.2 Saccharomyces cerevisiae as a Fermenting Yeast
Commercial distiller’s yeast has optimal reproduction temperature at 28°C or lower.
Even though the yeast can ferment well at 32-35°C, its metabolic activity is compromised at
high temperatures. Furthermore, the yeast’s ability to tolerate high temperature declines
dramatically when other stress conditions are introduced, such as high osmotic stress, low
pH, limited nutrients and low oxygen levels.
strains grow well at pH ranging from 3.8-6.0.
Most distiller’s yeast and brewing yeast
A yeast culture has an initial pH of 5.2-5.5,
and during fermentation, yeast cells keep excreting H+ ions which lowers the pH of broth to
4.0-4.5. Contamination by lactic acid bacteria in the late fermentation stage can lower the
pH further to ~3.8 (Russell, 2003).
Since the fermentation rate is much higher using yeast
cells in their growth phase, maintaining cells in their growth stage is very important for
maximum ethanol production.
Saccharomyces cerevisiae is able to utilize sugars in the presence of oxygen (aerobic)
and in conditions that are completely absent of oxygen (anaerobic). Aerobic catabolism
produces more energy and is called respiration, while anaerobic catabolism produces less
energy and is called fermentation.
Compared to most other yeast species, S. cerevisiae is
able to perform alcoholic fermentation, and can grow under strictly anaerobic conditions
(van Dijken et al., 1993).
It is one of the few yeast that is able to grow anaerobically
28
(Visser et al., 1990).
When S. cerevisiae is grown under conditions with abundant oxygen
but very limited amount of sugar, it produces very little or no ethanol. However, when the
amount of sugar exceeds 0.1%, it produces ethanol, regardless of the presence of oxygen
(Piskur et al., 2006). This is called the Crabtree effect.
Schizosaccharomyces pombe is
another example of Crabtree-positive yeast, which ferments in the presence of oxygen.
Yeast like Kluyveromyces lactis and Candidia albicans don’t produce ethanol in the
presence of oxygen, and thus are Crabtree-negative yeast.
Under aerobic conditions, after the depletion of glucose, the metabolism in S.
cerevisiae changes such that the accumulated ethanol is utilized for yeast propagation,
which is called the diauxic shift.
not be consumed.
However, under strict anaerobic conditions, ethanol will
Once sugars enter the yeast cell, they are processed through the
glycolytic pathway into pyruvate.
The fermentation pathway is involved in the conversion
of pyruvate to ethanol, producing ATP, CO2 and NAD+. During fermentation (aerobic or
anaerobic), yeast recycles NADH in the acetaldehyde-to-ethanol conversion step, which is
catalyzed by alcohol dehydrogenase and is readily reversible.
In the presence of oxygen,
ethanol can be converted back to acetaldehyde, which allows yeast to grow on ethanol after
depletion of sugars (Fig. 2.5).
Yeast other than S. cerevisiae also requires the supply of oxygen to trigger alcoholic
fermentation.
Since it is very difficult to achieve the right oxygen dose in large-scale
fermentation, yeast species that are sensitive to oxygen availability are not suitable for
large-scale fermentation.
Other factors, such as ethanol tolerance, glucose conversion rate,
vitality under acidic conditions, and public acceptance, also determine whether a yeast strain
can be used for commercial-scale ethanol production.
S. cerevisiae can tolerate high
ethanol concentrations up to 15%, has a glucose conversion rate of about 95% of the
theoretical maximum, can grow at low pH conditions, and has the property of GRAS (Vertes
et al., 2008).
Given its long history of use in industrial fermentation and deep knowledge
about its genetics and biochemistry, to date, S. cerevisiae is still superior to other yeast
species for commercial fermentation applications.
29
Figure 2.5. Aerobic and anaerobic catabolic pathways for glucose utilization in yeast.
Yeast is able to utilize glucose aerobically (respiration) and anaerobically (fermentation).
Glucose is converted into pyruvate via the glycolytic pathway, and then pyruvate is
converted into carbon dioxide and energy (with oxygen) or is converted into ethanol, carbon
dioxide and less energy (without oxygen).
However, some drawbacks of using S. cerevisiae have always been some concern for
industry.
Saccharomyces cerevisiae is sensitive to temperatures greater than 35°C and is
subject to lactic acid bacteria contamination and glucose repression (Verte et al., 2008).
Moreover, it cannot ferment on xylose, which is a major component in cellulosic biomass,
and is incapable of directly utilizing starch due to the lack of its own starch-hydrolyzing
enzymes.
Increased understanding over the last 30 years of yeast genetics and metabolism,
along with advances in DNA recombinant technologies, have enabled researchers to
construct a series of genetically modified yeast strains with improved industrial properties.
The application of biotechnology has significantly transformed the bioethanol industry.
30
2.5
Genetic Engineering of Saccharomyces cerevisiae for Bioethanol Production
2.5.1 An Overview of Using Genetically Engineered Yeast for Bioethanol Production
Metabolic engineering is a relative young scientific field (Branduardi et al., 2008)
and its definition was first stated in 1991 by Bailey as “the improvement of cellular
activities by manipulations of enzymatic, transport, and regulatory functions of the cell with
the use of recombinant DNA technology” (Bailey, 1991).
In general, metabolic
engineering is used to improve the production and productivity of metabolites by certain
microorganisms, extend the substrate range for cell growth and/or product formation,
over-express homologous or heterologous proteins, or improve stress tolerance (pH,
temperature etc.).
Eventually, the engineered properties can lead to the development of an
industrial process (Branduardi et al., 2008).
Random mutagenesis was traditionally used to improve the properties and
productivity of microorganisms by employing chemical mutagens (Branduardi et al., 2008).
Metabolic engineering is a more rational approach to engineering cellular properties and
function.
Its success is based on detailed understanding of the biochemistry and
metabolism of microorganisms, combined with various recombinant DNA techniques.
The
approaches can involve the introduction of one or several functional heterologous enzymes,
over-expression or deletion of homologous proteins, silencing of a transcription factor,
modification of transcription activation elements, and so on.
Creative screening methods
are also important to obtain the genetically engineered microorganisms with desired
properties.
Depending on the specific application conditions, different cellular properties are
favored, based on which metabolic engineering approach is employed. With the increasing
popularity of bioethanol as an alternative fuel, much effort has been made to improve strains
of S. cerevisiae for bioethanol production.
Guo et al. (2009) deleted the genes coding for
3-phosphate dehydrogenase from an industrial yeast strain.
By eliminating glycerol
synthesis, the recombinant strain showed increased ethanol production under anaerobic
fermentation compared to the parental strain.
By increasing the expression of a
transcription factor MSN2 during stationary phase, the recombinant strain showed improved
31
stress resistance during fermentation, and a higher fermentation rate was observed (Cardona
et al., 2007). Efficient utilization of maltose by the yeast is a desired property under many
fermentation conditions.
Maltose utilization requires a maltose permease and a maltase,
and both are induced in the presence of maltose. However, the expression of the two
enzymes is repressed by the presence of glucose, which results in incomplete utilization of
the sugars. By replacing the promoter of maltase and maltose permease with constitutive
promoters, increased maltose utilization was observed with the recombinant strain (Osinga
et al., 1988).
Saccharomyces cerevisiae cannot directly utilize starch for its vegetative growth due
to the lack of expression of amylolytic enzymes.
Starch has to be converted into glucose
by starch-hydrolyzing enzymes before it can be used by the yeast for fermentation.
However, the cost of starch-hydrolyzing enzymes is expensive, especially for raw starch
hydrolysis (Lang et al., 2001).
Development of a fermenting amylolytic yeast strain would
make bioethanol production more economically feasible.
There are over 150 naturally-occurring amylolytic yeast species, and some of those
yeast species produce multiple types of starch-hydrolyzing enzymes and are able to directly
utilize starch (even raw starch).
However, compared to S. cerevisiae, they have the
disadvantages of having low ethanol tolerance, slow growth rate, and are not suitable for
large-scale bioethanol production.
In order to develop an amylolytic yeast strain for
bioethanol production, there are two potential strategies.
One is to develop a genetically
modified strain from the naturally occurring amylolytic yeast species so that it will be able
to ferment ethanol with high efficiency. The second strategy is to engineer a superior
ethanol producing species, such as S. cerevisiae, so that it is able to directly utilize starch.
Due to the lack of genetic information for many naturally occurring amylolytic yeast species,
many research groups have opted for the second strategy, by attempting to introduce
heterologous genes coding for amylolytic enzymes into S. cerevisiae.
In order to construct a recombinant strain of S. cerevisiae that is capable of breaking
down starch, heterologous starch-hydrolyzing enzymes need to be expressed in their
functional forms at high levels in S. cerevisiae.
32
2.5.2 Heterologous Protein Expression in Saccharomyces cerevisiae
S. cerevisiae is an excellent host for the production for complex heterologous
proteins.
S. cerevisiae is superior to bacterial hosts because its secretion pathway
generates posttranslational modifications, which are required for full biological activity of a
large number of enzymes, and correct formation of disulfide bonds, which are generally
required for enzyme stability.
In addition, purification of secreted proteins is relatively
simple since S. crevisiae secretes a low level of endogenous proteins.
However, S. cerevisiae tends to over-glycosylate proteins, and inefficient secretion is
often encountered because of retention of the expressed proteins in the periplasmic space
(Branduardi et al., 2008).
These disadvantages have promoted the search for and study of
alternative yeast species for heterologous protein expression, such as Hansenula
polymorpha, Pichia pastoris, Kluyveromyces lactis and K. marxianus (van Dijken et al.,
1993). These yeast species have the advantages of having broader substrate specificity, no
alcoholic fermentation under aerobic conditions (alcoholic generation reduces the biomass
and products formation), and higher secretion efficiency.
None of these yeast have been
employed in large-scale fermentation, since their performance is sensitive to conditions such
as, imperfect mixing and gradients in substrate and oxygen, which cannot be avoided in
large-scale fermentation (van Dijken et al., 1993).
Conversely, with our deep understanding on the genetics and biochemistry of S.
cerevisiae, combined with recombinant DNA techniques, high levels of heterologous protein
expression can still be achieved without severely affecting the microorganism’s performance
on fermentation.
2.5.3 Recombinant DNA Techniques for Efficient Heterologous Protein Expression in
Saccharomyces cerevisiae
Heterologous genes are commonly introduced into S. cerevisiae through designed
genetic carriers.
The yeast genetic carriers, which are commonly known as yeast
expression vectors or plasmids, can be used to transform yeast and be recognized by the
yeast as part of its genetic information.
Inside the yeast cells, the yeast expression
plasmids can either exist and replicate independently from the yeast genome DNA
33
(episomal plasmids) or integrate into certain loci of the yeast genome (integration plasmids)
through homologous recombination. The destination of the plasmids depends on certain
genetic elements that the plasmids carry.
2.5.3.1 Use of Yeast Episomal Plasmids for Expression of Heterologous Genes
Commonly used yeast episomal plasmids include the ARS (autonomously
replicating sequence)/CEN (centromere) plasmid and the 2µ based plasmid. ARS/CEN
plasmid is very stable in yeast, but only exists in 1 or 2 copies per cell. Thus, it is only
used for low level expression. The most commonly employed yeast episomal plasmid is
the 2µ based plasmid (YEp). Yeast episomal plasmids generally exist in 20-100 copies per
cell, and have been widely used for high-level expression of heterologous proteins.
The backbone of a yeast expression episomal plasmid is a bacteria-yeast shuttle
vector, which is composed of both bacterial and yeast elements for the selection and
replication of the plasmid in bacterial and yeast, respectively. The bacterial components of
the plasmid consists of a bacteria selection marker (usually an antibiotic resistance gene),
and the bacterial origin of replication, which is responsible for the replication of the plasmid
in E. coli.
The yeast elements are composed of a yeast selection marker, which could be an
antibiotic resistance gene (auxotrophic selection marker) or could be an amino acid
synthesis gene (prototrophic selection marker), and the yeast origin of replication (for
replication of the plasmid in yeast).
On the shuttle vector backbone, the gene of interest
can be inserted at the designated multiple cloning sites (MCS), which is located between a
yeast promoter and transcription termination site.
The promoter regulates the transcription
of the inserted gene of interest.
When the auxotrophic markers (such as Trp-, Ura- or Leu-) are used for selection
purposes, the yeast strain transformed with the plasmid has to be an auxotrophic mutant of
the corresponding amino acid.
This means that the strain is deficient for synthesizing that
particular amino acid and is unable to survive without the amino acid supplied in the
medium or transformed with the plasmid containing the auxotrophic marker.
However,
almost all auxotrophic mutants of S. cerevisiae are haploid laboratory strains, which are not
suitable for industrial applications.
Yeast strains selected for brewing and distilling
34
industries are mostly polyploidy strains, which are much more tolerant to various stress
conditions, such as high ethanol, low pH and high osmotic environments, which are
commonly encountered in industrial environments. However, constructing an auxotrophic
mutant from an industrial polyploid yeast strain is very complicated and time-consuming,
and it would be very inconvenient to produce all of the auxotrophic mutants for every
industrial yeast strain available and to screen the best ones for bioethanol production.
In order to select yeast strains with no auxotrophic mutants, yeast episomal plasmids
containing dominant (or prototrophic) selection markers can be used.
The commonly used
dominant selection markers are blasticidine and geneticine resistance genes, which encode
resistance to blasticidin S (Bsd) and Geneticin 418 (G418), respectively.
When a dominant
selection marker is used for the selection purpose, the corresponding antibiotic has to be
added into the culture medium.
However, the addition of antibiotics adds extra operational
costs to fermentation plants and use of a large amount of antibiotic is considered hazardous
to the environment.
Another issue related to using episomal plasmids for heterologous protein expression
is plasmid stability (or segregational instability).
The instability is mainly caused by
unequal separation of plasmids from the mother cell to daughter cells and eventually cells
carrying no plasmids will appear in the cultural medium.
Even under selective conditions,
episomal plasmids are mitotically unstable during long-term cultivation.
For heterologous
protein expression, high plasmid stability is desired, since higher value of the plasmid
stability corresponds to a higher ratio of the number of plasmid-bearing cells over the
number of plasmid-free cells in the fermentor at the checked time point.
Plasmid stability
is normally measured as a ratio of the number of plasmid-bearing cells over that of the
plasmid-free cells at a certain time point during the fermentation (Filho et al., 1986;
Ruohonen et al., 1987; Kondo et al., 2002). Since the selection pressure is lost as a result
of the degradation of the antibiotic, the plasmid-free cells grow whenever free fermentable
sugars are available in the fermentor. The plasmid-free cells almost always overgrow the
plasmid-bearing cells under the conditions where selection pressure is lacking (Altintas et
al., 2001; Nakamura et al., 2002).
It was reported that without selection pressure, plasmid-free cells can quickly
35
overgrow plasmid-bearing cells (Alintas et al., 2001). Plasmid stability was studied based
on data from starch fermentation using recombinant S. cerevisiae strain YPB-G, which
expresses and secretes B. subtilis -amylase and A. awamori glucoamylase as a fusion
protein.
The recombinant yeast were pre-cultured in minimal medium, and used to
inoculate a 2.5-litre bioreactor containing 30 g L-1 soluble starch at 30C. A mathematical
model was used to simulate plasmid stability in the fermentation broth and then actual
experimental data were obtained to confirm the prediction from the model. Based on
experimental
observations,
during
the
first
generation,
plasmid-free
cells
and
plasmid-bearing cells grew at the same rate. However, starting with the second generation,
the growth rate of plasmid-free cells surpassed plasmid-bearing cells and reached twice the
growth rate of plasmid-bearing cells. Their results indicated that as more fermentable
sugars became available, plasmid-free cells tend to outgrow plasmid-bearing cells, with the
biomass of plasmid-free cells and plasmid-bearing cells becoming 2.27 g L-1 to 0.51 g L-1,
respectively. Based on the data, the authors suggested that high copy number plasmids are
relatively stable under selection conditions but not suitable for large-scale fermentation in
long-term culturing conditions. They further showed using an integrative plasmid to insert
the DNA sequence into the yeast chromosome, that this approach could solve the plasmid
stability problem.
Genes integrated into the yeast genome are generally very stable, even
under non-selective conditions.
2.5.3.2 Use of Yeast Integration Plasmids for Expression of Heterologous Genes
The use of yeast integration plasmids is another effective way for introducing
heterologous genes into S. cerevisiae.
Integration of genes into yeast chromosomes
eliminates the segregational instability related to episomal plasmids, and offers an
alternative for stable expression of heterologous genes. Yeast integration plasmids (YIp)
have the same composition as episomal plasmids except that the yeast replication origin is
replaced with a segment of DNA which is homologous to the yeast genome sequence where
the plasmid is integrated.
The integration plasmid is normally linearized at the
homologous DNA sequence before transformation, since linearized plasmids are
transformed at a much higher efficiency than circular ones.
36
Traditionally, heterologous
genes are integrated at gene sequences involved in amino acid synthesis, such as URA3,
TRP1, or HIS3, via homologous recombination.
However, only one or a few tandem
copies can be inserted at these targeting locations (Romanos et al., 1992). Compared to the
copy numbers of episomal plasmids that a single yeast cell can maintain, the copy numbers
of genome integrated genes markedly limits the expression levels of heterologous proteins.
In order to obtain multiple integrations of the heterologous gene into the yeast
genome, studies have shown that several repetitive chromosomal DNA sequences, such as
Ty1 elements (Kingsman et al., 1985), the ribosomal DNA (rDNA) cluster (Lopes et al.,
1991; 1996), or  sequences (Sakai et al., 1991; Lee and Silva, 1997) can be employed as
the target location for high-copy-number integration of heterologous genes.
The S. cerevisiae  sequences are found to exist in close association with the long
terminal repeats of retrotransposons Ty1 and Ty2 (Boeke, 1989; Boeke and Sandmeyer,
1991), or found scattered in the yeast genome as isolated and independent elements (Boeke
1989).
The  sequences are dispersed throughout the genome at approximately 100 copies
(Cameron et al., 1979; Boeke, 1989), and have been employed as targets for the integration
of a variety of cloned genes into the host chromosomes (Shuster et al., 1990; Sakai et al.,
1990, 1991; Wang et al., 1995; Lee and Silva et al., 1997).
Wang et al. (1995)
demonstrated multiple-copy integration of a G418-resistance gene into the  sequences of a
haploid laboratory strain of S. cerevisiae using this approach.
A set of randomly selected
transformants showed 10-43 copies of the integrated gene cassette in the yeast genome that
were structurally stable for 70-100 generations in non-selective medium.
However,
inclusion of a SUC2 gene under the control of the MF1 promoter, in the integration
cassette resulted in a significant decline in structural stability.
Transformants showed
lower copy numbers of the integrated cassette (2-16 copies), and of the ones with five or
more integrated copies, 20-75% of the cells lost all integrated SUC2 cassettes during 70-100
generations of non-selective growth.
Lee and Solva (1997) employed a similar -sequence integration plasmid to
over-express the E. coli lacZ gene in a haploid strain of S. cerevisiae, however structural
instability and low integration copy number of the -sequence integrated gene cassette was
observed.
By employing a double--sequence integration plasmid, which carries two 
37
sequences flanking the integration gene cassette instead of single  sequence in the previous
designs, they found that the transformants had increased copy numbers (more than double)
and higher structural stability of the insertions.
In the double--sequence integration
plasmid, the unneeded bacterial sequences were removed, which reduced the size of the
integration plasmid from ~8 kb to ~5 kb.
The double--sequence integration system
showed higher expression level of the inserted gene with no negative effect on cell.
The other commonly used multiple-integration plasmid targets the yeast rDNA
sequences.
Yeast rDNA region consists of about 150-200 tandem repeats on chromosome
XII (for a haploid yeast cell), and each repetitive unit is about 9 kb in size (Lopes et al.,
1991).
Each rDNA encodes two primary transcripts, one for 5S rRNA and one for 37S
pre-rRNA, separated by two nontranscribed spacers both of which are about 1 kb long
(NTS1 and NTS2) (Skryabin et al., 1984).
Lopes et al. (1989) constructed an rDNA
integration plasmid containing a deficient LEU2d gene as the auxotrophic selection marker.
Targeting the plasmid into the rDNA of a haploid strain of S. cerevisiae resulted in
transformants carrying 100-200 inserts per cell.
The cells retained 80-100% of the
integrated plasmid copies after 70 generations of batch growth under nonselective
conditions.
They found that, by using this integration plasmid, expression levels of
heterologous genes were comparable to that using an episomal plasmid. The deficient
LEU2d gene contains a deficient promoter of the wild type LEU2 gene, which causes a
minimal level of leucine synthesis in the transformants.
Lopes et al. (1991) showed that
the deficient promoter region in the selection marker gene was the key for high copy
number integration, since switching to the wild type LEU2 gene as the selection marker
dramatically reduced the copies of inserts.
Initially, the integration plasmid only integrated
in low copy numbers, and it was the strong selection pressure posed by the deficient
selection gene that amplified the integrated copy numbers dramatically (Lopes et al., 1991).
In addition, factors that impacted the structural stability of the integrated cassette were also
studied.
They found that the size of the integrated gene cassette should be within 9 kb,
which is the size of a single rDNA tandem unit; otherwise the structural stability of the
insert drops dramatically.
Also, incorporating rDNA segments that contain the Pol I
transcription-termination region or the TOP1 site, or avoiding the Pol I transcription
38
initiation region in the integration plasmid seem to increase the stability of the inserts
(Lopes et al., 1996).
In summary, the use of a multiple integration system for heterologous protein
expression is advantageous compared to episomal plasmids, especially for large-scale
fermentation, because of the high mitotic stability of heterologous genes and high
expression levels of the genes integrated.
2.5.3.3 High-Level Expression of Heterologous Proteins in Saccharomyces cerevisiae
Many strategies can be used to optimize the expression of heterologous proteins in S.
cerevisiae.
Yeast constitutive promoters, such as GAPDH (or TDH3), ADH1 (or ADC1) and
PGK1, have been widely used to optimize the transcription levels of heterologous genes.
Even though many non-yeast promoters have been shown to be functional in yeast systems,
the activity is relatively low compared to yeast promoters.
Inducible promoters have also
been used for the expression of toxic proteins.
In addition to promoters, high-level protein expression can be a result of the
stabilization of ribosome binding on the mRNA during translation initiation. An optimal
AUG context, (A/U)A(A/C)AA(A/C)AUGUC(U/C), was reported to increase protein
expression levels up to 10-20% in yeast (Hamilton et al., 1987; Miyasaka et al., 1999).
Extreme codon bias was reported for highly expressed S. cerevisiae genes (Bennetzen and
Hall, 1982).
Given that DNA synthesis is becoming cheap and efficient, synthesizing a
heterologous gene composed of all yeast-favored codons is now possible. However, many
studies have shown that optimizing the codon sequences in the heterologous gene is not
always necessary for obtaining high-level expression (Juge et al., 1993).
With respect to amylolytic enzyme over-expression and secretion into the culture
medium, secretion signal peptides have also been employed for efficient secretion of the
amylolytic enzymes.
Yeast native secretion signal peptides were shown to direct efficient
heterologous protein secretion (Notano and Shishido, 1988).
R. oryzae glucoamylase and
barley -amylase 1 were shown to be secreted efficiently from S. cerevisiae using their
native secretion signal peptide (Ashikari et al., 1986; Juge et al., 1993; Wong et al., 2002).
39
However, wheat -amylase and barley -amylase 2 showed markedly reduced secretion
levels from S. cerevisiae using their native secretion signal peptides (Rothstein et al., 1984;
Juge et al., 1993).
Expression of proteins that assist in protein folding or secretion were also found to
increase heterologous protein expression and secretion. Over-expression of S. cerevisiae
Sso protein, which is involved in the fusion of secretory vesicles to the plasma membrane,
were found to increase the secretion of B. amyloliquefaciens -amylase (Ruohonen et al.,
1997). Over-expression of an unfolded-protein response (UPR) pathway regulator, HAC1,
produced a 70% increase in B. amyloliquefaciens -amylase secretion (Valkonen et al.,
2003).
Environmental factors, such as agitation, aeration, temperature, pH, and medium
composition can also affect heterologous protein expression dramatically.
highly medium-dependent.
S. cerevisiae is
Rothstein et al. (1987) observed that use of rich medium can
increase the expression and secretion of wheat -amylase from S. cerevisiae compared to
minimal medium.
Increased expression levels of certain proteins were observed when
more amino acids (Castelli et al., 1994) or a higher amount of glucose (Ichikawa et al., 1989)
A significant increase in barley -amylase
was supplemented into the yeast medium.
expression and secretion by S. cerevisiae was observed when glucose was substituted with
glycerol in YPD medium (Wong et al., 2002).
Studies using other yeast species have also
provided valuable information. Casamino acids (1%) were shown to increase heterologous
protein expression and plasmid stability in Kluyveromyces lactis (Macreadie et al., 1993).
It is clear that a combination of several factors is necessary to achieve optimal level
of expression.
There is likely no “best” combination that can be universally applied to one
specific circumstance.
2.5.4 Recombinant Saccharomyces cerevisiae that Secrete Heterologous Amylolytic
Enzymes
Genes coding for amylolytic enzymes from various organisms have been introduced
into S. cerevisiae and expressed in their fully functional forms.
These include the
-amylase gene from wheat (Rothstein et al., 1984; 1987), mouse salivary g Land
40
(Thomsen, 1983), mouse pancreas (Filho et al., 1986), human salivary g Land (Nakamura et
al., 1986), human lung carcinoid tissue (Shiosaki et al., 1990), Lipomyces kononenkoae
(Eksteen et al., 2003), B. subtilis (Rouhonen et al., 1984), B. amyloliquefaciens (Kovaleva
et al., 1989; Pretorius et al., 1988; Ruohonen et al., 1987), B. stearothermophilus (Nonato
and Shishido, 1988), Schwanniomyces occidentalis (Wang et al., 1989) and the
glucoamylase genes from A. awamori (Cole et al., 1988; Inlow et al., 1988; Innis et al.,
1985), R. oryzae (Ashikari et al., 1986; 1989), Saccharomycopsis fibuligera (Yamashita et
al., 1985) and S. diastaticus (Erratt and Nasim, 1986; Pardo et al., 1986; Pretorius et al.,
1986). These studies demonstrated that it is feasible to express and secrete highly active
heterologous amylolytic enzymes in S. cerevisiae.
However, most of the early studies
focused only on expression and characterization of the expressed enzymes, while starch
assimilation and fermentative ability of the recombinant amylolytic yeast were not assessed.
Moreover, early studies employed 2µ-based yeast episomal plasmids for
heterologous amylolytic enzyme expression.
Even though desirable expression levels were
achieved, during prolonged fermentation, the expression levels were often observed to drop
because of mitotic instability of the episomal plasmids (Wang et al., 1989).
In order to
cope with this issue, many research groups switched to integration plasmids for amylolytic
enzyme expression in S. cerevisiae.
Integrated genes have been shown to be mitotically
stable even under non-selective conditions.
Nakamura et al. (1996) reported a
recombinant strain of S. cerevisiae that stably expressed a glucoamylase gene from S.
diastaticus. The integrated gene was mitotically stable for 200 h during fermentation.
In
addition, instead of expressing one type of amylolytic enzymes, Steyn et al. (1991) found
that a higher starch hydrolysis rate can be achieved by integrating both glucoamylase and
-amylase.
The recombinant strains secreting S. diastaticus glucoamylase, B.
amyloliquefaciens -amylase or both enzymes showed the ability to hydrolyze soluble
starch with an efficiency of 69%, 84% and 93%, respectively over 120 h, showing that the
one expressed both enzymes hydrolyzed the starch most efficiently.
Furthermore, all three
recombinant strains showed higher starch hydrolysis rates than the wild type S. diastaticus,
which is a naturally-occurring amylolytic yeast, suggesting that S. cerevisiae can be
engineered to hydrolyze starch with higher efficiency than the naturally-occurring
41
amylolytic yeast.
Given that S. cerevisiae is superior to the naturally-occurring amylolytic
yeast on high ethanol tolerance and producing ability, genetically engineered S. cerevisiae
has great potential in its application in industrial bioethanol production from starch
materials.
However, traditional integration plasmids only insert one to few copies of the
integrated gene in the yeast genome (Romanos et al., 1992), which is low compared to the
copy numbers of episomal plasmids can be found in a yeast cell.
In order to increase the
integrated gene copy numbers, it was shown that repetitive gene sequences in the yeast
genome could be targeted for integration of an expression gene cassette.
By targeting the
integrated genes into the repetitive sequences in yeast genome such as rDNA or  sequences,
the copy numbers of the integrated genes up to 100-150 have been reported (Lopes et al.,
1991; Wang et al., 1996; Lee et al., 1997; Nieto et al., 1999).
Heterologous genes coding
for amylolytic enzymes have also been integrated in high copy numbers in the yeast genome.
Lin et al. (1998) integrated the gene of A. awamori glucoamylase into the rDNA loci of the
yeast genome.
By employing a deficient promoter of URA3, 140 copies of the integrated
gene were achieved in one of the isolated yeast clones.
As expected, the clone with the
highest amylolytic activity also had the highest copy number of glucoamylase genes.
The
inserts were shown to be mitotically stable for over 50 generations when cultivated on
non-selective medium.
With such high integrated gene copy numbers, the expression level
of the amylolytic enzyme can be comparable to episomal plasmids are used (Nieto et al.,
1999).
Since integrated genes are more mitotically stable than episomal plasmids,
integrating genes coding for amylolytic enzymes in the yeast genome is more practical for
industrial application than employing episomal plasmids.
Haploid laboratory yeast strains were widely used in early studies because they are
easy to manipulate at the genetic level.
However, haploid yeast strains have very little use
in industrial applications due to their low stress tolerance.
Most brewing and distiller’s
yeast strains are polyploid, which are superior to the haploid laboratory strains with respect
to ethanol production and stress tolerance.
As mention in previous sections, since
auxotrophic mutants of polyploid strains are difficult to obtain, a dominant selection marker
needs to be used for the selection of polyploid strains of S. cerevisiae.
42
By employing the
dominant selection marker AUR1-C, which encodes resistance to aureobasidin A, Kang et al.
(2003) successfully expressed Schwanniomyces occidentalis -amylase in a polyploid
industrial strain of S. cerevisiae. The expression gene cassette was integrated in multiple
copies in the  sequences of the yeast genome. The integrated genes showed 100% mitotic
stability over 100 generations on non-selective medium.
In order to improve the
starch-hydrolyzing ability of this strain, Ghang et al. (2007) co-expressed Debaryomyces
occidentalis glucoamylase with S. occidentalis -amylase in the recombinant yeast.
Unexpectedly, the integrated genes showed low mitotic stability, resulted in lower
amylolytic activity compared to that of the previous strain.
It was speculated that the
over-sized integration plasmid, which contained both glucoamylase and -amylase genes
caused the mitotic instability of the integrated genes.
By reducing the size of the
integration plasmid by deleting the bacterial originated sequences in the plasmid, higher
mitotic stability was achieved (Ghang et al., 2007).
Similar findings that over-sized
integrated plasmids could result in low mitotic stability were also reported when genes were
integrated into the rDNA sequences, and the mitotic stability was improved by decrease the
size of the plasmids (Lopes et al., 1996).
Instead of incorporating both glucoamylase and -amylase genes in the same
integration plasmid as performed by Ghang et al. (2007), Wong et al. (2010) used two
separate integration plasmids for each gene. Designing the plasmid in this way avoided an
over-sized integration plasmid that can cause mitotic instability.
Wong et al. (2010)
successfully co-expressed both Lentinula edodes glucoamylase and barley -amylase in a
strain of S. cerevisiae with high mitotic stability under non-selective conditions.
The two
integration plasmids containing the glucoamylase and -amylase genes were used to
co-transform the yeast strain.
The -sequence was targeted as the integration locus for
recombination to ensure multiple-copy integration of the glucoamylase and -amylase
genes into the yeast genome.
Over a 6-day period, the recombinant strain co-expressing
both glucoamylase and -amylase hydrolyzed 77% of the starch, showing the highest starch
conversion rate compared to 44% and 67% starch hydrolysis using the recombinant strains
expressed only -amylase and glucoamylase, respectively.
However, they observed that
both exo-cleavage activity and endo-cleavage activity in the co-expressing recombinant
43
strain was lower than that of the one only expressing glucoamylase or -amylase,
respectively.
This was speculated to be due to the competition between glucoamylase and
-amylase genes for space in the  sequences when co-integrated.
Thus, this strategy is
much less constrained by the size of the integration plasmid, but results in reduced
expression of both enzymes.
When different types of amylolytic enzymes are co-expressed in the recombinant
yeast, targeting the integrated genes into different repetitive gene sequences is another
effective strategy.
Kim et al. (2010) constructed a recombinant yeast strain that expressed
three amylolytic enzymes including a glucoamylase from A. awamori, an -amylase, and a
glucoamylase with debranching activity from Debaryomyces occidentalis.
Both
-sequence and rDNA integration systems were employed to achieve multiple integration of
the three genes into the genome of an industrial strain of S. cerevisiae.
The genes encoding
for A. awamori glucoamylase and D. occidentalis -amylase were cloned into an integration
vector based on -sequence integration, and the gene encoding for D. occidentalis
glucoamylase was cloned into another integration vector based on rDNA integration system.
The bacterial originated sequences were deleted to reduce the size of the integration
plasmids.
The recombinant strain expressing all three genes was able to completely
hydrolyze 2% soluble starch in 24 h, and high mitotic stability of the integrated genes was
observed under non-selective conditions.
Thus, the studies described above indicate that polyploidy industrial strains of S.
cerevisiae can be genetically engineered to perform efficient starch assimilation for
fermentation by employing multiple high-copy-number integration systems and a
combination of different amylolytic enzymes.
2.5.5 Recombinant Yeast with Surface-Displayed Amylolytic Activity
2.5.5.1 Cell Wall Anchored Proteins and the GPI Anchor
Many studies have shown that it is possible to use the features derived from native
surface proteins to display heterologous proteins on various cell surfaces.
Proteins have
been displayed on the surface of bacteria, mammalian cells and yeast (Little et al., 1993;
44
Georgious et al., 1993; Dustin et al., 1987; Hiebert et al., 1988; Vijaya et al., 1988).
It has
been suggested that compared to intracellularly-expressed or secreted proteins, cell
surface-displayed proteins have the advantages of stabilizing the anchored enzymes and
simplifying the protein purification since the enzymes can be collected together with the
cells.
Many different cell wall proteins have been identified, such as cell wall protein 1
(Cwp1), cell wall protein 2 (Cwp2), flocculation protein 1 (Flo1) and -agglutinin (AG1)
(Vaart et al., 1995; Schreuder et al., 1996; van der Vaart et al., 1997).
Among these
proteins, -agglutinin is the one that has been studied the most (Lipke et al., 1989, 1992;
Cappellaro et al., 1994; Chen et al., 1995; Zou et al., 2000; Shen et al., 2001; Zhao et al.,
2001; Huang et al., 2003;).
The cell wall-anchoring mechanism of -agglutinin involves many subcellular
compartments in the yeast secretory pathway and several components in the cell wall.
The
two major components of the cell wall of S. cerevisiae are glucan and mannoproteins, which
are present in roughly equal amounts. The mannoproteins, which form the outer layer of
the cell wall, are generally heavily glycosylated, and determine most of the surface
properties of the cell.
Lipke et al. (1998) suggested that mannoproteins are covalently
linked to glucan, because they are resistant to extraction in hot sodium dodecyl sulfate (SDS)
but can be cleaved from the wall by -1,3- and -1,6-glucanase.
Glucan, which is
composed of -1,3 and -1,6 linked glucose, is complexed with chitin to provide
mechanical strength to the cell wall.
-1,3 glucan forms a fibrous network, while -1,6
glucan is highly branched.
-Agglutinin was originally identified as a mannoprotein involved in the sexual
adhesion of S. cerevisiae mating type  cells with mating type a cells (Lipke et al., 1989,
1992).
-Agglutinin consists of 650 amino acid residues, including a N-terminal secretion
signal, N-terminal binding domain (consisting of three different domains that are involved
in the interaction with a-type S. cerevisiae cells), a C-terminal Serine/Threonine rich domain
and a C-terminal glycosylphosphatidylinositol (GPI) addition signal that is involved in cell
wall anchorage (Chen et al., 1995).
Studies have indicated that -agglutinin is covalently
linked to the cell wall glucan (Lipke et al., 1989, 1992; Cappellaro et al., 1994).
45
In
addition, studies showed that the C-terminal Serine/Threonine rich domain is extensively
O-glycosylated, which suggests that this domain might have a rod-like conformation that
acts as a spacer to extend the N-terminal binding domain to the cell surface (Lipke et al.,
1989, 1992; Cappellaro et al., 1994; Chen et al., 1995; Zou et al., 2001; Shen et al., 2001).
Schreuder et al. (1993) and Lu et al. (1995) showed that the 320 amino acids at the
C-terminal half of -agglutinin are responsible for the cell wall anchoring ability.
A
well-accepted explanation of the anchoring process of -agglutinin is that the GPI anchor
attachment signal is recognized in the yeast endoplasmic reticulum (ER).
Then, the
hydrophobic attachment signal is replaced by a GPI anchor by a trans-peptide reaction in the
ER.
Following the attachment of the GPI anchor, -agglutinin is transferred to the outer
leaflet of the plasma membrane. The GPI anchor is then cleaved at its C-terminal glycan
position and the remnant forms a glycosidic linkage with the branched -1,6 glucan in the
cell wall (Lipke et al., 1989, 1992; Huang et al., 2003; Chen et al., 1995).
Even though -agglutinin is the most widely used cell wall anchoring protein in
yeast, many studies have shown reduced or eliminated enzyme activity after being anchored
on the cell wall by -agglutinin. This promoted a search for alternative strategies, and
several other alternative cell wall anchoring proteins have been used for displaying enzymes
on the cell surface, such as Flo1p (Shigechi et al., 2004).
Depending on the particular
enzyme, different anchoring proteins may be chosen for optimal enzyme activity.
2.5.5.2 Displaying Amylolytic Enzymes on the Yeast Cell Wall
The accumulating studies on yeast cell wall structure and cell wall anchored proteins
suggested that heterologous proteins could be immobilized on the yeast cell wall by being
expressed as fusion proteins with one of the yeast’s surface anchoring proteins (Schreuder et
al., 1996).
Murai et al. (1997) first anchored R. oryzae glucoamylase on the cell surface of S.
cerevisiae by fusing the R. oryzae glucoamylase cDNA with the 3’ half of -agglutinin
cDNA, which is the part that has the GPI anchor attachment signal and is involved in
anchoring proteins on the yeast cell wall.
The enzyme is an exo-type amylolytic enzyme
and is able to cleave both -1,4 and -1,6-linkages in starch.
46
They also attached the
coding region for the secretion signal of the glucoamylase precursor protein to the 5’ end of
the fusion gene.
They hypothesized that when the fusion protein reached the cell
membrane, R. oryzae glucoamylase would be covalently linked to glucan in the cell wall
through the GPI anchor, with the enzyme facing away from the cell wall.
To test if the
enzymes were covalently anchored on the cell wall, they first extracted the cell wall with
hot SDS, which removed non-covalently bound proteins or proteins bound through disulfide
bridges.
Then, they treated the hot SDS-extracted cell wall with -1,3-glucanase.
Their
results showed that 93% of the total extractable glucoamylase was covalently anchored on
the cell wall. The recombinant yeast cells were aerobically cultivated on 1% soluble starch
as the sole carbon source for over 100 h. They were able to show that the recombinant
yeast could grow on starch and that cell growth was comparable to that on 1% glucose. The
activity of cell wall-anchored glucoamylase was comparable to secreted forms of the
enzyme.
The same plasmid was then used to transform a flocculation strain of S. cerevisiae,
which is a diploid strain with tryptophan auxotrophy.
Flocculation is a reversible,
non-sexual aggregation of yeast cells at the end of fermentation.
The process simplifies
the separation of cells from the medium at the end of fermentation. The fermentative
ability of the recombinant yeast on soluble starch was studied in batch, repeated-batch and
fed-batch fermentations.
The recombinant yeast was first aerobically cultivated using
soluble starch as the carbon source, then the harvested cells were used for batch,
repeated-batch and fed-batch fermentations on soluble starch under anaerobic conditions.
An ethanol production rate of 0.71 g h-1L-1 was reported with batch fermentation, and
ethanol concentration reached 25 g L-1 after 30 h (60 g L-1 of soluble starch plus 5 g L-1 of
glucose as carbon sources) and the same rate was maintained during repeated-batch
fermentation over 300 h.
In the fed-batch fermentation over approximately 120 h, an
ethanol concentration of 50 g L-1 was reported.
Additionally, a high plasmid stability of
over 80% was observed during the repeated-batch fermentation (Kondo et al., 2002).
Interesting Ly, they were unable to detect glucose in the medium during the fermentation.
Kondo et al. (2002) proposed that since the glucoamylase was anchored on the yeast cell
wall, only those starch molecules close to the yeast cells were degraded by the enzymes, and
47
the released glucose was readily taken up by these yeast cells as a result. This suggests
that there would be no accumulation of large quantities of glucose in the fermentation
medium over time, consistent with their observations.
If no free glucose was available in
the medium, plasmid-free cells would not be able to proliferate. Thus, starch degradation
ability can potentially play the role of a selection agent.
By using the recombinant yeast
with cell surface-anchored glucoamylase during starch fermentation, contamination caused
by bacteria would be prevented, since no free glucose would accumulate in the fermentation
medium.
The recombinant strain’s ability to utilize and ferment on starch was further
improved by co-expressing both glucoamylase and -amylase.
Murai et al. (1999)
developed several recombinant yeast strains that displayed R. oryzae glucoamylase and/or
-amylase from B. stearothermophilus on the surface of the cell wall. The enzymes were
anchored with -agglutinin. They found that the recombinant strains displaying either
glucoamylase alone or both glucoamylase and -amylase were able to grow on starch as the
sole carbon source.
In contrast, the recombinant strain displaying -amylase alone was
unable to grow on starch. Their results indicated a dramatic activity reduction for cell
surface-anchored B. stearothermophilus -amylase.
To improve the cell surface display activity of B. stearothermophilus -amylase,
another anchoring protein, flocculation 1 protein (Flo1p), was employed to anchor the
enzyme in the cell wall.
Flocculation 1 protein is involved in cell-cell adhesion between
yeast cells during flocculation (Miki et al., 1982; Straveret et al., 1994; Bony et al., 1997).
The protein is 1,536 amino acids long, and has a N-terminal secretion signal sequence.
Flocculation 1 protein also contains a spacer domain that is rich in Serine and Threonine
residues, and a hydrophobic GPI-anchor attachment signal sequence at its C-terminus.
-Amylase anchored with -agglutinin showed little activity; however, when anchored with
Flo1p, much higher activity was observed.
They proposed that since the C-terminus of B.
stearothermophilus -amylase is involved in starch binding, that when the C-terminal of the
enzyme was fused to -agglutinin, the anchoring might have hindered the enzyme’s
accessibility for its substrate.
In contrast, Flo1p, when attached to the N-terminal of B.
stearothermophilus -amylase, allowed the enzyme’s C-terminus to extend freely into the
48
medium.
Different from -agglutinin, the N-terminal of Flo1p anchors itself on the cell
wall by non-covalently attaching to the mannoproteins of the outer layer of the cell wall
(Shigechi et al., 2002).
A similar study done by Matsumoto et al. (2002) showed that R.
oryzae lipase, which also has a substrate-binding site located in the C-terminus and showed
no detectable activity when anchored in the cell wall with -agglutinin, showed increased
activity when anchored with Flo1p.
However, the recombinant strain co-displaying R. oryzae glucoamylase and B.
stearothermophilus -amylase was unable to utilize raw starch (Shigechi et al., 2004). By
replacing B. stearothermophilus -amylase with the -amylase from Streptococcus bovis
148, which was shown to have a strong ability to hydrolyze and be absorbed onto raw corn
starch, the new recombinant yeast strain co-displaying R. oryzae glucoamylase and S. bovis
148 -amylase was able to directly utilize raw starch for fermentation. Moreover, the
C-terminal-half region of -agglutinin and the flocculation functional domain of Flop1 were
used as the anchor proteins for R. oryzae glucoamylase and S. bovis 148 -amylase,
respectively.
In a 72-h fermentation, this recombinant strain produced 61.8 g L-1 ethanol
from raw corn starch (86.5% of the theoretical yield) (Shigechi et al., 2004).
The use of recombinant yeast with cell surface-displayed amylolytic activity for
fermentation process has only been recently developed. However, this field has already
shown promising results and the potential to be used in industrial applications.
2.6
Hypothesis and Objectives
2.6.1 Hypothesis
Currently, starch hydrolysis through high temperature cooking is commonly used in
bioethanol production.
in reduced energy input.
Use of cold starch hydrolysis eliminates the cooking step, resulting
Barley -amylase has been found to be superior to bacterial and
fungal -amylases for cold hydrolysis of wheat starch based on its enzymatic kinetic
properties and its pH optimum which is compatible with the pH of yeast culturing
conditions (Textor et al., 1998). However, much larger amounts of starch-hydrolyzing
49
enzymes are generally required for efficient raw starch hydrolysis, which make bioethanol
production by cold starch hydrolysis economically unfeasible.
By employing DNA
recombinant techniques, S. cerevisiae can be genetically engineered to over-express barley
-amylase to levels that would support fermentation on starch.
2.6.2 Rationale
Our lab has previously constructed a recombinant yeast strain that was able to
express cell surface-anchored barley -amylase.
An industrial ethanol-tolerant strain of S.
cerevisiae, NRRL Y-132, was transformed with a yeast expression plasmid pAnchored (Bo
Liao, Master’s thesis, University of Saskatchewan, 2008). The plasmid contains a fusion
gene, which is composed of a barley -amylase (-1, 4 glucan glucanohydrolase, EC
3.2.1.1, type VIII-A) cDNA that had an oligonucleotide coding for a secretion signal
sequence (to excrete the enzyme through the cell membrane) attached to its 5’ end and its 3’
end fused to the 3’ half of the -agglutinin (a mannoprotein involved in the sexual adhesion
of mating type  - S. cerevisiae cells with mating type a - S. cerevisiae cells) cDNA.
The
recombinant yeast expressed barley -amylase with its C-terminus anchored on the yeast
cell wall.
The recombinant yeast showed the ability to hydrolyze soluble starch under batch
fermentation conditions.
However, the recombinant yeast was not able to produce
sufficient -amylase activity, resulting in inefficient starch hydrolysis and no ethanol
production.
Several major factors were speculated to contribute to the low starch
hydrolysis rate, and possible strategies to address these are provided below.
The cell wall-anchored barley -amylase displayed low enzyme activity.
The
reduction in activity was speculated to be a result of steric hindrance caused by the
anchoring process.
In order to improve the activity of the anchored barley -amylase, a
flexible peptide linker can be inserted between the barley -amylase and the anchoring
protein. The linker may provide flexibility between the enzyme and the anchoring protein,
reducing the steric hindrance caused by the anchoring process.
Alternatively, barley
-amylase can be anchored through its N-terminus instead of the C-terminus. Since the
enzyme’s C-terminal domain plays an important role in starch binding (Robert et al., 2005),
50
anchoring barley -amylase through its N-terminus could free the C-terminal domain for
more efficient substrate binding.
Flo1p can be used to perform such an anchoring process
(Matsumoto et al., 2002; Shigechi et al., 2004).
Instead of expressing barley -amylase as the cell wall anchored form, the enzyme
can be secreted.
Although cell wall-anchored enzymes have advantages such as being
more thermostable and reutilizable, they are physically restrained in accessing the substrates
compared to their free counterparts.
-amylase
directly into
starch-hydrolyzing ability.
the
Thus, a recombinant yeast strain that secretes barley
medium
might
be
expected
to
show
improved
Comparison of the two expression systems will provide
insights as to which system is more efficient for starch hydrolysis.
Furthermore, the 2µ-based episomal plasmids are mitotically unstable without the
presence of adequate selection pressure; this instability reduces the expression level of
barley -amylase.
Continuous addition of large amounts of antibiotics can keep the
plasmid stability high during fermentation; however, it is not applicable for bioethanol
production at the industrial scale.
An alternative method is to integrate the barley
-amylase gene into the yeast genome through homologous recombination.
In theory, the
integrated gene should be mitotically stable even in the absence of selection pressure.
Moreover, high copy numbers of the expressed gene can be achieved by integrating the gene
into repetitive sequences in the yeast genome.
2.6.3 Objectives
The goal of this project was therefore to optimize the starch-hydrolyzing ability of
an engineered recombinant yeast so that it is able to produce sufficient -amylase activity
for raw starch hydrolysis.
Both genetic and environmental factors were studied and
combined to optimize the performance of the recombinant yeast under batch fermentation
conditions.
The specific objectives of this thesis were:
1.
To improve the activity of cell wall-anchored barley -amylase.
2.
To express barley -amylase in a secreted form.
3.
To compare the starch-hydrolyzing ability of recombinant yeast expressing the
51
anchored and secreted forms of barley -amylase.
4.
To improve on the expression level of barley -amylase by modifying environmental
and genetic factors.
5.
To stabilize and increase the expression of barley -amylase under non-selective
fermentation conditions by integrating the expression cassette into the yeast genome.
52
3.0
MATERIALS AND METHODS
3.1
Reagents
The names of the reagents and their suppliers are listed in Table 3.1. The addresses of the
individual suppliers are given in Table 3.4.
Table 3.1: Lists of reagents and suppliers.
Reagent
Supplier
Absolute Ethanol
BDH
Agarose
Bio-Rad
Dimethylsulfoxide (DMSO)
BDH
Ethidium Bromide
Sigma-Aldrich
Ethylene-Diamine Tetraacetic Acid Disodium Salt (EDTA)
BDH
Glacial Acetic Acid
EMD
D-Glucose
BDH
Glycerol
BDH
Hydrochloric Acid (HCl)
BDH
Isopropanol
BDH
Sodium Hydroxide (NaOH)
BDH
Ampicillin
ICN
Blasticidin
Invitrogen
Bacto-Agar
DIFCO
Bacto-Tryptone
DIFCO
Bacto-Yeast Extract
DIFCO
Peptone
DIFCO
53
Table 3.2: List of commercial kits.
Commercial Kit
Supplier
QIAquick Gel Extraction Kit
Qiagen
TOPO TA Cloning Kit (pCR2.1 TOPO)
Invitrogen
Plasmid pTEF1/Bsd
Invitrogen
Plasmid Miniprep DNA Kit (catalog# 13300)
Norgen
Table 3.3: List of oligonucleotides for PCR.
Note: restriction sites are under-lined; mutations are bolded.
Primer
Primer Sequences
Names
Primer 1
5’-GGTGTGTGGGCGCGAGCTAGCGATATCGC-3’
Primer 2
5’-GCGATATCGCTAGCTCGCGCCCACACACC-3’
Primer 3
5’-CGGTACCCGGGGATCCCTCGAGCGTC-3’
Primer 4
5’-GACGCTCGAGGGATCCCCGGGTACCG-3’
Primer 5
5’-GCCGCGGCAACACTCTAGCGTCCC-3’
Primer 6
5’-GGGACGCTAGAGTGTTGCCGCGGC-3’
Linker-1
5’-(phos)-TCGAGGGTTCTTCTGGTGGTTCTGGTGGTTCTGGTGGTTCTGGTGGTTCTGGTTCTC-3’
Linker-2
5’-(phos)-TCGAGAGAACCAGAACCACCAGAACCACCAGAACCACCAGAACCACCAGAAGAACCC-3’
Primer 7
5’-CTTGTCACCAGAATTCAGAACATCC-3’
Primer 8
5’-GTTGGTATCGATGACGGTGGTC-3’
Primer 9
5’-GCAAGCTGAATACCCGGGGATCCTCTAGAGTCG-3’
Primer 10
5’-CGACTCTAGAGGATCCCCGGGTATTCAGCTTGC-3’
Primer 11
5’-TTATCAAGCATGCCATTTCAAAGAATACG-3’
Primer 12
5’-GCAACTCTGAATTCTTTGTTTGTTTATGTG-3’
Primer 13
5’-CATCAACCACCGCGCCGCCGACTACAAGG-3’
Primer 14
5’-CCTTGTAGTCGGCGGCGCGGTGGTTGATG-3’
Primer 15
5’-CGGGGCCTAGTTTAGAGAGAA-3’
(continues on next page)
54
Primer 16
5’-ATTCGCACTCTGCAGCTGCACT-3’
Primer 17
5’-AGCCGGGGGCTAGCTTAGAGAGAAGTAGAC-3’
Primer 18
5’-CCTGCACTAACATTTTGCCGCATTACAC-3’
Primer 19
5’-GTGTAATGCGGCATGCTGTTAGTGCAGG-3’
Primer 20
5’-CAACTGTAGTTAAGCTTGTAAGAGCCTGA-3’
dTEF1-1
5’-TTTCTCGAGCTCCGCGCATC-3’
dTEF1-2
5’-TTTCTTCCTCGAGGGTGTCGT-3’
dTEF1-3
5’-CCCGTACTCGAGGTTTGGAAAAG-3’
dTEF1-4
5’-CCGCCTCGAGTCTTTTTCTTCG-3’
dTEF1-5
5’-TTCTCTCTCGAGGACCTCCCA-3’
dTEF1-6
5’-CTCGAGTTTCAGTTTCATTTTTCTTGTTC-3’
dTEF1-R
5’-TGCCAAGCTTGCATGCCT-3’
Table 3.4: Names and addresses of suppliers.
Supplier
Address
BDH
501-45th Street West, Saskatoon, SK., Canada
Bio-Rad
5671 McAdam Road, Mississauga, Ont., Canada
DIFCO
7 Loveton Circle, Sparks, MI 48232-7058, USA
EMD Biosciences, Inc.
10394 Pacific Center Court, San Diego, CA 92121, USA
ICN
12 Morgan, Irvine, CA 92618-2005, USA
Invitrogen
100 Faraday Avenue, Carlsbad, CA 92008, USA
Sigma-Aldrich
2149 Winston Park Drive, Oakville, Ont., Canada
Qiagen
2800 Argentia Road, Unit 7, Mississauga, Ont., Canada
NEB
1815 Ironstone Manor, Unit 6, Pickering, Ont., Canada
Norgen
3430 Schmon Parkway, Thorold, Ont., Canada
55
3.2
Bacteria Strain, Yeast Strains and Media Preparations
E.coli NM522 (F' proA+B+ lacIq Δ(lacZ)M15/ Δ(lac-proAB) glnV thi-1
Δ(hsdS-mcrB)5, New England Biolabs) was used as the host for the propagation of plasmids.
Yeast strain S. cerevisiae NRRL Y-132 was obtained from the Northern Regional Research
Laboratories, USDA, Peoria, IL.
Terrific Broth (TB) medium was used to cultivate and propagate E. coli NM522.
TB consisted of 1.2% (w/v) bacto-tryptone, 2.4% (w/v) bacto-yeast extract, and 0.4% (v/v)
glycerol. The solution was autoclaved for 20 min at 15 lb/sq. in., and then completed by
adding 10% (v/v) sterile 0.17 M KH2PO4 and 0.72 M K2HPO4 to the cooled medium.
solution was placed at 4C for long-term storage.
The
Luria-Bertani (LB) medium and
Luria-Bertani Ampicillin (LBA) plates were used to cultivate and select transformed
bacterial cells.
LB medium consisted of 1% (w/v) tryptone, 0.5% (w/v) yeast extract and
0.5% (w/v) sodium chloride.
The solution was autoclaved and place at 4C for storage.
LBA consists of 1% (w/v) tryptone, 0.5% (w/v) yeast extract, 0.5% (w/v) sodium chloride
and 1.5% (w/v) agar.
After autoclaving, ampicillin was added to a final concentration of
100 g mL-1 into the cooled medium.
The medium was then mixed and poured into petri
dishes. The LBA plates were placed at 4C for long-term storage.
YPD (1% (w/v) yeast extract, 2% (w/v) peptone and 2% (w/v) D-glucose) medium
was used primarily for yeast pre-cultivation and propagation during competent yeast
preparation.
The appropriate amount of blasticidin was added into the medium when the
recombinant yeast strains were cultivated.
YPS (1% (w/v) yeast extract, 2% (w/v) peptone
and 2% (w/v) soluble potato starch) medium was used for most batch fermentation studies.
All media were autoclaved before use, and placed at 4C for long-term storage.
3.3
Reagents for Fermentation and Starch Concentration Analysis
Glucoamylase (amyloglucosidase; 1, 4, -D-glucan glucanohydrolase, EC 3.2.1.3,
catalog #A-7255) from Rhizopus was obtained from Sigma-Aldrich.
Soluble potato starch
(catalog #A-2004), unmodified regular corn starch (catalog #S4126) and waxy corn starch
(catalog #S9679) were also purchased from Sigma-Aldrich.
56
Wheat Starch 4 was donated
by CSP Mills, Saskatoon (manufactured by Archer-Daniels-Midland/Ogilvie in Montreal).
Wheat starch 4 is a class of A grade starch which consists of particles ranging from 18 to 20
m in size.
Iodine colour reagent solution for starch concentration analysis contained 0.5%
(w/v) iodine (I2, Sigma-Aldrich, catalog #229695), 5% (w/v) potassium iodide (KI,
Sigma-Aldrich, catalog #204102) and 0.5 N HCl.
3.4
DNA Subcloning
Protocols in this section are based on those described in Sambrook et al. (1989).
3.4.1 Restriction Digestion of Plasmid DNA and PCR-Generated DNA Fragments
All of the restriction enzymes and the corresponding buffers were purchased from
New England Biolabs (NEB).
Digestion of plasmid DNA with restriction enzymes was
carried out with 0.5- 1 g DNA, 1X NEB buffer, and 1-2 U of each restriction enzyme in a
final volume of 20 L.
Digestions were performed at 37C for 1-2 h.
3.4.2 Agarose Gel Electrophoresis and Gel Purification of DNA Samples
DNA samples (plasmid DNA or PCR-generated DNA fragments) were subjected to
agarose gel electrophoresis in a 1% agarose gel containing 40 mM Tris-Acetate and 1 mM
EDTA (TAE) at pH 8.0 and 1 g mL-1 ethidium bromide in TAE running buffer. DNA
samples were mixed with an appropriate volume of 5X NEB agarose gel sample buffer
before being loaded onto the gel.
Electrophoresis was carried out at 100 volts until the
necessary resolution was achieved.
For downstream ligation reactions, DNA samples were extracted and purified from
the agarose gel using the QIAquick Gel Extraction Kit (Qiagen).
3.4.3 Ligation of Digested Plasmid DNA
Each reaction was set up as follows: 3-30 ng of vector DNA, 9-90 ng insert DNA
and 1 unit of T4 DNA ligase were used for each 20 µl ligation reaction and the reactions
were incubated overnight at room temperature.
57
3.4.4 Competent Bacteria Preparation
A single colony of E. coli NM522 was picked from a LB plate and inoculated into 2
mL of LB medium, then incubated with shaking at 180 rpm at 37ºC overnight.
The 2 mL
overnight culture was transferred into another 200 mL of sterilized LB medium, followed by
an additional 1-2 h of incubation with shaking at 200 rpm and 37ºC.
The bacterial
cultivation was stopped when the O.D.590 reached 0.375. The cell culture was transferred
into pre-chilled sterile 50 mL tubes, and centrifuged at 7,000X g at 4ºC for 7-10 min to
collect the cells.
The cell pellet was resuspended in 10 mL of cold CaCl2 solution (60 mM
CaCl2, 15% glycerol, and 10 mM PIPES pH 7.0) and re-centrifuged. This washing step
was repeated twice and the cell suspension was kept on ice for 30 min followed by a further
centrifugation step to re-pellet the cells. The cell pellet was resuspended in 4-8 mL of cold
CaCl2 solution. The concentrated cell suspension was aliquoted into 1.5 mL sterile tubes
and stored at -80ºC.
3.4.5 Escherichia coli Transformation
One hundred L of competent E. coli NM522 cells were thawed on ice, and then
mixed and aliquoted into pre-chilled 14-mL BD Falcon polypropylene round-bottom tubes.
Two to five µL of DNA ligation mixture were mixed with competent cells by gently
swirling the tube several times.
The tubes were incubated on ice for 30 min, then
subjected to a heat-pulse at 42ºC for 30 sec. The tube was kept on ice for 2 min after the
heat-pulse. One mL of LB media was transferred into the tube and incubated at 37ºC for 1
h. Two hundred µL of the transformation mixture were spread onto a LBA plate and
incubated at 37ºC overnight.
3.4.6 Mini-Preparation of Plasmid DNA
Mini-preparation of plasmid DNA was performed according to the manufacturer’s
instructions (Plasmid MiniPrep DNA Kit, Norgen).
58
3.5
Plasmid Construction
The plasmid maps are presented in Results section.
The software that was used for
plasmid drawing is Plasm (http://biofreesoftware.com).
3.5.1 Construction of Plasmid pSecreted
Plasmid pSecreted was constructed by deleting the -agglutinin domain from
pAnchored (Bo Liao, Master’s Thesis, University of Saskatchewan, 2008).
To achieve this,
an additional XhoI restriction site in pAnchored was first silenced by performing
site-directed mutagenesis using Primers 1 and 2.
The PCR reaction for site-directed
mutagenesis was set up as follows (in a total reaction volume of 50 L): 1X reaction buffer
(20 mM Tris-HCl (pH 8.8), 10 mM (NH4)2SO4, 10 mM KCl, 2 mM MgSO4, 0.1% (v/v)
Triton X-100, 0.1 mg mL-1 BSA), 0.2 mM dNTPs, 5 µL of DMSO, 125 ng each of Primers
1 and 2, 50 ng of plasmid DNA (diluted in double distilled water) and 2.5 U of PfuTurbo
DNA polymerase (Stratagene, catalog#200519).
PCR thermal cycling conditions were set
up as follows: (1) initial denaturation step at 95C for 30 sec; (2) denaturation step at 95C
for 30 sec; (3) annealing step at 55C for 1 minute; (4) extension step at 68C for 10 min; (5)
repeat 18 cycles from step (2) to step (4). The PCR products were treated with DpnI for 1
h and then used to transform XL-10 Gold Supercompetent E. coli (Stratagene,
catalog#200519) according to the QuikChange Site-Directed Mutagenesis Kit Instruction
Manual (Revision#124008p).
The -agglutinin cDNA had a KpnI site at 5’ end and a XhoI site at 3’ end. An
additional XhoI restriction site was generated downstream of the KpnI site by performing
site-directed mutagenesis using Primers 3 and 4. The PCR reaction conditions were the
same as the one described above.
After the two sequential site-directed mutagenesis
modifications, the -agg Lutnin cDNA was deleted from pAnchored by XhoI restriction
digestion. The plasmid was re-circularized by performing a ligation reaction.
Finally, a stop codon was generated at the 3’ end of the barley -amylase cDNA by
mutating a single nucleic acid at the XhoI site using Primers 5 and 6.
conditions were the same as that described above.
The PCR reaction
This final plasmid was named
pSecreted, which allows for expression and secretion of the barley -amylase isozyme 1.
59
The sequence of plasmid pSecreted was confirmed by DNA sequencing.
3.5.2 Construction of Plasmid pLinker
Plasmid pLinker was constructed by inserting a 57 nucleotide-long double-stranded
DNA fragment (Linker) with 3’ and 5’ flanking XhoI restriction digestion sites into
pAnchored.
The
linker
encodes
for
a
peptide
with
the
sequence
GSSGGSGGSGGSGGSGS (G: Glycine, S: Serine). The two complementary DNA strands
of Linker (Linker-1 and Linker-2) were annealed as follows: 50 µM of each
deoxyoligonucleotide were mixed in a solution of 10 mM Tris-HCl and 1 mM EDTA (pH
8.0), then the solution was boiled in a water bath for 15 min and slowly cooled to room
temperature. After the annealing process, 2 µL of the solution was subjected to a ligation
reaction to insert the Linker into the XhoI site between the barley -amylase cDNA and the
-agglutinin cDNA of plasmid pAnchored.
The extra XhoI site in pAnchored was
previously silenced by site-directed mutagenesis as described in Section 3.5.1 and then the
plasmid was linearized by XhoI restriction digestion prior to the ligation reaction. The
final plasmid was named pLinker. The proper in-frame ligation of Linker in pLinker was
confirmed by DNA sequencing.
3.5.3 Construction of Plasmid pFLO1
The cDNA that contained the secretion signal and the mannose binding domain of
flocculation protein 1 (FLO1) was ligated into plasmid pSecreted between the EcoRI and
ClaI restriction sites to generate plasmid pFLO1.
The cDNA of FLO1 was amplified from
the plasmid pBisou, which contains the full-length of FLO1 cDNA (a gift from Dr. Blondin).
The PCR reaction was set up as follows: 1X reaction buffer (10 mM Tris-HCl (pH 9.0),
1.5 mM KCl, 2 mM MgCl2, 0.1% (v/v) Triton X-100), 0.2 mM dNTPs, 125 ng each of
Primers 7 and 8, 50 ng of yeast genomic DNA that was isolated and purified from S.
cerevisiae NRRL Y-132 using Qiagen Genomic DNA Extraction Kit, and 2 U of EconoTaq
DNA polymerase (Lucigen, catalog#30031-1).
PCR thermal cycling conditions were set
up as follows: (1) initial denaturation step at 95C for 2 minutes; (2) denaturation step at
95C for 30 sec; (3) annealing step at 55C for 30 sec; (4) extension step at 72C for 1
60
minutes; (5) repeat 30 cycles from step (2) to step (4); (5) final extension step at 72C for 10
minutes.
The amplified PCR products of FLO1 were first ligated into the TOPO TA Cloning
vector (Invitrogen, TOPO TA Cloning Kit (pCR2.1 TOPO)) and then the ligation reaction
was used for E. coli transformation according to the manufacturer’s instructions. The
DNA fragments of FLO1 were isolated from the TOPO TA Cloning vector by performing a
double digestion with EcoRI and ClaI, and then the isolated FLO1 fragment was ligated into
plasmid pSecreted. Prior to this, the second EcoRI site in plasmid pSecreted was silenced
by performing site-directed mutagenesis using Primers 9 and 10 under the same conditions
as described in Section 3.5.1. This plasmid was then linearized with EcoRI and ClaI prior
to the ligation reaction.
The final plasmid was named pFLO1. Proper in-frame ligation
of pFLO1 was confirmed by DNA sequencing.
3.5.4 Construction of Plasmid pSecreted (TDH3)
Yeast episomal plasmid pSecreted (TDH3) was constructed by replacing the ADH1
(~400 bp) promoter with yeast promoter TDH3 (~650 bp).
Yeast promoter TDH3 was amplified from the yeast genome DNA by PCR.
Yeast
genomic DNA was isolated and purified from S. cerevisiae NRRL Y-132 using Qiagen
Genomic DNA Extraction Kit. The PCR reaction was set up using Primers 11 and 12
under the same conditions as that described in Section 3.5.3.
The amplified PCR product of TDH3 was first ligated into the TOPO TA Cloning
vector (Invitrogen, TOPO TA Cloning Kit (pCR2.1 TOPO)) and then the ligation reaction
was directly subjected to E. coli transformation according to the manufacturer’s instructions.
The DNA fragments of TDH3 were isolated from the TOPO TA Cloning vector by
performing a double digestion with SphI and EcoRI, and then the isolated TDH3 fragment
was ligated into the SphI and EcoRI sites of pSecreted. The final plasmid was named
pSecreted (TDH3).
3.5.5 Construction of Plasmid pSecreted-C95A
Yeast episomal plasmid pSecreted-C95A was obtained by performing a double
61
nucleotide mutation in the barley -amylase cDNA in plasmid pSecreted. The double
mutation changed the codon for cysteine 95 (TGC) to a codon for alanine (GCC).
This
mutation was performed by site-directed mutagenesis using Primers 13 and 14 as described
in Section 3.5.1. The sequence of pSecreted-C95A was confirmed by DNA sequencing.
3.5.6 Construction of Integration Plasmids pIR-1 and pIR-2
Yeast ribosomal DNA (rDNA) integration plasmid pIR-1 was constructed by
combining plasmid pTEF1/Bsd; the fusion gene containing the ADH1 promoter, barley
-amylase cDNA and the CYC1 transcription termination sequence; and part of the
nontranscribed-spacer region (NTS) of rDNA (~980 bp) (Valenzuela et al., 1977).
The NTS DNA fragment was amplified from NRRL Y-132 genomic DNA by PCR.
The PCR reaction was set up using Primers 15 and 16 under the same conditions as the one
described in section 3.5.3.
The amplified PCR products were first ligated into the TOPO
TA Cloning vector (Invitrogen, TOPO TA Cloning Kit (pCR2.1 TOPO)) and then the
ligation reaction was directly subjected to E. coli transformation according to the
manufacturer’s instructions. The NTS DNA fragments were isolated from the TOPO TA
Cloning vector by performing a double digestion with EcoRI and PstI, then the isolated NTS
fragment was ligated into the EcoRI and PstI sites of pTEF1/Bsd.
The fusion gene, which
was directly isolated from pSecreted-C95A by performing a double digestion with SphI and
PstI, was ligated between the SphI and PstI sites of the plasmid composed of pTEF1/Bsd
and the previously inserted NTS region of rDNA. The final plasmid was named pIR-1.
The plasmid was linearized by HpaI digestion before it was used to transform yeast.
DNA fragments NTS-1 (~720 bp) and NTS-2 (~410 bp) were amplified by PCR
from yeast genomic DNA using Primers 17 and 18, and Primers 19 and 20, respectively.
The PCR reaction was carried out under the same conditions as the one described in Section
3.5.3. The amplified PCR products NTS-1 and NTS-2 were first ligated into the TOPO TA
Cloning vector, and then the ligation reaction was directly used for E. coli transformation
according to the manufacturer’s instructions. The NTS-1 DNA fragment was isolated from
the TOPO TA Cloning vector by performing a double digestion with NotI and NheI, and
then the isolated NTS-1 DNA fragment was ligated into plasmid pTEF1/Bsd into the same
62
sites.
The NTS-2 DNA fragment was similarly isolated from the TOPO vector by
performing a double digestion with SphI and HindIII, and then the isolated NTS-2 DNA
fragment was ligated into the same sites of the plasmid composed of pTEF1/Bsd and the
NTS-1 DNA fragment.
Then the fusion gene, which was directly isolated from
pSecreted-C95A by performing a SphI/PstI double digestion, was ligated into the same sites
of the plasmid composed of pTEF1/Bsd, NTS-1 and NTS-2 of rDNA.
The final plasmid
was named pIR-2. The plasmid was linearized with HindIII before being used to transform
yeast.
3.5.7 Construction of Integration Plasmids pdIRs
Integration plasmids with deficient TEF1 promoters (pdIR-1, 2, 3, 4, 5, and 6) were
constructed by replacing the original TEF1 promoter in pIR-1 with sequentially truncated
TEF1 promoters. Six sequentially truncated promoters of TEF1 were generated by PCR
using six different forward primers: Primer dTEF1-1, dTEF1-2, dTEF1-3, dTEF1-4,
dTEF1-5 and dTEF1-6 in conjunction with one shared reverse primer, Primer dTEF1-R.
The PCR reaction was carried out under the same conditions as the one described in Section
3.5.3.
The amplified primers were subjected to TOPO TA Cloning, respectively, as
described in the previous section and then isolated from the TOPO TA Cloning vector by
performing a XhoI and EcoRI double digestion. The isolated truncated promoters of TEF1
were ligated into the XhoI and EcoRI sites of pIR-1 to replace the original TEF1 promoter.
The final plasmids were named pdIR-1, pdIR-2, pdIR-3, pdIR-4, pdIR-5 and pdIR-6. The
plasmids were linearized with HpaI before being used to transform yeast.
3.6
Preparation of Competent Yeast and Yeast Transformation
Preparation of competent yeast and yeast transformation were performed according
to the lithium transformation method from Invitrogen (catalog #V510-20, version F
5-20-2003).
63
3.7
-Amylase Activity Assays
3.7.1 Detection of -Amylase Activity on Starch-Containing YPD Plates
Transformed yeast were incubated on YPD agar plates containing 2% (w/v) soluble
potato starch (Sigma-Aldrich, catalog #S4251) at 30C for 2-3 days.
In order to prepare
these plates, soluble potato starch powder was first added into the YPD-agar media and then
autoclaved. The starch containing YPD-agar solution was cooled to 55°C and poured into
plates with the appropriate amount of blasticidin (final concentration of 100 g mL-1). The
agar plates were solidified at room temperature and stored at 4°C. Transformed yeast cells
were plated on starch-containing YPD plates at a dilution that gave colonies that were
well-separated from each other after 3 days of incubation at 30C.
-Amylase activity was directly visualized by halo formation around the yeast
colonies on starch-containing YPD plates after iodine vapour staining.
The staining
procedure was performed using the method described by Rothstein et al. (1984).
Iodine
crystals, which vaporize at room temperature, were placed into a beaker in a fume hood,
with the circumference of the beaker matching that of the plate. The plate was inverted
onto the beaker, exposing the colonies to the vaporized iodine. Staining was considered
complete when the medium on the plate turned a dark purple colour.
Pictures of the
stained plates were then taken with a digital camera.
3.7.2 Quantification of Secreted -Amylase Activity in Liquid Medium
The starch solution (1% w/v) used as the substrate for the -amylase reaction was
prepared by adding 1 g of soluble potato starch powder to 100 mL water and then boiled for
15 min with constant stirring.
After the solution was cooled to room temperature, water
was added to bring the solution up to 100 mL.
The solution was stable for 3 days under
room temperature with constant stirring.
A yeast overnight culture was used to inoculate 100 mL of fresh sterile YP medium
(1% w/v yeast extract and 2% w/v peptone) containing 0.5% w/v of glucose to give an
initial OD600 = 0.1. When recombinant yeast strains were used, blasticidin was added to
the overnight culture media to a final concentration of 100 g mL-1. The batch cultures
64
were carried out in 250 mL shake flasks in an incubator at 30C with an agitation rate of 180
rpm. A small amount of culture was transferred into 1.5 mL microcentrifuge tubes at
different time points and stored at 4 C. The supernatant of the aliquoted culture samples
was then measured for -amylase activity assay within 48 h.
The -amylase activity assay was performed by incubating 1 mL of culture
supernatant with 1 mL of starch solution at 45 C for 20 min.
The reaction was stopped
by aliquoting 50 µL of the reaction solution into 2 mL of iodine color reagent solution (see
Reagent section) at room temperature.
The -amylase activity was measured as the
absorbance change at 580 nm (A mg biomass-1). Proper dilutions were made to ensure
that the measured absorbance was in the linear range of a previously constructed standard
curve (the measured absorbance is linear from 0 to 3.2 at 580 nm, R2=0.9946).
With slight modifications, the same method was used to screen the -amylase
activity generated by the recombinant yeast clones that had the barley -amylase cDNA
integrated into the yeast genome. The modifications were as follows: an overnight culture
was used to inoculate 10 mL of YPD medium to give an initial OD600 = 0.1. The cultures
were grown to an OD600 ≥ 11 and stored at 4C.
-amylase activity within 48 h.
The culture supernatant was assayed for
The -amylase activity was measured as the absorbance
change at 580 nm (A mg biomass-1).
3.8
Batch Fermentation
All fermentations were carried out in 250 mL shake flasks (with sponge stoppers at
the openings) at 30C with the shaking rate at 180 rpm. Frozen yeast cell cultures were
used to inoculate 10 mL of YPD medium and allowed to grow overnight to give an OD600 =
7-8. When recombinant yeast strains were used, blasticidin was added to the overnight
culture to a final concentration of 100 g mL-1. The appropriate amount of overnight
culture was used to inoculate fresh sterile 100 mL YPS medium to give an initial OD600 =
0.1.
65
3.8.1 Preparation of Frozen Cultures
Frozen yeast cultures were prepared by picking a single colony with a sterilized
needle, which was used to inoculate 10 mL of YPD medium. The culture was incubated
with shaking at 180 rpm and 30C overnight to give an OD600 = 4-5. Then, 930 L of the
overnight culture was transferred into a 1.5 mL sterilized microcentrifuge tube along with
70 L of DMSO. The tubes were stored at -80C.
3.8.2 Biomass Analysis
A 1 mL sample was collected from the fermentation broth and measured at a
wavelength of 600 nm using a spectrophotometer (SmartSpecTM3000, BIORAD).
Dilutions were made using distilled water when the OD600 exceeded 0.4.
OD600 was
converted to cell dry weight (g L-1) using a previously constructed standard curve (from 0 to
10 g L-1).
3.8.3 Ethanol Analysis
Samples for determining ethanol concentration were collected from the fermentation
broth and filtered through a 0.22 m syringe filter (catalog#144666, Whatman Inc.) to
produce a clear filtrate.
Filtrates were stored in 1.5 mL vials (part#5182-0557, Agilent
Technologies) with screw caps at -20C.
Ethanol concentrations were determined by gas chromatography using a flame
ionization detector and a 30 m, 0.25 mm ID poly (5% phenyl, 95% dimethyl) siloxane
capillary column. The oven temperature was 60C and 1-butanol was used as an internal
standard (Lang et al., 2001). Frozen samples were thawed at room temperature. Samples
were placed on the sample region of the detector in order and injections were automatically
performed by the detector and analyzed. For each sample, three separate injections were
analyzed by the detector.
Standards of ethanol samples (from 0.5 to 3 g L-1) were prepared
in de-ionized water using volumetric flasks.
The theoretical maximum yield of ethanol from 100 parts of glucose is 51.1
(Ingledew, 1993).
For example, 100 g L-1 of glucose theoretically yields 51.1 g L-1 of
ethanol through the fermentation process.
The fermentation efficiency can be calculated
66
as:
% fermentation efficiency = (weight of ethanol produced  100)/(theoretical weight of
ethanol from produced glucose)
3.8.4 Starch Concentration
A 1 mL fermentation sample was collected from the fermentation broth and stored at
-20 C. Samples were thawed at room temperature and boiled in a water bath for 30 min
to re-solubilize the starch. After cooling to room temperature, 300 µL samples were mixed
with 2 mL of iodine color reagent solution and diluted with 9 mL of water. The final
solution was measured at 580 nm using a spectrophotometer (SmartSpecTM3000, BIORAD).
Starch concentrations were determined by using a standard curve (from 0 to 1 g L-1).
Dilutions were made to ensure the readings were within the linear portion of the standard
curve.
3.8.5 Plasmid Stability
For yeast strains transformed with episomal plasmids, cell samples were collected
from the fermentor during batch fermentation, and diluted to the appropriate cell density
with sterilized deionized water. Equal amounts of cell suspension were plated on both
YPD plates and blasticidin-containing YPD plates. For each sample set, three different
dilutions were prepared to ensure that one dilution would give the appropriate cell density
such that the colonies on the plates were separate from each other for accurate counting.
Each dilution was plated in triplicate. Plates were incubated at 30C for 1 to 2 days.
After counting the number of colonies, the ratio of the number of colonies on
blasticidin-containing YPD plates over that on YPD plates was recorded as relative plasmid
stability.
For yeast strains transformed with integration plasmids, diluted cell samples were
directly plated on starch-containing YPD plates.
After incubation at 30C for 1 to 2 days,
the percentage of the number of colonies showing halo formation over the total number of
colonies was recorded as integrated plasmid stability.
67
3.8.6 Fermentable Sugar Analysis
Samples for measuring fermentable sugar (glucose and maltose) concentration were
collected from the fermentation broth and filtered through a 0.22 m syringe filter (catalog#
144666, Whatman Inc.) to produce a clear filtrate.
Filtrates were stored in 1.5 mL vials at
-20C.
Fermentable sugar concentrations were determined by High Performance Liquid
Chromatography (HPLC) (Agilent 1100 series,) using a G1362A Refractive Index Detector
(RID) and a 6.5 mm × 300 mm column (Sugar-pakTM1, Waters). The mobile phase was
water with a flow rate of 0.5 mL min-1. The column temperature was 80C. Frozen
samples were thawed at room temperature. Samples were placed on the sample region of
the detector in order and injections were automatically performed by the detector. Samples
were injected at a volume of 20 µL. Standards for fermentable sugars (from 0.5 to 10 g L-1)
were prepared in de-ionized water.
68
4.0
RESULTS
4.1
Barley -Amylase (BA)-Secreted Shows Higher -Amylase Activity and
Fermentation Ability Compared to BA-Anchored
4.1.1 Structure of Expression Plasmid pSecreted
Previously, cell wall-anchored barley -amylase was shown to have low enzyme
activity, which was speculated to be a result of steric hindrance by the anchoring process
(Bo Liao, Master’s Thesis, University of Saskatchewan, 2008).
Thus, we generated a
recombinant yeast strain that secretes barley -amylase into the medium by transforming
yeast with expression plasmid pSecreted.
pSecreted was constructed by deleting the
-agglutinin domain from pAnchored, which was used previously to express cell
wall-anchored barley -amylase (Fig. 4.1).
The yeast strain used in this study, S. cerevisiae NRRL Y-132, is an industrial,
ethanol-tolerant strain, and no auxotrophic mutants of this strain exist. A dominant marker
gene which conferred resistance to blasticidin, which is an antibiotic that inhibits the growth
of a wide range of prokaryotic and eukaryotic cells by interfering with protein synthesis
(Izumi et al., 1991), was used for selection. Antibiotic sensitivity experiments indicated
that the recombinant yeast could be cultured in media containing 100 µg mL-1 blasticidin
while non-transformed yeast showed no growth at this concentration (Bo Liao, Master’s
Thesis, University of Saskatchewan, 2008).
4.1.2 Amylolytic Activity of BA-Secreted
In order to assess whether functional barley -amylase was expressed in yeast,
NRRL Y-132 yeast were transformed with pSecreted, then plated and cultured on YPD agar
plates containing blasticidin and 2% w/v soluble starch. The plates were then exposed to
iodine vapour.
Iodine forms complexes with starch molecules which produces a dark
purple colour.
Formation of clear halo zones around the yeast colonies is indicative of
amylolytic activity. As shown in Figure 4.2, halos were detected around each yeast colony
harbouring pSecreted (Fig. 4.2A) and pAnchored (Fig. 4.2B), while wild type yeast colonies
69
showed no halo formation (Fig. 4.2C).
It was also noted that yeast colonies harbouring pSecreted generated larger halos
than the one harbouring pAnchored, which is likely due to the ability of the secreted enzyme
to diffuse. The results indicated that pSecreted-transformed yeast cells express functional
barley -amylase.
B
A
AmpR
AmpR
Figure 4.1. Structure of plasmids pSecreted and pAnchored. pSecreted (A) was
constructed by deleting the gene encoding for -agglutinin (AG) from pAnchored (B).
The transcription of the barley -amylase cDNA (AMY1) and the barley -amylase -agglutinin fusion gene were under the control of the constitutive promoter ADH1
(pADH1), and the transcription termination sequence of CYC1 (CYC1). Bsd, blasticidin
resistance gene; pTEF1, constitutive promoter TEF1.
A
C
B
Figure 4.2. Iodine vapour assay for the detection of amylolytic activity by the
recombinant yeast. -Amylase activity was directly visualized by halo formation around
the yeast colonies on starch-containing YPD plates after iodine vapour staining. (A)
Enlarged view of a yeast colony harbouring pSecreted, (B) pAnchored, and (C) wild type
yeast stained with iodine vapour. The size of the colonies ranges from 0.5 to 0.8 cm.
70
4.1.3 Batch Fermentation Studies Using BA-Secreted or BA-Anchored
4.1.3.1 Utilization of Soluble Starch in Batch Fermentation
Strain BA-Anchored was previously studied in a 2.5-L bioreactor (New Brunswick
Scientific model 2.5 L Bioflo 310 fermentor) and showed the ability to hydrolyze soluble
starch under batch fermentation conditions (Bo Liao, Master’s Thesis, University of
Saskatchewan, 2008).
In order to study the ability of BA-Secreted to grow and ferment on
soluble starch as the sole carbon source, small-scale (in 250 mL shake flask) batch
fermentations were performed.
In addition, both BA-Anchored and the wild type yeast
were studied under the same experimental conditions as the BA-Secreted for comparison.
As shown in Figure 4.3, no starch hydrolysis was observed for batch fermentations
performed using the wild type strain on 20 g L-1 soluble starch.
BA-Anchored and
BA-Secreted hydrolyzed starch at rates of 0.088 and 0.24 g L-1h-1, respectively, over the first
24 h (Fig. 4.3). No ethanol production was detected for any of the three strains under these
fermentation conditions (data not shown).
4.1.3.2 Spiking with Glucose Improves the Starch-Hydrolyzing Ability of the Recombinant
Yeast
When soluble starch was used as the sole carbon source, batch fermentations
performed with BA-Secreted showed a 2.7 times higher starch hydrolysis rate than
BA-Anchored (Fig. 4.3).
However, neither recombinant strain was able to hydrolyze
starch to completion during the fermentation experiments, and no ethanol was generated.
It was suspected that the low starch hydrolysis rates by the recombinant yeast strains were
due to insufficient enzyme activity in the fermentor.
71
wild type
BA-Anchored
BA-Secreted
Starch concentration (g L-1)
30
25
20
15
10
5
0
0
10
20
30
40
50
60
70
Time (hours)
Figure 4.3. Starch-hydrolyzing ability of wild type and recombinant yeast.
Fermentations were performed using soluble potato starch (20 g L-1) as the sole carbon
source. Cells were pre-cultured in 10 mL YPD medium overnight and then was used to
inoculate 100 mL fresh YPS to give an initial OD600 = 0.1. Samples were taken at the
indicated time points and assayed for starch concentration. Wild type (triangle),
BA-Anchored (square), and BA-Secreted (circle). The data shown are means ± SD of
three replicate experiments (some of the error bars are too small to be shown on the graph).
Since there was no consumable carbon sources i.e. free sugars, available at the start
of the fermentations, it was likely that there was an insufficient energy supply to allow the
yeast to synthesize adequate levels of the enzyme to provide for effective starch hydrolysis.
In order to increase the expressed enzyme activity, a small amount of glucose (5 g L-1) was
added to the medium at the beginning of the fermentation. Under
this
condition,
BA-Anchored and BA-Secreted showed improved starch hydrolysis rates of 0.38 g L-1h-1 for
BA-Anchored (Fig. 4.4B) and 0.70 g L-1h-1 for BA-Secreted (Fig. 4.4C) at 12 h
post-inoculation. The starch hydrolysis rates for BA-Anchored and BA-Secreted represent
increases of 6.3- and 2.9-fold, respectively, compared to the fermentations without glucose
spiking. Moreover, all three strains generated a small amount of ethanol from the spiked
glucose within the first 24 h of fermentation.
For both the wild type strain (Fig. 4.4A) and
BA-Anchored (Fig. 4.4B), the ethanol generated was gradually consumed over the next 24 h.
However, for BA-Secreted, the ethanol was consumed at a much slower rate and was
depleted after 80 h (Fig. 4.4C). The results demonstrate that by spiking with a small
amount of glucose, improvement in starch hydrolysis rates could be achieved.
72
A
Starch(wild type)
Ethanol (wild type)
5
25
4
20
3
15
2
10
Ethanol (g L-1)
1
5
0
0
0
20
40
60
Time (hours)
80
100
B
C
5
25
4
20
3
15
2
10
1
5
0
0
0
20
40
60
80
100
Starch(BA-Secreted)
Ethanol (BA-Secreted)
30
Starch concentration (g L-1)
B
30
Ethanol (g L-1)
Starch concentration (g L-1)
Starch (BA-Anchored)
Ethanol (BA-Anchored)
120
25
4
20
3
15
2
10
5
1
0
0
0
Time (hours)
5
20
40
60
80
Ethanol (g L-1)
Starch concentration (g L-1)
30
100
Time (hours)
Figure 4.4. The effect of glucose spiking on the starch-hydrolyzing and fermentative
ability of the wild type and recombinant yeast. Cells were pre-cultured in 10 mL YPD
medium overnight and then was used to inoculate 100 mL fresh YPS containing 5 g L-1 of
glucose to give an initial OD600 = 0.1. Samples were taken at the indicated time points and
assayed for starch concentration (circle) and ethanol (square).
Wild type (A),
BA-Anchored (B), and BA-Secreted (C). The data shown are means ± SD of three
replicate experiments (some of the error bars are too small to be shown on the graph).
73
4.1.3.3 The Addition of Exogenous Glucoamylase Improves Ethanol Fermentation
It has been reported that the rate of fermentable sugar release is the rate-limiting
factor for ethanol production during starch fermentations (de Moraes et al., 1995).
While
spiking with a small amount of glucose provided enhanced starch hydrolysis, there was no
significant increase in ethanol production from soluble starch. This suggested that the rate
of starch hydrolysis remained inadequate.
It was reported that barley -amylase initially
converts starch molecules primarily into short oligosaccharides consisting of 5-7 glucose
units (G5-G7), and subsequently converts the oligosaccharides into maltose albeit at a
slower rate (Textor et al., 1998). Our batch fermentation results showed that NRRL Y-132
is capable of directly fermenting on maltose as efficiently as on glucose (data not shown).
Based on these observations, we suspected that most of the oligosaccharides were not
efficiently converted into maltose and remained as G5-G7, which cannot be utilized by the
recombinant yeast during fermentation.
In order to address this issue, a small amount of glucoamylase (5 U L-1) was added
into the batch cultures at the 24 h time point.
Glucoamylase attacks starch and
oligosaccharides from the non-reducing ends to produce exclusively glucose. As shown in
Figure 4.5A for the wild type strain, the addition of exogenous glucoamylase resulted in
mild starch hydrolysis at a rate of 0.11 g L-1h-1 over the first 24 h. However, there was no
increase in ethanol production compared to the fermentation performed without the addition
of glucoamylase (compare Fig. 4.4A with Fig. 4.5A). For the recombinant yeast, there was
no increase in the overall starch hydrolysis rates as expected.
BA-Anchored and
BA-Secreted hydrolyzed starch at rates of 0.39 g L-1h-1 and 0.72 g L-1h-1, which are similar
to the starch hydrolysis rates of 0.38 g L-1h-1 and 0.70 g L-1h-1 from the fermentations
without glucoamylase.
However, the ethanol produced by BA-Anchored and BA-Secreted
increased to 1.3 g L-1 and 3.5 g L-1, respectively (Figs. 4.5B and 4.5C). The results
confirmed that the low ethanol production in the previous fermentation experiments was due
to inefficient fermentable sugar conversion from G5-G7 oligosaccharides.
In addition,
ethanol production by the recombinant yeast can be increased by adding exogenous
glucoamylase, which enhances the rate of fermentable sugar release.
74
A
30
5
25
4
20
3
15
2
10
5
1
0
0
0
20
40
60
80
100
Ethanol (g L-1)
Starch concentration (g L-1)
Starch (wild type)
Ethanol (wild type)
120
Time (hours)
B
C
5
25
4
20
3
15
2
10
5
1
0
0
0
20
40
60
80
30
5
25
4
20
3
15
2
10
5
1
0
0
0
100 120 140 160
Time (hours)
20
40
60
80
100
Ethanol (g L-1)
30
Starch concentration (g L-1)
Starch (BA-Secreted)
Ethanol (BA-Secreted)
Ethanol (g L-1)
Starch concentration (g L-1)
Starch (BA-Anchored)
Ethanol (BA-Anchored)
120
Time (hours)
Figure 4.5. The effect of exogenous glucoamylase on the starch-hydrolyzing and
fermentative ability of the wild type and recombinant yeast. Cells were pre-cultured in
10 mL YPD medium overnight and then was used to inoculate 100 mL fresh YPS containing
5 g L-1 glucose to give an initial OD600 = 0.1. Glucoamylase (5 U L-1) was added at the 24
h point. Samples were taken at the indicated time points and assayed for starch
concentration (circle) and ethanol (square). Wild type (A), BA-Anchored (B), and
BA-Secreted (C). The data shown are means ± SD of three replicate experiments (some of
the error bars are too small to be shown on the graph).
75
Moreover, these experiments demonstrated that the presence of both glucoamylase
and -amylase activity provided more efficient starch conversion to fermentable sugars
compared to when only -amylase was present (compare Figs. 4.5B and 4.5C with 4.4B,
4.4C).
In order to further confirm that ethanol production can be increased by increasing
the fermentable sugar releasing rate, batch fermentations using BA-Secreted were
performed with an increased amount of glucoamylase.
As shown in Figure 4.6, the
addition of 4X the amount of glucoamylase had little effect on the overall starch hydrolysis
rate (compare Fig. 4.5C and Fig. 4.6), however, BA-Secreted was able to produce ethanol
concentration of 7.0 g L-1 from 20 g L-1 of starch in 48 h, with an ethanol production rate of
0.23 g L-1h-1. When 16X the amount of exogenous glucoamylase (80 U L-1) was added at
the 24 h point, ethanol accumulated to 9.4 g L-1 in 36 h, which is 92% of the theoretical
maximum yield (data not shown).
Starch (BA-Secreted)
Ethanol (BA-Secreted)
10
25
8
20
6
15
4
Ethanol (g L-1)
Starch Concentration (g L-1)
30
10
2
5
0
0
0
20
40
60
80
100
Time (hours)
Figure 4.6. The effect of a 4X greater amount of exogenous glucoamylase on the
starch-hydrolyzing and fermentative ability of BA-Secreted. Cells were pre-cultured in
10 mL YPD medium overnight and then was used to inoculate 100 mL fresh YPS containing
5 g L-1 glucose to give an initial OD600 = 0.1. Glucoamylase was added at the 24 h point.
Samples were taken at the indicated time points and assayed for starch concentration (circle)
and ethanol (square). The data shown are means ± SD of three replicate experiments
(some of the error bars are too small to be shown on the graph).
76
It was also examined whether increasing the starch concentration had an effect on
As shown in Figure 4.7, an ethanol concentration of 17 g L-1 with a
the ethanol yield.
production rate of 0.41 g L-1h-1 was obtained when 50 g L-1 starch was used with a
proportional increase in the amount of glucoamylase added (40 U L-1).
The ethanol yield
was 66% of the theoretical maximum yield, which is comparable to 69% obtained when
performed on 20 g L-1 starch (Fig. 4.7).
Starch (BA-Secreted)
Ethanol (BA-Secreted)
18
16
60
14
50
12
40
10
30
8
6
Ethanol (g L-1)
Starch concentration (g L-1)
70
20
4
10
2
0
0
0
20
40
60
80
100
120
140
Time (hours)
Figure 4.7. The effect of starch concentration on the starch-hydrolyzing and
fermentative ability of BA-Secreted. Cells were pre-cultured in 10 mL YPD medium
overnight and then was used to inoculate 100 mL fresh YPS containing 5 g L-1 glucose to
give an initial OD600 = 0.1. Fermentations were performed with 50 g L-1 of soluble starch
with the addition of 40 U L-1 of glucoamylase at the 24 h point. Samples were taken at the
indicated time points and assayed for starch concentration (circle) and ethanol (square).
The data shown are means ± SD of three replicate experiments (some of the error bars are
too small to be shown on the graph).
77
4.1.3.4 Fermentation Studies Using Highly Branched Starch
Since branched starches are encountered in most industrial applications, the
fermentative ability of BA-Secreted on various branched starches was evaluated. Batch
fermentations were performed on 20 g L-1 of corn starch (73% amylopectin and 27%
amylose) and waxy corn starch (>98% amylopectin), respectively, with the same amount of
exogenous glucoamylase added (20 U L-1) at the 24 h point. As shown in Figure 4.8,
ethanol concentrations of 6.5 and 6.4 g L-1 were obtained from corn starch and waxy corn
starch, respectively, with the same ethanol production rate of 0.18 g L-1h-1 for both.
The
ethanol yields were about 63-64% of the theoretical maximum yield, which are comparable
to the 69% obtained when performed on non-branched potato starch (≥99% amylose).
Starch hydrolysis rates were not determined due to technique difficulties in measuring the
highly-branched starch concentration.
However, similar starch hydrolysis rates should be
expected since comparable ethanol yields were obtained from both non-branched and
branched starch under the same fermentation conditions.
The results demonstrated that
BA-Secreted is able to ferment on branched starches as efficiently as on high-amylose
starch.
Corn Starch (BA-Secreted)
10
Ethanol (g L-1)
8
6
4
2
0
0
20
40
60
80
100
120
Time (hours)
Figure 4.8. The fermentative ability of BA-Secreted on highly branched starch.
Cells were pre-cultured in 10 mL YPD media overnight and then was used to inoculate 100
mL fresh YPS containing 5 g L-1 glucose to give an initial OD600 = 0.1. Glucoamylase (20
U L-1) was added at the 24 h point. Samples were taken at the indicated time points and
assayed for ethanol (square: Corn starch; circle: Waxy corn starch). The data shown are
means ± SD of three replicate experiments (some of the error bars are too small to be shown
on the graph).
78
4.2
Attempts to Improve Cell Wall-Anchored -Amylase Activity
The previous results showed that BA-Anchored is less effective in performing starch
hydrolysis during batch fermentation than BA-Secreted. One possibility for the low starch
hydrolysis rate was a hindering effect caused by the anchoring process on the C-terminal
domain of barley -amylase.
The C-terminal domain of barley -amylase was suggested to be involved in starch
binding (Sogaard et al., 1991).
Thus, fusing this domain to the anchoring protein
-agglutinin may reduce the ability of the enzyme to bind starch molecules, a result either
of the fusion process itself and/or due to reduced access of starch to the binding site as a
result of the enzyme being in close proximity to the cell wall.
Based on these concerns, two strategies were considered that could potentially
improve the anchored barley -amylase activity.
One was to employ a different anchoring
protein, which would anchor barley -amylase through its N-terminal domain. Another
was to employ a flexible peptide linker, which was inserted between the C-terminal domain
of barley -amylase and the anchoring protein which would extend the enzyme away from
the cell wall.
4.2.1 Replacing -Agglutinin with Flo1p as the Anchoring Protein
4.2.1.1 Structure of the Yeast Expression Plasmid pFLO1
Flocculation protein 1 (Flo1p) is a cell surface protein, which is involved in the
flocculation ability of certain yeast strains (Matsunoto et al., 2002). Structural studies
have shown that Flo1p has two distinct domains that have cell wall-anchoring properties:
the GPI anchoring domain at its C-terminus and mannoprotein binding domain at its
N-terminus.
It has been reported that the N-terminal mannoprotein binding domain binds
non-covalently to mannoproteins on the outer layer of the yeast cell wall (Matsumoto et al.,
2002; Shigechi et al., 2004). By deleting the C-terminal GPI anchoring domain of Flo1p,
its N-terminal mannoprotein binding domain can be used to anchor barley -amylase on the
cell wall.
In this way, barley -amylase is anchored through its N-terminus, with its
C-terminal starch-binding domain presumably having free access to the substrates. The
79
yeast episomal plasmid pFLO1 was constructed from pAnchored by replacing barley
-amylase/-agglutinin coding region with the FLO1/ barley -amylase coding region.
A
B
R
Amp
Figure 4.9. Structure of Flo1p and plasmid pFLO1. (A) Flo1p consists of four
functional domains, and the domains employed for anchoring barley -amylase are
indicated. (B) pFLO1 was constructed by inserting the gene coding for the secretion signal
and the mannoprotein binding domain (FLO1) of Flo1p into plasmid pSecreted. FLO1
was fused to the 5’ end of barley -amylase cDNA (AMY1) and in-frame with it.
80
4.2.1.2 Batch Fermentation Studies Using BA-FLO1
To study the fermentative ability of BA-FLO1, batch fermentations using 20 g L-1 of
soluble potato starch as the sole carbon source were performed. Starch hydrolysis rate was
measured and compared with fermentations performed using BA-Anchored. As shown in
Figure 4.9, BA-FLO1 showed starch hydrolysis rates of 0.081 g L-1h-1, which is comparable
to the rate of 0.088 g L-1h-1 by BA-Anchored (compare Figs. 4.3 and 4.10). The finding
suggests that the orientation of the cell wall-anchored barley -amylase is not the major
reason for the low activity.
Starch concentration (g L-1)
30
25
20
15
10
5
0
0
10
20
30
40
50
60
70
Time (hours)
Figure 4.10. The starch-hydrolyzing ability of BA-FLO1. Cells were pre-cultured in
10 mL YPD medium overnight and then was used to inoculate 100 mL fresh YPS to give an
initial OD600 = 0.1. Samples were taken at the indicated time points and assayed for starch
concentration. The data shown are means ± SD of three replicate experiments (some of the
error bars are too small to be shown on the graph).
4.2.2 Use of A Flexible Linker to Improve Cell Wall-Anchored -Amylase Activity
4.2.2.1 Structure of the Yeast Expression Plasmid pLinker
The studies above indicate that a mechanism other than enzyme orientation relative
to the cell wall is contributing to the low activity.
Since with either -agglutinin or Flo1p
anchoring, the -amylase remains in close proximity to the anchoring protein, which causes
81
steric hindrance between the two proteins.
We next examined whether extending the
enzyme away from the anchoring protein would improve the enzyme activity.
Short
peptides composed of repetitive glycine and serine residues can serve as a flexible peptide
linker between the anchored enzyme and the anchoring protein, resulting in increased
enzyme activity (Robinson and Sauer, 1998; Washida et al., 2001). The yeast expression
plasmid pLinker was modified from pAnchored by inserting a 57 nucleotide long DNA
fragment between the barley -amylase and -agglutinin cDNAs (Fig. 4.11). The inserted
DNA fragment encoded for the linker composed of repetitive glycine and serine residues
(GSSGGSGGSGGSGGSGS).
The starch plate assay showed that pLinker-transformed
recombinant yeast were able to generate halos on iodine-stained starch agar plates, which
indicated that functional -amylase was expressed (data not shown).
Amp
R
Figure 4.11. Structure of plasmid pLinker. A double-stranded 57 nucleotide long
DNA fragment (underlined) was inserted at the XhoI site (italicized) between the barley
-amylase cDNA (AMY1) and -agglutinin cDNA (AG). The linker (not including the
XhoI restriction sites) encodes for a peptide GSSGGSGGSGGSGGSGS (G, Glycine; S,
Serine).
82
4.2.2.2 Batch Fermentation Studies Using BA-Linker
To study the fermentative ability of BA-Linker, batch fermentations using 20 g L-1 of
soluble potato starch were performed.
Starch hydrolysis and ethanol production were
measured and compared with fermentations performed using BA-Anchored. As shown in
Figure 4.12, without the addition of exogenous glucoamylase, BA-Linker showed starch
hydrolysis rates of 0.52 g L-1h-1 (Fig. 4.12A).
A small amount of ethanol was generated
from the spiked glucose within the first 24 h of fermentation, and was gradually consumed
to depletion at 60 h (Fig. 4.12B). The addition of exogenous glucoamylase (5 U L-1) had
little effect on the overall starch hydrolysis rate (0.50 g L-1h-1), however, it did produce a
second ethanol peak (1.8 g L-1 at 100 h), which was slowly consumed to depletion by 140 h
(Fig. 4.12B).
BA-Linker showed a 1.3 times increase in the starch hydrolysis rate as well
as a 1.4X increase in the ethanol production compared to BA-Anchored (Table 4.1). The
results show that BA-Linker was able to hydrolyze starch at a higher rate and accumulate
more ethanol than BA-Anchored.
However, this strain remained relatively inefficient with
respect to ethanol production, achieving only 18% of the theoretical yield.
A
B
Without glucoamylase
With glucoamylase
35
2.5
30
Ethanol (g L-1)
Starch concentration (g L-1)
Without glucoamylase
With glucoamylase
25
20
15
10
5
2
1.5
1
0.5
0
0
0
20
40
60
80
0
100 120 140 160
Time (hours)
20 40 60 80 100 120 140 160
Time (hours)
Figure 4.12. The starch-hydrolyzing and fermentative ability of BA-Linker. Cells
were pre-cultured in 10 mL YPD medium overnight and then was used to inoculate 100 mL
fresh YPS containing 5 g L-1 of glucose to give an initial OD600 = 0.1. Glucoamylase (5 U
L-1) was added at the 24 h point. Samples were taken at the indicated time points and
assayed for (A) starch concentration and (B) ethanol concentration. The data shown are
means ± SD of three replicate experiments (some of the error bars are too small to be shown
on the graph).
83
Table 4.1 Comparison of the ethanol concentration and starch hydrolysis rate
between BA-Anchored and BA-Linker
Batch fermentation experiments were performed on 20 g L-1 soluble potato starch.
Glucoamylase (5 U L-1) was added into the fermentation broth at the 24 h point.
Ethanol Concentration (g L-1)
Yeast strains
-Glucoamylase
+Glucoamylase
(at 100 h)
Starch hydrolysis rate (g L-1h-1)
-Glucoamylase
+Glucoamylase
BA-Anchored
ND
1.3 (0.70)
0.38 (0.036)
0.39 (0.048)
BA-Linker
ND
1.8 (0.37)
0.52 (0.049)
0.50 (0.049)
ND: not detectable
4.3
Attempts to Improve Secreted -Amylase Activity
The results from batch fermentation studies reported earlier in this thesis showed
that the most efficient strain for generating starch hydrolysis activity was BA-Secreted.
However, the amylolytic activity generated by this strain was still not sufficient to hydrolyze
raw starch (data not shown).
It has been reported that the amount of enzyme activity
required for raw starch hydrolysis is about 200-1000 times more than the amount required to
hydrolyze the same concentration of soluble starch (Robertson et al., 2006).
This
suggested that the amount of secreted barley -amylase by BA-Secreted was not adequate
to initiate raw starch hydrolysis.
In order to further increase the secreted amylolytic
activity, we employed two different approaches. The first approach was to generate a
mutant of barley -amylase that had greater intrinsic starch hydrolysis activity under
fermentation conditions.
The second approach was to increase the expression level of
barley -amylase by employing a different constitutive promoter.
4.3.1 Mutating of Cysteine 95 to Alanine in Barley -Amylase
4.3.1.1 Structure of Expression Plasmid pSecreted-C95A
It was previously reported that barley -amylase can be glutathionylated when the
84
enzyme is expressed in S. cerevisiae, resulting in a decrease in the secreted enzyme activity
(Sogaard et al., 1993).
The study also showed that mutating cysteine 95 to alanine
eliminated the glutathionylated forms of secreted barley -amylase and improved the
enzyme activity.
Based on this study, a single mutation (cysteine 95 to alanine) was
introduced into plasmid pSecreted to generate pSecreted-C95A.
4.3.1.2 Batch Fermentation Studies Using BA-Secreted-C95A
Batch fermentations were performed using BA-Secreted-C95A on YPS medium.
Starch hydrolysis and ethanol production were analyzed and compared with those
performed under the same batch fermentation conditions using BA-Secreted. As shown in
Figure 4.13, starch was hydrolyzed to completion within 24 h and a second ethanol peak of
2.2 g L-1 (the first ethanol peak within 24 h was generated from the spiked glucose) was
observed at 36 h. Batch fermentations performed using BA-Secreted-C95A showed 1.6
times increase in starch hydrolysis rate as well as 1.8 times increase in ethanol prodcution
compared to BA-Secreted (Table 4.2). Since glucoamylase was not added during the
fermentations,
a higher ethanol production indicates a higher conversion rate from
oligosaccharides into maltose.
A higher starch hydrolysis rate could be a result of
improved enzyme activity, higher secretion efficiency, or both (Sogaard et al. 1993). The
results show that BA-Secreted-C95A out-performs BA-Secreted with respect to the starch
conversion rate.
85
Starch (BD-SecretedC95A)
Ethanol (BD-SecretedC95A)
5
25
4
20
3
15
2
10
Ethanol (g L-1)
Starch concentration (g L-1)
30
1
5
0
0
0
20
40
60
Time (hours)
80
100
120
Figure 4.13. The starch-hydrolyzing and fermentative ability of BA-Secreted-C95A.
Cells were pre-cultured in 10 mL YPD media overnight and then was used to inoculate 100
mL fresh YPS containing 5 g L-1 of glucose and 20 g L-1 soluble starch to give an initial
OD600 = 0.1. Samples were taken at the indicated time points and assayed for starch
concentration (square) and ethanol (triangle). The data shown are means ± SD of three
replicate experiments (some of the error bars are too small to be shown on the graph).
Table 4.2
Comparison of the ethanol concentration and starch hydrolysis rate
between BA-Secreted and BA-Secreted-C95A
Batch fermentation experiments were performed on 20 g L-1 soluble potato starch.
Ethanol concentration (g L-1)
(at 36 h)
Starch hydrolysis rate
Yeast strains
BA-Secreted
1.2 (0.23)
0.70 (0.068)
BA-Secreted-C95A
2.2 (0.031)
1.1 (0.28)
(g L-1h-1)
4.3.2 Replacing the ADH1 Promoter with TDH3
In the plasmid pSecreted, the transcription of the barley -amylase gene is under the
control of a truncated ADH1 promoter (~400 bp), which is considered to be constitutive
(Ruohonen et al., 1995). Another commonly used constitutive yeast promoter is TDH3
(triosephosphate dehydrogenase isozyme 3).
Both promoters have been shown to provide
robust transcription activity for heterologous protein expression in yeast, although there are
86
no reported studies in the literature that have directly compared their transcriptional
efficiencies. Therefore, we decided to compare both promoters in our episomal plasmid
constructs to determine which one gave higher -amylase expression.
Figure 4.14 shows the starch hydrolysis and ethanol production during batch
fermentations performed on 20 g L-1 soluble starch using BA-Secreted (TDH3). The starch
hydrolysis rate of 0.68 g L-1h-1 (Fig. 4.14A) was comparable to the rate of 0.70 g L-1h-1
generated by BA-Secreted.
A small amount of ethanol from the spiked glucose was
produced within the first 24 h of fermentation, and the ethanol was gradually consumed to
depletion at 80 h (Fig. 4.14B). With the addition of exogenous glucoamylase, another
distinct ethanol peak (2.1 g L-1) was observed at 48 h point, and the generated ethanol was
slowly consumed to depletion at 140 h (Fig. 4.14B).
Without exogenous glucoamylase, BA-Secreted (TDH3) showed comparable
starch-hydrolyzing and fermentative ability as BA-Secreted (Table 4.3).
With the addition
of glucoamylase, both recombinant yeast generated ethanol at comparable levels for the first
48 h, however, BA-Secreted was able to continue ethanol accumulation until 100 h while
the ethanol production by BA-Secreted (TDH3) leveled off after 48 h (compare Figs. 4.4C
and 4.14B). The ethanol generated by BA-Secreted was twice the amount generated by
BA-Secreted (TDH3) at 100 h (Table 4.3). The results showed that use of the ADH1
promoter provides for superior ethanol production from soluble starch relative to the TDH3
promoter, even though the difference in starch hydrolysis rates is modest.
Ethanol production is determined by the fermentable sugar releasing rate, which is
limited by the amount of glucoamylase activity and the available non-reducing ends of
starch at a given time point. Given that the same amount of glucoamylase was used in the
batch fermentations for both recombinant strains, the amount of non-reducing ends
generated by -amylase activity determines the fermentable sugar releasing rate. Although
both recombinant strains hydrolyzed starch to completion in 36 h (compare Figs. 4.5C and
4.14A), the difference in the conversion of long oligosaccharides further into fermentable
sugars after the 36 h point cannot be determined by examining the starch hydrolysis rate.
Thus, we performed an assay to measure whether there is a difference in the secreted
-amylase activity after 36 h between the two recombinant strains.
87
B
Without glucoamylase
With glucoamylase
30
Without glucoamylase
With glucoamylase
2.5
25
2
20
Ethanol (g L-1)
Starch concentration (g L-1)
A
15
10
5
0
1.5
1
0.5
0
0
20
40
60
80
100
0
Time (hours)
40
80
120
160
Time (hours)
Figure 4.14. The starch-hydrolyzing and fermentative ability of BA-Secreted (TDH3).
Cells were pre-cultured in 10 mL YPD medium overnight and then was used to inoculate
100 mL fresh YPS containing 5 g L-1 of glucose to give an initial OD600 = 0.1.
Glucoamylase (5 UL-1) was added at the 24 h point. Samples were taken at the indicated
time points and assayed for (A) starch concentration and (B) ethanol. The data shown are
means ± SD of three replicate experiments (some of the error bars are too small to be shown
on the graph).
Table 4.3
Comparison of ethanol concentrations and starch hydrolysis rates of
BA-Secreted and BA-Secreted (TDH3)
Batch fermentation experiments were performed on 20 g L-1 soluble potato starch and
exogenous glucoamylase (5 U L-1) added into the fermentation broth at the 24 h point.
Yeast
Ethanol concentration (g L-1)
Starch hydrolysis rate (g L-1h-1)
strains
-Glucoamylase
+Glucoamylase
1.2 (±0.23)
2.6 (±0.019) (at 48 h)
(at 36 h)
3.4 (±0.073) (at 100 h)
1.3 (±0.070)
2.1 (±0.10) (at 48 h)
(at 36 h)
1.7 (±0.37) (at 100 h)
pSecreted
pSecreted
(TDH3)
88
-Glucoamylase +Glucoamylase
0.70 (0.068)
0.72 (0.13)
0.68 (0.14)
0.66 (0.17)
Batch fermentations were performed on YP medium containing 5 g L-1 of glucose.
Culture supernatants were collected and assayed for amylolytic activity.
Figure 4.15A
shows that the secreted -amylase activity by BA-Secreted reached 0.056 (A mg biomass-1)
in 48 h, and gradually dropped to zero at 130 h. For BA-Secreted (TDH3), the secreted
-amylase activity reached 0.036 (A mg biomass-1) at 36 h, and gradually dropped to zero
at 100 h. The levels of the secreted -amylase activity by both recombinant strains were
comparable until around 28 h, then the recombinant strain with promoter ADH1 surpassed
the one with promoter TDH3 with respect to the levels of secreted -amylase activity (Fig.
4.15A), resulting in total of 1.6 times higher amounts of -amylase activity generated by
BA-Secreted compared to BA-Secreted (TDH3).
The results confirmed that the two
recombinant strains generated different levels of secreted -amylase activity. The higher
-amylase activity secreted by the recombinant strain with promoter ADH1 presumably
provided a higher conversion of long oligosaccharides into fermentable sugars.
Both recombinant strains showed comparable biomass production during the batch
fermentation experiments (Fig. 4.15B).
A small ethanol peak was observed for both strains
at 12 h and was gradually consumed to zero at 60 h (data not shown). The differences in
the levels of secreted -amylase activity is likely to be a result of growth stage shift such as
entering diauxic growth stage.
Diauxic growth stage is when yeast growth is shifting from
one carbon source to another after the more favourable carbon source in the media was
completely consumed, in this case, from glucose to ethanol.
The results show that
although both promoters are generally considered to be constitutive, the strength of the
promoters can be regulated during certain growth stage shifts (from glucose to ethanol
consumption stage).
89
A
BA-Secreted
BA-Secreted(TDH3)
Amylase Activity (A mg biomass-1)
0.07
0.06
0.05
0.04
0.03
0.02
0.01
0
0
20
40
60
80
100
120
140
Time (hours)
B
BA-Secreted
BA-Secreted(TDH3)
3
Biomass (g L-1)
2.5
2
1.5
1
0.5
0
0
20
40
60
80
100
120
Time (hours)
Figure 4.15. -Amylase activity and biomass production of BA-Secreted and
BA-Secreted (TDH3). Cells were pre-cultured in 10 mL YPD medium overnight and then
was used to inoculate 100 mL fresh YP media containing 5 g L-1 of glucose to give an initial
OD600 = 0.1. (A) Samples were taken at the indicated time points and assayed for
-amylase activity. (B) Biomass was measured by optical density (OD600) and converted
into gram per litre using a previously constructed standard curve. The data shown are
means ± SD of three replicate experiments (some of the error bars are too small to be shown
on the graph).
90
4.4
Genome Integration of the Barley -Amylase gene
4.4.1 Structure of Integration Plasmids pIR-1 and pIR-2
As stated in the previous section, to increase secreted amylolytic activity in the batch
culture, the second approach was to increase the expression level of barley -amylase.
An
effective way to achieve this is to increase the copy number of the barley -amylase gene in
yeast. All of the previous yeast expression plasmids described in this thesis were episomal
plasmids, which exist in 50-200 copies in transformed yeast and replicate independently
from the yeast chromosome.
However, plasmid stability data showed that in batch
fermentation experiments, on average only 20% of the yeast population in the batch cultures
maintain detectable -amylase expression (data not shown).
The loss of plasmid
expression from the transformed yeast cells was considered to be due to segregational
instability.
Higher plasmid stability can be achieved by maintaining high selection
pressure in the fermentor, however, this requires the addition of more antibiotic, which is
not economical in an industrial setting.
To cope with the drawbacks of using episomal plasmid systems, as well as to
increase the gene copy number of barley -amylase, we designed an integration plasmid to
insert the barley -amylase gene into the yeast genome at multiple sites. Genes that are
integrated into the yeast genome can be stably maintained in yeast without any requirements
for antibiotic selection pressure, however, generally only one copy of the gene can be
introduced into a targeted locus in the yeast genome (Romanos et al., 1992). Previous
studies showed that integration of 150-200 copies of a given gene can be achieved by
targeting the ribosomal DNA loci in the yeast genome, which number 150-200 in haploid
yeast, and therefore many more loci in NRRL Y-132 which is polyploidy (Lopes et al., 1989,
1991, Lin et al., 1998, Nieto et al., 1999). Moreover, only a small portion of ribosomal
DNA are actively transcribed. Thus, integration of foreign genes into ribosomal DNA
locus should not significantly affect the expression of ribosomal RNA in yeast (Yokoyama
and Suzuki, 2008).
Both integration plasmids pIR-1 and pIR-2 contain the blasticidin resistance gene
and the barley -amylase gene cassette.
pIR-1 contains a 980 bp DNA fragment from the
91
NTS region in rDNA (Fig. 4.16), which is used to insert the plasmid into the corresponding
location in the yeast genome by homologous recombination.
NTS region is a
non-transcribed spacer region between rDNA repeats (Valenzuela et al., 1977).
Since
linearized integration plasmids show higher integration efficiency than their circular
counterparts (Orr-Weaver et al., 1981).
A unique restriction site HapI in the NTS insert
was used to linearize pIR-1 priori to yeast transformation (Fig. 4.17).
In addition, Ghang
et al. (2007) showed that reducing the size of the integration plasmid could increase the
copy number of the integrated gene cassette, which can be performed by removing the
bacterial components from the plasmid.
Thus, for pIR-2, the cloned NTS region was
inserted as two separate parts (720 and 410 bp) into the plasmid (Fig. 4.16), flanking the
blasticidin resistance gene and the barley -amylase gene cassette (Fig. 4.17).
In this case,
pIR-2 was linearized by restriction digestion by HindIII, resulting in removal of the
bacterial components consisting the ampicillin resistance gene and the replication origin in
E. coli prior to yeast transformation (Fig. 4.17).
Both integration plasmids were
transformed into yeast strain NRRL Y-132, and clones with stable integrated genes were
selected for blasticidin resistance. Moreover, since the recombinant yeast that secretes the
C95A form of barley -amylase showed a higher rate in starch conversion, all of the
integration strains were engineered to express the mutant.
Figure 4.16. Structure of rDNA and the regions used for constructing integration
plasmids pIR-1 and pIR-2. The corresponding regions in NTS (non-transcribed spacer
region) which were cloned and inserted into pIR-1 (NTS, 980 bp) and pIR-2 (NTS-1, 720 bp
and NTS-2, 410 bp) are indicated. The restriction sites used for subcloning and
linearization of the plasmids are also indicated.
92
A
Amp
R
B
AmpR
Figure 4.17. Structure of pIR-1 and pIR-2. The transcription of the barley -amylase
cDNA (AMY1) was under the control of the constitutive promoter ADH1 (pADH1), and the
transcription termination sequence of CYC1 (CYC1). pIR-1 (A) contains the NTS insert,
and a unique restriction site HpaI in the NTS insert was used to linearize pIR-1 prior to
yeast transformation. pIR-2 (B) contains NTS-1 and NTS-2 inserts, and was linearized by
restriction digestion by HindIII, resulting in removal of the ampicillin resistance gene
(AmpR) and pUC origin prior to yeast transformation. The restriction sites used to ligate
the inserts into the plasmids are indicated.
93
4.4.2 Stable Expression of the Integrated Barley -Amylase Gene
The initial selection was based on blasticidin resistance. Selected clones were then
transferred onto YPD plates containing 2% (w/v) soluble starch. After 24 h incubation at
30 C, clones with visible halos were selected and transferred to fresh YPD plates
containing 2% (w/v) soluble starch. This process was repeated for 7 days. The clones
that were still showing halos on starch containing YPD plates were considered stable and
cultured for fermentation experiments.
Before the selected clones were subjected to batch fermentation studies, each clone
was screened for its amylolytic activity.
As shown in Figure 4.18, clones of BA-IR-1 (Fig.
4.18A) and clones of BA-IR-2 (Fig. 4.18B) demonstrated variations in secreted amylolytic
activity.
Clones of BA-IR-1 generally show higher amylolytic activity than clones of
BA-IR-2. For clones of BA-IR-1, 6 out of 9 clones showed amylase activity higher than
0.05 (A mg biomass-1), and two clones showed activity lower than 0.04 (A mg biomass-1).
For clones of BA-IR-2, one of the 10 clones showed amylase activity higher than 0.05 (A
mg biomass-1), and most clones showed activity lower than 0.04 (A mg biomass-1). After
linearization, pIR-2 is approximately 1.5 kb shorter than pIR-1.
The results show that the
removal of bacterial DNA from the region integrated does not improve expression of
-amylase.
Although Ghang et al. (2007) showed that reducing the size of the integration
plasmid increase the copy number of the integrated gene cassette, our results did not agree
with this, suggesting that the size of the integration plasmid was not the only factor that
contributed to the integration efficiency.
94
-Amylase activtiy (A mg biomass-1)
A
0.09
0.08
0.07
0.06
0.05
0.04
0.03
0.02
0.01
0
-Amylase activtiy (A mg biomass-1)
B
0.09
0.08
0.07
0.06
0.05
0.04
0.03
0.02
0.01
0
Figure 4.18. The -amylase activity of the clones of BA-IR-1 and BA-IR-2. Clones
from (A) BA-IR-1 or (B) BA-IR-2 were pre-cultured in 5 mL YPD medium containing 100
µg mL-1 of blasticidin for 24 h and then used to inoculate 10 mL fresh YPD media to give an
initial OD600 = 0.1. At OD600 ≥ 11, 1 mL of culture supernatant were mixed with 1 mL of 1%
(w/v) soluble starch and incubated at 45 C for 20 min. The -amylase activity was
measured as the absorbance change at 580 nm (A mg biomass-1) after the addition of
iodine color reagent. The indicated activity was normalized by the amount of biomass.
The data shown are means ± SD of three replicate experiments.
95
4.4.3 Comparison of the Amylolytic Activity of Yeast Clones that Transformed with
Integration Plamids pIR-1 and pIR-2
To study whether the differences in measured amylase activity among each clone
affected the recombinant yeasts’ ability to hydrolyze soluble starch or produce ethanol under
batch fermentation conditions, 9 different clones of BA-IR-1 were subjected to batch
fermentation study.
As shown in Figure 4.19, biomass (Fig. 4.19A), starch hydrolysis rate (Fig. 4.19B)
and ethanol production (Fig. 4.19C) were measured at the indicated time points for clones
#2, 6, 7, and 8 of BA-IR-1.
Although all of the clones were subjected to batch
fermentation studies, the results showed that clones with -amylase activity higher than
0.05 (A mg biomass-1), which are clones #1, 2, 3, 4, 5, and 9, generated very similar
patterns for biomass, starch hydrolysis rate and ethanol production. Therefore, to reduce
the complexity of the graphs in the figures, only clones #2 (which is representative of clones
#1, 3, 4, 5, and 9), 6, 7, and 8 are presented in Figure 4.19.
Clone #2 of BA-IR-1 is representative of clones with -amylase activity higher than
0.05 (A mg biomass-1), clone #7 is representative of clones with -amylase activities
between 0.03 and 0.05, and clone #6 and 8 are representative of clones with -amylase
activities lower than 0.03 (Figure 4.18A). All clones showed similar biomass production
(Fig. 4.19A), clones #6 and 8 showed lower starch hydrolysis rates compared to other clones
(Fig. 4.19B), while clones # 6, 7 and 8 showed reduced ethanol production compared to
clones #2 (Fig. 4.19C).
Table 4.4 summarizes the ethanol concentrations and starch
hydrolysis rates from batch fermentation studies performed using different clones of
BA-IR-1 and compare them with BA-Secreted and BA-Secreted-C95A.
Compared to the
performance of BA-Secreted and BA-Secreted-C95A, clone #2 of BA-IR-1 showed 2.7 and
1.7 times higher starch hydrolysis rates, and 1.9 and 1.2 times higher concentrations of
ethanol produced, respectively (Table 4.4).
Importantly, no antibiotics were added to the fermentation broth when BA-IR-1 was
used. At the end of the fermentation, yeast cells were diluted with water and plated on
starch containing YPD plates. All of the colonies showed halo formation after 2 days of
incubation at 30 C (data not shown). The results indicate that the integrated genes were
96
still stable and expressing after over 100 h of fermentation.
A
B
Clone#2
Clone#7
Clone#2
Clone#7
Clone#6
Clone#8
Starch concentration (g L-1)
6
Biomass (g L-1)
5
4
3
2
1
0
Clone#6
Clone#8
30
25
20
15
10
5
0
0
20
40
60
80
100
120
0
Time (hours)
20
40
60
80
Time (hours)
C
Clone#2
Clone#7
Clone#6
Clone#8
3
Ethanol (g L-1)
2.5
2
1.5
1
0.5
0
0
20
40
60
80
100
120
Time (hours)
Figure 4.19. The fermentative ability of different clones of BA-IR-1. Cells were
pre-cultured in 10 mL YPD medium overnight and then was used to inoculate 100 mL fresh
YPS containing 5 g L-1 of glucose to give an initial OD600 = 0.1. (A) Biomass was
measured by optical density (OD600) and converted into gram per litre using a previously
constructed standard curve. Samples were taken at the indicated time points and assayed for
(B) starch concentration and (C) ethanol. Each fermentation was only performed once.
97
Table 4.4
Comparison of ethanol concentrations and starch hydrolysis rates of
-amylase secreting yeast strains
Batch fermentation experiments were performed on 20 g L-1 soluble potato starch.
Ethanol concentration (g L-1)
Yeast strains
Starch hydrolysis rate (g L-1h-1)
BA-Secreted
1.3 (0.13) (at 24 h)
0.70 (0.068)
BA-Secreted-C95A
2.0 (0.028) (at 24 h)
1.1 (0.28)
BA-IR-1 Clone #2
2.5 (at 24 h)
1.9
BA-IR-1 Clone #6
1.3 (at 24 h)
1.4
BA-IR-1 Clone #7
2.1 (at 24 h)
1.1
BA-IR-1 Clone #8
1.4 (at 24 h)
1.1
Fermentation was only performed once for BA-IR-1 Clone #2, 6, 7 and 8, thus, SD was not
calculated for these strains.
4.4.4 Batch Fermentation Studies Using BA-IR-1 on Soluble Starch
The previous results showed that ethanol production is positively correlated with the
amount of fermentable sugar present in the fermentation media. To increase fermentable
sugar release rate from starch hydrolysis, it was found that the addition of a small amount of
glucoamylase was very effective and this in turn increased the ethanol production at the end.
However, in the previous studies, glucoamylase was always added at the 24 h point, which
was generally the time point when half of the starch has been hydrolyzed in the fermentation
broth using BA-Secreted.
To study the effect of the timing of the exogenous glucoamylase spiking on ethanol
yield in starch fermentation, batch fermentations were performed on 20 g L-1 soluble starch
using BA-IR-1.
In this study, three parallel batch fermentations were performed under the
same conditions except that, for every fermentation experiment, the glucoamylase was
added at different time points.
As shown in Figure 4.20, biomass (Fig. 4.20A), starch
hydrolysis rate (Fig. 4.20B) and ethanol concentration (Fig. 4.20C) were measured at the
indicated time points for batch fermentations performed using clone #2 of BA-IR-1.
98
A
B
8.5h
12.5h
24.5h
8.5h
Starch concentration (g L-1)
9
Biomass (g L-1)
8
7
6
5
4
3
2
1
0
12.5h
24.5h
30
25
20
15
10
5
0
0
20
40
60
80
100 120 140 160
0
Time (hours)
20
40
60
80 100 120 140 160 180
Time (hours)
C
8.5h
12.5
24.5
4.5
4
Ethanol (g L-1)
3.5
3
2.5
2
1.5
1
0.5
0
0
20
40
60
80 100 120 140 160 180
Time (hours)
Figure 4.20. The effect of the timing of exogenous glucoamylase on the fermentative
ability of the recombinant yeast. Cells were pre-cultured in 10 mL YPD medium
overnight and then was used to inoculate 100 mL fresh YPS containing 5 g L-1 of glucose
and 20 g L-1 soluble starch to give an initial OD600 = 0.1. Glucoamylase (5 U L-1) was
added at 8.5 (circle), 12.5 (square) and 24.5 (triangle) h points. (A) Biomass was
measured by optical density (OD600) and converted into gram per litre using a previously
constructed calibration curve. Samples were taken at the indicated time points and assayed
for (B) starch concentration and (C) ethanol. The data shown are means ± SD of three
replicate experiments (some of the error bars are too small to be shown on the graph).
99
For the three fermentation experiments performed, the same amount of
glucoamylase was added into the fermentation broth at 8.5, 12.5 and 24.5 h points,
respectively. All three batch fermentations showed similar biomass production (Fig 20A)
and starch hydrolysis rates (Fig. 4.20B). However, ethanol productions were different (Fig.
4.20C). When glucoamylase was added into the fermentation broth at the 12.5 h point,
ethanol accumulated up to 4.0 g L-1 at 72 h, and the concentration was 1.2 and 1.3 times
higher than the ones obtained from fermentations that had glucoamylase added at 8.5 and
24.5 h points, respectively.
In addition, HPLC data showed that when glucoamylase was
added at the 12.5 h point, higher amounts of glucose were present in the fermentation broth
compared with the other two time-points (data not shown). The results demonstrated that
adjusting the time point when glucoamylase is added into the fermentation broth is another
factor to consider when attempt to maximize the ethanol production.
4.4.5 Batch Fermentation Studies Using BA-IR-1 on Raw Starch
Cold starch hydrolysis is a method which uses starch that is dissolved at low
temperature (lower than the gelatinization temperature of starch) and raw starch granules
directly for fermentation. This method requires much lower energy input, and has been
commercially used in many industrial plants for ethanol production. However, the amount
of enzymes required for cold starch hydrolysis is high compared to high temperature cooked
starch (Robertson et al., 2006). Thus, a genetically engineered yeast strain that is capable of
raw starch hydrolysis would significantly reduce the cost of bioethanol production.
Since BA-IR-1 showed higher amylolytic activity than BA-Secreted, the latter which
was unable to hydrolyze raw starch (data not shown), it was of interest to study whether
BA-IR-1 was able to ferment on raw starch. Batch fermentations were performed on 20 g
L-1 of raw wheat starch, and 10 mM CaCl2 were added into the fermentor with the yeast to
start the fermentation.
CaCl2 is considered essential for barley -amylase stability,
especially during raw starch hydrolysis (Bush et al., 1989). Parallel fermentations were
performed using the wild type yeast for comparison.
As shown in Figure 4.21, after 150 h of fermentation, only the batch fermentation
containing both the recombinant yeast and glucoamylase was able to produce ethanol and
100
the only one that showed complete hydrolysis of raw starch particles.
In order to confirm
the completion of raw starch hydrolysis in each fermentation, the collected starch samples
were first boiled for 30 min, then stained by iodine solution.
The completion of raw starch
hydrolysis was determined based on the observation of complete disappearance of dark
purple color in the boiled sample. The result showed that the raw starch was completely
hydrolyzed at 133 h (data not shown). However, this method was unable to quantify the
raw starch concentration in the collected samples, since the boiling process did not generate
a homogenous starch solution, and thus no consistent measurements are possible.
The
results showed that the amount of -amylase activity expressed by BA-IR-1 was able to
initiate raw starch hydrolysis, and complete hydrolysis of raw starch can be achieved by
adding a small amount of glucoamylase (5 U L-1).
wild type
BA-IR-1 (clone#2)
wild type + glucoamylase
BA-IR-1 (clone#2) + glucoamylase
2.5
Ethanol (g L-1)
2
1.5
1
0.5
0
0
20
40
60
80
100
120
140
160
180
200
220
Time (hours)
Figure 4.21. The fermentative ability of BA-IR-1 on raw wheat starch. Cells were
pre-cultured in 10 mL YPD medium overnight and then was used to inoculate 100 mL fresh
YP media containing 5 g L-1 of glucose, 20 g L-1 raw wheat starch and 10 mM CaCl2 to give
an initial OD600 = 0.1. Glucoamylase (5 U L-1) was added at the 24 time point. Samples
were taken at the indicated time points and assayed for ethanol. Each fermentation was
only performed once.
101
5.0
DISCUSSION
Development of recombinant yeast strains that can efficiently convert raw starch into
bioethanol requires three key elements: 1) an industrial strain of S. cerevisiae that is
resistant to various stress conditions that are often encountered in industrial fermentation
processes; 2) a starch-hydrolyzing enzyme that not only can perform raw starch hydrolysis,
but do so efficiently at the pH and temperatures that exist in the industrial fermentation
environment; and 3) a genetic carrier that enables stable and high copy numbers of the
expressed gene inside the yeast cells. Selection of each key element requires careful
examination and comparison.
However, due to the diversity of the biological world and
complexity of the problem itself, it is almost impossible to study every aspect to generate a
“best” combination.
Based on the findings of previous studies, our project was an attempt
to offer alternative strategies, broaden our knowledge in this field, and hopefully inspire
others to advance this research field further.
First, we have engineered a polyploid distiller’s yeast strain which is a potent
ethanol-producing strain of S. cerevisiae. Secondly, the selection of which enzyme to
express was based on a study by Textor et al. (1996), in which they compared the activity of
starch-hydrolyzing enzymes commonly used in industrial raw starch processing. Textor et
al. (1996) reported that barley -amylase showed an increase in catalytic activity when the
pH is dropped from 5.5 to 4.5, whereas the -amylase from Bacillus lost all activity. The
other advantage that barley -amylase displayed in this study is that much lower levels of
the enzyme activity is required to perform similar rates of starch granule hydrolysis
compared with its bacterial counterpart.
Thus, barley -amylase is a good candidate to be
expressed by recombinant yeast for raw starch hydrolysis under fermentation conditions.
It has also been reported that barley -amylase 1 can be expressed in S. cerevisiae in its
fully functional form (Sogaard et al., 1993; Wong et al., 2001).
It was our interest to
explore whether a recombinant yeast strain could be engineered that produced barley
-amylase, allowing it to directly utilize raw starch for ethanol production.
The industrial strain NRRL Y-132 was genetically engineered to either secrete barley
-amylase into the medium or express it in a cell surface-anchored form.
Both episomal
plasmids and integration plasmids were used to express the cloned barley -amylase gene.
102
Several major issues relating to the use of recombinant yeast for efficient starch hydrolysis
and fermentation will be discussed in the following sections of the thesis.
5.1
Strategies to Improve the Cell Surface-Anchored Barley -Amylase Activity
Our lab has previously constructed a recombinant yeast strain that expresses cell
surface-anchored barley -amylase (Bo Liao, Master’s thesis, University of Saskatchewan,
2008). The enzyme was expressed as a fusion protein with its C-terminus fused to the cell
wall anchoring domain of -agglutinin. After the enzyme was transported across the cell
membrane, the C-terminus of -agglutinin is covalently attached to the yeast cell wall
structure through its GPI anchoring signal peptide. This recombinant yeast strain was able
to directly convert soluble starch into fermentable sugars in vitro at a pH of 4.5 and
temperature of 45°C.
However, under batch fermentation conditions (pH ~6, 30°C), the
starch hydrolysis rate by the recombinant yeast was insufficient to support ethanol
production.
Enzyme immobilization is a common strategy used in industry.
Immobilization
methods include entrapment and surface anchoring. Considering the fact that enzymes are
expensive, the use of immobilized enzymes has the advantages of being easily purified and
reutilizable.
In addition, immobilized enzymes show increased thermostability and pH
stability (Shuler and Kargi, 2002). However, low activity or complete loss of activity has
been reported when enzymes are immobilized (Shuler and Kargi, 2002).
Many factors
could affect the immobilized enzyme’s ability to interact with their substrates such as: the
orientation of the enzyme’s catalytic site; the nature of the substrate (molecular weight or
solubility); or the microenvironment of the immobilized enzyme, which can result in
reduced diffusion rate of substrates towards the enzyme, altered enzyme structure, or steric
hindrance during the immobilization process.
In our case, barley -amylase was anchored
on the surface of a living yeast cell; however, the immobilization process could affect the
enzyme activity by similar mechanisms. For cell wall anchored barley -amylase, two
major factors were speculated to cause the reduced enzyme activity. Since soluble starch is
a long chain of linked glucose monomers, cell surface anchored barley -amylase has
103
limited accessibility towards such a high-molecular-weight substrate compared to the free
enzyme.
In addition, expressing barley -amylase as a fusion protein with -agglutinin,
the latter having a rod-shaped structure, may have altered the enzyme structure, resulting in
reduced enzyme activity.
Structural studies provided some insights into the second factor that we speculated
was causing reduced enzyme activity through the cell wall anchoring process.
Barley
-amylase is composed of three distinct domains arranged into a barrel-shaped (/)8
structure.
Domain A, which is located at the N-terminus, is the catalytic domain.
Domain B protrudes from domain A and functions as a nearby starch binding site.
It was
recently suggested that the C-terminal domain of barley -amylase, termed domain C, not
only functions as an extra starch binding site but can also be involved in disentang Ling the
-helical structure of the starch molecules and positioning it in the correct orientation for
the catalytic site in domain A (Robert et al., 2005). However, domain C does not maintain
its starch binding ability when it is isolated from the other two domains (Tibbot et al., 2002).
In addition, partial deletions of domain C resulted in low or no measurable enzyme activity
(Tibbot et al., 2002). Tibbot et al. (2002) suggested that different regions of domain C are
not only involved in starch binding, but also interact with domain A and play a critical role
in the enzyme’s overall structural stability.
Thus, it is reasonable to consider that
successful catalysis by the enzyme is achieved by properly coordinated actions between
each domain.
However, the anchoring process may physically limit such cooperation
between domain C and the other domains, resulting in low enzyme activity.
In order to address this issue, a flexible peptide linker consisting of 11 amino acids
(composed of repetitive glycine, G and serine, S residues with a sequence of
GSSGGSGGSGGSGGSGS) was inserted between barley -amylase and -agglutinin.
Peptide linkers rich in glycine residues are more flexible than ones composed of non-glycine
residues with the same length, and insertion of serine residues between the glycine residues
can reduce the instability caused by introducing too many glycine residues in a region
(Robinson and Sauer, 1998).
It was further shown that linkers composed of 11 residues or
more are required for biological activity such as catalytic activity (Robinson and Sauer,
1998). Under batch fermentation conditions, the recombinant yeast that expressed the
104
linker-containing cell wall anchored barley -amylase showed increased starch hydrolysis
rate (0.52 g L-1h-1) compared to that of the recombinant yeast expressing the anchored
without a linker (0.38 g L-1h-1) as well as higher ethanol generation upon glucoamylase
addition (1.8 g L-1 compared to 1.3 g L-1 at 100 h).
These results demonstrated that
insertion of the flexible peptide linker between barley -amylase and -agglutinin had a
positive effect on the anchored enzyme activity towards soluble starch substrate. Similarly,
Washida et al. (2001) reported the use of glycine and serine based peptide linkers of various
lengths to improve cell surface anchored R. oryzae lipase activity.
In conclusion, insertion
of a flexible peptide linker between the anchored enzyme and the anchoring protein could
increase the activity of cell surface anchored barley -amylase.
In order to completely unrestrain the C-terminal domain of barley -amylase, the
enzyme was expressed as a fusion protein with its N-terminus fused to the mannose binding
domain of Flo1p.
In contrast to -agglutinin, the mannose binding domain of Flo1p
non-covalently attaches to the mannose of structural proteins in the cell wall (Matsumoto et
al., 2004). Under the same batch fermentation conditions, the recombinant yeast that
expresses N-terminus anchored barley -amylase (through Flo1p) did not show significant
improvement of the starch hydrolysis rate compared to that of the recombinant yeast
expressing C-terminus anchored barley -amylase using -agglutinin.
domain of barley -amylase is located at its N-terminus.
The catalytic
It is possible that fusing the
catalytic domain to the anchoring protein reduced its accessibility towards the starch
substrate since the catalytic site now faced towards the cell wall instead of towards the
medium.
While in this case the C-terminal starch binding domain has a better chance of
accessing the starch substrates, it seems that the effect is cancelled by having a catalytic
domain directed towards the cell wall.
Other similar studies showed different results.
Using Flo1p as the anchoring
protein, Matsumoto et al. (2004) obtained a recombinant yeast strain that expressed cell wall
anchored R. oryzae lipase with greater enzyme activity compared to a strain that R. oryzae
lipase was anchored with -agglutinin.
In their case, the catalytic domain of the lipase was
located at the C-terminus, so anchoring the enzyme through the N-terminus exposed the
catalytic domain, which presumably was the basis for the increased enzyme activity.
105
Shigechi et al. (2002) found a similar increase in the cell wall anchored B.
stearothermophilus -amylase activity towards raw starch when the anchoring protein
-agglutinin (C-terminal anchoring) was replaced by Flo1p (N-terminal anchoring). Even
though the catalytic domain of B. stearothermophilus -amylase was suggested to be
located at its N-terminus, a free C-terminus, which has strong raw starch binding ability,
seemed to have more impact on the enzyme activity. However, in both studies cited above,
a long spacer domain of the Flo1p was employed to anchor the enzyme.
Similar to the
spacer domain of -agglutinin, the spacer domain of Flo1p is rich in Ser/Thr residues and
has a rod-shaped structure (Watari et al., 1994). Sato et al. (2002) employed various
lengths of Flo1p to anchor R. oryzae glucoamylase, and showed that those with longer
anchors had higher reactivity towards antibodies, suggesting better accessibility towards
substrates.
However, other studies have challenged the notion that longer spacers enhance the
biological activity of anchored proteins.
van der Vaart et al. (1997) anchored
-galactosidase in the yeast cell wall with various cell wall proteins including Cwp1p,
Cwp2p, Ag1, Tip1p, and Flo1p, and found that larger anchoring protein did not necessarily
generate higher enzyme activity. Their results showed that among all of the anchored
-galactosidases, the one anchored with Cwp2p, which is only 67 amino acids long, showed
the highest cell wall anchored -galactosidase activity and was most accessible to large
substrates. Smaller anchoring proteins have the advantage of penetrating through the cell
wall more easily than larger anchoring proteins, which may get trapped (van der Vaart et al.,
1997). Sato et al. (2002) also noted a decrease in protein expression levels when the
length of the anchoring protein was increased.
It seems that larger proteins are more
vulnerable to misfolding and more likely to be retained in the intracellular space.
In conclusion, there appear to be no absolute rule as to which anchoring protein is
the most suitable for anchoring enzymes on the yeast cell wall.
Depending on the
particular enzyme, substrates, and the microenvironment between the enzyme and the
anchoring protein, the choice of which anchors to use may need to be determined
empirically.
Certainly, some general strategies can be applied, such as changing the
orientation of the catalytic domain, introducing flexible peptide linkers, or adjusting the
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length of the anchoring protein’s spacer domain.
Moreover, the use of cell surface
anchored enzymes may not be practical under every circumstance.
In the future, protein
structure studies as well as the application of random mutagenesis should provide more
insights into designing cell wall anchored enzymes with improved activity.
5.2
Comparison between Secreted and Cell Wall Anchored Enzyme Systems for
Starch Fermentation
Since the recombinant yeast strains that expressed cell wall anchored barley
-amylase were not able to generate sufficient activity to perform efficient starch hydrolysis,
we were interested to see how much activity the recombinant yeast was able to achieve
when the enzyme was secreted into the medium.
Since the secreted enzyme is not
physically restrained by the anchoring process, higher activity might be expected.
Indeed, the recombinant yeast strain that secreted barley -amylase out-performed
the anchored strains in all batch fermentation experiments (Figs. 4.3, 4.4 and 4.5).
Depending on the fermentation conditions, the recombinant strain that secreted barley
-amylase showed 1.8 to 2.7 times higher starch hydrolysis rates than the anchored strain
(Figs. 4.3, 4.4B and 4.4C).
In the batch fermentations performed with the addition of
exogenous glucoamylase, the ethanol production by the secreting strain was 2.6 times
higher than that of the anchored strain (Figs. 4.5 B and 4.5C).
A number of other reports have performed similar studies in which amylolytic
enzymes have been expressed in either secreted or cell surface anchored forms in S.
cerevisiae.
Both strategies have provided promising results.
Naturally-occurring
amylolytic bacterial and fungal species secrete starch-hydrolyzing enzymes into the medium,
so most attempts have engineered recombinant S. cerevisiae to secrete enzymes into the
medium.
However, the choice to secrete the enzyme was not based on careful examination
and side-by-side comparisons of the efficacy of both strategies.
As discussed in the previous section, surface-immobilized enzymes can be easily
collected and re-cycled for additional rounds of use, and have shown enhanced
theromostability and pH stability (Shuler and Kargi, 2002). Enzymes immobilized on the
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cell surface should share similar advantages.
It was shown that cell wall anchored
-galactosidase is much more stable than the secreted variant (Schreuder et al., 1996).
The secreted -galactosidase was gradually deactivated after 30 h, however, the cell wall
anchored versions showed stable activity levels for over 50 h.
In addition, cell surface
anchored enzymes can be re-cycled with the yeast cells and re-utilized. Kondo et al. (2002)
demonstrated that by using a recombinant yeast strain expressing cell surface anchored
glucoamylase, they observed no time lag for starch hydrolysis and ethanol production for
repeated batch fermentations.
Moreover, they observed stable starch hydrolysis and
ethanol production rates for seven batch fermentations by repeated utilization of the
recombinant yeast cells.
However, cell surface anchored enzymes can also display reduced activity (as
discussed in the previous section).
Schreuder et al. (1989) showed that cell surface
anchored Humicola lanuginose lipase showed low activity towards p-nitrophenyl butyrate
and no activity towards an emulsion of olive oil.
Shigechi et al. (2002) reported reduced
activity of Bacillus stearothermophilus -amylase when anchored by -agglutinin in the
cell wall of yeast.
In addition, depending on the degree of the exposure of the anchored
enzymes out of the cell wall, larger substrates can be more difficult to hydrolyze compared
to smaller, more soluble substrates (van der Vaart et al., 1997).
It was suggested that the
hydrophobic nature of the yeast cell wall may further reduce the chances of the long
hydrophobic carbon chains of lipids to make contact with the anchored enzymes (van der
Vaart et al., 1997). Furthermore, considering that each yeast cell has a limited surface area,
there is a physical limit as to how many enzymes can be anchored on the yeast cell wall.
Moreover, a crowded cell surface may disrupt the cell wall structure, which is critical for
cell survival under high osmotic fermentation environments.
Secreted enzymes are not physically restrained in binding their substrates.
As long
as there are carbon sources, it is unlikely that there is a physical limitation to the amount of
enzyme that can be secreted.
Thus, in theory, secreting strain should be able to achieve
higher so greater enzyme concentrations relative to anchored strains. Our results showed a
quick deactivation of secreted barley -amylase after the depletion of carbon sources in the
culture medium (Fig. 4.15A). However, barley -amylase activity increased exponentially
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as long as there were carbon sources available.
In addition, the time lag for starch
hydrolysis during batch fermentation could be shortened by increasing the inoculated cell
concentration (data not shown). Khaw et al. (2002) engineered yeast to have glucoamylase
anchored on the cell surface, and then further engineered the strain to either secrete or
anchor a bacterial -amylase.
Consistent with our results, they showed that the
recombinant strain that secreted -amylase showed higher starch hydrolysis rates and
ethanol productions compared to that of the recombinant strain that expressed cell wall
anchored -amylase.
However, Khaw et al. (2002) maintained that the strain that
expressed cell wall anchored amylolytic enzymes had more potential in industrial
applications because of the reusability of cells and the anchored enzymes.
Based on our studies, we believe that it is more practical to express barley
-amylase in the secreted form for several reasons. First, both ethanol concentration and
the production rate were improved by the higher -amylase activity that was achieved with
the enzyme-secreting strain.
Studies have shown that starch hydrolysis is the rate-limiting
step for ethanol production (de Moraes et al., 1995; Robertson et al., 2006). Since
glucoamylase only attacks the non-reducing ends of starch molecules, the available
non-reducing ends limit the saccharification rate.
Thus, it is the liquefaction (catalyzed by
-amylase) rate that determines how fast fermentable sugars can be generated during
saccharification (catalyzed by glucoamylase).
Our batch fermentation results are
consistent with this. With the same amount of glucoamylase present, strains that generated
the highest -amylase activity showed the highest concentration of ethanol produced (Fig.
4.5).
Even though cell wall anchored -amylase can be recycled, the low ethanol
production of this strain would not be practical for industrial applications. Furthermore,
for raw starch hydrolysis, even higher quantities of barley -amylase are required and have
been estimated to be about 1000X of that required for hydrolyzing soluble starch (Textor et
al., 1998).
Given that the amount of -amylase activity that is generated by the
recombinant yeast is the rate-limiting factor for starch hydrolysis, optimizing -amylase
activity should be a priority when designing amylolytic strains of yeast for bioethanol
production. Our studies characterizing anchored versus secreting strains in a side-by-side
comparison suggest that secreting the enzyme is the design of choice.
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The mechanism of raw starch hydrolysis is another factor that argues in favour of
secreting strains.
Digesting raw starch requires the attachment of the starch-hydrolyzing
enzyme on the starch granule surface, and successful catalysis requires that the enzyme be
able to access the interior of the starch granule, mostly through cracks or small openings on
the granule surface.
Then, the starch granules are digested from the inside out by the raw
starch-hydrolyzing enzymes (Textor et al., 1998).
Thus, it would be physically impossible
for cell surface anchored enzymes to perform this task efficiently.
In conclusion, given that -amylase activity is the rate-limiting factor and its
catalytic mechanism towards starch granules does not favour the anchored form, we
conclude that -amylase should be expressed in the secreted form rather than the anchored
form when designing amylolytic yeast for bioethanol production.
5.3
Strategies to Improve the Secreted Barley -Amylase Activity
Since the secreted barley -amylase activity by the recombinant yeast is the rate
limiting factor for starch hydrolysis, factors that could potentially improve the -amylase
activity were studied.
This included changing fermentation conditions, testing different
promoters to drive gene expression, or introducing mutations into barley -amylase to
improve its performance.
5.3.1 Spiking the Culture Medium with Glucose
Expressed -amylase activity can be improved by changing the fermentation
conditions.
For both recombinant yeast strains that expressed secreted and cell wall
anchored barley -amylase, the batch fermentations using starch as the sole carbon source
resulted in less than 30% of the starch being hydrolyzed (Fig. 4.3).
We found that the
starch hydrolysis rate was improved by spiking the medium with a small amount of glucose.
The rationale for this observation is that a supply of utilizable sugars at the beginning of the
fermentation helps to generate higher biomass, which in turn generates higher amounts of
-amylase activity, resulting in higher starch hydrolysis rates.
Figure 4.15A shows that the
secreted -amylase activity started to drop quickly following the depletion of available
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carbon sources in the medium, further suggesting our contention that the recombinant yeast
requires constant carbon sources for heterologous enzyme production.
5.3.2 Introduction of Single Amino Acid Mutation into Barley -Amylase
A mutant of barley -amylase was also generated which showed improved starch
hydrolysis rate and ethanol production compared to the wild type counterpart.
The
mutation incorporated was based on a study by Sogaard et al. (1993). They showed that S.
cerevisiae expresses four recombinant isoforms of barley -amylase, and two showed
dramatically reduced activity. The isoforms that had reduced activity were found to be
glutathionylated on cysteine 95, which reduces its structural stability, and as a result is
secreted less efficiently.
(Sogaard et al., 1993).
Mutating cysteine 95 to alanine restores the enzyme activity
Glutathionylation is generally involved in redox balancing in yeast
under stress conditions, and over-expression of heterologous proteins can result in elevated
levels of glutathionylation activity (Sogaard et al., 1993). However, Wong et al. (2001)
found that three recombinant isoforms of barley -amylase were expressed in a different
strain of S. cerevisiae, and no reduction in the overall enzyme activity was observed. This
suggests that there are strain differences as to what isoforms of the enzyme are expressed.
Juge et al. (1996) showed that when barley -amylase is expressed in P. pastoris, only a
minor fraction of the isoforms was glutathionylated, and no significant reduction in the
overall enzyme activity was observed. Our results suggest that the yeast strain we used
expresses a significant amount of glutathionylated barley -amylase, since the C95A mutant
showed increased starch hydrolysis and higher ethanol production. The increase in starch
hydrolysis rate could result from both improved enzyme activity and a higher enzyme
secretion efficiency.
These findings emphasize that mutants of barley -amylase,
generated either through random or rational, targeted approaches based on structure/function
studies, should be a part of any strategy to engineer an amylolytic strain of yeast.
It should be mentioned that the terminal two amino acids (Arginine-Serine) of barley
-amylase were deleted for the convenience of cloning. When expressed in S. cerevisiae,
the C-terminal end of barley -amylase is normally processed by a membrane-bound
carboxypeptidase, which removes these residues (Sogaard et al., 1993).
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This
post-translational process does not affect enzyme activity.
In addition, a previous study
showed that the last 10 amino acids are not involved in either enzyme activity or overall
structural stability (Tibbot et al., 2002).
5.3.3 Selection of Promoters
Constitutive promoters are employed to optimize barley -amylase expression at the
transcription level.
Both yeast constitutive promoters ADH1 and TDH3 were studied, and
their effect on secreted barley -amylase activity was compared.
Both promoters are
commonly used for high level heterologous expression, however, our results showed that
ADH1 generates higher barley -amylase activity than TDH3.
Interesting Ly, both
recombinant strains produced comparable levels of barley -amylase activity during the
glucose consumption stage, but the recombinant yeast strain with the promoter ADH1
generated 1.6X higher amounts of barley -amylase activity after entering the ethanol
consumption stage. Although both promoters are generally considered as constitutive,
reports have shown that they can be slightly regulated under certain conditions (McAlister
and Holland, 1985; Pavlovic and Horz, 1988; Ruohonen et al., 1995).
TDH3 has been
reported to be less active when yeast cells enter the stationary phase (Pavlovic and Horz,
1988).
ADH1, which is actually a short version of the full promoter, was found to
become much more active once the cells enter the ethanol consumption stage (Ruohonen et
al., 1995), which likely explains the higher levels of barley -amylase that were achieved
by this promoter.
In summary, a suitable promoter needs to be selected based on the
particular application, and in the current application, ADH1 generated higher levels of
barley -amylase activity than TDH3.
The factors discussed in this section to improve enzyme activity are not an exclusive
list.
Studies in this area are continuously demonstrating the importance of other factors.
For example, supplementing the medium with 1% casamino acids was shown to
significantly minimize proteolytic degradation of recombinant barley -amylase in P.
pastoris (Juge et al., 1996). The addition of Tween 80 has been shown to have positive
effects on extracellular enzyme production by various microorganisms (Kamande et al.,
2000; Shi et al., 2006; Zeng et al., 2006). Conversely, a high initial glucose concentration
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can repress expression of heterologous proteins (Meijer et al., 1998).
With respect to
genetic factors, a constitutive promoter PGK1 (3-phosphog Lycerate kinase) (Hitzeman et
al., 1983) and an inducible promoter MAL61-62 (Finley Jr. et al., 2002) have been shown to
be effective for high-level expression of heterologous proteins in yeast (also see Literature
review 2.5.3.3). Each factor needs to be studied specifically for its particular applications,
since no strategy is universally applicable.
In summary, through engineering and
adjustment of the genetic and environmental factors, the expression level of heterologous
proteins can be improved significantly.
5.4
Glucoamylase Requirement for Bioethanol Production by Recombinant Yeast
In industry, it is common to add both glucoamylase and -amylase to the fermentor
to maximize sugar release.
Glucoamylase is an exo-type amylolytic enzyme, and it acts by
releasing single glucose residues from the non-reducing ends of starch chains and
oligosaccharides.
Generally, glucoamylase activity is limited by the availability of
non-reducing ends in the fermentation broth.
-Amylase, on the other hand, is able to
generate large numbers of oligosaccharides from starch by its random endo-type attack,
which generates more non-reducing ends for glucoamylase to subsequently attack.
Thus,
the two enzymes act synergistically to hydrolyze starch to free sugars.
Many -amylases are able to convert oligosaccharides further into fermentable
sugars, such as glucose and maltose.
not be necessary.
If the conversion rate is sufficient, glucoamylase may
Barley -amylase is an endo-type amylolytic enzyme, cleaving
-1,4-linkages of starch chains in a random fashion to generate primarily short
oligosaccharides.
It then acts secondarily on the oligosaccharides to convert them to
maltose (Textor et al., 1998). Using our initial secreted strain, Figure 4.4C shows that
while starch can be completely liquefied in 36 h, the amount of ethanol generated was neg
Ligible.
It needs to be clarified that the ethanol peak present within the first 24 h was not
derived from the starch, but from fermentation on the glucose that was initially present in
the medium (Fig. 4).
Compared to the wild type and cell wall anchored strains, the
secreting strain showed a prolonged presence of ethanol during batch fermentation.
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However, the amount that accumulated in the medium was low and was entirely consumed
by the yeast by 80 h (Fig. 4C). Use of the mutant barley -amylase strain resulted in
improved ethanol production due to a higher maltose conversion rate (Fig. 4.13), however,
the majority of liquefied starch was still not fermented to ethanol. These results strong Ly
suggest that although barley -amylase is able to convert starch into maltose, which NRRL
Y-132 yeast are capable of fermenting to ethanol (data not shown), the conversion from
oligosaccharides to maltose was too slow to support fermentation.
Textor et al. (1998)
showed that the majority of the initial products generated from the hydrolysis of starch
particles by barley -amylase are oligosaccharides consisting of 5 to 7 glucose residues, and
that high amounts of barley -amylase are required to convert the oligosaccharides to
fermentable sugars.
Thus, the low ethanol production in our case was likely due to the
inefficient conversion of oligosaccharides into fermentable sugars.
To confirm this, a
small amount of exogenous glucoamylase (5 U L-1) was added into the fermentation broth.
As expected, this significantly enhanced the amount of ethanol produced, and
increasing the amount of glucoamylase to 20 U L-1, led to an ethanol yield from starch that
was 70% of the theoretical maximum.
It is of interest to note that the increase in ethanol
yield that was observed upon addition of exogenous glucoamylase occurred in the absence
of any change in the overall starch hydrolysis rate. This is consistent with our hypothesis
that glucoamylase is acting primarily on the oligosaccharides produced by barley -amylase
to produce glucose and maltose, which can be readily fermented.
A number of related
studies have constructed recombinant yeast strains that co-express both -amylase and
glucoamylase.
The amylolytic enzymes were either co-displayed on the cell surface,
co-secreted into the medium, combination of two, or expressed as a single bifunctional
fusion protein enzyme (Shigechi et al., 2004; Wong et al., 2010; Khaw et al., 2006 and de
Moraes et al., 1995).
We decided on a strategy to express only -amylase and add glucoamylase
exogenously for several reasons.
First, of the two enzymes, glucoamylase is much cheaper
than -amylase (Lang et al., 2001), thus providing an economic basis for selecting
-amylase for over-expression. This reason, however, becomes irrelevant if both enzymes
can be co-expressed, but only if the over-expression of the more expensive -amylase is
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unaffected by the co-expression of the glucoamylase.
Since the total copy number of
plasmid is maintained between 50 and 100 by the 2 µm origin of replication, regardless of
what the plasmid codes for, it is predictable that the copy number of the plasmid expressing
-amylase would be reduced by 50% if a plasmid expressing glucoamylase was
co-transformed into the yeast.
This could lead to a corresponding decrease in the
expression level of -amylase. This would likely also be the case with integrated yeast
strains, as there is a limitation on how many copies of a given gene that can be inserted into
the yeast genome.
In support of this, Wong et al. (2010) showed that a recombinant yeast
strain that had both glucoamylase and -amylase genes co-integrated had decreased
expression of both enzymes compared with the strains that had either gene integrated alone.
Given our findings and that of others that -amylase activity is the rate-limiting factor for
raw starch hydrolysis, coupled with the relatively cheap cost of glucoamylase relative to
-amylase (Lang et al., 2001), we suggest that our approach to over-express only the
-amylase and to add glucoamylase exogenously to be the most cost effective manner in
which to perform starch fermentation.
A second advantage of adding glucoamylase
exogenously is that it provides flexibility in controlling the timing of the saccharification
step. Figure 4.20 shows that controlling the glucoamylase addition time point can have a
significant impact on ethanol concentrations. This same level of control is not possible to
achieve when both enzymes are co-expressed.
In conclusion, glucoamylase activity is an absolute requirement for generating the
high rates of starch hydrolysis to fermentable sugars that is required for ethanol production.
By adding glucoamylase exogenously, not only are we able to maximize expression of the
rate-limiting -amylase, but ethanol production can be further enhanced by optimizing the
timing of glucoamylase addition.
5.5
Use of Integration Plasmids to Achieve Stable, High Level Expression of Barley
-Amylase
Our initial strains of yeast that expressed barley -amylase were performed with
2µ-based plasmids that remained episomal.
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These plasmids are widely used for
heterologous protein expression in S. cerevisiae. Episomal expression plasmids have many
convenient properties such as high transformation efficiency, and maintenance of high copy
numbers in each cell.
However, recombinant yeast lose their plasmids very quickly if the selection
pressure is removed (Bo Liao, Master’s thesis, University of Saskatchewan 2008). Besides
environmental factors, such as stress conditions and the presence of certain chemicals, one
major cause for this plasmid loss is that the dividing cells can separate the plasmids
unequally into the two daughter cells during cell division (known as segregational
instability), which results in the generation of plasmid-free cells in the culture (Shuler and
Kargi, 2002). The plasmid-free cells, because they no longer have the burden off carrying
the plasmids and expressing heterologous proteins, can quickly dominate the cultivated
population, resulting in a dramatic drop in the population of plasmid-carrying cells (Altintas
et al., 2001). While relatively high plasmid stability can be maintained under selection
conditions, this is not usually practical for industrial applications. Our results showed that
during fermentation with our recombinant yeast strains, only 20-30% of the yeast were
expressing the plasmid (data not shown), even in the presence of antibiotics, which certainly
would have a negative effect on the expression level of barley -amylase. The use of
recombinant yeast selected by auxotrophic markers could effectively avoid having to use
expensive antibiotics. Kondo et al. (2002) reported high plasmid stability of over 85%
during prolonged fermentation using recombinant yeast selected with an auxotrophic marker.
The problem with using auxotrophic selection markers is that an auxotrophic mutant of the
yeast to be used is required.
However, most of the available mutants are haploid
laboratory strains of yeast, which are not suitable for industrial ethanol fermentation
applications.
Therefore, we decided to pursue integration of our expression cassette into the yeast
genome. A gene cassette can be inserted into the yeast genome at a target location by
homologous recombination.
Integrated heterologous genes are segregationally stable even
under non-selective conditions.
The use of integrated recombinant yeast would allow
fermentations to be carried out in the absence of antibiotics, and also ensures that all of the
yeast cells express the enzyme. Traditionally, the selected sites of integration only allow
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for the integration of a few copies of the gene.
Based on the evidence provided above that
-amylase activity is required at very high levels to achieve raw starch hydrolysis at rates
sufficient to allow for ethanol production, it was clear that a large copy number of the barley
-amylase gene would need to be integrated.
Achieving high copy number requires a targeted genetic locus that is repetitively
present in the yeast genome in high copy numbers. Fortunately, the rDNA and -sequence
exist in the yeast genome in sufficient copy numbers for this purpose.
For example, in a
haploid yeast cell, there are 50-200 copies of rDNA genes. Since integration disrupts the
targeting gene element, and rDNA is essential for cell survival, the yeast integration plasmid
was designed to target the non-transcribed spacer region (NTS) of rDNA (Valenzuela et al.,
1977). The integrated clone (pIR-1 clone #2) showed 3 times higher secreted -amylase
activity than that of the episomal plasmid strain (BA-Secreted) (data not shown).
Moreover, 100% mitotic stability of the integrated barley -amylase expression gene
cassette was observed for over 200 h without any selection pressure present in the medium
(data not shown). Additionally, one of the isolated clones that showed the highest barley
-amylase activity was capable of directly fermenting on raw starch fermentation (Fig.
4.21). These results demonstrated that the integration strain out-performed the episomal
plasmid strain with respect to the expression level of -amylase.
For the episomal plasmid strains, among different clones from a single
transformation, there was no significant difference in the expressed -amylase activity (data
not shown). However, clones of the integrated strains showed a large variation in the
expressed -amylase activity (Fig. 4.18).
This suggests that there is the potential to
optimize the integrated copy numbers of a given gene cassette in the yeast genome. This
was attempted using two different strategies. The first strategy was to reduce the size of
the integration plasmid by deleting all of the bacterial-originated components in the original
plasmid.
Kang et al. (2003) reported that a shorter integration plasmid lacking bacterial
sequences had about four times higher integration efficiency than that of the long one.
However, our results showed otherwise.
The short integration plasmid had a 1.5 kb
decrease in size, but on average the clones generated using the short integration plasmid
showed lower -amylase activity compared to those generated with the long one (Fig.18).
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Clearly, factors that contribute to integration efficiency are not well understood.
The second strategy was to use a deficient promoter to express the selectable marker
gene, Bsd, which provides for blasticidin resistance. The reasoning for using deficient
promoters is that by reducing the transcriptional efficiency of the TEF1 promoter that drives
Bsd expression, only cells with high copy numbers of the integrated cassette would be able
to produce sufficient levels of the selectable marker gene to deactivate the antibiotic, and
thus survive on Bsd. Since the barley -amylase gene is linked to the Bsd gene on the
same plasmid, the selected cells with deficient TEF1 promoter should also have higher copy
number of the barley -amylase gene, theoretically resulting in higher -amylase activity.
Our results showed that the clones carrying the shorter deficient TEF1 promoters were more
likely to show higher amylolytic activity, however, none of the clones had higher activity
than the yeast strains that carried the original integration plasmid (data not shown). The
results suggested that either the integration efficiency is already maximized or that even the
shortest deficient promoter constructed so far was able to produce sufficient level of
blasticidin resistance.
To explore this further, future experiments could either try to
construct even shorter versions of the TEF1 promoter or increase the blasticidin
concentration used to select the transformed yeast.
In summary, the use of integration plasmids can achieve high copy number
integration of a given gene cassette in the yeast genome.
In the present study, the
integrated strain out-performed the episomal plasmid strain with respect to the amount of
-amylase activity and the mitotic stability of the integrated gene, and was able to directly
ferment raw starch. Further improvement of the integration efficiency requires a better
understanding of the integration mechanism.
5.6
Conclusions
An industrial strain of S. cerevisiae was genetically engineered to express cell wall
anchored or secreted barley -amylase.
The recombinant yeast strains were capable of
hydrolyzing soluble starch under batch fermentation conditions, however, they differed
dramatically in starch hydrolysis efficiency. The recombinant strain that secreted barley
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-amylase hydrolyzed starch at a much higher rate, and was able to generate more ethanol
with the addition of exogenous glucoamylase.
Our results indicate that secreted barley
-amylase performs starch hydrolysis more efficiently than the cell wall anchored
counterpart.
Although barley -amylase is able to convert starch into maltose, which is a
fermentable sugar, the conversion occurs too slowly to support fermentation.
Since barley
-amylase is incapable of converting the majority of oligosaccharides into maltose,
complete starch hydrolysis can only be achieved in the presence of glucoamylase activity.
With the proper amount of exogenous glucoamylase, the barley -amylase secreting strain
was able to ferment 2% soluble starch into ethanol up to 92% of the theoretical yield in 36 h.
Moreover, providing a small amount of utilizable carbon source at the start of fermentation
was shown to be critical to allow for biomass production of the recombinant yeast cells and
therefore to achieve adequate expression of -amylase activity.
Furthermore, a mutant of
barley -amylase (Sogaard et al., 1993) was employed in our study, and was shown to
out-perform the original enzyme in starch hydrolysis under fermentation conditions.
Although the 2µ-based episomal plasmid is generally maintained at high copy
numbers in yeast cells, it suffers low mitotic stability even under selection conditions.
To
explore a strategy to stably express the barley -amylase gene in the recombinant yeast, a
yeast integration plasmid was employed in the study. By integrating the barley -amylase
gene into the highly repetitive rDNA locus, strain was developed which showed a higher
expression level of barley -amylase compared to the strain employed the 2µ-based
episomal plasmid.
In addition, the integrated gene showed 100% mitotic stability under
non-selective conditions.
Thus, integration of the gene cassette is a more reliable
expression system and should be used to over-express amylolytic enzymes in recombinant
yeast for bioethanol production.
Moreover, the integrated strain that we generated was
capable of expressing sufficient amounts of barley -amylase to initiate raw starch
hydrolysis.
With the addition of exogenous glucoamylase, raw starch was hydrolyzed to
completion, however, at a slow rate.
Further development on the strain could improve its
efficiency on raw starch hydrolysis.
Our study has avoided drawbacks present in previous studies.
119
Instead of using
common haploid laboratory yeast strains, we employed an industrial yeast strain, which has
more potential for ethanol production. To avoid the use of antibiotics and stabilize the
expressed gene, we integrated the barley -amylase gene in the yeast genome.
Although
both glucoamylase and -amylase are required for efficient starch hydrolysis, our results
and related findings (Wong et al., 2010) support the strategy of optimizing the expression of
-amylase instead of expressing both.
To maximize the expressed -amylase activity, we
only expressed the -amylase in yeast, and added glucoamylase exogenously.
Considering
the cheap price of glucoamylase, this strategy is economically feasible.
Comparing our strain with recently reported studies: Wong et al. (2010) showed a
recombinant yeast co-expressed both glucoamylase and -amylase converted 77% of lintner
(soluble) starch in over a 6-day period.
The conversion of starch increased to 86% and 90%
when the initial starch concentration was increased to 2% and 4% from 1%, respectively.
However, no ethanol production was reported.
Kim et al. (2010) engineered a recombinant
yeast strain that simultaneously expresses three amylolytic enzymes including a
glucoamylase from A. awamori, an -amylase, and a glucoamylase with debranching
activity from Debaryomyces occidentalis.
They showed ethanol concentration of 46.3 g
L-1 from 100 g L-1 (~ 91% of the theoretical yield) soluble starch in 7 days of fermentation.
Few studies reported raw starch-hydrolyzing ability by recombinant yeast.
A recombinant
yeast expressing the cell wall anchored R. oryzae glucoamylase and S. bovis 148 -amylase
produced 61.8 g L-1 ethanol from raw corn starch (86.5% of the theoretical yield) in a 72 h
fermentation (Shigechi et al., 2004).
However, the initial cell density used was very high,
suggesting that their approach may be yet an additional factor to consider when developing
yeast strains for raw starch fermentation.
Our recombinant yeast that secreted barley
-amylase showed higher efficiency on utilizing and fermenting on soluble starch compared
to related studies.
For raw starch hydrolysis, higher efficiency can be achieved by
increasing the biomass inoculated or by increasing the expression or activity of barley
-amylase, which will be discussed in the next section.
120
5.7
Future Directions
The current integrated yeast strain was able to perform raw starch hydrolysis, but at
a relatively slow rate.
Future studies could focus on optimizing the secreted barley
-amylase activity for efficient raw starch hydrolysis under fermentation conditions.
Some specific suggestions are provided bellow:
One aspect to consider would be to improve the expression level of barley
-amylase by increasing the integrated copy number of the -amylase gene.
Since each
haploid yeast cell has 150-200 copies of rDNA, more copies of rDNA exist in polyploid
yeast such as NRRL Y-132. Performing an additional round of homologous recombination
to integrate more copies of the gene should allow for an increase in the copy number.
alternative is to target another repetitive sequence in the yeast genome.
An
The -sequence is
a very good candidate for this approach.
The second aspect to consider is to improve the raw starch-hydrolyzing ability of the
secreted barley -amylase.
This could possibly be achieved by fusing an additional raw
starch binding domain to barley -amylase.
A previous study showed that by fusing the
starch binding domain of A. niger glucoamylase to the C-terminus of barley -amylase, the
engineered enzyme was able to hydrolyze barley starch granules 15-fold faster than the
original form (Juge et al., 2006).
The third aspect to consider is to introduce mutations into barley -amylase that
may improve its activity under fermentation conditions.
By using random mutagenesis,
Wong et al. (2004) have generated large amounts of mutants of barley -amylase, and some
showed improved specific and total activity.
Employment of these mutants of barley
-amylase may improve the raw starch-hydrolyzing ability of our engineered strain.
121
6.0
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