Hindawi Publishing Corporation
International Journal of Carbohydrate Chemistry
Volume 2013, Article ID 624967, 14 pages
http://dx.doi.org/10.1155/2013/624967
Review Article
Production Methods for Hyaluronan
Carmen G. Boeriu,1,2 Jan Springer,1 Floor K. Kooy,1,3
Lambertus A. M. van den Broek,1 and Gerrit Eggink1
1
Wageningen UR Food & Biobased Research, P.O. Box 17, 6700 AA Wageningen, The Netherlands
Universitatea “Aurel Vlaicu,” Str. E. Dragoi nr. 2, 310330 Arad, Romania
3
Genencor International B.V., P.O. Box 218, 2300 AE Leiden, The Netherlands
2
Correspondence should be addressed to Carmen G. Boeriu;
[email protected]
Received 29 November 2012; Accepted 21 January 2013
Academic Editor: Thomas J. Heinze
Copyright © 2013 Carmen G. Boeriu et al. This is an open access article distributed under the Creative Commons Attribution
License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly
cited.
Hyaluronan is a polysaccharide with multiple functions in the human body being involved in creating flexible and protective layers
in tissues and in many signalling pathways during embryonic development, wound healing, inflammation, and cancer. Hyaluronan
is an important component of active pharmaceutical ingredients for treatment of, for example, arthritis and osteoarthritis, and
its commercial value far exceeds that of other microbial extracellular polysaccharides. Traditionally hyaluronan is extracted from
animal waste which is a well-established process now. However, biotechnological synthesis of biopolymers provides a wealth of new
possibilities. Therefore, genetic/metabolic engineering has been applied in the area of tailor-made hyaluronan synthesis. Another
approach is the controlled artificial (in vitro) synthesis of hyaluronan by enzymes. Advantage of using microbial and enzymatic
synthesis for hyaluronan production is the simpler downstream processing and a reduced risk of viral contamination. In this paper
an overview of the different methods used to produce hyaluronan is presented. Emphasis is on the advancements made in the field
of the synthesis of bioengineered hyaluronan.
1. Introduction
Hyaluronic acid, also known as hyaluronan, is a linear polysaccharide composed of a repeating disaccharide unit of
𝛽(1,4)-glucuronic acid (GlcUA)-𝛽(1,3)-N-acetylglucosamine
(GlcNAc) (Figure 1). Both individual carbohydrate residues
in hyaluronan adopt the stable chair conformation which
determines the conformation of the polymer in solution that
is described as an overall random coil structure that may
have also highly flexible regions. Nevertheless, in terms of
chemical structure, hyaluronan is a simple linear polymer
with high molecular mass and exceptional rheological properties. Hyaluronan is a member of the glycosaminoglycans
family that includes chondroitin/dermatan sulfate, keratan
sulfate, and heparin/heparan sulfate, each with a characteristic disaccharide-repeating structure of an amino sugar, either
glucosamine or galactosamine, and a hexose, either galactose,
glucuronic acid, or iduronic acid, which can be carboxylated
or sulfated [1]. Hyaluronan is the only glycosaminoglycan
member that is not sulfated and is not covalently bound to
a proteoglycan core protein.
Research on hyaluronan expands over more than one
century (Table 1). The first report that can be linked to
hyaluronan dates from 1880, when the French chemist
Portes observed that the mucin in the vitreous body, which
he named “hyalomucine,” behaved differently from other
mucoids in cornea and cartilage. Fifty-four years later Meyer
and Palmer described a procedure for isolating a novel glycosaminoglycan from the vitreous of bovine eyes, which they
named hyaluronic acid based on hyaloid (vitreous) and
uronic acid [3]. Since in vivo the polymer exists in an ionized
form as polyanion, it is generally referred to as hyaluronan.
In living organisms, hyaluronan is produced by hyaluronan synthase enzymes, which synthesize large, linear polymers of the repeating disaccharide 𝛽(1,4)-GlcUA-𝛽(1,3)GlcNAc by alternate addition of GlcUA and GlcNAc to
the growing chain using their activated nucleotide sugars
(UDP-glucuronic acid and UDP-N-acetylglucosamine) as
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International Journal of Carbohydrate Chemistry
Table 1: Important events in research on hyaluronan products.
Time
1880
1934
30s–50s
50s
40s–70s
1979
90s–00s
1993
1996
2003
Event
Portes reported that mucin from the vitreous
body differs from other mucoids in cornea and
cartilage and named it hyalomucine [2].
Meyer and Palmer isolated and identified the
polysaccharide from the vitreous body and named
it hyaluronic acid [3].
Hyaluronan from many different tissues of
vertebrates was isolated, identified, and
characterized. A few pathogenic bacteria were
found that produce hyaluronan and use it to
encapsulate their cells.
The chemical structure of hyaluronan was
elucidated by Karl Meyer and his team. They used
hyaluronidase to produce overlapping
oligosaccharides that were structurally analyzed
by conventional techniques [4].
Interest emerged to use hyaluronan in eye surgery
as a substitute of the vitreous body.
Extraction processes from animal tissues were
optimized to remove protein and to minimize
hyaluronan degradation. First studies on
hyaluronan production through bacterial
fermentation and chemical synthesis were
initiated.
First patent on ultrapure hyaluronan isolated from
rooster combs [5]. This was the starting of the
industrial manufacturing of hyaluronan from
animal sources for human applications. In 1980,
using the methods of Balasz Pharmacia (Sweden)
introduced Healon, a product used in cataract
surgery.
Revival of studies on bacterial fermentation to
produce hyaluronan of high molecular weight.
Emphasis on controlling polymer size and
polydispersity.
The gene encoding for a single enzyme that
polymerizes UDP-GlcNAc and UDP-GlcUA into
hyaluronan is isolated by DeAngelis and
coworkers from Streptococcus pyogenes.
Hyaluronan synthases from other
microorganisms were identified and characterized
[6, 7].
The largest hyaluronan fragment, an octamer, was
chemically synthesized through controlled
addition of disaccharide units [8].
Research on the enzymatic synthesis of
hyaluronan and monodisperse hyaluronan
oligosaccharides with defined length [9, 10].
substrates. The number of repeat disaccharides, 𝑛, in a completed hyaluronan molecule is approximately 10,000 but
chains of hyaluronan with up to 25,000 disaccharide units
have been reported. The size of the polymers (𝑀𝑤 from
5,000 Da to 20 million Da) in vivo varies with the type of
tissue. For example, hyaluronan in the human umbilical cord
has a molecular mass of 3-4 million Da, whilst the molecular
O
O
HO
OH
OH
O
O
HO
O
O
NH
OH
C
O
Glucuronic acid
𝑁-Acetylglucosamine
𝑛
Figure 1: Repeating unit of hyaluronan.
mass of hyaluronan from human synovial fluid is 6 million Da. However, the reported molecular weight of isolated
hyaluronan polymers may be underestimated, depending on
the isolation and analysis method used. Hyaluronan is not
monodisperse in molecular weight and the experimentally
determined polydispersity (𝑀𝑤 /𝑀𝑛 ) of the polymer depends
as well on both the method of extraction and the method of
analysis used. For example, the values of the polydispersity
of a hyaluronan sample determined by high-performance
liquid chromatography varied with the type of column, the
detection method, and the flow rate used [11–13]. This results
in a big variation of the values reported for the 𝑀𝑤 and
polydispersity of hyaluronan between laboratories and shows
the importance for standardization of the methodology
for characterization and reporting of hyaluronan molecular
properties.
Hyaluronan is present in all vertebrates. High concentrations are found in the umbilical cord, the synovial fluid
between joints, skin, and the vitreous body of the eye. It
was estimated that in the body of a person of 70 kg, about
15 g hyaluronan is found in different tissues, of which one
third is turned over every day. The human skin contains
over 50% of the hyaluronan in the body [14, 15]. Rooster
combs contain the highest concentrations of hyaluronan
(7.5 mg g−1 ) ever reported for animal tissues [16]. Hyaluronan
is also present in the capsule of a small number of microbial pathogens such as Pasteurella multocida and group A
and C streptococci among which Streptococcus pyogenes (a
human pathogen) and the animal pathogens Streptococcus
equi and Streptococcus uberis, that quite likely pirated the
enzymatic machinery from vertebrate hosts for its synthesis.
These microorganisms use hyaluronan to encapsulate their
cells, forming a perfect disguise against the animal defense
system and facilitating the adhesion and colonization of the
bacterial cells [17]. Since the hyaluronan polymer produced
in animals and the aforementioned bacteria is identical,
the host immune defense is not triggered to repel the
pathogenic bacteria contrary to other bacteria with a different
extracellular capsule. Intriguingly, the nonimmunogenic and
noninflammatory properties of bacterial hyaluronan benefit
mammals currently too, since bacterial hyaluronan forms
an excellent source for medical grade hyaluronan. Initially
hyaluronan was believed to be an inert compound having
no specific interaction with other macromolecules. However,
International Journal of Carbohydrate Chemistry
Table 2: Biomedical and pharmacologic applications of hyaluronan.
Area
Biomedical
Application
Example
Orthopedic surgery
Rheumatology
Ophthamology
Arthritis
Osteoarthritis
Eye surgery
Treatment of the
vocal fold
Otolaryngology
Dermatology and plastic
surgery
Wound healing and
dressing
Tissue engineering
Dermal filler
Diabetic ulcer, skin
burns
Pharmacology Drug delivery
from the discovery of the interaction of hyaluronan with
cartilage proteoglycans by Hardingham and Muir [18] a large
number of reports have been published on the role of hyaluronan in cellular activity, migration, mitosis, inflammation,
cancer, angiogenesis, fertilization, and so on (see for more
information [19, 20]).
In addition, hyaluronan and its derivatives have been
largely studied and applied in the biomedical field (Table 2)
[21–23]. Due to the high level of biocompatibility, hyaluronan
is extensively used in viscosurgery, in the treatment of
arthritis to supplement the lubrication of arthritic joints, as
microcapsule for targeted drug delivery and in cosmetics as a
hydrating and antiaging material.
The biological functions of hyaluronan are strongly
depending upon its size. High molecular weight hyaluronan
polymers (𝑀𝑤 > 5 × 105 Da) are space filling, antiangiogenic,
and immunosuppressive; medium size hyaluronan chains
(𝑀𝑤 between 2 × 104 –105 Da) are involved in ovulation,
embryogenesis, and wound repair; oligosaccharides with
15–50 repeating disaccharide units (𝑀𝑤 between 6 × 103 –
2 × 104 Da) are inflammatory, immuno-stimulatory, and
angiogenic while small hyaluronan oligomers (𝑀𝑤 from 400
to 4000 Da) are anti-apoptotic and inducers of heat shock
proteins [24]. Lower molecular weight hyaluronan and small
oligosaccharides are produced by controlled depolymerization of high molecular weight hyaluronan using physical
treatment (thermal treatment, pressure), irradiation (electron
beam, gamma ray, microwave), acid treatment, ozonolysis,
metal catalyzed radical oxidation, and enzymatic hydrolysis
with hyaluronidase (EC 3.2.1.35).
The commercial value of hyaluronan far exceeds that
of other microbial extracellular polysaccharides. With an
estimated world market value of $US 500 million, it is sold for
up to $US 100,000 per kilogram [25]. On top of this, several
groups have reported effects of hyaluronan oligosaccharides
on cellular behavior that could imply the use of these
molecules in cancer treatment or in wound healing [26]. It is
clear that hyaluronan is much more than a space-filling inert
component of the extracellular matrix and will become more
and more important as a pharmaceutical component and
indeed is a multifunctional megadalton stealth molecule [27].
3
In this paper the different methods to produce hyaluronan
will be discussed as depicted in Figure 2.
2. Biosynthetic Pathways for Hyaluronan in
Living Organisms
The overall reaction of hyaluronan synthesis is as follows:
𝑛 UDP-GlcUA + 𝑛 UDP-GlcNAc
→ 2𝑛 UDP + [GlcUA + GlcNAc]𝑛 .
(1)
The reaction is catalyzed by a single enzyme utilizing both
sugar substrates to synthesize hyaluronan [28]. In contrast to
other glycoaminoglycans which are synthesized in the Golgi
network, hyaluronan is synthesized at the plasma membrane
[29, 30]. Hyaluronan synthases (EC 2.4.1.212) are predicted
to have multiple membrane spanning domains with a large
intracellular loop on the plasma membrane’s inner face.
The only known exception is the P. multocida hyaluronan
synthase which has a membrane attachment domain near the
carboxyl terminus. Hyaluronan molecules are extruded into
the extracellular matrix in coordination with their synthesis
[31], but they also exist intracellularly. In the extracellular
matrix hyaluronan exists in a number of different forms.
For example, in vertebrates it can be intercalated within a
proteoglycan complex, referred to glycocalyx if it is a more
delicate pericellular matrix, or it can be bound to membrane
receptors of the cell surface. It can interact with binding
proteins so-called hyaladherins, which is also the case for
intracellular hyaluronan [15]. The mechanisms regulating
extracellular versus intracellular location of hyaluronan are
not known yet. Recently, the presence of hyaluronan cables
has been reported that was the result of the incorporation of
the heavy chain of interalpha-trypsin inhibitor with hyaluronan [32]. Depending on the type of tissue, different hyaluronan concentrations and molecular weight evoke different
cell responses, for example, during embryonic development,
healing processes, inflammation, and cancer. How the cell
response is influenced by the different molecular weight of
hyaluronan is still an intriguing question [15, 24].
In 1996 the first reports were published concerning the
cloning of mammalian hyaluronan synthases [33, 34]. Since
then in human cells three hyaluronan synthase genes have
been found encoding enzymes having distinct enzymatic
properties with respect to hyaluronan size and size distribution [35] which are differentially inducible by growth factors
and cytokines [36, 37].
In 1993 the first gene encoding the enzyme responsible
for hyaluronan synthesis, denoted HA synthase or HAS,
was identified from group A streptococci [6, 7]. This gene
is part of an operon containing the hasA gene encoding
hyaluronan synthase, the hasB gene encoding UDP-Glucose
dehydrogenase, and the hasC gene encoding UDP-Glucosepyrophosphorylase [38–40]. A hydropathy plot of the hasA
gene predicts three transmembrane segments which is in
accordance with the early findings of Markovitz and coworkers who showed that the hyaluronan synthase activity is
located in the cell membrane of streptococci [41]. Cloning of
4
International Journal of Carbohydrate Chemistry
Production methods for
hyaluronan
Extraction from
Bacterial
animal tissues
production
Rooster combs
In vitro production
Enzymes from
Streptococci
Human umbilical cords
Streptococcus pyogenes
Bovine synovial fluid
Pasteurella multocida
Vitreous humor of cattle
Non pathogenic
micro organisms
Energy and carbon resources
Process conditions
Batch versus chemostat
Strain improvement
Enterococcus faecalis
Escherichia coli
Agrobacterium sp.
Lactococcus lactis
Bacillus subtilis
Figure 2: Production methods for hyaluronan.
the group A and later the group C streptococcal hyaluronan
synthase genes [42] allowed researchers to confirm that only
one gene product (the HA-synthase protein) is required for
hyaluronan biosynthesis.
The first vertebrate gene identified as being HA synthase
was the Xenopus laevis gene DG42 [43]. Since then more
vertebrate HA synthase genes were identified [44] and in
addition an HA synthase from an algal virus was discovered
[17, 45]. The streptococcal, vertebrate, and viral HA synthases
show protein sequence similarity and appear to have a single
glycosyl transferase 2 (GT2) family module [46] with some
similarities to chitin synthase. The last extension of organisms
in which a HA synthase gene was identified is Cryptococcus
neoformans. The CPS1 gene from this pathogenic yeast
encodes a HA synthase with high homology to the previously
mentioned HA synthase proteins [47]. An HA synthase that
differs in protein sequence and in predicted topology from all
other HA synthases was identified and cloned from type A P.
multocida, an animal pathogen [48].
Unlike the other HA synthases (Table 3; grouped as Class
I HA synthases) which are integral membrane proteins and
elongate hyaluronan chains at the reducing end (Class I-R) or
at the nonreducing end [50], the P. multocida HA synthase has
a membrane attachment domain near the carboxyl terminus,
has two GT2 modules, and elongates the HA chains at the
nonreducing end. It is therefore called a Class II HA synthase
[17]. So far the P. multocida HA synthase is the only known
member of the Class II HA synthases [49, 51, 52].
The regulation of hyaluronan synthesis is much more
complex in vertebrates than in bacteria because hyaluronan
fulfills various functions in the mammalian body depending
Table 3: Hyaluronan synthase classification system as proposed by
Weigel and DeAngelis [49].
Class I
Members
Class II
Reducing
Nonreducing
Streptococcus
pyogenes, S.
equisimilis, S.
uberis, humans,
and mice
Xenopus laevis,
algal virus
Pasteurella
multocida
Nonreducing end
Nonreducing
end
Hyaluronan
chain
Reducing end
growth
on the tissue involved and the hyaluronan chain length
required. For the three mammalian HA synthase isozymes
different expression patterns are observed between adult and
embryonic tissues [54, 55]. Several regulatory factors are
known to be involved in controlling HA synthase transcription, such as morphogenesis, cytokines, growth factors, and
antisense mRNA [56, 57]. Furthermore, hyaluronan synthesis
seems to be controlled through translation regulation, since
a latent pool of HA synthase exits within the cell interior, in
the endoplasmic reticulum-Golgi compartments, that upon
insertion in the cell membrane becomes activated [53]. HA
synthase activity can be modulated by posttranslational modification, such as phosphorylation and N-glycosylation [58].
Intermediate and small oligosaccharides are probably provided through hyaluronan cleavage by hyaluronidases, since
mammalian HA synthase isozymes synthesize hyaluronan
International Journal of Carbohydrate Chemistry
5
Glycolysis
Hexokinase
hasE (Phosphoglucoisomerase)
Fructose-6-P
Glucose-6-P
Pgm
GlmS
(Amidotransferase)
(Phosphoglucomutase)
Cell wall
Glucosamine-6-P
Glucose-1-P
polysaccharides hasC
GlmM
(UDP-glucose pyrophosphorylase)
(Mutase)
Glucosamine-1-P
UDP-glucose
Glucosamine-1-P
hasD
hasB
(Acetyltransferase)
(UDP glucose dehydrogenase)
UDP-glucuronic acid
𝑁-Acetylglucosamine-1-P
hasD
(Pyrophosphorylase)
UDP-N-acetylglucosamine
Glucose
Hyaluronan
hasA (Hyaluronan synthase)
Peptidoglycan
Figure 3: Schematic overview of the hyaluronan synthesis pathway in S. zooepidemicus adapted from [53] 2013 The American Society for
Biochemistry and Molecular Biology.
higher than 105 Da [14]. Due to the high complexity of the
regulation of hyaluronan synthesis in vertebrates a cohesive
view on the relation between the regulatory components is
not available yet.
Recently, the metabolic route of hyaluronan formation
in S. zooepidemicus was elucidated. The streptococcal HA
synthase is found in operons also encoding one or more
enzymes involved in biosynthesis of the activated sugars [59].
The has operon in S. zooepidemicus encodes for five genes
(Figure 3): hyaluronan synthase (hasA), UDP-glucose dehydrogenase (hasB), UDP-glucose pyrophosphorylase (hasC),
a glmU paralog encoding for a dual function enzyme acetyltransferase and pyrophosphorylase activity (hasD), and a pgi
paralog encoding for phosphoglucoisomerase (hasE) [60].
HasB and hasC are involved in UDP-GlcUA synthesis, while
hasD and hasE are involved in the synthesis of UDP-GlcNAc.
The hyaluronan biosynthesis is a costly process for bacteria
in terms of carbon and energy resources and competes with
cell growth because of the large pathway overlap with the
cell wall biosynthesis (Figure 3). Other streptococci strains
have has operons that contain besides hyaluronan synthase,
only the hasB and hasC genes. This suggests that the superior
hyaluronan synthesis observed for S. zooepidemicus relates to
the availability of UDP-sugar precursors. Understanding the
metabolic routes for hyaluronan synthesis had a significant
role in the optimization of the microbial production of
hyaluronan, allowing the control of the polymer chain length
and increasing the product yield.
3. Production of Hyaluronan
Industrial manufacturing of hyaluronan is based on two main
processes, the extraction from animal tissues and microbial fermentation using bacterial strains. Both technologies
produce polydisperse high molecular weight hyaluronan
(𝑀𝑤 ≥ 1 × 106 Da, polydispersity ranging from 1.2 to 2.3)
for biomedical and cosmetic applications [12, 61, 62]. The
first process, to be applied at industrial scale, was the
extraction of hyaluronan from animal waste which is still
an important technology for commercial products, but is
hampered by several technical limitations. One drawback
in the extraction process is the inevitable degradation of
hyaluronan, caused by (a) the endogenous hyaluronidase
activity in animal tissues, breaking down the polymer chain
through enzymatic hydrolysis, and (b) the harsh conditions
of extraction. Extraction protocols have been improved over
the years, but still suffer from low yields, due to the intrinsic
low concentration of hyaluronan in the tissue, and from high
polydispersity of polymer products due to both the natural
polydispersity of hyaluronan and to the uncontrolled degradation during extraction. As in any process for the production
of therapeutic compounds from animal sources, there is a
potential risk of contamination with proteins and viruses,
but this can be minimized by using tissues from healthy
animals and extensive purification. Nevertheless, concerns
on viral (particularly avian) and protein (particularly bovine)
contamination increased the interest in the biotechnological
production of hyaluronan.
Production of hyaluronan by bacterial fermentation
evolved to a mature technology in the last two decades of the
20th century. In the early developmental stages of bacterial
fermentation using group A and C streptococci, optimization
of culture media and cultivation conditions along with strain
improvement were used to increase hyaluronan yield and
quality. Hyaluronan yields reached 6-7 g L−1 , which is the
upper technical limit of the process caused by mass transfer
limitation due to the high viscosity of the fermentation
broth. Bacterial fermentation produces hyaluronan with high
molecular weight and purity, but risk of contamination
with bacterial endotoxins, proteins, nucleic acids, and heavy
metals exists. The identification of the genes involved in
the biosynthesis of hyaluronan and of the sugar nucleotide
6
International Journal of Carbohydrate Chemistry
Table 4: A comparative view of technologies for manufacturing hyaluronan.
Extraction from animal
materials
Bacterial production
Advantages
Disadvantages
(i) Well-established technology
(ii) Available raw material at low costs
(iii) Product with very high 𝑀𝑤 up to
20 MDa
(iv) Natural product
(i) Variation in product quality
(ii) Risk of polymer degradation
(iii) Risk of contamination with protein, nucleic acids, and
viruses
(iv) Extensive purification needed
(v) Low yield
(i) Mature technology
(i) Use of genetically modified organism (GMO)
(ii) Risk of contamination with bacterial endotoxins,
proteins, nucleic acids, and heavy metals
(ii) High yield
(iii) Product with high 𝑀𝑤 (1–4 MDa)
Enzymatic synthesis
(i) Versatile technology
(ii) Control of the 𝑀𝑤 of the products.
Products up to 0.55–2.5 MDa and also
defined oligosaccharides can be
obtained
(iii) No risks of contamination
(i) Emerging technology in development stage
(ii) Technological and economic viability must still be
demonstrated
(iv) Constant product quality
precursors in the 90s opened the way towards hyaluronan
production using safe, nonpathogenic recombinant strains.
In the past decade a new technology emerged using
isolated HA synthase to catalyze the polymerization of the
UDP-sugar monomers. This novel enzymatic technology for
hyaluronan synthesis is very versatile allowing producing
both high molecular weight hyaluronan and hyaluronan
oligosaccharides with defined chain length and low polydispersity. Production of monodisperse hyaluronan oligosaccharides on mg-scale using two single-action mutants of the
P. multocida HA synthase was demonstrated by the group of
DeAngelis [9], but large scale production was not achieved
yet. A comparative analysis of established and emerging technologies for high molecular weight hyaluronan production is
given in Table 4.
3.1. Extraction from Animal Tissues. Extraction of hyaluronan from animal tissues was initially used for laboratory
investigations in order to identify and characterize the polymer and to elucidate its biological potential and biomedical
application. Hyaluronan from almost all tissues of vertebrates
including the vitreous body of the eye, umbilical cord,
synovial fluid, pig skin, the pericardial fluid of the rabbit,
and the cartilage of sharks was isolated and described [63].
Recently, the extraction of hyaluronan from fish eyes as an
alternative resource outside animal husbandry was reported
[64]. Nevertheless, the most accessible sources for large scale
production high molecular weight hyaluronan are rooster
combs (1.2 × 106 Da), the human umbilical cords (3.4 ×
106 Da), the vitreous humor of cattle (7.7 × 104 –1.7 × 106 Da),
and the bovine synovial fluid (14 × 106 Da).
Despite the numerous methods for isolation and purification of hyaluronan reported in the early literature, it was only
in 1979 that a method became available for the production of
pharmaceutical grade hyaluronan by extraction from animal
tissues. Balazs [5] developed an efficient procedure to isolate
and purify hyaluronic acid from rooster combs and human
umbilical cords that set the basis of the industrial production
of hyaluronan from rooster combs for medical application.
Hyaluronan is soluble in water, but extraction of highly
pure, high molecular weight hyaluronan from animal tissues
is difficult, since in biological materials hyaluronan is usually
present in a complex with other biopolymers including proteoglycans [65]. Methods to release hyaluronan from these
complexes comprise the use of proteolytic enzymes (i.e.,
papain, pepsin, pronase, and trypsin), hyaluronan ion-pair
precipitation (with e.g., cetylpyridinium chloride), precipitation with organic solvents, nonsolvent precipitation, detergents, and so forth [21]. Ultrafiltration and chromatography
are used to remove the degradation products and other
contaminants. Sterile filtration is used to remove all microbial
cells prior to alcohol precipitation, drying, and conditioning
of the end product.
Despite extensive purification, animal hyaluronan can be
contaminated with proteins and nucleic acids. The quantity
and nature of contaminants differ with the source, as was
shown for hyaluronan isolates from human umbilical cord
and bovine vitreous humor, containing elevated protein and
nucleic acid levels compared to those from rooster comb and
bacterial capsule isolates [12]. Diseases as bovine spongiform
encephalopathy (BSE) made us aware of the risk of crossspecies viral infection. Consequently, hyaluronan from animal sources has to be extensively purified to get rid of these
contaminants and extraction and purification processes are
continuously improved to fulfill the high quality standards
for medical applications. To date, animal waste is still the
most important source for the industrial manufacturing of
hyaluronan for medical application and provides up to several
tones of medical-grade hyaluronan preparations per year.
Pharmacia (Sweden), Pfizer (USA), Genzyme (USA), and
International Journal of Carbohydrate Chemistry
Diosynth (The Netherlands) were among the first companies
that produced hyaluronan from animal waste at industrial
scale. Commercially available hyaluronan preparations from
animal sources have a molecular weight ranging from several
hundred thousand up to 2.5 million Da [66].
3.2. Bacterial Production. The development of hyaluronan
production through bacterial fermentation started in the 60s,
when it was acknowledged that animal derived hyaluronan
sources can contain undesired proteins causing possible
allergic inflammation responses [67]. Since the hyaluronan
polymer produced in animals and bacteria is identical,
bacterial hyaluronan is not immunogenic and therefore is
an excellent source for medical grade hyaluronan. Extracting
hyaluronan from microbial fermentation broth is a relatively
simple process with high yields. An additional and important advantage of microbial hyaluronan production is that
microbial cells can be physiologically and/or metabolically
adapted to produce more hyaluronan of high molecular
weight. Therefore, microbial hyaluronan production using
either pathogenic streptococci or safe recombinant hosts,
containing the necessary hyaluronan synthase, is nowadays
more and more preferred.
3.2.1. Production of Hyaluronan with Streptococci Strains.
The first reported hyaluronic acid isolation from group A
hemolytic streptococci resulted in 60–140 mg L−1 hyaluronan
[68]. Since then, many different attempts have been made
to increase the amount of hyaluronan, including traditional
techniques as optimizing the extraction process, adapting the
culture media, and selecting strains with high hyaluronan
productivity. In thirty years, the hyaluronan yields using
batch fermentation increased from 300–400 mg L−1 [69] to 67 g L−1 [70], which is the practical limit due to mass transfer
limitations [25]. Group C streptococci which are not human
pathogens and have superior hyaluronan productivity are
frequently used for hyaluronan synthesis instead of Group
A streptococci or instead the animal pathogenic bacterium
P. multocida. The most used strains are S. equi subsp. equi
and S. equi subsp. zooepidemicus. From dairy food products
S. thermophilus strains with high productivity of useful
exopolysaccharides including hyaluronan were isolated [71],
offering a safe hyaluronan producing organism. Recently, a
recombinant S. thermophilus was constructed that was able
to produce 1.2 g L−1 hyaluronan and the average molecular
weight was comparable with the wild type strain [72].
Streptococci strains for hyaluronan production generally
use glucose as carbon source. The yield of hyaluronan on glucose under aerobic fermentation conditions varies between 5
and 10% [25, 73, 74], which is significantly higher than typical
yields for complex polysaccharides in lactic acid bacteria.
Other carbon sources like starch [75], lactose, sucrose, and
dextrin [76] can be used for hyaluronan productivity similar
to glucose. The use of carbon sources like starch and lactose
which are largely available at lower costs is important in view
of the process economics.
Research on optimization of hyaluronan production
through fermentation of streptococci strains addressed the
7
improvement of the hyaluronan yield, the increase of the
molecular weight, and the reduction of the polymer polydispersity, parameters which are essential for hyaluronan
applications. The main question was if the molecular weight
could be further regulated either through strain improvements, optimized process conditions, or through metabolic
engineering. These topics will be discussed below.
Regulation within Bacterial Cells. Energy and Carbon Resources. Hyaluronan biosynthesis in streptococci requires a
large amount of energy and competes with the bacterial
cell growth for glucose as energy provider or as UDP-sugar
precursor. Indeed, when unlimited glucose is available, the
highest bacterial growth was observed at optimal cultivation
conditions (pH 7 and 37∘ C) whilst the highest hyaluronan
productivity and molecular weight were obtained at suboptimal growth conditions, since when cells are growing
slowly, the carbon and energy resources are available for
other processes [77]. Under glucose-limiting conditions first
hyaluronan productivity and then molecular weight declines
[25]. Decrease of the initial glucose concentrations from
60 to 20 g L−1 decreased the hyaluronan concentration and
molecular weight from 4.20 g L−1 and 3.0 million Dalton, to
1.65 g L−1 and 2.65 million Dalton, respectively [77].
Aerobic culturing conditions increased the hyaluronan productivity by 50% as well as hyaluronan molecular
weight whereas cell growth was unaffected [77–79]. Under
aerobic fermentation conditions streptococci change their
metabolism from producing lactate into producing acetate,
formate, and ethanol. This generates four ATP molecules
per hexose for the acetate production instead of two ATP
molecules formed in lactate production [80]. It was assumed
that the increase of hyaluronan production is due to the
increased levels of ATP or to increased levels of NADH
oxidase which removes the excess of NADH in the presence of oxygen. However, studies designed to improve ATP
formation by increased acetate production through maltose
metabolism [80] or increasing NADH oxidase levels by
metabolic engineering [81] revealed that hyaluronan synthesis was not limited by energy resources.
Studies have shown that only a critical level of dissolved
oxygen of 5% is needed to increase the hyaluronan yield
during cell growth, above this value the yield is constant
[79]. Interestingly, the expression of HA synthase in S. zooepidemicus is nine times higher under aerobic conditions than
under anaerobic conditions [74], explaining the increase of
hyaluronan synthesis. Furthermore, oxygen induces enzymes
involved in the production of UDP-GlcNAc (hasD) and
acetoin recycling, which offers an additional ATP molecule
and acetyl-CoA that can be used in hyaluronan production
[82].
Based on these results, we can conclude that elevated HA
synthase concentrations and UDP-GlcNAc availability seem
to be the main reason for the increase in hyaluronan synthesis
under aerobic conditions.
Influence of Process Conditions on Hyaluronan Yield and
Molecular Weight. Streptococcal fermentation to produce
hyaluronan is, like other fermentations, influenced by
8
medium composition, pH, temperature, aeration, and agitation. Especially aerobic conditions and the initial glucose
concentration had a large influence on hyaluronan yield and
molecular weight, as was discussed above. Both agitation
and fermentation modus have a rather complex effect on
hyaluronan production and are, therefore, in greater detail
described, below.
The effect of agitation on the yield and the molecular weight of hyaluronan in Streptococcus fermentation is
complex. Increasing agitation increases the hyaluronan yield
under both anaerobic and aerobic conditions, most probably
due to an enhanced mass transfer induced by the reduced
viscosity of the broth [77, 79, 83, 84]. However, hyaluronan
molecular weight increases at moderate impeller speed due
to mass transfer enhancement, but decreases at high impeller
speed most probably due to degradation by the reactive oxygen species that are formed under aerobic conditions at high
impeller speed [85]. Oxidative degradation of hyaluronan can
be prevented by the addition of oxygen scavengers such as
salicylic acid in the culture media resulting in an increase of
the hyaluronan molecular weight [85, 86].
Batch versus Chemostat. In batch fermentation, the hyaluronan production is limited to 6-7 g L−1 due to high viscosity
of the broth and mass transfer constraints. Further improvement of the hyaluronan yield is therefore not possible through
high-cell-density fermentation or through a high-yield strain.
A continuous culture is the best strategy to improve the
volumetric productivity of hyaluronan synthesis for four
reasons. First, cell growth of streptococci can be maintained
at the exponential phase. Extension of the exponential phase
could lead to an increased amount of hyaluronan, since it
was shown that hyaluronan synthesis stops in the stationary
phase [40, 87] and the excretion of the HA synthase and other
cell wall proteins which takes place in the stationary phase
is prevented [88]. Second, suboptimal growth conditions can
be imposed by nutrient limitation in order to decrease the
resource competition between cell growth and hyaluronan
synthesis [77, 89]. Third, a turnaround phase is avoided by
using a continuous culture [90]. Finally, the viscosity of the
broth can be controlled by controlling the hyaluronan concentration, thus enhancing the mass transfer in the reactor.
Chemostat cultivation of S. zooepidemicus by controlling the
medium conditions has been reported [75, 90, 91] and this
opens the way for further improvement of the hyaluronan
production process.
Strain Improvement. Improved hyaluronan producing strains
have been obtained by random mutagenesis and selection of
mutant strains oriented to the reduction of the hyaluronandepolymerising enzyme responsible for the decrease of
molecular weight of hyaluronan during fermentation (hyaluronidase-negative strains) and to the reduction of streptolysin, the exotoxin responsible for the 𝛽-hemolysis (nonhemolytic strains). Fermentation of nonhemolytic and
hyaluronidase-free mutants produced hyaluronan with a
molecular weight varying from 3.5 to 5.9 million Da and
yields around 6-7 g L−1 [70, 76, 92]. Random mutagenesis has
also resulted in Streptococci strains that produce hyaluronan
International Journal of Carbohydrate Chemistry
of extraordinary high molecular weight ranging from 6 to 9
million Da and with moderate yields of 100 to 410 mg L−1 [93].
Further improvement of strains requires genetic engineering of the producer microorganisms to engineer the
hyaluronan metabolic pathway. Metabolic engineering in
S. zooepidemicus and in recombinant hosts has demonstrated
that the ratio between UDP-GlcNAc and UDP-GlcUA and
the ratio between HA synthase and substrates are the most
important factors to influence hyaluronan molecular weight.
Chen et al. individually overexpressed each of the five genes
in the has operon in S. zooepidemicus. They discovered that
overexpression of hasA, involved in hyaluronan biosynthesis,
and hasB and hasC, involved in UDP-GlcUA biosynthesis,
decreased molecular weight, while overexpression of hasE,
involved in UDP-GlcNAc biosynthesis, greatly enhanced
molecular weight. Overexpression of hasD had no effect,
but molecular weight was further increased when combined
with hasE overexpression [60]. Upregulation of hasD in S.
zooepidemicus has been observed at aerobic conditions [82]
but also in the presence of an empty plasmid [94], both
accompanied by a production of higher 𝑀𝑤 hyaluronan.
Marcellin et al. [94] suggested that regulation of hasD is at the
translational or posttranslational level and that the plasmid
burden is sufficient to remove the hasD encoded step as a
limiting step. In summary, these results indicate that, with
a proper balance between the synthesis pathways to UDPGlcNAc and UDP-GlcUA, the hyaluronan molecular weight
can be further increased and controlled.
In addition to the ratio between the two precursors, the
importance of the ratio between precursor UDP-GlcUA and
HA synthase also influences hyaluronan molecular weight.
This was demonstrated for the heterologous host L. lactis, two
plasmids were constructed containing either hasA or hasB
from S. zooepidemicus with the inducible expression promoters NICE and lacA. Hyaluronan molecular weight increased
significantly at hasA/hasB ratios below 2, indicating that
higher UDP-GlcUA availability per HA synthase enhances
hyaluronan molecular weight [95].
To conclude, production of hyaluronan by fermentation
of streptococci strains is a mature technology, affording
high molecular weight, highly pure polymers suitable for
medical, pharmaceutical and cosmetic applications. Several
tons per year of bacterial hyaluronan are currently produced
for medical and cosmetic applications. Companies that produce hyaluronan through streptococcal fermentation are, for
instance, Q-Med (Uppsala, Sweden), Lifecore Biomedical
(Chaska, USA), and Genzyme (Cambridge, USA).
3.2.2. Hyaluronan from Recombinant Nonpathogenic Microorganisms. To avoid the risk of exotoxin contamination in hyaluronan products from pathogenic streptococci
strains, safe organisms have been genetically engineered into
hyaluronan producers, by introducing HA synthase enzymes
from either streptococci or P. multocida. Using this approach,
hyaluronan producing strains of Enterococcus faecalis [7],
E. coli [7, 48, 96, 97], Bacillus subtilis [98], Agrobacterium
sp. [99], and L. lactis [95, 100–102] were obtained and
exploited for hyaluronan production. Heterologous expression of HA synthase (hasA gene in streptococci) was sufficient
International Journal of Carbohydrate Chemistry
to induce production of hyaluronan. Since the hyaluronan
biosynthesis has partially a common pathway with cell wall
biosynthesis (Figure 3), production of hyaluronan by the host
goes, however, on the expense of cell growth, due to the
depletion of sugar precursors. Better hyaluronan producing
heterologous strains with improved intracellular availability
of sugar precursors were obtained by coexpression of the
HA synthase hasA gene derived from S. equi or P. multocida
with hasB homologue (UDP-glucose dehydrogenase) or hasB
+ hasC homologues (UDP-glucose dehydrogenase + UDPglucose pyrophosphorylase) from E. coli. A recombinant
E. coli strain obtained using this strategy produced 2 g L−1
hyaluronan, and the yield was increased to 3.8 g L−1 when the
culture media was supplemented with glucosamine [99].
Using a similar strategy, namely the coexpression of the
HA synthase hasA gene from P. multocida with hasB homologue (UDP-glucose dehydrogenase) from E. coli, into the
food-grade microorganisms Agrobacterium sp. [99] and L.
lactis [101], these strains were able to produce hyaluronan at
levels up to 0.3 g L−1 for engineered Agrobacterium sp. and
0.65 g L−1 for the recombinant L. lactis. Although hyaluronan
production yields were low, the hyaluronan production from
these food-grade microorganisms has high potential for food
and biomedical applications.
Without overexpression of hasB or an analog encoding
for UDP-glucose dehydrogenase, UDP-GlcUA levels are often
limiting in heterologous hosts [7, 97, 98, 101], whereas UDPGlcNAc seems to be adequately available in the hosts since
it is a main component for bacterial cell wall synthesis.
Attempts to overexpress both synthetic pathways to UDPGlcNAc and UDP-GlcUA in heterologous hosts have been so
far unsuccessful, but overexpression of hasA, hasB, and hasC
has led to considerable increases in hyaluronan synthesis
varying between 200 and 800% depending on the host [96,
100].
A process for the production of ultrapure sodium hyaluronate by the fermentation of a novel, nonpathogenic
recombinant strain of B. subtilis was developed by the biotech
company Novozymes [103]. A major advantage of using B.
subtilis as host microorganism is that it is cultivable at large
scale, does not produce exo- and endotoxins, and does not
produce hyaluronidase. To build up the recombinant Bacillus
strain that produces hyaluronan, expression constructs utilizing the hasA gene from S. equisimilis in combination with
overexpression of one or more of the three native B. subtilis
precursors genes—tuaD (hasB homologue, encoding UDPglucose dehydrogenase), gtaB (hasC, encoding UDP-glucose
pyrophosphorylase), and gcaD (hasD, encoding UDP-Nacetyl glucosamine pyrophosphorylase). The recombinant B.
subtilis strain was able to produce up to 5 g L−1 of hyaluronan
with a molecular weight of 1–1.2 million Da, when cultivated
on a minimal medium based on sucrose at pH 7 and 37∘ C
[94]. The hyaluronan is secreted into the medium and is not
cell-associated, which simplifies the downstream processing
significantly, making the use of water-based solvents possible
for hyaluronan recovery. The Bacillus hyaluronan shows
very low levels of contaminants (i.e., proteins, nucleic acids,
metal ions, and exo- and endotoxins) and thus has a high
9
safety profile, including no risks of viral contamination or of
transmission of animal spongiform encephalopathy.
As an alternative to prokaryotic HA production, hyaluronan can be produced by infecting green algae cells of the
genus Chlorella with a virus [45, 104], although the reported
yields were low (0.5–1 g L−1 ).
3.3. In Vitro Production of Hyaluronan Using Isolated HA
Synthase. Synthesis of hyaluronan using isolated HA synthase becomes relevant when hyaluronan polymers of defined
molecular weight and narrow polydispersity are needed.
Isolated HA synthase is able to catalyze in vitro at well-defined
conditions the same reaction as it catalyzes in vivo, namely,
the synthesis of hyaluronan from the nucleotide sugars UDPGlcNAc and UDP-GlcUA. Preparative enzymatic synthesis of
hyaluronan using the crude membrane-bound HA synthase
from S. pyogenes was demonstrated, although the yield was
low, around 20% [105]. The hyaluronan yield was increased to
90% when the enzymatic hyaluronan synthesis was coupled
with in situ enzymatic regeneration of the sugar nucleotides
using UDP and relatively inexpensive substrates, Glc-1-P and
GlcNAc-1-P in a one-pot reaction. The average molecular
weight of the synthetic hyaluronan was around 5.5 × 105 Da,
corresponding to a degree of polymerization of 1500.
High molecular weight monodisperse hyaluronan polymers with 𝑀𝑤 up to 2.500 kDa (∼12,000 sugar units) and
polydispersity (𝑀𝑤 /𝑀𝑛 ) of 1.01–1.20 were obtained by enzymatic polymerization using the recombinant P. multocida HA
synthase, PmHAS, overexpressed in E. coli [106]. PmHAS
uses two separate glycosyl transferase sites to add GlcNAc and
GlcUA monosaccharides to the nascent polysaccharide chain.
Hyaluronan synthesis with PmHAS was achieved either by de
novo synthesis from the two UDP-sugars precursors (1) and
by elongation of an hyaluronan-like acceptor oligosaccharide
chain by alternating, repetitive addition of the UDP-sugars as
follows:
𝑛 UDP-GlcUA + 𝑛 UDP-GlcNAc + z[GlcUA-GlcNAc]𝑥
→ 2𝑛 UDP + [GlcUA + GlcNAc]𝑥+𝑛 .
(2)
The control of the chain length and polydispersity of
the hyaluronan polymer is determined by the intrinsic
enzymological properties of the recombinant PmHAS, (i)
the rate limiting step of the in vitro polymerization appears
to be the chain initiation, and (ii) in vitro enzymatic
polymerization is a fast nonprocessive reaction. Therefore,
the concentration of the hyaluronan acceptor controls the
size and the polydispersity of the hyaluronan polymer in
the presence of a finite amount of UDP-sugar monomers
[106]. Using this synchronized, stoichiometrically-controlled
enzymatic polymerization reaction, low molecular weight
hyaluronan (∼8 kDa) with narrow size distribution was
synthesized. One important feature of the PmHAS is that
chain elongation occurs at the nonreducing end of the
growing chain and this makes the use of modified acceptors as substrates possible and consequently the synthesis
of hyaluronan polymers with various end-moieties. The
elongation of a hyaluronan tetrasaccharide labeled at the
reducing end with the fluorophore 2-aminobenzoic acid
10
using PmHAS and the quantitative formation of fluorescent hyaluronan oligomers and polymers was demonstrated
[10, 107].
These studies have shown the high potential of the in vitro
enzymatic synthesis of hyaluronan polymers and oligomers
with low polydispersity, but for the large scale application
further developments are needed. Technological bottlenecks
that must be resolved are (a) production of robust enzymes
for hyaluronan synthesis, (b) selective separation of the UDP
byproduct from the reaction mixture, to prevent enzyme
inactivation and to allow UDP recycling for the synthesis
of UDP-sugar substrates, (c) development of efficient processes for the production of the hyaluronan-like template
for elongation, and (d) the development of simple, low cost
technologies for the synthesis of sugar nucleotide substrates
starting from bulk carbohydrates. Since sugar nucleotides are
expensive substrates their regeneration is a crucial step for the
development of an economic process for the production of
hyaluronan.
For in vitro production of hyaluronan the Class I HA
synthases are less suitable due to the fact that they are integral
membrane proteins. Research with these HA synthases is
mostly performed on membrane fractions but this system is
less suitable for larger scale production. An alternative could
be the immobilization of the enzyme in the cell wall of for
instance yeast. Production of various human oligosaccharides
has shown to be feasible with yeast cells in which glycosyl
transferases are expressed and anchored in the cell wall
glucan [108].
A far more promising enzyme for in vitro production of
hyaluronan is the P. multocida Class II HA synthase. This
enzyme is not an integral membrane protein and the analysis
of the subcellular location and enzyme activity has shown that
a recombinant truncated version of the protein lacking a 216
carboxy terminal amino acids retained HA synthase activity
but was located in the cytoplasm when expressed in E. coli
[109]. Furthermore, it was shown that two distinct transferase
activities are located on the P. multocida HA synthase each
of which can be inactivated by mutations in the DXD amino
acid motif present in the active site while retaining the other
transferase activity [110].
UDP-Recycling and Sugar Nucleotide Synthesis (Enzymatic
Cascade, Bacterial Production). The general metabolic routes
for incorporation of sugar units into glycosaminoglycans
are their prior conversion into sugar nucleotides such as
UDP-GlcA and UDP-GlcNAc for hyaluronan synthesis. The
biosynthetic reactions are known as LeLoir pathways and
these pathways can be different in microorganisms [111]. For
example, the pathways for the synthesis of UDP-GlcNAc in
eukaryotes and prokaryotes are different. Production of sugar
nucleotides needs also cofactor regeneration for preparative
applications, because the cofactors are too expensive to be
used as stoichiometric agents. Nowadays, several cofactors
can be effectively regenerated using enzymes, for example,
NAD+ by glutamate dehydrogenase with 𝛼-ketoglutarate,
or whole cell based methods, for example, Corynebacterium ammoniagenes that converts orotic acid into UTP
[112–115].
International Journal of Carbohydrate Chemistry
4. Future Perspectives
Hyaluronan, with its exceptional properties and multiple and
diverse applications, has proved to be an exceptional material for medical, pharmaceutical, and cosmetic applications.
Tremendous developments have been achieved in the past
decades in all hyaluronan-related fields, covering the whole
chain from the production of hyaluronan polymers and
oligomers, to the development of advanced materials for clinical applications. Latest accomplishments in hyaluronan production and in particular the elucidation of the biosynthetic
pathways in hyaluronan producing microorganisms opens
new premises for the further optimization of biotechnological
process for hyaluronan production with safe hosts. Further
understanding of the in vivo regulation of hyaluronan molecular weight will benefit the biotechnological production of
defined hyaluronan products with narrow polydispersity. We
believe that bacterial fermentation using nonpathogenic, safe
heterologous hosts will become the main source of high
molecular weight hyaluronan, and metabolic engineering
will be used to improve and control molecular weight.
Stimulated by the developments of production techniques
and the growing knowledge on the biological function of
hyaluronan, research to enhancing existing medical products
and to validating new concepts in medical therapies will be
set forth.
Abbreviations
BSE:
HA synthase or HAS:
𝑀𝑤 :
𝑀𝑛 :
GlcAU:
GT:
UDP:
GlcNac:
GMO:
Bovine spongiform encephalopathy
Hyaluronan synthase
Weight average molecular mass
Number average molecular mass
Glucuronic acid
Glycosyl transferase
Uridine diphosphate
N-acetyl glucosamine
Genetically modified organism.
References
[1] P. L. DeAngelis, “Glycosaminoglycan polysaccharide biosynthesis and production: today and tomorrow,” Applied Microbiology
and Biotechnology, vol. 94, no. 2, pp. 295–305, 2012.
[2] H. G. Garg and C. A. Hales, Chemistry and Biology of Hyaluronan, Elsevier, 2004.
[3] K. Meyer and J. W. Palmer, “The polysaccharde of the vitreous
humor,” The Journal of Biological Chemistry, vol. 107, no. 3, pp.
629–634, 1934.
[4] B. Weissmann and K. Meyer, “The structure of hyalobiuronic
acid and of hyaluronic acid from umbilical cord,” Journal of the
American Chemical Society, vol. 76, no. 7, pp. 1753–1757, 1954.
[5] E. A. Balazs, “Ultrapure hyaluronic acid and the use thereof,”
U.S. Patent, US4141973, 1979.
[6] P. L. DeAngelis, J. Papaconstantinou, and P. H. Weigel, “Molecular cloning, identification, and sequence of the hyaluronan
synthase gene from group A Streptococcus pyogenes,” The
Journal of Biological Chemistry, vol. 268, no. 26, pp. 19181–19184,
1993.
International Journal of Carbohydrate Chemistry
[7] P. L. DeAngelis, J. Papaconstantinou, and P. H. Weigel, “Isolation of a Streptococcus pyogenes gene locus that directs
hyaluronan biosynthesis in acapsular mutants and in heterologous bacteria,” The Journal of Biological Chemistry, vol. 268, no.
20, pp. 14568–14571, 1993.
[8] G. Blatter and J. C. Jacquinet, “The use of 2-deoxy-2trichloroacetamido-D-glucopyranose derivatives in syntheses
of hyaluronic acid-related tetra-, hexa-, and octa-saccharides
having a methyl 𝛽-D-glucopyranosiduronic acid at the reducing
end,” Carbohydrate Research, vol. 288, pp. 109–125, 1996.
[9] P. L. DeAngelis, L. C. Oatman, and D. F. Gay, “Rapid chemoenzymatic synthesis of monodisperse hyaluronan oligosaccharides with immobilized enzyme reactors,” The Journal of Biological Chemistry, vol. 278, no. 37, pp. 35199–35203, 2003.
[10] F. K. Kooy, Enzymatic Production of Hyaluronan Oligo- and
Polysaccharides, Wageningen University, Wageningen, The
Netherlands, 2010.
[11] B. Porsch, R. Laga, and C. Konak, “Batch and size-exclusion
chromatographic characterization of ultra-high molar mass
sodium hyaluronate containing low amounts of strongly scattering impurities by dual low angle light scattering/refractometric
detection,” Journal of Liquid Chromatography and Related Technologies, vol. 31, no. 20, pp. 3077–3093, 2008.
[12] A. Shiedlin, R. Bigelow, W. Christopher et al., “Evaluation of
hyaluronan from different sources: streptococcus zooepidemicus, rooster comb, bovine vitreous, and human umbilical cord,”
Biomacromolecules, vol. 5, no. 6, pp. 2122–2127, 2004.
[13] R. Mendichi and L. Soltes, “Hyaluronan molecular weight and
polydispersity in some commercial intra-articular injectable
preparations and in synovial fluid,” Inflammation Research, vol.
51, no. 3, pp. 115–116, 2002.
[14] R. Stern, “Devising a pathway for hyaluronan catabolism: are we
there yet?” Glycobiology, vol. 13, no. 12, pp. 105R–115R, 2003.
[15] M. Erickson and R. Stern, “Chain gangs: new aspects of
hyaluronan metabolism,” Biochemistry Research International,
vol. 2012, Article ID 893947, 9 pages, 2012.
[16] T. C. Laurent and J. R. E. Fraser, “Hyaluronan,” The FASEB
Journal, vol. 6, no. 7, pp. 2397–2404, 1992.
[17] P. L. DeAngelis, “Hyaluronan synthases: Fascinating glycosyltransferases from vertebrates, bacterial pathogens, and algal
viruses,” Cellular and Molecular Life Sciences, vol. 56, no. 7-8,
pp. 670–682, 1999.
[18] T. E. Hardingham and H. Muir, “The specific interaction of
hyaluronic acid with cartilage proteoglycans,” Biochimica et
Biophysica Acta, vol. 279, no. 2, pp. 401–405, 1972.
[19] V. C. Hascall, “Hyaluronan, a common thread,” Glycoconjugate
Journal, vol. 17, no. 7–9, pp. 607–616, 2000.
[20] N. Afratis, C. Gialeli, D. Nikitovic et al., “Glycosaminoglycans:
key players in cancer cell biology and treatment,” FEBS Journal,
vol. 279, no. 7, pp. 1177–1197, 2012.
[21] C. Schiraldi, A. La Gatta, and M. De Rosa, “Biotechnological
production and application of hyaluronan,” in Biopolymers, M.
Elnashar, Ed., pp. 387–412, InTech, Rijeka, Croatia, 2010.
[22] F. Freitas, V. D. Alves, and M. A. M. Reis, “Advances in bacterial exopolysaccharides: from production to biotechnological
applications,” Trends in Biotechnology, vol. 29, no. 8, pp. 388–
398, 2011.
[23] B. D. Ulery, L. S. Nair, and C. T. Laurencin, “Biomedical
applications of biodegradable polymers,” Journal of Polymer
Science B, vol. 49, no. 12, pp. 832–864, 2011.
11
[24] R. Stern, A. A. Asari, and K. N. Sugahara, “Hyaluronan
fragments: an information-rich system,” European Journal of
Cell Biology, vol. 85, no. 8, pp. 699–715, 2006.
[25] B. F. Chong, L. M. Blank, R. Mclaughlin, and L. K. Nielsen,
“Microbial hyaluronic acid production,” Applied Microbiology
and Biotechnology, vol. 66, no. 4, pp. 341–351, 2005.
[26] S. Ghatak, S. Misra, and B. P. Toole, “Hyaluronan oligosaccharides inhibit anchorage-independent growth of tumor cells
by suppressing the phosphoinositide 3-kinase/Akt cell survival
pathway,” The Journal of Biological Chemistry, vol. 277, no. 41,
pp. 38013–38020, 2002.
[27] J. Y. Lee and A. P. Spicer, “Hyaluronan: a multifunctional,
megaDalton, stealth molecule,” Current Opinion in Cell Biology,
vol. 12, no. 5, pp. 581–586, 2000.
[28] P. L. DeAngelis and P. H. Weigel, “Immunochemical confirmation of the primary structure of streptococcal hyaluronan
synthase and synthesis of high molecular weight product by the
recombinant enzyme,” Biochemistry, vol. 33, no. 31, pp. 9033–
9039, 1994.
[29] P. Prehm, “Hyaluronate is synthesized at plasma membranes,”
Biochemical Journal, vol. 220, no. 2, pp. 597–600, 1984.
[30] A. A. E. Chavaroche, L. A. M. van den Broek, and G. Eggink,
“Production methods for heparosan, a precursor of heparin and
heparan sulfate,” Carbohydrate Polymers, vol. 93, no. 1, pp. 38–
47, 2013.
[31] N. Itano, “Hyaluronan biosynthesis: a multifaceted process,”
Connective Tissue, vol. 33, no. 3, pp. 221–226, 2001.
[32] C. A. de la Motte and J. A. Drazba, “Viewing hyaluronan:
imaging contributes to imagining new roles for this amazing
matrix polymer,” Journal of Histochemistry and Cytochemistry,
vol. 59, no. 3, pp. 252–257, 2011.
[33] N. Itano and K. Kimata, “Molecular cloning of human hyaluronan synthase,” Biochemical and Biophysical Research Communications, vol. 222, no. 3, pp. 816–820, 1996.
[34] A. P. Spicer, M. L. Augustine, and J. A. McDonald, “Molecular
cloning and characterization of a putative mouse hyaluronan
synthase,” The Journal of Biological Chemistry, vol. 271, no. 38,
pp. 23400–23406, 1996.
[35] N. Itano, T. Sawai, M. Yoshida et al., “Three isoforms of
mammalian hyaluronan synthases have distinct enzymatic
properties,” The Journal of Biological Chemistry, vol. 274, no. 35,
pp. 25085–25092, 1999.
[36] P. Heldin, “Molecular mechanisms that regulate hyaluronan
synthesis,” Gene Therapy and Molecular Biology, vol. 3, pp. 465–
474, 1999.
[37] A. Jacobson, J. Brinck, M. J. Briskin, A. P. Spicer, and P. Heldin,
“Expression of human hyaluronan syntheses in response to
external stimuli,” Biochemical Journal, vol. 348, no. 1, pp. 29–35,
2000.
[38] B. A. Dougherty and I. Van de Rijn, “Molecular characterization
of hasB from an operon required for hyaluronic acid synthesis
in group A streptococci. Demonstration of UDP-glucose dehydrogenase activity,” The Journal of Biological Chemistry, vol. 268,
no. 10, pp. 7118–7124, 1993.
[39] B. A. Dougherty and I. Van de Rijn, “Molecular characterization
of hasA from an operon required for hyaluronic acid synthesis
in group A streptococci,” The Journal of Biological Chemistry,
vol. 269, no. 1, pp. 169–175, 1994.
[40] D. L. Crater, B. A. Dougherty, and I. Van de Rijn, “Molecular characterization of hasC from an operon required for
12
[41]
[42]
[43]
[44]
[45]
[46]
[47]
[48]
[49]
[50]
[51]
[52]
[53]
[54]
[55]
[56]
International Journal of Carbohydrate Chemistry
hyaluronic acid synthesis in group A streptococci. Demonstration of UDP-glucose pyrophosphorylase activity,” The Journal of
Biological Chemistry, vol. 270, no. 48, pp. 28676–28680, 1995.
A. Markovitz, J. A. Cifonelli, and A. Dorfman, “The biosynthesis
of hyaluronic acid by group A Streptococcus. VI. Biosynthesis
from uridine nucleotides in cell-free extracts,” The Journal of
Biological Chemistry, vol. 234, pp. 2343–2350, 1959.
K. Kumari and P. H. Weigel, “Molecular cloning, expression,
and characterization of the authentic hyaluronan synthase from
Group C Streptococcus equisimilis,” The Journal of Biological
Chemistry, vol. 272, no. 51, pp. 32539–32546, 1997.
P. L. DeAngelis and A. M. Achyuthan, “Yeast-derived recombinant DG42 protein of Xenopus can synthesize hyaluronan in
vitro,” The Journal of Biological Chemistry, vol. 271, no. 39, pp.
23657–23660, 1996.
P. H. Weigel, V. C. Hascall, and M. Tammi, “Hyaluronan
synthases,” The Journal of Biological Chemistry, vol. 272, no. 22,
pp. 13997–14000, 1997.
P. L. DeAngelis, W. Jing, M. V. Graves, D. E. Burbank, and J.
L. Van Etten, “Hyaluronan synthase of chlorella virus PBCV-1,”
Science, vol. 278, no. 5344, pp. 1800–1803, 1997.
P. M. Coutinho, E. Deleury, G. J. Davies, and B. Henrissat,
“An evolving hierarchical family classification for glycosyltransferases,” Journal of Molecular Biology, vol. 328, no. 2, pp. 307–317,
2003.
A. Jong, C. H. Wu, H. M. Chen et al., “Identification and characterization of CPS1 as a hyaluronic acid synthase contributing
to the pathogenesis of Cryptococcus neoformans infection,”
Eukaryotic Cell, vol. 6, no. 8, pp. 1486–1496, 2007.
P. L. DeAngelis, W. Jing, R. R. Drake, and A. M. Achyuthan,
“Identification and molecular cloning of a unique hyaluronan
synthase from Pasteurella multocida,” The Journal of Biological
Chemistry, vol. 273, no. 14, pp. 8454–8458, 1998.
P. H. Weigel and P. L. DeAngelis, “Hyaluronan synthases:
a decade-plus of novel glycosyltransferases,” The Journal of
Biological Chemistry, vol. 282, no. 51, pp. 36777–36781, 2007.
S. Bodevin-Authelet, M. Kusche-Gullberg, P. E. Pummill, P. L.
DeAngelis, and U. Lindahl, “Biosynthesis of hyaluronan: direction of chain elongation,” The Journal of Biological Chemistry,
vol. 280, no. 10, pp. 8813–8818, 2005.
P. E. Pummill, T. A. Kane, E. S. Kempner, and P. L. DeAngelis,
“The functional molecular mass of the Pasteurella hyaluronan
synthase is a monomer,” Biochimica et Biophysica Acta, vol. 1770,
no. 2, pp. 286–290, 2007.
P. L. DeAngelis, “Microbial glycosaminoglycan glycosyltransferases,” Glycobiology, vol. 12, no. 1, pp. 9R–16R, 2002.
K. Rilla, H. Siiskonen, A. P. Spicer, J. M. T. Hyttinen, M. I.
Tammi, and R. H. Tammi, “Plasma membrane residence of
hyaluronan synthase is coupled to its enzymatic activity,” The
Journal of Biological Chemistry, vol. 280, no. 36, pp. 31890–31897,
2005.
M. Nardini, M. Ori, D. Vigetti, R. Gornati, I. Nardi, and R.
Perris, “Regulated gene expression of hyaluronan synthases
during Xenopus laevis development,” Gene Expression Patterns,
vol. 4, no. 3, pp. 303–308, 2004.
J. Y. L. Tien and A. P. Spicer, “Three vertebrate hyaluronan
synthases are expressed during mouse development in distinct
spatial and temporal patterns,” Developmental Dynamics, vol.
233, no. 1, pp. 130–141, 2005.
M. Suzuki, T. Asplund, H. Yamashita, C. H. Heldin, and P.
Heldin, “Stimulation of hyaluronan biosynthesis by plateletderived growth factor-BB and transforming growth factor-𝛽1
[57]
[58]
[59]
[60]
[61]
[62]
[63]
[64]
[65]
[66]
[67]
[68]
[69]
[70]
[71]
involves activation of protein kinase C,” Biochemical Journal,
vol. 307, no. 3, pp. 817–821, 1995.
H. Chao and A. P. Spicer, “Natural antisense mRNAs to
hyaluronan synthase 2 inhibit hyaluronan biosynthesis and cell
proliferation,” The Journal of Biological Chemistry, vol. 280, no.
30, pp. 27513–27522, 2005.
D. Vigetti, A. Genasetti, E. Karousou et al., “Modulation of
hyaluronan synthase activity in cellular membrane fractions,”
The Journal of Biological Chemistry, vol. 284, no. 44, pp. 30684–
30694, 2009.
L. M. Blank, P. Hugenholtz, and L. K. Nielsen, “Evolution of
the hyaluronic acid synthesis (has) operon in Streptococcus
zooepidemicus and other pathogenic streptococci,” Journal of
Molecular Evolution, vol. 67, no. 1, pp. 13–22, 2008.
W. Y. Chen, E. Marcellin, J. Hung, and L. K. Nielsen,
“Hyaluronan molecular weight is controlled by UDP-Nacetylglucosamine concentration in Streptococcus zooepidemicus,” The Journal of Biological Chemistry, vol. 284, no. 27, pp.
18007–18014, 2009.
L. Soltes, R. Mendichi, D. Lath, M. Mach, and D. Bakoš, “Molecular characteristics of some commercial high-molecular-weight
hyaluronans,” Biomedical Chromatography, vol. 16, no. 7, pp.
459–462, 2002.
P. S. Harmon, E. P. Maziarz, and X. M. Liu, “Detailed characterization of hyaluronan using aqueous size exclusion chromatography with triple detection and multiangle light scattering
detection,” Journal of Biomedical Materials Research B, vol. 100,
no. 7, pp. 1955–1960, 2012.
E. U. Ignatova and A. N. Gurov, “Principles of extraction and
purification of hyaluronic acid,” Khimiko-Farmatsevticheskii
Zhurnal, vol. 24, no. 3, pp. 42–46, 1990.
I. Amagai, Y. Tashiro, and H. Ogawa, “Improvement of the
extraction procedure for hyaluronan from fish eyeball and the
molecular characterization,” Fisheries Science, vol. 75, no. 3, pp.
805–810, 2009.
M. O’Regan, I. Martini, F. Crescenzi, C. De Luca, and M.
Lansing, “Molecular mechanisms and genetics of hyaluronan biosynthesis,” International Journal of Biological Macromolecules, vol. 16, no. 6, pp. 283–286, 1994.
G. Kogan, L. Šoltés, R. Stern, and P. Gemeiner, “Hyaluronic
acid: a natural biopolymer with a broad range of biomedical and
industrial applications,” Biotechnology Letters, vol. 29, no. 1, pp.
17–25, 2007.
J. C. Thonard, S. A. Migliore, and R. Blustein, “Isolation
of hyaluronic acid from broth cultures of Streptococci,” The
Journal of Biological Chemistry, vol. 239, pp. 726–728, 1964.
F. E. Kendall, M. Heidelberger, and M. H. Dawson, “A serologically inactive polysaccharide elaborated by mocoid strains
of group A hemolytic Streptococcus,” The Journal of Biological
Chemistry, vol. 118, no. 1, pp. 61–69, 1937.
B. Holmstrom and J. Ricica, “Production of hyaluronic acid by
a Streptococcal strain in batch culture,” Applied Environmental
Microbiology, vol. 15, no. 6, pp. 1409–1413, 1967.
J. H. Kim, S. J. Yoo, D. K. Oh et al., “Selection of a Streptococcus
equi mutant and optimization of culture conditions for the
production of high molecular weight hyaluronic acid,” Enzyme
and Microbial Technology, vol. 19, no. 6, pp. 440–445, 1996.
N. Izawa, T. Hanamizu, R. Iizuka et al., “Streptococcus thermophilus produces exopolysaccharides including hyaluronic
acid,” Journal of Bioscience and Bioengineering, vol. 107, no. 2,
pp. 119–123, 2009.
International Journal of Carbohydrate Chemistry
[72] N. Izawa, M. Serata, T. Sone, T. Omasa, and H. Ohtake,
“Hyaluronic acid production by recombinant Streptococcus
thermophilus,” Journal of Bioscience and Bioengineering, vol. 111,
no. 6, pp. 665–670, 2011.
[73] H. J. Gao, G. C. Du, and J. Chen, “Analysis of metabolic fluxes
for hyaluronic acid (HA) production by Streptococcus zooepidemicus,” World Journal of Microbiology and Biotechnology, vol.
22, no. 4, pp. 399–408, 2006.
[74] X. J. Duan, H. X. Niu, W. S. Tan, and X. Zhang, “Mechanism
analysis of effect of oxygen on molecular weight of hyaluronic
acid produced by Streptococcus zooepidemicus,” Journal of
Microbiology and Biotechnology, vol. 19, no. 3, pp. 299–306,
2009.
[75] J. Zhang, X. Ding, L. Yang, and Z. Kong, “A serum-free
medium for colony growth and hyaluronic acid production by
Streptococcus zooepidemicus NJUST01,” Applied Microbiology
and Biotechnology, vol. 72, no. 1, pp. 168–172, 2006.
[76] J. H. Im, J. M. Song, J. H. Kang, and D. J. Kang, “Optimization
of medium components for high-molecular-weight hyaluronic
acid production by Streptococcus sp. ID9102 via a statistical
approach,” Journal of Industrial Microbiology and Biotechnology,
vol. 36, no. 11, pp. 1337–1344, 2009.
[77] D. C. Armstrong, M. J. Cooney, and M. R. Johns, “Growth and
amino acid requirements of hyaluronic-acid producing Streptococcus zooepidemicus,” Applied Microbiology and Biotechnology, vol. 47, no. 3, pp. 309–312, 1997.
[78] M. J. Cooney, L. T. Goh, P. L. Lee, and M. R. Johns, “Structured model-based analysis and control of the hyaluronic acid
fermentation by Streptococcus zooepidemicus: physiological
implications of glucose and complex-nitrogen-limited growth,”
Biotechnology Progress, vol. 15, no. 5, pp. 898–910, 1999.
[79] W. C. Huang, S. J. Chen, and T. L. Chen, “The role of
dissolved oxygen and function of agitation in hyaluronic acid
fermentation,” Biochemical Engineering Journal, vol. 32, no. 3,
pp. 239–243, 2006.
[80] B. F. Chong and L. K. Nielsen, “Aerobic cultivation of Streptococcus zooepidemicus and the role of NADH oxidase,”
Biochemical Engineering Journal, vol. 16, no. 2, pp. 153–162, 2003.
[81] B. F. Chong and L. K. Nielsen, “Amplifying the cellular reduction
potential of Streptococcus zooepidemicus,” Journal of Biotechnology, vol. 100, no. 1, pp. 33–41, 2003.
[82] T. F. Wu, W. C. Huang, Y. C. Chen, Y. G. Tsay, and C. S.
Chang, “Proteomic investigation of the impact of oxygen on
the protein profiles of hyaluronic acid-producing Streptococcus
zooepidemicus,” Proteomics, vol. 9, no. 19, pp. 4507–4518, 2009.
[83] M. R. Johns, L. T. Goh, and A. Oeggerli, “Effect of pH, agitation
and aeration on hyaluronic acid production by Streptococcus
zooepidemicus,” Biotechnology Letters, vol. 16, no. 5, pp. 507–
512, 1994.
[84] S. Hasegawa, M. Nagatsuru, M. Shibutani, S. Yamamoto, and S.
Hasebe, “Productivity of concentrated hyaluronic acid using a
Maxblend (R) fermentor,” Journal of Bioscience and Bioengineering, vol. 88, no. 1, pp. 68–71, 1999.
[85] X. Zhang, X. J. Duan, and W. S. Tan, “Mechanism for the effect of
agitation on the molecular weight of hyaluronic acid produced
by Streptococcus zooepidemicus,” Food Chemistry, vol. 119, no.
4, pp. 1643–1646, 2010.
[86] M. Cazzola, F. O’regan, and V. Corsa, “Culture medium and
process for the preparation of high molecular weight hyaluronic
acid,” Fidia Advanced Biopolymers S.R.L., 2003.
[87] I. Van De Rijn, “Streptococcal hyaluronic acid: Proposed mechanisms of degradation and loss of synthesis during stationary
13
[88]
[89]
[90]
[91]
[92]
[93]
[94]
[95]
[96]
[97]
[98]
[99]
[100]
[101]
[102]
[103]
[104]
phase,” Journal of Bacteriology, vol. 156, no. 3, pp. 1059–1065,
1983.
A. Mausolf, J. Jungmann, H. Robenek, and P. Prehm, “Shedding
of hyaluronate synthase from streptococci,” Biochemical Journal, vol. 267, no. 1, pp. 191–196, 1990.
D. C. Ellwood, C. G. T. Evans, G. M. Dunn et al., “Production
of hyaluronic acid,” Fermentech Medical Limited, 1996.
W. C. Huang, S. J. Chen, and T. L. Chen, “Production of
hyaluronic acid by repeated batch fermentation,” Biochemical
Engineering Journal, vol. 40, no. 3, pp. 460–464, 2008.
L. M. Blank, R. L. McLaughlin, and L. K. Nielsen, “Stable
production of hyaluronic acid in streptococcus zooepidemicus
chemostats operated at high dilution rate,” Biotechnology and
Bioengineering, vol. 90, no. 6, pp. 685–693, 2005.
H. Y. Han, S. H. Jang, E. C. Kim et al., “Microorganism producing hyaluronic acid and purification method of hyaluronic
acid,” p. 26, 2004.
S. Stahl, “Methods and means for the production of hyaluronic
acid,” US6090596, 2000.
E. Marcellin, W. Y. Chen, and L. K. Nielsen, “Understanding
plasmid effect on hyaluronic acid molecular weight produced
by Streptococcus equi subsp. zooepidemicus,” Metabolic Engineering, vol. 12, no. 1, pp. 62–69, 2010.
J. Z. Sheng, P. X. Ling, X. Q. Zhu et al., “Use of induction
promoters to regulate hyaluronan synthase and UDP-glucose6-dehydrogenase of Streptococcus zooepidemicus expression
in Lactococcus lactis: a case study of the regulation mechanism
of hyaluronic acid polymer,” Journal of Applied Microbiology,
vol. 107, no. 1, pp. 136–144, 2009.
H. Yu and G. Stephanopoulos, “Metabolic engineering of
Escherichia coli for biosynthesis of hyaluronic acid,” Metabolic
Engineering, vol. 10, no. 1, pp. 24–32, 2008.
Z. Mao, H. D. Shin, and R. Chen, “A recombinant E. coli
bioprocess for hyaluronan synthesis,” Applied Microbiology and
Biotechnology, vol. 84, no. 1, pp. 63–69, 2009.
B. Widner, R. Behr, S. Von Dollen et al., “Hyaluronic acid
production in Bacillus subtilis,” Applied and Environmental
Microbiology, vol. 71, no. 7, pp. 3747–3752, 2005.
Z. Mao and R. R. Chen, “Recombinant synthesis of hyaluronan
by Agrobacterium sp,” Biotechnology Progress, vol. 23, no. 5, pp.
1038–1042, 2007.
S. B. Prasad, G. Jayaraman, and K. B. Ramachandran,
“Hyaluronic acid production is enhanced by the additional coexpression of UDP-glucose pyrophosphorylase in Lactococcus
lactis,” Applied Microbiology and Biotechnology, vol. 86, no. 1, pp.
273–283, 2010.
L. J. Chien and C. K. Lee, “Hyaluronic acid production by
recombinant Lactococcus lactis,” Applied Microbiology and
Biotechnology, vol. 77, no. 2, pp. 339–346, 2007.
S. B. Prasad, K. B. Ramachandran, and G. Jayaraman, “Transcription analysis of hyaluronan biosynthesis genes in Streptococcus zooepidemicus and metabolically engineered Lactococcus lactis,” Applied Microbiology and Biotechnology, vol. 94, no.
6, pp. 1593–1607, 2012.
A. Sloma et al., “Recombinant expression of bacterial hyaluronan synthase operon genes in Bacillus and hyaluronic acid
production,” p. 218, 2003.
M. V. Graves, D. E. Burbank, R. Roth, J. Heuser, P. L. Deangelis,
and J. L. Van Etten, “Hyaluronan synthesis in virus PBCV-1infected chlorella-like green algae,” Virology, vol. 257, no. 1, pp.
15–23, 1999.
14
[105] C. De Luca, M. Lansing, I. Martini et al., “Enzymatic synthesis of
hyaluronic acid with regeneration of sugar nucleotides,” Journal
of the American Chemical Society, vol. 117, no. 21, pp. 5869–5870,
1995.
[106] P. L. DeAngelis, “Monodisperse hyaluronan polymers: synthesis
and potential applications,” Current Pharmaceutical Biotechnology, vol. 9, no. 4, pp. 246–248, 2008.
[107] F. K. Kooy, M. Ma, H. H. Beeftink, G. Eggink, J. Tramper,
and C. G. Boeriu, “Quantification and characterization of
enzymatically produced hyaluronan with fluorophore-assisted
carbohydrate electrophoresis,” Analytical Biochemistry, vol. 384,
no. 2, pp. 329–336, 2009.
[108] Y. I. Shimma, F. Saito, F. Oosawa, and Y. Jigami, “Construction of
a library of human glycosyltransferases immobilized in the cell
wall of Saccharomyces cerevisiae,” Applied and Environmental
Microbiology, vol. 72, no. 11, pp. 7003–7012, 2006.
[109] W. Jing and P. L. De Angelis, “Dissection of the two transferase
activities of the Pasteurella multocida hyaluronan synthase: two
active sites exist in one polypeptide,” Glycobiology, vol. 10, no. 9,
pp. 883–889, 2000.
[110] W. Jing and P. L. DeAngelis, “Analysis of the two active sites
of the hyaluronan synthase and the chondroitin synthase of
Pasteurella multocida,” Glycobiology, vol. 13, no. 10, pp. 661–671,
2003.
[111] S. Milewski, I. Gabriel, and J. Olchowy, “Enzymes of UDPGlcNAc biosynthesis in yeast,” Yeast, vol. 23, no. 1, pp. 1–14, 2006.
[112] H. Zhao and W. A. Van Der Donk, “Regeneration of cofactors
for use in biocatalysis,” Current Opinion in Biotechnology, vol.
14, no. 6, pp. 583–589, 2003.
[113] A. Ruffing and R. R. Chen, “Metabolic engineering of microbes
for oligosaccharide and polysaccharide synthesis,” Microbial
Cell Factories, vol. 5, p. 25, 2006.
[114] W. Liu and P. Wang, “Cofactor regeneration for sustainable
enzymatic biosynthesis,” Biotechnology Advances, vol. 25, no. 4,
pp. 369–384, 2007.
[115] C. A. G. M. Weijers, M. C. R. Franssen, and G. M. Visser,
“Glycosyltransferase-catalyzed synthesis of bioactive oligosaccharides,” Biotechnology Advances, vol. 26, no. 5, pp. 436–456,
2008.
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