Download Genetic engineering of lactic acid bacteria to produce

Genetic engineering of lactic acid bacteria to produce
optically pure lactic acid and to develop a novel cell
immobilization method suitable for industrial fermentations
Kari Kylä-Nikkilä
Doctoral Programme in Microbiology and Biotechnology
Division of Microbiology and Biotechnology
Department of Food and Environmental Sciences
Faculty of Agriculture and Forestry
Division of Microbiology and Epidemiology
Department of Veterinary Biosciences
Faculty of Veterinary Medicine
University of Helsinki
Finland
ACADEMIC DISSERTATION
To be presented, with the permission of the Faculty of Agriculture and Forestry of the
University of Helsinki, for public criticism in the lecture hall 2402 at the University of
Helsinki, Biocenter 3, Viikinkaari 1 on May 27th, 2016 at 12 noon.
Helsinki 2016
Supervisors
Professor Per Saris
Department of Food and Environmental Sciences, University of Helsinki, Finland
Professor Airi Palva
Department of Veterinary Biosciences, University of Helsinki, Finland
Reviewers
Associate Professor Alexander Frey
Department of Biotechnology and Chemical Technology, Aalto University, Finland
Adjunct Professor Ville Santala
Department of Chemistry and Bioengineering, Tampere University of Technology,
Finland
Opponent
Dr. Antti Nyyssölä, Senior Research Scientist,
VTT Technical Research Centre of Finland Ltd., Finland
Custos
Professor Per Saris
Department of Food and Environmental Sciences, University of Helsinki, Finland
ISBN 978-951-51-2209-4 (Paperback)
ISBN 978-951-51-2210-0 (PDF)
ISSN 2342-5423 (Print)
ISSN 2342-5431 (Online)
Unigrafia Oy
Helsinki 2016
Cover page photo:
Trade card for Lactart Milk Acid, Avery Lactate Company,
Boston, Massachusetts, 1884.
To my family
Table of contents
Table of contents
4
List of original publications
6
Abbreviations
8
Abstract
11
1.
13
Introduction
1.1
Lactic acid bacteria – interesting observations from the history
13
1.2
Current definion of LAB
15
1.3
The Gram-positive cell envelope
16
1.3.1 Structure of peptidoglycan backbone and interconnecting peptide
crosslinks
17
1.3.2 Secondary cell wall polymers
18
1.3.3 Lipoteichoic acids
21
1.3.4 S-layers
22
1.4
Cell retention mechanisms for secreted and excreted proteins in LAB
24
1.4.1 Protein secretion mechanisms among LAB
24
1.4.2 Autolysins and excretion of cytosolic proteins
25
1.4.3 Various cell envelope retention mechanism for proteins in LAB
26
1.5
Fermentation pathways in LAB
37
1.5.1 Fermentation and respiration in LAB
37
1.5.2 Typical LAB fermentations
37
1.5.3 Other fates of pyruvate
40
1.5.4 Classification of LAB according to isomer of lactic acid produced
41
1.6
Lactic acid –usage, applications and production
42
1.6.1 Synthesis of PLA
44
1.6.2 Conventional LAB based industrial scale fermentation processes for
lactic acid
45
1.6.3 Modern technologies in industrial bacterial fermentations–
immobilization of cells for extracellular production of metabolites
48
1.7
Genetic engineering of Gram-positive bacteria to achieve whole cell
immobilization with industrial potential
53
1.7.1 Immobilization on collagen surfaces by non-covalent binding
55
1.7.2 Immobilization on starch surfaces by non-covalent binding
55
1.7.3 Immobilization on cellulose surfaces by non-covalent binding
55
4
1.7.4Immobilization on chitin surfaces by non-covalent binding
56
1.7.5 Immobilization to inorganic or synthetic surfaces by non-covalent
binding
57
1.8
Genetic engineering of LAB to improve optical purity of the produced
lactic acid
58
1.8.1 Various biosynthetic routes leading to D(-)- and L(+)-lactic acid
formation in LAB
58
1.8.2 Use of genetic engineering to produce lactic acid with enhanced optical
purity
62
2.
Aims of the study
67
3.
Materials and methods
68
4.
3.1
Plasmids
68
3.2
Bacterial strains, growth conditions and media
69
3.3
Oligonucleotides
71
3.4
Methods used in this study
73
Results and discussion
4.1
76
Production of PLA grade L(+)-lactic acid by genetically modified
homofermentative Lactobacillus strain (publication I)
77
4.1.1 Construction of D-LDH negative L. helveticus strain by inactivation of
ldhD by gene replacement
77
4.1.2 Construction of D-LDH negative L. helveticus strain by replacing the
structural gene of ldhD with that of ldhL
77
4.1.3 Growth and fermentation characteristics of the wild type strain and DLDH negative strains
78
4.2
Development of whole cell immobilization method for LAB by surface
engineering
81
4.2.1 Studying the potential of S-layer protein (SlpA) of L. brevis for anchoring
peptides for cell surface display (publication II)
81
4.2.2 Indicating the potential of whole cell immobilization of L. lactis by cell
surface displayed binding domain (publication III and IV)
83
5.
Conclusions and future prospects
90
6.
Acknowledgements
92
7.
References
93
Publications I-IV
5
List of original publications
This thesis is based on the following publications, which are referred to in the text by their
Roman numerals. The original articles are reprinted with the kind permission of the
publishers. In addition, some unpublished data are presented.
I
Kylä-Nikkilä K., Hujanen, M., Leisola, M. and Palva, A. (2000) Metabolic
engineering of Lactobacillus helveticus CNRZ32 for production of pure L(+)-lactic acid. Applied and Environmental Microbiology, 66(9), 3835-3841.
II
Åvall-Jääskeläinen, S., Kylä-Nikkilä, K., Kahala, M., Miikkulainen-Lahti,
T. and Palva, A. (2002) Surface display of foreign epitopes on the
Lactobacillus brevis S-layer. Applied and Environmental
Microbiology, 68(12), 5943-5951.
III
Kylä-Nikkilä K., Alakuijala U. and Saris P.E.J. (2010) Immobilization of
Lactococcus lactis to cellulosic material by cellulose-binding domain of
Cellvibrio japonicus. Journal of Applied Microbiology, 109(4), 1274-1283.
IV
Liu S., Kylä-Nikkilä K. and Saris P.E.J. (2011) Cell immobilization studies
using a cellulose-binding domain fused to PrtP in Lactococcus lactis.
Bioengineered Bugs, 2(3), 160-162.
The author’s contribution in the above mentioned publications:
Publication I
- construction of the recombinant strains and verification of the resulted strains
(except sequencing)
- enzymatic assays
- transcriptional analysis
- Biostat B2 fermentations
- interpretation of the results and writing the publication in collaboration with the
other authors
Publication II
- construction of pKTH5006, pKTH5007 and pKTH5008 plasmids
- transformation of these plasmids to Lactococcus lactis and Lactobacillus brevis in
co-operation with the 1st author
- detection of the surface exposure of the VP1 epitope by whole cell ELISA in the
Lactobacillus brevis strains carrying the above mentioned plasmids in cooperation with the 1st author
6
Publication III
- all genetic constructs
- major part of the adhesion testing and the method development
- major responsibility in the interpretation of the results and writing the publication
in collaboration with the other authors
Publication IV
- all genetic constructs
- developing the adhesion testing method
- minor contribution in the interpretation of the results and writing the publication
7
Abbreviations
ABC
CBD
CBM
ChBD
CPS
CWBD
DEAE
DHAP
ECM
ECP
EFSA
EPS
FDP (or FBP)
FEA
FPE
GAP
GAPDH
GLP
HePS
HicDH
HoPS
iLDH
LAB
LDH
LTA
MCS
MG
MGS
MLF
nLDH
PDLA
PET
PG
PLA
PLLA
PS
PTS
ATP binding cassette
cellulose-binding domain
cellulose- binding module
chitin-binding domain
capsular polysaccharide
cell wall binding domain
diethylaminoethyl
dihydroxyacetone phosphate
extracellular matrix
excretion of cytosolic enzymes
European food safety authority
exopolysaccharide (sugar)
fructose-1,6-diphosphate (or fructose-1,6-biphosphate)
flagella export apparatus
fimbrilin-protein exporter (or pseudopilin export pathway)
glyceraldehyde-3-phosphate
glyceraldehyde-3-phosphate dehydrogenase
glycerol intrinsic proteins (glycerol facilitators)
heteropolysaccharides
2-hydroxyisocapronate dehydrogenase
homopolysaccharides
NAD-independent LDH
lactic acid bacteria
lactate dehydrogenase enzyme
lipoteichoic acid
multiple cloning site
methylglyoxal
methylglyoxal synthase
malolactic fermentation
NAD dependent LDH
poly(D-lactic acid)
polyethylene terephthalate
peptidoglycan layer
poly(lactic acid)
poly(L-lactic acid)
polystyrene
phosphoenolpyruvate-dependet phosphotransferase system
8
QPS
R-PCR
S-layer
SBD
scPLA
SCWP
Sec pathway
SLH
TA
Tat
TMD
TMS
TUA
WPS
Wss
WTA
qualified presumption of safety
recombinant PCR
surface layer
starch-binding domain
stereocomplex PLA
secondary cell wall polymers
the general secretory pathway
surface layer homology domain
teichoic acid
twin-arginine translocation pathway
transmembrane domain
transmembrane segment
teichuronic acid
cell wall polysaccharides
WXG100 secretion system
wall teichoic acid
9
10
Abstract
In this study Lactobacillus helveticus CNRZ32 was genetically engineered to produce
pure L(+)-lactic acid. Naturally, this organism produces mixture of D(-)- and L(+)-lactic
acid during fermentation of sugars. Production of D(-)- and L(+)-lactic acid is based on
the activity of LdhD and LdhL, respectively. In this work, LdhD enzyme activity was
removed by using two different genetic engineering approaches. In the first approach,
ldhD gene expression and thus the LdhD activity was prevented by deleting the ldhD
promoter region. In the second approach, the structural gene of ldhD was replaced with
that of ldhL of CNRZ32. In both cases, a stable chromosomal mutant, neither requiring
any additional selection for maintenance of the mutation nor carrying any foreign DNA in
its genome, was achieved. Based on the fermentation tests, these two strains were
demonstrated to be potential candidates for production of L(+)-lactic acid for the
manufacturing of high melting temperature biodegradable poly(lactic acid) based plastics,
as the optical purity of the L(+)-lactic acid produced by these strains exceeded the criteria
(optical purity > 95 %) recommended for this process.
Also, the surface display and anchoring of heterologous peptides and cellulose binding
domain were studied with Lactobacillus brevis and Lactococcus lactis, respectively. In the
Lactobacillus brevis study, the main interest was to demonstrate the surface display of
poliovirus epitope VP1 by using the L. brevis S-layer as the anchoring system. In
Lactococcus lactis, different types of anchors were studied to facilitate the whole cell
immobilization of Lactococcus lactis with unmodified cheap cellulosic carrier. Most
typically, continuous fermentation with immobilized micro-organism is the preferred
choice, whenever possible, to run an industrial-scale fermentation in which the product is
secreted to the surrounding medium. In this work, the whole cell immobilization of
lactococcal cells on chemically unmodified cellulosic material was demonstrated by using
two different cell wall anchors in combination with cellulose binding domain of Cellvibrio
japonicus.
11
12
1.
Introduction
1.1 Lactic acid bacteria – interesting observations from the
history
Lactic acid bacteria (hereafter referred as LAB), plants and animals (including also Homo
sapiens) share a long and common history and interdependent relationship. The Neolithic
revolution, so transition from nomadic way of living (hunting, gathering) to more
sedentary way of living with agriculture and pastoralism, seems also to be a starting point
for the development of many food fermentations (Teuber, 2000; Douillard and de Vos,
2014). Old evidences found in Mesopotamia supports that dairying was highly developed
already about 6000 B.C. (Prajapati and Nair, 2008). Furthermore, from the era right after
the invention of writing (about 3200 B.C by Sumerians of Mesopotamia), also written
evidences have been found that contains signs for dairy products, such as milk, butter, fat,
and cheese (Teuber, 2000).
About five thousands years after (1780) from the first documented evidences of food
fermentations, Swedish chemist Carl Wilhelm Steele refined and studied lactic acid from
sour milk (Teuber, 1993). About three decades later, 1808, another Swedish scientist, Jöns
Jacob Berzelius found lactic acid in meat and other animal products (Benninga, 1990).
One motive for his research with lactic acid was to prove its individual nature – at that
time especially some French chemists were unwilling to acknowledge the individuality of
this acid discovered by Berzelius famous fellow countryman, Scheele (Jorpes, 1970).
Indeed, this individuality question remained open for a long time. In 1833, French
scientists Gay-Lussac and Pelouzed tackled to this same question in their work and made
and investigated 15 different salt of lactic acid. During these experiments, they found
sublimation of pure lactic acid and found that the sublimate had almost completely lost its
acid taste but regains acidic properties after prolonged boiling in water. Indeed, this was
the first document record of a polymeric derivate of lactic acid (polylactide), although
these scientists did not realize that in their time (Benninga, 1990). About 100 years later
(1932), a process to polymerize lactide (cyclic di-ester of lactic acid) was developed and
later patented by DuPont (1954) (Auras et al., 2010). However, it took several decades
before the first large-scale production of this new biodegradable plastic started (Taskila
and Ojamo, 2013).
In 1848 Engelhardt noticed in his study with calcium and zink salts, that the racemic
lactate from sour milk and lactate originated from muscle had different solubility, water of
crystallization, and decomposition points. Later, the concept of isomerism in lactate was
introduced in 1873 by Wislicenus after he evaluated the optical rotation of the various
lactate salts and concluded that structure of lactic acid needs to be Į-hydroxypropionic
acid in order to comply with isomerism property of lactic acid molecule (Benninga, 1990).
13
He also stated that this geometrical isomerism must somehow be explained by “the
different arrangements of their atoms in space”, and his discoveries were also promoting
the invention of “asymmetric carbon atom” by another scientist, van Hoff (1874) (Rocke,
2010).
In the middle of 19th century, Louis Pasteur, another scientist with passion to study crystal
structures of compounds and their optical activity, got interested about fermentations.
Pasteur was convinced that only living processes could produce optically active
compounds (Simon, 2013). He also started to study the spoilage of wine and beer and
discovered new form of yeast that was responsible of the sour taste in beverages. In 1857
Pasteur coined this yeast as lactic yeast, because it had a different size and shape when
compared to commonly known brewer’s yeast and the yeast involved in alcoholic
fermentation. In addition, Pasteur’s studies with alcoholic and lactic fermentations
confirmed his original hypothesis - ‘the ferment is life’, meaning that the fermentation
takes place due to the activity of living organisms (Debré, 1998; Barnett, 2003). Later
(1873), Lister was the first scientist who isolated a pure culture of bacteria by using liquid
laboratory techniques. He coined this species as Bacterium lactis (currently known as
Lactococcus lactis). In this work, the original driving force for Pasteur was to model the
theory that infectious diseases of humans are the result of the growth of specific living
microscopic organisms in the human host. To support this theory, he constructed a model
system for which he isolated pure culture of bacterium from milk sample by dilution
method and inoculated that into pre-heated milk, resulting in lactic acid fermentation
(Santer, 2010).
Industrial scale lactic acid production started in 1883 in USA by Avery Lactate Co.
(Boston) and couple of years later (1885) by Albert Boehringer Chemische Fabrik in
Ingelheim, Germany (van Velthuijsen, 1994; Benninga, 1990). At that time, most of the
lactic acid produced was used by dyeing, leather, textile, and food industry (Anonymous,
2016b). As reviewed by Josephsen and Jespersen (2006), commercial production of
various starter cultures was initiated during the late 1880s by the Danish pharmacist Chr.
D.A. Hansen and the use of dairy pure cultures in the milk fermentations began in late 19th
century almost simultaneously in Denmark, Germany and United States. As reviewed by
Porto de Souza Vandenberghe et al. (2013), in the long run the progress with the dairy
starter cultures contributed to the development of the whole dairy chain and led to rise of
various specified standard products to fulfill different demands of market.
Interestingly, association of lactic acid fermentation products to beneficial health impacts
was used in marketing already in the early days of industrial production of lactic acid (see
cover page picture: Trade card for Lactart Milk Acid, Avery Lactate Company, Boston,
Mass., 1884). Couple of decades later, more scientific publications came out from
scientist originated from Russia, Elie Metchnikoff. During the first decade of new century,
he theorized that the human health could be enhanced and also senility delayd, by
manipulating the intestinal microbiome with host-friendly bacteria found in yogurt. One of
the things behind this theory was his observations that long-lived Bulgarian citizens
consumed Lactobacillus-fermented dairy products (Mackowiak 2013; Bibel 1988). His
theories got a lot of publicity and his texts were also sometime exaggerated in different
public papers or magazines – like using phrase ‘prolongation of life’ in headlines. As
reviewed by Bibel (1988), this kind popularization of science also effected to the growing
demand for different kind lactobacillus cultures and fermented milk products in the early
14
20th century. Notably, this early enthusiasm towards fermented milk products allowed the
word yoghourt to enter in the common language (Baglio, 2014).
Metchnikoff’s hypothesis for the connection between senility and intestinal flora must
have gained a lot of scientific attention in this era, because still four decades later (1949),
famous Danish scientist Orla-Jensen in his late days re-visited this theory with his
colleagues and found that it is not the undesirable flora found in the aged which causes
impaired digestion, but rather the impaired digestion resulting in undesirable intestinal
flora (Olsen, 1950). However, in the scientific literature Orla-Jensen is best known from
his work for classification of lactic acid bacteria. As reviewed by his contemporary
(Heineman, 1920) and later by Murray and Holt (2006), he made an extensive study with
various types of lactic acid-producing bacteria and delimited genera and species on the
basis of characteristics such as fermentation of various sugars, by-products from these
fermentations, morphology and temperature ranges for growth. In his work published in
1942, he described LAB as Gram-positive, non-motile, non-spore-forming, rod- or
coccus-shaped organisms that ferment carbohydrates and higher alcohols to form mainly
lactic acid (Franz and Holzapfel, 2011). Orla-Jensen’s work with classification was farreaching, because his basic scheme for the classification remained useful for a long time.
After the 2nd World War and the development of modern biochemical and molecular
methods, it was found out that this traditional indentification scheme was not anymore
completely correlating with phylogenetic relationships learned by using these new tools
(Stackebrandt and Teuber, 1988). Pot and Tsakalidou (2009) have reviewed the
development of the classification of the lactic acid bacteria (and especially that of the
genus Lactobacillus) by reviewing the most popular phenotypic fermentation
characteristics that were in use for the classification of LAB before the development of
modern genotyping tools and the current polyphasic approach for the taxonomic
classification of LAB.
1.2 Current definion of LAB
Although no unequivocal definition exists for LAB, some usual characteristics for a
typical member of LAB can be listed (Axelsson, 2004). The general description for lactic
acid bacteria refers typically to group of non-sporulating, Gram-positive bacteria which
are devoid of functional cytochromes, non-respiring but aerotolerant anaerobes, catalasenegative (some may have pseudo-catalase), acid tolerant fastidious cocci, coccobacilli or
rods with low G+ C content of DNA (less than 50 mol %) and which produce lactic acid
as the major end product during the fermentation of carbohydrates (Axelsson, 2004;
Vandamme et al., 2014).
Due to their fastidious nature and other typical characteristics, LAB are usually associated
with nutrient rich niches like plant and animal raw materials, environments and products
in which fermentation of these materials occurs. Also, some species of LAB may occur in
the respiratory, intestinal and genital tracts of humans and animals (Giraffa, 2014).
The traditional definition of LAB is not a taxonomic one, but rather a definition for
functional characteristics that food microbiologists have used to describe harmless bacteria
15
that produce lactic acid and which occurs spontaneously in traditional lactic acid
fermented foods (Molin, 2008). As presented by Axelsson (2004), LAB comprise around
20 genera, out of which the main genera of food related, ‘principal’, LAB are: Aerococcus,
Carnobacterium, Enterococcus, Lactobacillus, Lactococcus, Leuconostoc, Oenococcus,
Pediococcus, Streptococcus, Tetragenococcus, Vagococcus and Weissella. As summarized
in more recent review by Vandamme et al. (2014), the genuine members of LAB belong to
the order Lactobacillales, in class Bacilli in phylum Firmicutes. Under this order, there are
six families: Aerococcaceae, Carnobacteriaceae, Enterococcaceae, Lactobacillaceae,
Leuconostocaceae and Streptococcaceae. Taxonomy under these families is evolving
constantly – in 2014, total 38 genera and about 400 species were classified under the order
Lactobacillales.
1.3 The Gram-positive cell envelope
In general, Gram-positive bacteria carry only one membrane (plasma membrane) around
the cell, but have a relatively thick peptidoglycan layer (30-100 nm) covering the plasma
membrane. Instead, Gram-negative bacteria have relative thin peptidoglycan layer, but
these organisms possess an additional membrane (outer membrane) on top of the
peptidoglycan layer. In some cases, bacteria may also carry an S-layer, a monomolecular
layer composed of identical subunits, which shows as the outermost layer in the cell
envelope. The combined structure that is formed by the above mentioned cellular
components is known as cell envelope. The major structures in the envelope,
membrane(s) and peptidoglycan layer, are decorated with various molecules, like proteins,
polymers and other macromolecules, which are important contributors not only for the
envelope structure and function (Silhavy et al., 2010), but ultimately for the survival of the
whole cell, too. Also, S-layers may function as a scaffold to display functional proteins
and/or carbohydrate moieties on the cell surface (Fagan and Fairweather, 2014; Schuster
and Sleytr, 2015). An artist’s view of the cell envelope of a Gram-positive bacterium is
presented in Fig. 1.
The main function of the peptidoglycan layer (or peptidoglycan macromolecule) is to
maintain cell integrity by withstanding the internal osmotic pressure. It has also important
role in maintaining the defined cell structure and it is intimately involved in the cell
division process (Nanninga, 1998). This peptidoglycan (PG) layer (sacculus) formed
around the cell is the structural equivalent of the exoskeleton of insects. Actually, one of
the best term to describe the PG layer is that of a fisherman’s net - so a structure that
wraps the bacterial cellular contents.
16
Figure 1. Simplified view of the cell envelope components of a Gram-positive bacterium
(modified Delcour et al., 1999). The cytoplasmic membrane is surrounded by mutilayered
peptidoglycan (sacculus), decorated with neutral polysaccharides, lipoteichoic acids and
wall teichoic acids. Peptidoglycan layers are covalently staggered by radial halfcrosslinks. Lipoteichoic acids are anchored into cytoplasmic membrane via glycolipid
anchor. For some Gram-positive bacteria, the peptidoglycan layer is covered by S-layer
proteins.
In this net structure, the mesh is formed by two segments of parallel, rather inextensible
glycan threads, held together by two small elastic peptide crosslinks. These crosslinks
allow the net to expand or shrink. In Gram-positive bacteria, several concentric PG layers
are covalently staggered by radial half-crosslinks, expanding the 3D architecture of the
peptidoglycan.
1.3.1 Structure of peptidoglycan backbone and interconnecting peptide
crosslinks
In LAB, like in all eubacteria, the glycan thread in the peptidoglycan structure is a
polymer of the disaccharide N-acetyl-glucosamine-ȕ(1ĺ4)-N-acetyl-muramic acid, so it
has alternating N-acetyl-muramic acid (MurNAc) and N-acetyl-glucosamine subunits
(GlcNAc) (Delcour et al., 1999). There are only minor variations in the chemistry of the
glycan chains among all the bacteria. Instead, more considerable variation is found in the
composition of stem peptides which are linked to the carboxyl group of the MurNac
(Scheffers and Pinho, 2005). Typically, the consensus amino acid sequence of this stem
peptide among LAB is L-Ala-Ȗ-D-Glu-X-D-Ala (see Fig. 2). The third amino acid (X) is a
di-amino acid, most often L-Lys (like in L. lactis and most lactobacilli) but can also be
mesodiaminopimelic acid (mDAP) or lornithine (Chapot-Chartier and Kulakauskas,
2014). Interestingly, although the occurrence of amino acids with the D configuration
(like those found in stem consensus sequence) in nature is quite rare, review published by
Martínez-Rodríguez et al. (2010) reveals that D-amino acids are often, but not always,
17
found in various self-defence related molecules, like those of antibiotics, venoms,
bacteriocins or defensive structures like cell wall peptidoglycan layer. Another interesting
discovery related to the structure of stem peptides of some LAB, is the presence of D-Lac
as the last residue in the stem peptide among the vancomycin resistance LAB species, like
Lactobacillus plantarum or Lactobacillus casei. As pointed out by Chapot-Chartier and
Kulakauskas (2014), most often D-Ala predominates at this position in the newly
synthesized PG of a typical (and vancomycin sensitive) LAB strain.
Variation exists also in the interpeptide bridge connecting the stem peptides elongating
from different glycan strands. Quite often, the interpeptide bridge is composed out of
single D-amino acid (most typically D-Asp or D-Asn), as it the case with many
lactobacilli (Chapot-Chartier and Kulakauskas, 2014), but it is also possible that it
contains several L-amino acids as is the case with Streptococcus thermophilus and with
many other streptococci, too, as reported by Schleifer and Kandler (1972). Indeed, these
two researchers (1972) developed specific nomenclature and rules to classify different
variations found in the PG structure. According this system, the primary structure of PG is
divided into two main classes depending on the anchorage point of the crossbridge in the
stem peptide, and further divided into several subclasses depending on the presence/type
of the connecting interpeptide bridge and the amino acid in the third position of the stem
peptide (Šimelyte et al., 2003).
Figure 2. Schematic structure of peptidoglycan typically found among many LAB
presented with some possible variations (modified from Chapot-Chartier, 2014).
1.3.2 Secondary cell wall polymers
According to classification developed during 1980s, the cell wall polysaccharides of
Gram-positive organisms can be classified under three distinct structural related groups:
(i) teichoic acids, (ii) teichuronic acids, and (iii) other neutral or acidic polysaccharides not
classified in the former groups (Schäffer and Messner 2005, Araki ja Ito 1989, Munson
and Glaser, 1981). As all of these compounds are not primarily related to the main
18
function of the cell wall, the term of secondary cell wall polymers (SCWP) has been
assigned to them. The first two groups of these polysaccharides are often referred as
‘classical’ SCWPs. Although, the exact biological functions of these ‘classical’ SCWPs
are not fully understood, several different functions have been attributed to them, like the
following ones reported by Schäffer and Messner (2005):
(i)
(ii)
(iii)
(iv)
(v)
(vi)
(vii)
binding of divalent cations
role in the balance of metal ions for membrane functionality
binding of proteins
role in folding of extracellular metallo-proteins
providing a source of phosphate under phosphate starvation
conditions
interaction with cell wall lytic enzymes
formation of a barrier to prevent diffusion of nutrients and
metabolites
More information for the biological functions of various cell wall glycopolymers in LAB
is available in the recent review by Chapot-Chartier and Kulakauskas (2014). In the
following section the main focus is on introducing the prevalence and basic structure of
these molecules among LAB.
1.3.2.1 Classical secondary cell wall polymers
1.3.2.1.1 Teichoic acids
Different polyanionic cell wall carbohydrates are found to be important determinants to
bacterial electrostatic charge and hydrophobicity and play pivotal role in bacterial
attachment to abiotic surfaces and eukaryotic cells (Theilacker and Hübner, 2009). For
Gram-positive bacteria, teichoic acids (TAs) represent the most abundant polyanionic
compounds in the cell envelope (Fischer, 1994). The polyanionic characteristic of TAs is
due to the negatively charged phosphodiester bonds located in the polyol-phosphate
backbone of these molecules (Theilacker and Hübner, 2009). Indeed, most teichoic acids
have also zwitterionic properties. This is due to the presence of negatively charged
phosphate groups and modifications with free amino groups that are contained in residues
such as D-alanine (Neuhaus and Baddiley, 2003; Fischer, 1997; Weidenmaier and Peschel,
2008). The cell envelope related TAs are usually divided into two different categories:
wall teichoic acids (WTA) and lipoteichoic acids (LTA). As lipoteichoic acids are not cell
wall bound but attached to the cell membrane via glycolipid anchor, those will be
introduced in more detail in other section in this work.
WTAs are covalently bound to peptidoglycan. Most commonly, the structure of WTA is
composed by glycerol or ribitol groups that are connected by phosphodiester bonds. More
complex repeating units, consisting of trioses or hexoses or glycosylpolyolphosphate
monomers, exists also, e.g in Streptococcus pneumoniae and in Enterococcus faecalis
(Fischer et al., 1993; Theilacker and Hübner, 2009). WTAs are not found in every species;
L. casei and Lactoccus lactis subsp. cremoris being known examples among LAB that
19
lack these compounds (Delcour et al., 1999). However, in some bacterial species WTAs
may constitute up to half of cell wall total dry weight (Weidenmaier and Peschel, 2008;
Chapot-Chartier and Kulakauskas, 2014).
1.3.2.1.2 Teichuronic acids
Many Gram-positive bacteria produce other polyanionic cell wall glycopolymers which
lack phosphate groups in their polymer backbone and are not, therefore, classified as TAs.
One compound belonging to this group is teichuronic acid (TUA) (Weidenmaier and
Peschel, 2008). Structurally TUAs are consisted of glycosidically linked sugar monomers
and uronic acid residues. The negative charge of TUA is due to the carboxyl groups of the
uronic acid residues (Delcour et al., 1999). If grown under phosphate starvation some
species like Bacillus subtilis replaces the phosphate containing WTA with TUA (Neuhaus
and Baddiley, 2003). Furthermore, under these conditions a phosphate loss from the walls
has been detected, proposing the usage of the WTA bound phosphate for the necessary
cell metabolism (Grant, 1979). It is speculated that this is an indication from the
physiological need to have an anionic polymer present in the cell wall (Neuhaus and
Baddiley, 2003). The interdependency between WTA and teichuronic acid synthesis
observed by Grant (1979), seems to ensure a constant level of anionic cell wall charge and
reserve phosphate source for the cell. So far, the presence of cell wall related TUA have
not been described for any LAB. However, as the phylogenetic distance between species
of the genera Bacillus and LAB is rather short, the existence of teichuronic acids in the PG
is possible in LAB, too (Delcour et al., 1999). In fact, part of the genes needed for TUA
synthesis were detected during a whole genome sequencing project of L. lactis (Bolotin et
al., 2001). Also, according to Poxton (2014) the presence of teichuronic acid in
Streptococcus bovis has been reported in early 1970s.
1.3.2.2 Other secondary cell wall related polymers
The last third group of SCWPs (so the group ‘others’) contains different neutral and acidic
polysaccharides. This group is usually divided into two other subclasses: capsular
polysaccharides (CPSs), which form a thick outermost shell around the bacteria and are
often covalently bound to the cell wall, and sensu stricto group of cell wall
polysaccharides (WPS), which decorate the envelope and are either covalently bound or
loosely associated with the peptidoglycan (Delcour et el., 1999).
Some LAB strains, like L. lactis and Streptococcus agalactiae, are covered by pellicle.
Pellicles are polysaccharide structures covalently linked to peptidoglycan and are therefore
qualified as WPS (Chapot-Chartier et al., 2010; Bessaurt et al., 2014). In L. lactis this
structure has predicted to confer a protective barrier against host phagocytosis by murine
macrophages whereas the pellicle detected in S. agalactiae seems to allow access to
underlying peptidoglycan. Interestingly, in some lactococci these pellicle polysaccharides
seem to act as a receptor for certain phages of this species, too (Mahony et al., 2013). In
general, as reviewed by Chapot-Chartier (2014), WPS in LAB appear to be omnipresent
components of the cell surfaces of these organisms and seems to display high structural
variety even in a strain level.
20
As mentioned above, CPSs are typically bound covalently to PG and form a thick outer
layer surrounding the cell. In addition to these polymers, there is distinct group of bacterial
polysaccharides found outside the cell, exopolysaccharides (EPSs), which do not
permanently remain attached to cell walls but remain loosely related to the cell or are
released into the medium as a slime (Sutherland, 1972). Differentiation and proper
nomenclature between CPS and EPS may sometimes be rather complicated, as it is
possible under specific conditions that CPSs are released in the growth medium and EPS
are associated with the cell surface (Cescutti, 2009). Also, in many occasions microbial
exopolysaccharides (EPS) are referred to be either capsular or slime associated (de Vuyst
and Degeest, 1999), which further blur the border between CPS and EPS.
According to Chapot-Chartier (2014), members of all above mentioned cell wall
polysaccharides (WPS, CPS and EPS) may be produced by the same bacterium, although
it may be sometime difficult to make difference between the members of these three
groups. Most often, these polysaccharides are neutral, but some may be acidic due the
branching with anionic substituents (Delcour et al., 1999; Kleerebezem et al., 2010).
Structurally, CPS and EPS can be divided into homopolysaccharides (HoPS) and
heteropolysaccharides (HePS), depending whether the structure in their repeating unit is
composed of polymers with single or multiple type of monosaccharides, respectively
(Cescutti, 2009; Ryan et al., 2015). Because of their abundant presence on the outer
surface of the cell wall, it is expected that extracellular and cell-wall associated
polysaccharides determine to a large extent the surface properties of microorganisms
(Schaer-Zammaretti and Ubbink, 2003). Production of HoPS is typical for many
Streptococcus, Leuconostoc and Pediococcus strains, whereas HePS are typically
produced by mesophilic or thermophilic LAB strains that are often utilized in dairy
industry, where the EPS producing ‘‘ropy’’ strains are important in developing the proper
consistency of fermented milks and yoghurts (de Vuyst and Degeest, 1999, Ruas-Madiedo
and Reyes-Gavilán, 2005). In addition to beneficial properties in food industry, it seems
that microbial EPS possess promising therapeutic potential in terms of anti-oxidation,
hypocholesterolemia, immunomodulation and promotion of a functional digestive tract
through prebiotic activity (Ryan et al., 2015).
Interestingly, Schäffer and Messner (2005) proposed that the glycoconjugates that mediate
the non-covalent attachment of some glycosylated S-layer proteins to the underlying cell
wall in Bacillaceae should be classified under the ‘other’- group of secondary cell wall
polymers. Furthermore, authors coined this type of SCWPs as ‘non-classical’ SCWPs to
differentiate those from teichoic acids and teichuronic acids (classical SCWPs). Also the
same authors divided the ’non-classical’ SCWPs into three groups according to common
features found among the representatives of these groups.
1.3.3 Lipoteichoic acids
Interestingly, the most common type of membrane-anchored anionic polymers,
lipoteichoic acids (LTAs), were first extracted from a lactic acid bacterium strain, namely
Lactobacillus fermentum NCTC6991 (formerly Lactobacillus fermenti NCTC6991)
(Wicken and Knox 1970; Delcour et al., 1999). These membrane bound polymers are
21
usually constituted of glycerol-phosphate repeating units connected to glycolipid and are
usually less diverse in structure than cell wall glycopolymers due to their peculiar
biosynthetic pathway (Weidenmaier and Peschel, 2008). However, more complex
structures have been presented for some strains, including Lactococcus garviae as an
example among LAB (Greenberg et al., 1996). In LAB, LTAs have been detected at least
in enterococci, lactobacilli, lactococci, leuconostocci, and streptococci (Fischer, 1994). In
fact, LTAs are quite abundant in some LAB species. It has been estimated that glycolipid
anchor of LTAs in L. lactis constitutes every fifth lipid molecule of the outer leaflet on the
cytoplasmic membrane of this strain (Fischer, 1981).
1.3.4 S-layers
S-layers are monomolecular isoporous crystalline lattice layers, that are usually found as
the most outermost cell envelope structure in many prokaryotic cells and represent the
most simplest form of biological membranes developed during evolution. Most typically,
these structures are composed of a single protein or glycoprotein species and the formed
S-layer covers the surface of the whole cell (Sleytr et al., 2001). As a result, if present, Slayers are among the most abundant proteins in a given cell, and represent typically 10-15
% of total cellular proteins (Boot and Pouwels, 1996). Indeed, the biosynthesis of the
lattice subunits needs to be very effective, because a typically rod-shaped prokaryotic cell
with generation time of 20 min, requires synthesis of 500 subunits in a second to keep the
whole cell covered (Sleytr et al. 1999).
The S-layer subunits are held together by non-covalent interactions. Morphological units
found in different S-layer lattices may consist of one or two (obliques), four (squares), or
three or six (hexagonals) identical protein subunits and isolated S-layer (glyco)proteins
possess remarkably intrinsic property for recrystallization in suspension and at a broad
range of surfaces and interfaces (Egelseer et al., 2010; Sleytr et al., 2001). Most typically,
anchoring of S-slayer proteins to the cell takes place via non-covalent interactions with
cell wall structures; like lipopolysaccharides (LPSs) in Gram-negative bacteria and cell
wall polysaccharides in Gram-positive bacteria (Fagan and Fairweather, 2014). In addition
to domain (or region) responsible for the binding to the cell wall, another conserved
domain exists in S-layer protein which is related to the self-assembly of S-layer monomers
(Sleytr et al., 2014).
As S-layers are usually the most outermost envelope structure, it is not surprising that the
discovery of these structures was linked to the development of microscopy. Indeed, the
first observation from S-layers was made over sixty years ago when studying the electron
micrographs of shadowed preparations (Houwink, 1953; Sleytr and Messner, 1983). With
the development of instrumentation and procedures used in microscopy and in other
analytics (like gene sequencing), the number of organisms that are known to carry S-layer
protein is currently counted in hundreds (Sleytr and Messenr, 1983; Messner et al., 2010).
In Fig. 3A, protein dense S-layer from Lactobacillus crispatus is shown as a thin layer
around the cell. In Fig. 3B, examples of S-layers composed out of different morphological
units are presented.
22
Figure 3. Cellular location and fine structures of selected S-layers. A) S-layer localization
as an outermost cell envelope structure (modified from Schaer-Zammaretti and Ubbink,
2003). B) Examples of S-layers composed out of different morphological sub-units. The
number in p2, p4 and p6 refers to number of individual S-layer proteins needed to
compose one morphological S-layer subunit (modified from Messner et al., 2010).
It seems that S-layers are typically found only in one genus among LAB - in
Lactobacillus. In a recent review by Hynönen and Palva (2013), various S-layers of
lactobacilli are reviewed. Inside this genus, S-layers are present on many but not in all
species. Typical characteristics of the S-layers of lactobacilli are: protein monomers are
secreted through the general secretary pathway, S-layer proteins are usually small (25-71
kDa) and form lattices with oblique or hexagonal symmetry, and have high predicted
overall pI value (isoelectric point): 9,4-10,4. In contrast to most other S-layers,
lactobacillar S-layer proteins do not possess surface layer homology domain (SLH) that is
in many organism responsible for attaching the S-layer lattice to the peptidoglycan layer
by means of non-covalent binding. Instead, it seems that lactobacillar S-layers typically
possess two repeated amino acid motifs (showing high predicted pI) in their cell wall
binding area. Most typically, the cell wall binding domain is C-terminal, but at least in
Lactobacillus brevis and Lactobacillus hilgardii, it is located at the N-terminus (ÅvallJääskeläinen et al., 2008; Dohm et al., 2011). As the positive charge of lactobacillar Slayer is especially concentrated into same area where the cell wall binding region exists, it
has been proposed that electrostatic interaction occurs between the negatively charged cell
wall polymers and S-layer cell wall binding region (Hynönen and Palva, 2013; Antikainen
et al., 2002).
As reviewed by Dohm et al. (2011), various post-translational modifications, such as
linking of glycan strands, lipids, sulphate or phosphate groups with the S-layer protein
scaffold, have been reported, indicating adaptation of the corresponding S-layers to their
specific environment. One of these modifications is O-glycosylation, which has been
detected in few lactobacillar S-layers. In a recent study reported by Anzengruber et al.
(2014b), the O-glycosylation of S-layer and the presence of relateted protein Oglycosylation mechanism in Lactobacillus buchneri were proposed.
23
Although various functions for different S-layers have been proposed and demonstrated,
no common general function for these structures has been found, yet (Sleytr et al., 2014).
Among Lactobacillus, S-layers have often been considered to mediate bacterial adherence
to various targets, and currently most of the S-layer research among LAB is focused to
study the interactions between S-layers or S-layer possessing strains with human or animal
derived cells, receptors or macromolecules. Other putative roles suggested for lactobacillar
S-layers include: protection of cell towards lytic enzymes, stress conditions (e.g. tannic
acid and copper in wines) and role as putative phage receptor (Hynönen and Palva, 2013).
Also, it has been demonstrated that S-layers are able to defend host cell by binding metal
ions (Dohm et al., 2011, Schut et al., 2011; Gerbino et al., 2012, Gerbino et al., 2015).
Interestingly, murein hydrolase activity (e.g. towards cell wall peptidoglycan of
Salmonella enterica) has been detected in the C-terminal part of the S-layer protein SA of
L. acidophilus ATCC 4356 as reported by Prado-Acosta et al. (2008). The same research
group detected later the synergestic relationship between nisin and S-layer of this same
species in inhibition of the growth of pathogenic Gram-negative S. enterica and potential
pathogenic Gram-positive bacteria Bacillus cereus and Staphylococcus aureus (PradoAcosta et al., 2010).
Genetically engineered S-layers have a great potential in many applications. As recently
reviewed by Sleytr et al. (2014), recombinant S-layers proteins have been engineered to be
used in wide range of applications, like in vaccine development, biochip applications,
bioremediation, purification/downstream processing, building blocks of nanoparticle
arrays, biosensors, protective barriers, and as a platform for immobilized biocatalysts.
1.4 Cell retention mechanisms for secreted and excreted
proteins in LAB
1.4.1 Protein secretion mechanisms among LAB
In Gram-positive bacteria, proteins synthesized in cytoplasm may remain in this
compartment, or are sorted to the cytoplasmic membrane, to the cell wall or to the
surrounding medium (Tjalsma et al., 2000). Sorting of proteins to their relevant
subcellular compartment or surrounding environment is essential as proteins of different
final location might have very unrelated functions (Bendtsen and Woolbridge, 2009).
Most often, proteins which need to be transported to an extracytoplasmic location contain
an N-terminal signal peptide. With this signal, the newly synthesized protein from the
ribosome is targeted to certain protein transport pathway. After the translocation, specific
signal peptidases cleave the signal sequence, resulting in possible detachment of the
protein from the membrane. In Gram- positive bacteria (including LAB), the majority of
the secreted proteins are exported from the cytoplasm through the cytoplasmic membrane
via the general Secretory (Sec) pathway (Sibbald and van Dijl, 2009). According to
Desvaux et al. (2009) other characterized protein secretion systems translocating proteins
with N-terminal signal peptides are those of Twin-arginine translocation pathway (Tat)
and Fimbrilin-protein exporter (FPE, also known as pseudopilin export pathway).
24
Furthermore, small antibacterial peptides, bacteriocins, with N-terminal leader sequence
are typically secreted by specific ABC exporters (Bendtsen and Woolbridge, 2009;
Kleerebezem et al., 2010).
As reviewed by Sibbald and van Dijl (2009), it seems that the proportion of secreted
proteins without any known secretion signal varies per organism. When considering B.
subtilis, the model organism of Gram-positive secretion, this number is relatively low,
whereas for group A Streptococcus, it seems to be much higher. Some of the proteins
without any known export signal are actively transported while others are proposed to be
released during cell lysis. In Gram-positive bacteria, the former group includes at least
proteins which are transported via the Wss pathway, holin transport system, flagella export
apparatus (FEA) and in some cases via the accessory Sec system (Desvaux and Hébraud,
2006; Desvaux et al., 2009). In some case, part of the cell’s plasmamembrane may be
reformed into vesicles carrying proteins or other macromolecules (like DNA) inside.
Indeed, active lysis-independent vesicle formation and subsequent release of these vesicles
into external medium is a conserved phenomenon among Gram-negative bacteria,
including pathogenic and non-pathogenic species, but detected also among Gram-positive
bacteria (MacDonald, 2012), also including some streptococci (Liao et al., 2014; OlayaAbril et al., 2014).
1.4.2 Autolysins and excretion of cytosolic proteins
The very ultimate mechanism for any single cellular organisms, like LAB, is to deliver
proteins to surrounding medium by induced partial or complete autolysis. In bacteria,
autolysis or self-digestion is a process, which can be defined as a breakdown (or lysis) of
the cell, and which results from the cell’s own hydrolytic enzyme activity towards various
and specific bonds in its cell wall peptidoglycan (Shockman et al., 1996; Crouigneau et
al., 2000). Enzymes that are responsible for this lytic activity are called autolysins.
Although, the exact mechanism and regulation of the extend of autolysis among LAB is
not fully understood, yet, the functional effects of this phenomenoma, like the release of
internal peptidases inside starter cultures during cheese ripening, are widely acknowledged
and employed in modern dairy business. Probably the best know autolysin among the
dairy related species, is the major autolysin of L. lactis, AcmA, which is covered in more
detail in later in this study.
As reviewed by Chapot-Chartier (2010), autolysins have been proposed and demonstrated
to be involved in several different functions during the bacterial growth cycle:
-
Septation and separation of daughter cells
Cell expansion
Peptidoglycan turn-over
Protein secretion
Also, it has been speculated that they possess roles in more specific functions, which are
related to: competence for genetic transformation, flagellar morphogenesis, spore
formation and germination, biofilm formation, pathogenicity, waking up dormant bacteria
and autolysis and programmed cell death (Chapot-Chartier, 2010). The role of autolysins
25
in the regulated excretion of cytosolic enzymes (ECP), during which cell’s viability
remains secured, remains to be a hot topic in scientific debate until the beauty of this
putative non-classical excretion mechanism has been completely revealed (Götz et al.,
2015).
1.4.3 Various cell envelope retention mechanism for proteins in LAB
Desvaux et al. (2009) have reviewed the different fates of translocated proteins for Grampositive bacteria. According these authors, after translocation a single protein may remain
to be anchored to the membrane, associate covalently or non-covalently with cell-wall
components, assemble into macromolecular structures on the cell surface (e.g. flagellum,
pilus and S-layer), be injected into a host cell, or be released in the surrounding medium.
Here, the main interest is to study various cell envelope retention mechanisms that provide
surface display of the target protein (or domain).
In most cases, translocated (or excreted) proteins are secreted or diffused to the
surrounding medium unless they do not posses any retention signals (or domains) which
keep them bound with the cell envelope constituents. However, protein localization within
the cell envelope does not necessarily guarantee its surface exposure. Vice versa,
translocation of the whole protein is not always required for the surface display. Typical
example for this is membrane proteins which contains one or several hydrophobic
transmembrane domains (TMD).
As reviewed by Desvaux et al. (2006), four major types of cell surface displayed proteins
are generally recognized: proteins which are anchored to the cytoplasmic membrane by
hydrophobic transmembrane domain(s), lipoproteins which are covalently attached to
membrane lipids, proteins which are covalently linked to peptidoglycan via C-terminal
LPXTG-like motif, and proteins attaching to cell wall by specific cell wall binding
domains. Together, this subset of surface exposed proteins constitutes the surfaceome of
the bacterial cell (Cullen et al., 2005, Desvaux et al., 2006). As stated by Cullen et al.
(2005), identification of the members belogning to this group is critical to understanding
the interactions of bacteria with their environments.
In the following sections, the mechanism of various cell membrane and cell wall retention
mechanisms are presented among LAB. Specially, the main focus is on introduction of the
non-covalent retention mechanisms in LAB.
1.4.3.1 Binding of proteins to membrane
Intrinsic (or integral) membrane proteins are tightly associated with biomembranes and do
not detach from those by washing (change in pH or in ionic strength). Indeed, detachment
during this kind of washing is the characteristic feature for the extrinsic (or peripheral)
membrane proteins, which are membrane associated. Most typically, the binding of the
peripheral proteins take place via electrostatic and hydrophobic interactions, whereas
integral proteins remain membrane bound by one (bitopic proteins) or several (polytopic
26
proteins) transmembrane spans. In addition, monotopic integral proteins penetreate
membranes partially but do not span the entire membrane, or are anchored in the
membrane by covalent bonding with the membrane lipids (Luckey, 2014). In the
following sections, binding mechanisms of the transmembrane proteins and lipid anchored
proteins are introduced in more detail.
1.4.3.1.1 Anchoring by membrane spanning
Transmembrane regions in proteins are structurally either ȕ-barrels or Į-helical sheets, out
of which the latter type of structure is typically found also among the membrane spanning
proteins of Gram-positive bacteria. The average length of a transmembrane Į –helix is
more than 25 amino acids and is not typically oriented perpendicular to the membrane
plane but is more or less tilted (Schneider et al., 2007). Indeed, there exist several different
topological variants within the group of Į-helical membrane proteins. According to Higy
et al. (2004) these proteins may be divivided into sub categories depending on
presence/absence of cleavable signal sequence and subcellular location of the N- and Cterminus (when anchored). Also, classification may also be based on the number of passes
through the membrane.
The actual biogenesis of Į-helical membrane proteins is often described by the model
proposed by Popot and Engelman (2000). This model includes two energetically distinct
stages: formation and subsequent side-to-side association of independently stable
transbilayer helices. During the first stage, membrane proteins are synthesized by the
ribosomes and their transmembrane domains are integrated into the target membrane,
which leads to adoption of their topologies (topogenesis). During the second stage,
membrane proteins fold and adopt their native structures. The stability of the individual
domains is a consequence of the hydrophobic effect and main-chain hydrogen bonding,
whereas other interactions drive to side-to-side helix association. As this model is very
rough, it is not always possible to clearly separate the second stage from the topogenesis.
According to Zweers et al. (2008), there are enormous variation among the number,
hydrophobicity and the membrane topology of the transmembrane segments (TMSs)
between different membrane proteins. Most typically, the topology of membrane proteins
follows the positive-inside rule (von Heijne, 1986). Accroding this rule, the positively
charged residues are more likely found in the cytoplasmic loops rather than in periplasmic
loops. However, alteration of the charge distribution in the regions flanking a TMS may
result in reversion of the topology. Also, the length and the mean hydrophobicity of the
TMS affect the membrane insertion. Typically, the first TMS of a polytopic membrane
protein determines the orientation of subsequent TMSs, as there is need to alternate the
direction for the membrane spanning (Driessen and Nouwen, 2008).
1.4.3.1.2 Covalent anchoring of lipoproteins
Bacterial lipoprotein biogenesis was established by Wu and co-workers (Braun and Wu,
1994) in Gram-negative model organism Escherichia coli, in which the related enzymes
have also been found to be indispensable for survival (Hutchings et al., 2009). As
discovered in E. coli, prolipoproteins carry Type II signal peptides, which direct these
molecules through Sec- (or Tat-) system to the lipoprotein biogenesis machinery. The type
II signal peptides include a conserved motif (the ‘lipobox’), which is typically
27
L-3-[A/S/T]-2-[G/A]--1-C+1 with the +1 cysteine absolutely conserved in all bacterial
lipoproteins. Lipid anchoring is initiated by formation of a covalent linkage between the
membrane phospholipid derived diacylglyceride group and the indispensable cysteine
residue found in the lipoprotein signal peptide. This process is catalyzed by
prolipoprotein diacylglyceryl transferase (Lgt) enzyme. Interestingly, the Lgt of L. lactis
was the first Gram-positive prolipoprotein diacylglyceryl transferase that has been
biochemically characterized (Banerjee and Sankaran, 2013). The next step in the
lipoprotein biogenesis includes the cleavage of the signal peptide by a dedicated Type II
lipoprotein signal peptidase (Lsp) at the conserved cleavage site in the lipobox. As a
result, the lipid-modified cysteine remains as the first N-terminal amino acid in the mature
lipoprotein. Finally, the lipoprotein N-acyl transferase (Lnt) catalyzes the addition of third
fatty acid in an amide linkage to the free amino group of the lipidated cysteine. According
to Buddelmeijer (2015), the first two enzymes in the lipoprotein biogenesis, so Lgt and
Lsp, are conserved in all bacteria species, but Lnt has only been identified in
proteobacteria and actinomycetes. The general model for bacterial lipoprotein biogenesis
is presented in the Fig. 4.
Figure 4. Lipoprotein biogenesis pathway in bacteria (modified from Buddelmeijer,
2015). Abbreviations: C, cytoplasm; CM, cytoplasmic membrane; LB, lipobox; Lgt,
phosphatidylglycerol:prolipoprotein diacylglyceryl transferase; Lnt, apolipoprotein Nacyltransferase; Lsp, prolipoprotein signal peptidase; SP, signal peptide; PG,
phosphatidylglycerol; PE, phosphatidylethanolamine.
As reviewed by Buddelmeijer (2015), triacylated lipoproteins and new sub groups for
diacylated lipoproteins have recently been detected also in some firmicutes and terenicutes
(that include mollicutes), suggesting the presence of Lnt-like enzymes and/or novel noncanonical lipidation pathways in these organisms. One of the novel diacylated lipoprotein
structures that was discovered by Kurokawaka et al. (2012) was coined by the authors as
“lyso” form. Authors found this type of lipoprotein in some low-GC Gram-positive
28
species, including also LAB species of E. faecalis, Streptococcus sanguis and
Lactobacillus bulgaricus. As suggested by the authors, the lyso form of lipoprotein may
be well distributed among wider group of low-GC content Gram-positive bacteria.
1.4.3.1.3 Sec-attached proteins
Sec-attached proteins are special group of proteins carrying a predicted cleavage sites for
signal peptidase in their signal sequence, but which for some unknown reason avoid the
cleavage and remain N-terminally anchored to the cell membrane (Tjalsma and van Dijl,
2005). When studying the S-layer associated proteins of Lactobacillus acidophilus,
Johnson et al. (2013) predicted with Hidden Markov Model, which was developed by
using the sequences of putatively sec-attached bacillus proteins, that some of the S-layer
associated (extracted) proteins were capable for sec-attachment. However, this was not
confirmed by further experimental work and researchers finally classified these candidates
as extracellular proteins that are predicted to be sec-attached.
1.4.3.2 Binding of proteins to cell wall by covalent anchoring
In early 70s’ the first evidence for covalent anchoring of protein A to the peptidoglycan of
S. aureus was presented (Sjöquist et al., 1972). In late 80’s, Pancholi and Fischetti (1988)
studied the anchoring mechanism of the M protein of Streptococcus pyogenes and
suggested that enzyme mediated cleavage of the C-terminal anchor of this protein takes
place between proline- and glycine-rich region and hydrophobic region. A year later, this
same group reported that an endogenous membrane anchor cleaving enzyme is able to to
release M protein from isolated streptococcal protoplasts (Pancholi and Fischetti, 1989). In
the following study Fischetti et al. (1990) reported that the LPXTG hexapeptide,
preceding the hydrophobic C-terminal domain, is conserved in anchor region of several
surface proteins originated from Gram-positive cocci (Fischetti et al., 1990). Navarre and
Schneewind (1994) proposed couple of years later a novel hypothesis for the cell wall
linkage of surface proteins in Gram-positive bacteria. In this model, after the translocation
across the cytoplasmic membrane, the polypeptide chain with C-terminal cell wall sorting
signal (including the LPXTG-motif) was suggested to be recognized (or sorted by) by
sortase enzyme, following the subsequent cleavage of the precursor between threonine and
glycine of the LPXTG motif and linkage of the resulting N-terminal fragment to the cell
wall. The structural characteristics for the sortase A dependent proteins are shown in Fig.
5.
During the following years more evidence and details were gathered supporting the key
role of the sortase in this process. Mazmanian et al. (1999) managed to screen and
sequence the DNA fragment from S. aureus that seemed to be responsible for reducing the
accumulation of uncleaved precursor form of the reporter polypetide in this organism.
They also reported the presence of homologs (based on database search) of this gene in
several other Gram-positive bacteria. As reviewed by Navarre and Schneewind (1999) and
Maraffini et al. (2006), this sortase dependent cell wall sorting pathway (see Fig. 5) is
universal in many Gram-positive bacteria.
29
Pallen et al. (2001) studied the presence of sortase-like proteins by database search.
Genomic data available at that time indicated that sortase homologous were present in
almost all Gram-positives studied and also the one of the very first observations of the
presence of sortase like enzymes in Gram-negative bacteria and archae was reported.
Authors also reported that there are usully more than one sortase-like protein encoded in
each Gram-positive genome, and proposed that this could be an indication of specificity to
different substrates. Because of the increasing number of reports on the identification
and/or characterization of paralogous sortase genes during the very first years of this
millenium, the need for classification and nomenclature system for sortases was evident.
Figure 5. Sortase A (SrtA) dependent surface display of proteins (modified from
Schneewind and Missiakas, 2014). Structures described in picture: P1, full length protein
precursor (with signal peptide); P2, protein precursor harboring only the C-terminal
sorting signal; AI, an acyl enzyme intermediate; P3 protein precursor that resulted from
the linkage of the C-terminal threonine of acyl intermediate with Lipid II (after
nucleophilic attack by free amino group of Lipid II) and subsequent regeneration of
sortase; M, mature covalently anchored surface protein. MN and GN denote Nacetylmuramic acid and N-acetylglucosamine, respectively.
Dramsi et al. (2005) gave their proposal for classification by dividing different sortases to
four main classes (classes A, B, C and D). This classification proposal was based on
detailed analysis of sixty-one sortases from complete Gram-positive genomes. Out of
these classes, the sortases belogning into group A (SrtA) are required for anchoring the
majority or all of the LPXTG-containing proteins of a given bacteria. Without few
expections, genes expressing SrtA like housekeeping enzymes seem to be present as single
copy in genomes of all low GC% Gram-positive bacteria (Dramsi et. al., 2008). Sortases
in other classes (B, C and D) are accessory and appear to anchor more dedicated substrates
(Schneewind and Missiakas, 2014). Several, but not all sortases (via their substrates) in
class B are related to iron acquisition, whereas enzymes belogning to SrtC–class are
polymerizing enzymes that are involved in the formation of pili or fimbria in several
Gram-positive bacteria, and finally some members belogning to SrtD-class have shown to
30
posses role under specific developmental processes (mycelium formation in Streptomyces
coelicor, sporulation in Bacillus anthracis) (Dramsi et al., 2008; Schneewind and
Missiakas, 2014). Recent review of the relevance and application of sortases in LAB has
been published by Call and Klaenhammer (2013).
1.4.3.3 Most typical non-covalently attached cell wall domains among LAB
For LAB proteins and LAB related phages endolysins, many different non-covalent
binding mechanisms to the cell wall have been proposed. The most relevant and best
known binding domains have been characterized and assigned with a Pfam (Finn et al.,
2014) and InterPro (Mitchell et al., 2015) identification codes.
Table 1. The most relevant non-covalent cell wall binding domains in LAB (with Pfam
identification codes).
31
Table 1 continued.
32
Table 1 continued.
In literature, some good reviews exist that covers these domains among the main species
of industrial related LAB (Zadravec et al., 2015; Chapot-Chartier and Kulakauskas, 2014;
Sengupta et al., 2013; Tarahomjoo, 2013; Kleerebezem et al., 2010). Furthermore, in some
studies cell wall binding domains of specific strain is compared to broader range of LAB
strains - see studies published by Kankainen et al. (2009) and van Pijkeren et al. (2006) as
examples. Out of these proposed non-covalent binding domains in literature, the most
important for LAB are presented in the Table 1.
1.4.3.4 LysM domain
Among LAB, LysM is the most studied non-covalent cell wall binding domain. In the
literature, several excellent reviews covering the LysM domain and its potential
33
applications have been published. One of the most recent one of these is the review
published by Visweswaran et al. (2014).
Also, the bindig mechanism of LysM domain is one the best known among all the noncovalent binding domains. Steen et al. (2003) demonstrated that cA domain of AcmA of
L. lactis is responsible for binding this autolysin enzyme to peptidoglycan. This Cterminal cA domain contains three separate LysM domains which are separated by nonhomologous sequences. Steen et al. (2003) noticed that other cell wall constituents, most
probably lipoteichoic acid in Lactococcus, hindered the interaction between peptidoglycan
and cA domain. Binding test with S-layer bearing Lactobacillus helveticus ATCC 15009
demonstrated that also S-layer seemed to hinder efficient binding. It was also found that
these C-terminal LysM domains bind to specific sites on the bacterial surface, i.e. near the
poles and septum of L. lactis cells. In L. lactis SK110 these locations were demonstrated
to be typical binding sites for galactosyl-decorated LTAs that are typical for this phage
resistant strain (Sijtsma et al., 1990). In conclusion, Steen et al. (2003) suggested that the
repetitive disaccharide component in peptidoglycan, composed out of Nacetylglucosamine (GlcNAc) and N-acetylmuramic acid (MurNAc) molecules, is most
probably the ligand for the binding as it is the only common part in A- and B-type
peptidoglycans (which were both demonstrated to bind cA).
As reviewed by Buist et al. (2008), in some eukaryotes LysM modules are present in
chitinases, suggesting that LysM is also able to bind chitin (polymer of GlcNAc capable to
form hydrogen bonding between the adjacent polymer chains). According to Buist et al.
(2008) it remains unanswered whether GlcNAc is the sole moiety recognized by LysM.
Wong et al. (2014) proposed after experiments with LysM domains from CwlS (NlpC/P60
D,L-endopeptidase) of B. subtilis that bacterial LysM domains do not discriminate
between peptidoglycan and chitin GlcNAc polymers but recognize both of these substrates
with similar affinity, which is not usual within hydrolase activity related LysM domains in
plants. In this study, they indicated by microscale thermophoresis assay that at least two
LysM domains are necessary for efficient binding to long chitin polymers wheras three
LysM domains are required for proper binding to peptidoglycan fragments. For
comparison, study conducted by Steen et al. (2005) with AcmA of L. lactis MG1363
indicated that at least two domains are needed for peptidoglycan binding, but three
domains provides optimal peptidoglycan binding and biological functioning. More
recently, Mesnage et al. (2014) proposed molecular basis for bacterial peptidoglycan
recognition by LysM domains. This study was based on multimodular LysM domain from
AtlA from E. faecalis. The data from this study suggested that LysM module recognizes
GlcNAc-X-GlcNAc moieties, in which X is GlcNAc or MurNAc, and peptide stems act as
negative discriminants that modulate the binding but do not prevent it. In this same study,
the earlier observations (Steen et al., 2005, Wong et al., 2014) suggesting that individual
LysM modules bind in cooperative manner to long glycan chains present in cell wall was
confirmed. Most recently, Wong et al. (2015) proposed model for intermolecular
dimerization of P60_tth (NlpC/P60 endopeptidase containing two N-terminal LysM
modules) of Thermus thermophiles with peptidoglycan substrate.
34
1.4.3.5 Other, less characterized and/or less abundant cell binding mechanism among
LAB
The Lc-LysBD-domain was recently characterized in the C-terminal part of prophage
endolysins (Lc-Lys and Lc-Lys2) of L. casei BL23 (Regulski et al., 2013). Authors
demonstrated that this domain binds to PG and is highly specific for amidated D-Asp
cross-bridge present in the peptidoglycan of L. casei. So far, this binding domain has also
been reported to be found in L. lactis phage 949 endolysin and in L. casei phage
endolysins A2 and PL-1 (Regulski et al., 2013; Chapot-Chartier 2014).
In some cases, no anchor or binding domain has been detected or does not exist for a
specific protein although it seems obvious that the protein is attached to cell envelope.
Also, in some cases slow diffusion through peptidoglycan layer or inner wall zone
temporary reservoirs of proteins may lead to wrong conclusion when detecting cell
attached proteins. These last examples hint that diffusion through the peptidoglycan is not
necessarily an unregulated event. As reviewed by Forster and Marquis (2012), also protein
size and form with many other factors, like the average length of glycan chains, the level
of crosslinking between glycan chains, the presence or absence of bridges between the
crosslinking peptides, electrostatic interactions, and the mechanical tension imposed by
cell turgor pressure may influence the cell wall permeability.
The charge of the Gram-positive cell wall is most typically negative, due to presence of
polyanionic teichoic acids (TAs) in the cell envelope (Neuhaus and Baddiley, 2003).
Under physiological conditions, secreted proteins with high overall pI values remain
positively charged, which may make their journey through the peptidoglycan layer slow
(Forster and Marquis, 2012) or even bind transiently these proteins to the peptidoglycan
layer. Among LAB, an example of this kind of binding is that of the cystine-binding
protein (CuyC) of Lactobacillus reuteri BR11 (formerly referred as BspA of L. fermentum
BR11) (Turner et al., 1997). More recently, Anzenburger et al. (2014a) characterized
homologous N-acetylmuramidases from two different L. buchneri strains and
demonstrated their binding to L. buchneri cell walls in vitro. In both of these above
mentioned examples, as no obivious cell wall binding domains were detected and high pI
value was predicted for both of these proteins, authors proposed that the cell wall binding
was due to electrostatic interactions between positively charged proteins and negatively
charged cell wall constituents. Turner et al. (1997) also demonstrated that the electrostatic
attachment was abolished and BspA released, when whole cells of L. fermentum were
treated with LiCl-solution. Earlier, LiCl- treatment was demonstarted to release S-layer of
L. helveticus, another cationic high pI protein from the cell surface (Lortal et al., 1992).
Some surface displayed enzymes are called as moonlighting proteins. The peculiar name
of moonlighting proteins is due to their dual life-style –the performance of more than one
function by a single protein (Copley, 2012). As summarized by Jeffery (2003), these
enzymes are single proteins which have multiple functions, but this dualism is not due to
gene fusions, splice variants or multiple proteolytic fragments. As reviewed by
Kainulainen and Korhonen (2014), moonlighting proteins often perform their canonical
and moonlighting functions in separate cell compartments, and in some cases, like in
many proteomic studies, this localization limited more narrow criterium for identification
for bacterial moonlighting proteins has remained in use. In these cases, moonlighting
proteins have more than one function but these functions need also to be displayed in
35
different cell compartments. Most typically moonlighting proteins are canonically
cytoplasmic proteins, but are found also on the surface of the cell or in the surrounding
medium, although they do not possess any recognizable sequence motifs for secretion or
anchoring to the bacterial or to the eukaryotic cell surface (Copley, 2012; Kainulainen and
Korhonen 2014). First bacterial moonlighting proteins were found from group A
streptococci by Pancholi and Fischetti (1992). These researchers demonstrated that,
glyceraldehyde-3-phosphate dehydrogenase (GAPDH), one of the key enzymes in
cytosolic glucose metabolism, is found on the surface of group A streptococci and is
capable to bind to fibronectin. As stated by the authors, this property suggests a putative
role for GAPDH in the colonization of the pharynx. This type of dual role for the
glycolytic enzymes is not rare, as practically all enzymes in the bacterial EmbdenMayerhof glycolytic pathway have been assigned with putative adhesive moonlighting
function (Kainulainen and Korhonen, 2014). Indeed, adhesion to host ephitelial cells,
mucus and extracellular matrix (ECM) components, and interaction with circulating host
components and modulation of host immune responses, are the most typical moonlighting
functions found among bacteria (Kainulainen and Korhonen, 2014; Mangiapane et al.,
2015) – in good (supporting probiotic action) or in bad (supporting virulence).
As reviewed by Kainulainen and Korhonen (2014), it seems that ionic interactions with
lipoteichoic acids and cell division sites seems to be critical for the surface localization for
many moonlighting proteins. Also, authors speculated that moonlighting proteins in
commensal lactobacilli and in pathogenic species are functionally different because it
seems that the nature of anchoring to the bacterial surface is often different: several
pathogenic species are able to retain moonlighting proteins on cell surface at neutral pH,
whereas among lactobacilli these proteins are released in many cases. As an supporting
example for this generalization, Terrasse et al. (2015) demonstrated that peptidoglycan
serves as binding ligand for the GAPDH of S. pneumoniae and this binding is not
dependent on the presence of the teichoic acids component, which is the case with some
lactobacilli – GAPDH of L. crispatus as an example (Antikainen et al., 2007, Kainulainen
et al., 2012). Regardless of the nature of the binding, observations suggesting that some
moonlighting proteins may be released from a single bacterial species and then reassociate
onto the surface of another species, or bind directly after secretion/cell lysis with an
universal ligand (like peptidoglycan) of another species, represents a novel mechanism in
bacteria-bacteria interactions and may have importance in bacteria-host interactions
(Oliveira et al., 2012; Kainulainen and Korhonen, 2014; Terrasse et al., 2015).
Interestingly, WaĞko et al. (2014) suggested that S-layer proteins of L. helveticus T159
binds some moonlighting proteins, which makes the surface of this strain more
hydrophobic and improves the adhesion and aggregation capacity of this strain. Authors
also observed the S-layer associated presence of some technologically important nonmoonlighting proteins, such as L-lactate dehydrogenase, glucose-1-phosphate
adenylyltransferase, and glycerol-3-phosphate ABC transporter. Earlier, Johnson et al.
(2013) studied S-layer associated proteins (which they coined as SLAPs) of L. acidophilus
NCFM. In this study, 37 proteins were solubilized from the S-layer wash fraction and
closer study suggested that four of these proteins were putative moonlighting proteins and
17 were putative SLAPs. One of the SLAP was studied closer (LBA1029), and it it was
demonstrated that it contributes to a pro-inflammatory TNF-Į response from murine DCs.
Based on their observations authors suggested that SLAPs appear to impart important
surface display features and immunological properties to S-layer coated microbes.
36
Notably, SLAP term used here does not refer to pfam SLAP domain (Lactobacillus Slayer protein –domain), although some SLAPs reported by Johnson et al. (2013) may also
carry SLAP pfam domains.
1.5 Fermentation pathways in LAB
1.5.1 Fermentation and respiration in LAB
Lactic acid bacteria are chemotrophic organisms that gain the energy for their metabolism
by oxidation of chemical compounds (Ribéreau-Gayon et al., 2006). According to the final
acceptor of electrons, metabolism of a cell is classified either as fermentative or
respirative. During fermentation, the oxidation and phosphorylation (energy generation by
formation of ATP) are not coupled, as is the case with respiration. For fermentative
organisms, like LAB, the final reduced molecule is characteristically an endogenous
compound. For LAB, this molecule is typically pyruvate. However, depending on the
presence of alternative electron acceptors, growth conditions and metabolic balance,
alternative pyruvate-utilizing pathways are possible for some LAB strains (von Wright
and Axelsson, 2011).
In rare cases, functional oxidative respiration has been observed in some LAB species
under special circumstances (heme, or heme and menanquine present in the growth
medium). The best documented and understood respiration process is that of L. lactis.
Respiration in this species has demonstrated to have positive impact on bacterial biomass,
resistance to oxygen, and long-term survival (Pedersen et al., 2012).
1.5.2 Typical LAB fermentations
In general, the generation of metabolic energy among LAB primarly occurs through the
substrate level phosphorylation (Konings, 2002). In this process, the oxidation of substrate
generates energy rich intermediates, which subsequently can be used for ATP generation.
This process results in the formation of NADH from NAD+, which has to be regenerated
in order for cells to continue the fermentation. In the case of LAB, pyruvate is the most
typical final electron (or hydrogen) acceptor, resulting in formation of the reduced end
product, lactate (Axelsson, 2004).
Pyruvate + NADH + H+ ĺ lactate + NAD+
The above sum reaction can be divided as follows:
Oxidation (NADH loses e-): NADH + H+ ĺ NAD+ + 2H+ + 2eReduction (pyruvate gains e-): Pyruvate + 2H+ +2e- ĺ lactate
In general, LAB as a group exhibits an enormous capacity to utilize different
carbohydrates and related compounds. Furthermore, as LAB are able to adapt their
37
metabolism in response to various conditions, this may result to different end product
patterns even with the case of single substrate (Axelsson, 2004).
In the literature the most common metabolic pathway described for LAB is the
fermentation of glucose (a hexose sugar, see Fig. 6). However, also other hexose sugars
such as mannose, galactose, and fructose are readily fermented by many LAB, too.
Figure 6. Most relevant metabolic pathways of selected sugars among LAB (AbdelRahman et al., 2013; Li and Cui, 2010; Wang et al., 2015). Most important enzymes for
homolactic and heterolactic fermentation are identidfied with number: 1, fructose-1,6
diphosphate aldolase; 2, lactate dehydrogenase; 3, transketolase; 4, transaldolase; 5,
glucose-6-phosphate dehydrogenase; 6, 6-phosphogluconate dehydrogenase; 7,
phosphoketolase; 8, acetate kinase; 9, alcohol dehydrogenase. The numbers for the key
enzymes for homolactic and heterolactic fermentation are presented with yellow
background.
38
Depending of the specific uptake mechanism of a given sugar (permease-/PTS-system),
the sugar molecule may be intact or phosphorylated after entering into cell. If needed, the
sugar is further processed, most typically phosphorylated or isomerized, before joining the
glycolytic pathway usually at the level of the glucose-6-phosphate or fructose-6phosphate. However, deviations to the above described route exist and one of those is the
galactose metabolism in some LAB. As an example, the galactose uptake in L. lactis, E.
faecalis, and L. casei happens through the PTS system, resulting in simultaneous
phosphorylation of the sugar. The resulting galactose-6-phosphate in these species is
processed through tagatose-6-phosphate pathway which coincides with the glycolytic
pathway at the level of glyceraldehyde-3-phosphate (GAP, see Fig. 6).
Under the normal conditions (excess of sugar, limited access to oxygen), LAB rely on two
principal pathways for catabolism of hexose sugars: homolactic pathway and heterolactic
pathway (Fig. 6). Theoretically, in homolactic pathway sugar is completely converted to
lactic acid. For hexose sugars, this means production of two mol of lactate per one mol of
consumed hexose with the concomitant production of 2 mol of ATP. More practically,
fermentation is often regarded as homolactic when more than 90% of the hexoses are
converted to lactic acid and 2 mol of ATP is generated per mole of hexose (Hutkins,
2001). As lactic acid is virtually the only end product, this type of sugar metabolism is
referred to as homolactic fermentation (Axelsson, 2004). This pathway is typical among
genera of Carnobacterium, Aerococcus, Enterococcus, Lactococcus, Vagococcus,
Pediococcus, Streptococcus, Tetragenococcus and in some lactobacilli (Axelsson, 2004).
Characteristics for this pathway (see Fig. 6) is the formation of fructose-1,6-diphosphate
(FDP) and subsequent branching of the pathway (by FDP aldolase) to glyceraldehyde-3phosphate (GAP) and dihydroxyacetonephosphate (DHAP). In the course of conversion of
glyceraldehyde-3-phosphate to lactate, four moles of ATP per mole of hexose are
generated, resulting in net generation of 2 moles of ATP per mole of hexose over the
whole pathway (two moles of ATP were consumed before generation of FDP). The very
last enzymatic reaction, so the reduction of pyruvate to lactic acid by a NAD+ dependent
lactate dehydrogenase (nLDH) concomitantly converts NADH to NAD+, providing the
electron sink required to maintain the vital redox balance in cell’s metabolism (Hutkins,
2001). For pentoses, the characteristic feature in homolactic pathway is the conversion of
xylulose-5-P to fructose-6-P and glyceraldehyde-3-P through series of reactions catalyzed
by two key enzymes: transaldolase and transketolase. This branch of the homolactic
fermentation presented in Fig. 6 is known as pentose phosphate pathway (Endo and Dicks,
2014). By this route, pentoses are converted completely to lactic acid with the theoretical
yield of 1.67 mol of both lactic acid and ATP from one mole of pentose sugar (Tanaka,
2002). Homolactic pentose fermentation is not common among LAB, but has been
demonstrated with few strains – L. lactis IO-1 (Tanaka et al., 2002) and Lactobacillus vini
Mont 4T (Rodas et al., 2006) as examples.
In heterolactic pathway (or 6-phosphoglucanate/phosphoketolase-pathway, 6-PG/PK),
approximate equimolar amounts of lactate, acetate, ethanol and CO2 are generated with 1
mol of ATP per mole of hexose (Hutkins, 2001). As this metabolic pathway results in
multiple end products (usually in equimolar amounts) it is referred as heterolactic
fermentation. This fermentation pathway is typical among genera of Leuconostoc,
Oenococcus, Weissella and some lactobacilli. Characterics for this pathway are the initial
dehydrogenation steps with the formation of 6-phosphogluconate, followed by
decarboxylation reaction (release of CO2). After the decarboxylation, the remaining
39
pentose-5-phosphate is split by phosphoketolase enzyme, resulting in formation of
glyceraldehyde-3-phosphate and acetyl phosphate. The former of these intermediates is
then processed to lactate by the same pathway that is employed during the homolactic
fermentation, whereas the latter is reduced to ethanol via acetyl CoA and acetaldehyde
with concominant regeneration of NADH to NAD+ (Axelsson, 2004). When compared to
the homolactic pathway, less ATP (one mole per one mol of hexose) is generated from the
hexose substrate by this pathway. However, in some cases when an external electron
acceptor is present, LAB using the heterolactic fermentation may gain an additional ATP
by reducing an external electron acceptor instead of reducing acetyl-phosphate to ethanol.
As a result, acetate kinase is able to dephosphorylate acetylphosphate, resulting in
formation of acetate and concominant phosphorylation of ADP to ATP (Axelsson, 2004).
In the case of pentoses, heterolactic fermentation yields equimolar amounts of lactate and
acetate. Ethanol is not produced as the final product, because no NADH is formed during
the preceeding 5-carbon metabolism. As an extra benefit, one additional mol of ATP is
generated when compared to heterolactic hexose fermentation.
Additionally, members belonging to genus lactobacillus have been traditionally divided
into three sub-groups based on their glucose fermentation characteristics: (i) the obligately
homofermentative lactobacilli, (ii) the facultatively heterfermentative lactobacilli, and (iii)
the obligately heterofermentative lactobacilli (Hammes and Vogel, 1995). This
physiological classification relies on the presence or absence of fructose-1,6-diphosphate
aldolase and phosphoketolase, that are the key enzymes for the homo- and
heterofermentative sugar metabolism, respectively (Axelsson, 2004).
1.5.3 Other fates of pyruvate
Under certain conditions some homolactic LAB strains may shift to mixed-acid
metabolism during the fermentation on sugars. Transition to this metabolism is strain
specific, but typically factors that initiate it may include: carbon limitation, excess of
slowly metabolized sugars (such as galactose), presence of oxygen or respiratory lifestyle
(Mayo et al., 2010; Neves et al., 2005). Typical characteristic of this mixed-acid
metabolism is production of formate, acetate, ethanol and/or CO2 in addition of lactate.
The formation of these products is orginated from pyruvate by the key enzymes presented
in the Fig. 7. Out of these enzymes, pyruvate formate lyase is operational only under
anaerobic conditions, whereas pyruvate oxidase is oxygen dependent. Pyruvate
dehydrogenase has catabolic role under aerobic conditions, so backing up the pyruvate
formate lyase in the presence of oxygen. Under anaerobic conditions, pyruvate dehydrogenase
mainly serves anabolism by providing acetyl CoA for lipid biosynthesis (Axelsson, 2004).
40
Figure 7. Alternative reaction pathways for pyruvate (Lorquet et al., 2004, von Wright
and Axelsson, 2011). Most important enzymes in these pathways are identidfied with
number: 1, pyruvate formate lyase; 2, pyruvate oxidase; 3, pyruvate dehydrogenase; 4,
phosphotransacetylase.
In addition to the mixed acid fermentation, pyruvate may also be converted to alanine by
alanine dehydrogenase or serve as starting substrate for diacetyl production (Endo and
Dicks, 2014). Diacetyl is known as the butter aroma compound in dairy products and it is
typically produced in very low amounts during lactic fermentations, as most of the flux
from pyruvate through this alternative metabolic pathway is directed to form acetoin. This
type of metabolism requires a surplus of pyruvate relative to the need of NAD+
regeneration, and is typically found in leuconostocs and some lactococci that can gain that
this surplus by citrate metabolism in milk (von Wright and Axelsson, 2011). The formed
acetoin is chemically quite similar to diacetyl, but is does not give the desired butter
aroma. Acetoin is formed by decarboxylation of Į-acetolactate, which in turn is formed
from pyruvate by the acetolactate synthase activity. Some dairy strains lack functional Įacetolactate decarboxylase, which results in accumulation of Į-acetolactate and enables its
subsequent chemical conversion to diacetyl in the presence of oxygen (Hugenholtz and
Kleerebezem, 1999).
1.5.4 Classification of LAB according to isomer of lactic acid produced
As an end product of catabolic reactions from sugars, lactic acid with either a slight
positive or negative specific optical activity is formed. More commonly, these chiral
molecules are known as L(+)- and D(-)-lactic acid, respectively. The optical isomer(s) of
lactic acid produced depends on the presence of D-nLDH, L-nLDH and lactate racemase
enzymes in the cell. As reviewed by Goffin et al. (2005), L-lactate induced lactate
racemase, enzyme catalyzing the racemization of L(+)-lactic acid to D(-)-lactic acid, has
been detected only in few species, including Lactobacillus sakei, Lactobacillus curvatus,
and Lactobacillus paracasei ssp. paracasei (formerly L. casei ssp. pseudoplantarum) and
L. plantarum. Most typically, if both D(-)-lactic and L(+)-lactic acids are detected in the
41
cell, these end products are formed by the activity of single D-nLDH and L-nLDH
enzyme, respectively, without the contribution of lactate racemase. As a result of the
enzymatic diversity among LAB, differerent enantiomers of lactic acid can be produced
either exclusively (L(+)-lactic acid or D(-)-lactic acid, alone), approximately equal
amounts of both, or predominatly one form but measurable amounts of the other
(Axelsson, 2004). The characteristics pathway used for glucose fermentation and optical
isomer of lactic acid produced by different LAB genus are presented in Table 2.
Table 2. Fermentation pathway for glucose utilization and optical isomer of lactic acid
produced of main LAB genus (adapted from Axelsson, 2004).
1.6 Lactic acid –usage, applications and production
Lactic acid is a versatile molecule and is used in various applications in several industries
as demonstrated by the following examples (Vijayakumar et al., 2008, Wee et al., 2006):
x
x
x
food industry: acidulant, flavouring
agent, preservative
chemical industry: base chemical for
variety of chemical conversions, like
conversion to acrylic acid via
dehydration
pharmaceutical industry: electrolyte
in many parenteral/I.V. (intravenous)
solutions, ingredient in various
mineral preparations and
formulations (especially lactate salts)
x
x
x
cosmetic industry: moisturizing
agent, pH regulator
polymer industry: raw material for
the production of different grades of
poly(lactic acid) (PLA)
other functions in various industries:
dyeing of various textiles, mordant in
the printing of woolens, bating and
plumping of leathers, deliming of
hides, tanning of vegetables, flux for
soft soldering, green solvent in
several applications
Relevant chemical characteristics and most typical applications of the most commonly
known commercial lactic acid grades are presented in Table 3.
42
Table 3. Commercial available lactic acid grades and their main properties and usage
(Kubicek and Karaffa, 2006; Henton et al., 2006; Anonymous, 2016a).
According to recent market study (Anonymous, 2014), markets of lactic acid have been
steadily and rapidly growing since 2008 and the 800 000 ton/year milestone was reached
in 2013. According to this study, three major manufacturer dominated the markets
(PURAC, Cargill and Henan Jindan Lactic Acid Technology Co., Ltd), manufacturing
together about 505 000 ton of lactic acid. Production of PLAs was evaluated to be one of
the most potential applications of lactic acid, and during 2013 the global PLA production
was estimated to be about 320 ton/year, from which a single company (Natureworks)
represents almost half (45.2 %) of the total market share.
Although the chemical manufacturing of lactic acid started to expand already in early
1960s mostly due to demand for heat stable stearoyl lactylates in the baking industry
(Benninga, 1990), almost all lactic acid currently available on the market is produced by
fermentation (Groot et. al, 2010). One of the main reason for this is the fact, that chemical
synthesis yields always an racemic mixture of D(-)- and L(+)-lactic acid. In some cases,
manufacturing of thermostabile PLA grade as an example, this kind of end product
mixture is not the preferred choice for the further processing of the lactic acid (Lunt, 1998;
Södergård and Stolt, 2002).
Indeed, the promising market of PLAs has been the main driving force during the last
decades to develop microbial fermentation processes that are able to deliver pure
enantiomeric forms of lactic acid - either L(+) or D(-). The beauty of PLAs is that they
are bio-based, bioresorbable, and biodegradable under industrial composting conditions. In
addition, thermoplastic properties with rigidity and clarity, make some of the PLA grades
good alternative for many oil-based polymers. Indeed, because of these polymer properties
together with attractive price and commercial availability, PLA became the first massproduced bio-based polyester in the market and is the front runner in replacing the oilbased polymers with bio-based alternatives (Groot et al., 2010).
In many applications, it does not make any difference whether the optical structure of the
used lactic acid is D(-), L(+) or mixture of both enantiomers, but especially food and
pharmaceutical industries prefer to use pure L(+)-lactic acid, since elevated levels of the
D(-)-isomer are harmful to humans (Anonymous, 1967). However, the D(-)-lactic acid
43
plays also an important role in novel polylactic acid formulations, as the polymer blend of
pure L(+)- and D(-)-polylactic acids (stereocomplex, scPLA) seems to possess superior
temperature tolerance when compared to polymers made from pure L(+)- or D(-)poly(lactic acids) (Tsuji and Fukui, 2003). As the melting temperature of both pure
homopolymers falls between 170-180 °C, the melting temperature of the 50:50 (%) blend
is about 230 °C (Ikada et al., 1987). If the single polymer chain is not pure homopolymer,
pure poly(L(+)-lactic acid) polymer (PLLA) or pure poly(D(-)-lactic acid) polymer
(PDLA), but is made out of both enantiomers, the melting temperature will drop and
crystallization behavior will change dramatically. According to Groot et al. (2010),
already D(-)-content higher than 12-15 % in mainly L(+)-based PLA, is enough to make
the resulting PLA amorphous. According to Lunt (1998), in many aspects, the basic
properties of typical PLA polymers lie between those of crystal polystyrene (PS) and
polyethylene terephthalate (PET).
Before the start of mass production of PLAs, the most relevant use of these polymers was
typically focused to high value applications, like biomaterials in medicine (Lasprilla et al.,
2012), but currently various PLA formulations are used in much broader range - even in
single use packing applications (Hazarika et al., 2015). This indicates that the economics
related to PLA production has improved substantially.
1.6.1 Synthesis of PLA
Pure solution of lactic acid at equilibrium conditions is indeed a mixture of monomeric
lactic acid, dimeric lactic acid, higher oligomers of lactic acid and lactide (cyclic dimer).
The explanation for this kind of mixture lies in the nature of lactic acid molecule – it has a
hydroxyl and an acid functional group. Due to this specific chemistry, intermolecular and
intramolecular esterification reactions may happen. First, a linear dimer (lactoyl lactic
acid) is formed and this condensation reaction can further proceed to higher oligomers.
This reaction is promoted by removal of water. Also, lactide, a cyclic dimer, is formed in
small amounts by intramolecular breakdown of higher oligomers or esterification of
lactoyl lactic acid.
Although this kind of polycondensation process is possible without modern process
control and catalysts, as accidentally demonstrated by Gay-Lussac and Pelouze in the first
half of the 19th century (Benninga, 1990), the benefit gained by the usage of optimized
catalyst and polymerization conditions is crucial to extend the reaction and gain polyesters
with higher molecular weights (Hiltunen et al., 1997). Most typically, manufacturing of
PLA from the purified lactic acid solution takes place via one of the three alternative
routes described in the Fig. 8.
44
Figure 8. Various reaction pathways to yield high molecular weight PLA (modified from
Lasprilla et al., 2012).
Currently, the above described conventional polycondensation process is not the primary
mechanism for industrial high molecular weight PLA production, as in the early 1990s
Cargill Inc. developed a process based on ring opening reaction (ROP) of lactide for this
purpose. Indeed, ROP of cyclic esters was already invented before 2nd world war by
Carothers and later patended by Lowe (1954) for DuPont, but Cargill was the first
company that managed to adopt it into profitable large scale PLA process in mid 1990s
(Masutani and Kimura, 2014).
1.6.2 Conventional LAB based industrial scale fermentation processes for
lactic acid
Typical industrial scale microbial lactic acid process can be divided into three main
processing phases: fermentation, recovery and purification. These phases can further be
divided into other process steps or unit operations, as shown in example presented in Fig.
9. Depending on production site, this core process is integrated to longer value chain with
various ways. In literature, a good example of this is available in the report describing the
whole production chain of PLAs from cradle (starting from the cultivation of raw
materials for the fermentation) to gate in one of the PURAC’s manufacturing site (Groot
and Borén, 2010).
45
Figure 9. Traditional lactic acid production process – simplified block scheme (adapted
from Groot et al., 2010).
In lactic acid fermentation, among the most relevant decisions to be made are: choice of
the substrate, choice of the production organism, mode of the fermentation and
management of the cell density, and choice of the fermentation conditions (pH and
temperature). Indeed, from fermentor design point of view, conventional LAB based lactic
acid production processes are simple, as fermentations with these organisms do not
usually require aeration, gas exchange or gas mass transfer (Taskila and Ojamo, 2013) and
have low power and cooling needs (Datta et al., 1995). As reviewed by Taskila and Ojamo
(2013), typical lactic acid fermentation choices other than simple batch are repeated batch,
fed-batch and use of continuous fermentation either with free cell-recycling or with
immobilized cells. Success of fermentations is typically measured with parameters like:
titer (g lactic acid /dm3), productivity (g lactic acid/(dm3 x h)) and yield (g lactic acid/g
carbon substrate). Enantiometric purity of the lactic acid produced is also verified, if
necessary. Datta et al. (1995) proposed that, over 90% yield from the carbohydrate
substrates, final product concentration of at least 1M (90 g/l) and high stable productivity
(> 2 g/l h) are criteria for a successful fermentation process.
As reviewed by Idler et al. (2015), the most important lactic acid LAB producers and the
optical isomer(s) of lactic acid produced by these species are listed in the Table 4.
Table 4. Typical LAB species used for lactic acid production (adapted from Idler et al.,
2015).
L(+)-lactic acid producers:
D(-)-lactic acid producers:
Lactobacillus amylophilus
Lactobacillus casei
Lactobacillus paracasei ssp. paracasei
Lactobacillus rhamnosus
Lactobacillus manihotivorans
Enterococcus faecium
Lactococcus lactis
Streptococcus thermophiles
Lactobacillus delbrueckii
D(-)- and L(+)-lactic acid
producers:
Lactobacillus acidophilus
Lactobacillus helveticus
Lactobacillus brevis
Lactobacillus fermentum
Lactobacillus reuterii
Lactobacillus plantarum
Pediococcus acidilactici
Use of low cost non-edible raw materials for the fermentation is currently under research
in many companies. However, as stated by Groot et al. (2010) there are also some possible
challenges with these raw materials: raw materials may include some microbial growth
inhibitors, and/or purification of the lactic acid may be costly and cumbersome. Most
46
typically, the selection of a specific carbohydrate feedstock depends on the price and
purity of the various carbohydrate sources locally available. Traditionally, a wide variety
of carbohydrate sources, e.g. whey, molasses, corn syrup, dextrose and cane or beet sugar,
have been used. Typical nitrogen and nutrient sources are corn steep liquor, yeast extract,
soy hydrolysate, etc. (Datta and Henry, 2006). Most often, pH during the fermentation is
kept constant with addition of NH3, NH4OH, KOH, NaOH, NaHCO3, Ca(OH)2 or CaCO3
as a neutralizing agent. Most commonly used out of these are Ca(OH)2 and CaCO3. As the
fermentations are typically run at pH value above the pKA of lactic acid (3.8), this leads to
precipitation of lactic acid as lactate salt in high performance batch fermentations. This
kind of trapping of fermentation end product reduces the toxic effects of lactic acid on the
growth and morphology of microbial cells (Juturu and Wu, 2015). As the pKa of the lactic
acid is 3.8, the higher the fermentation pH is the more calcium lactate is generated due to
result of addition of pH adjustment chemical. In a typical homolactic lactobacillar lactic
acid batch fermentation temperature is around 40°C and pH varies between 5.4 and 6.4
(Domenek et al., 2011). Running the process under these conditions requires large
quantities of neutralizing agents and results in high amounts of insoluble lactate salt.
During the recovery, cells of production strain are removed and calcium lactate is
regenerated with addition of sulfuric acid (see Fig. 9), resulting in generation of free lactic
acid and formation of insoluble gypsum waste (Upadhyaya et al., 2014). Depending on the
final usage of the lactic acid, solution is also purified and concentrated with various
additional processing steps during the recovery and purification. Also, if significant
amounts of by-products are formed during the fermentation, those are separated during the
downstream processing and used for further processing, rejected or recycled. According to
Jem et al. (2010) most typical purification steps in lactic acid manufacturing include those
of chromatography, esterification, and/or distillation. In addition, membrane filtration,
crystallization and/or evaporation may also be employed to purify and/or concentrate
product to meet the desired end product specifications.
Good example of the product specifications is the requirement for the heat stability. As
stated by Groot et al. (2010), the formation of color upon heating prohibits the use of
crude acid in foods that need to undergo heating, like sterilization or pasteurization, during
the manufacturing process. Also, heat stability is crucial for the monomer grade lactic
acid, because during the step-growth polymerization processes, lactic acid is typically
heated to between 150-190°C (Södergård and Inkinen, 2011). Formation of the color
upon the heating is most often result of the presence of excess residual sugar and nitrogen
compounds in the solution. Thus, production of heat stable lactic acid requires some
additional processing steps to get rid of these compounds (Groot et al., 2010).
Another important criterium for plastic grade lactic acid is the optical purity of the lactic
acid. According to Jem et al. (2010), plastic grade lactic acid needs to be heat stable, and
typically the optical purity needs to be at least 98%. In the patent application filed by
Purac Biochem B.V (EP 2748256 A2, Noordegraaf and De Jong, 2012), optical purity of
at least 95 %, but preferable at least 99.5 % was set for qualification criteria for
manufacturing PLA grade plastics. As stated by Jem et al. (2010), also the processing
time with high temperatures should be minimized during recovery and purification, as it
has been observed that racemization of D(-)-lactic acid to L(+)-lactic acid, and vice versa,
is prone to progress under these conditions. As described in Cargill Dow LLC’s patent
application (WO2001038284 A1, Quarderer et al., 2000), high temperature alone (180°C
47
or higher) or the presence of suitable catalyst may start the racemization. As an example of
catalyst, salts of group I and group II were mentioned (e.g. sodium, potassium, calcium
and magnesium salts). These two factors operate also together, so the exposure of lactate
material to temperatures above 100°C with the catalyst present can cause significant
racemization.
1.6.3 Modern technologies in industrial bacterial fermentations– immobilization of cells
for extracellular production of metabolites
As defined in the literature by Karel et al. (1985), whole cell immobilization refers to the
localization of intact cells to defined region of space with the preservation of catalytic
activity, whereas Abbott (1977) pointed out that immobilization is physical confinement
or localization of a microorganism that permits the economical reuse of the
microorganisms. As reviewed by Abott (1977), one of the first examples of the usage of
immobilized whole cells was the “quick-vinegar” production process developed by
Schuetzenbach in 1823. In this process, wooden vat with perforated bottom was filled with
wood shavings and the substrate, ethanol solution, was trickled through this vat, resulting
in outflow of acetic acid solution. At that time, it was believed that the conversion of
ethanol to acetate was due to a chemical reaction. Only after a couple of decades later,
Pasteur found the connection between fermentations and the presence of micro-organisms.
End and Schöning (2004) have weighted the potential advantages and disadvantages of
use of immobilization technology as follows:
x
x
x
x
x
x
x
Potential advantages:
Ease of developing continuous
processes
Fast reaction rate due to high catalyst
concentration (for certain reactor types)
Higher activity by better availability of
catalytic centers
Repetitive use of biocatalysts
Higher resistance to shear stress and
contamination
Higher stability with regard to
temperature, pH, and catalyst poisoning
Easy separation from reaction media
(easier downstream processing)
x
x
x
Potential disadvantages:
The necessity of developing an
immobilization process
The existence of mass transfer
resistances (diffusion limitations)
Additional cost caused by support and
additional reagents
Methods for the whole cell immobilization have been reviewed by Pilkington et al. (1998)
and Kourkoutas et al. (2004) and those are usually categorized based on the physical
mechanism employed for the immobilization:
(a) attachment or adsorption on
solid carrier surfaces
(b) entrapment within a porous
matrix
(c) self-aggregation by flocculation
(natural) or with crosslinking
(d) cell containment
48
Figure 10 illustrates typical solutions and releated mechanisms how the above mentioned
four physical immobilization methods may be applied for various immobilization systems.
Also, simplified model for the biofilm formation and natural flocculation is presented.
Figure 10. Most typical whole cell immobilizations methods and dynamics of biofilm
formation and deformation. Adapted from Fukuda 1994; Kourkourtas et al., 2004; Ploux
et al., 2010; Hall-Stoodley et al., 2004; Wingender and Flemming 1999.
Adsorption is generally regarded as the intial attachment of cells to solid surface (or
deposition), whereas aggregation is one mechanism for the subsequent expansion of the
initial adhesion population. If aggregates are not attached to a surface, term ‘floc’ has
been used to describe these cellular aggregates (Hall-Stoodley et al., 2004). The
importance of autoaggregation, reversible aggregation of similar cells to form discrete
groups (Dube, 2005), in beer brewing was first understood by Pasteur 1876 when studying
alcoholic yeasts fermentations. In these fermentations, liquid suspended aggregates are
most typically formed on the top (surface yeast) or on the bottom (bottom yeast) of the
fermentation vessel. Bacterial cells may also be cross-linked together with chemicals.
This kind of chemical treatment leads to formation of artificial flocs. Cross-linking agents
can also be used to bind cells to an organic or inorganic support by creating covalent
bonds between cells and support material. However, covalent bonding is not the preferred
way to immobilize living cells, since many of the chemicals used in these methods are
highly toxic for the cells (Genisheva et al., 2014). Alternatively, cells can be artificially
bound to solid support by applying various bioaffinity binding strategies. Many of these
systems requires cumbersome and costly modifications (surface engineering of bacteria
and/or need of special functional groups attached to binding carrier) and are not viable for
large scale industrial productions. However, in the scientific literature there are some
promising reports available that are based on genetically modified bacteria which are able
to adhere by bioaffinity mechanism to cheap immobilization supports without any
significant pretreatment steps of the support material. As an example, Wang et al. (2001)
demonstrated the cellulose binding capacity of genetically engineered E. coli strain and
49
ùimúek et al. (2013) managed to construct genetically engineered L. lactis strain that is
capable to bind to chitin.
Immobilization of whole cells is also possible by enclosing bacteria inside a layer which
permits the necessary nutrient and product flow for the biocatalysis. Genisheva et al.
(2014) have reviewed the positive and negative aspects of this technology in their recent
publication. Among LAB, the most widely used immobilization method in various food
fermentations is based on cell entrapment using natural polymers like alginate, agarose,
carrageenan, chitosan, and pectin. These natural gelling polysaccharides are non-toxic,
biocompatible, cheap, and offer versatile techniques for biocatalyst preparation. The most
common drawbacks of this immobilization method are the limited cell proliferation and
activity (due to mass-transfer limitations in matrice) and lack of regeneration method for
this kind of immobilization (Kosseva et al., 2011).
Adsorption of a bacterial cell to solid surface and the following putative biofilm formation
is a complex process, which involves several steps and is affected by several variables.
The initial bacterial adsorption to a surface is a reversible process, allowing also
desorption from the surface if the net repulsive forces are greater than the net attractive
forces (Garret et al., 2008). As summarized by Araújo et al. (2010), adhesion of bacteria to
a solid surface is dependent upon van der Waals, electrostatic, and acid–base interactions.
Furthermore, the physiochemical properties of surface and the characteristics of the
bacterial surface, like hydrophobicity, surface charge, and electron donor–electron
acceptor properties, are influencing the above mentioned interactions.
First task for a planktonic cell (a single cell floating in a bulk phase) is to get closer to the
surface. This task can be initially fueled by Brownian motion, sedimentation due to
differences in specific gravity between the bacteria and the bulk liquid, or by movement of
the bulk fluid (convective mass transport). Also, some bacteria may move actively using
flagella to move toward target surface (Palmer et al., 2007). When getting closer to the
surface, typical long range forces like van der Waals and electrostatic interactions, but
later also and hydrophobic/hydrophilic interactions may effect the movement of bacterial
cells towards the surface. If the bacterial cell is able to overcome the energetic barrier
surrounding the binding substrate, it may get a step closer to irreversible adsorption. In
this battle, the above mentioned forces are taken into account, but also short range
interactions, such as covalent and hydrogen bonding interactions are part of the equation.
These interactions are receptor/ligand-type of interactions and are due to specific
properties present on the bacterial cell surface but also include the specific properties of
the binding substrate. Examples of specific bacterial surface properties which may
promote adhesion with the target surface are bacterial flagella and pili (Ploux et al., 2010).
In some cases, irreversible binding never happens and bacteria detach from the surface and
return back to the planktonic form of life. This was also pointed out in the review written
by Kosseva et al. (2011), as the low binding force between the cell and immobilization
substrate was considered to be one of the weaknesses of the adsorption based
immobilization systems. However, the other side of the coin in this matter is the relative
easy regeneration of the carrier.
If irreversible adsorption is initiated, in some bacteria it is enhanced by cell’s capability to
sense the surface (“surface sensing”) and to adapt rapidly its adhesive arsenal (e.g. by
modifying the cell wall composition by expressing adhesive appendages or receptors)
50
(Ploux et al., 2010). Soon, this initial adhesion of first bacteria to cell surface is followed
by colonization of the immobilization support, leading to the potential formation of a
biofilm (Junter and Jouenne, 2004).
According to Saxena (2015), most typically the cellular growth in artificial immobilization
systems is restricted. However, as reviewed by Junter and Jouenne (2004), in some cases
growth in immobilized system may be ever faster than in the corresponding free cell
system. In most cases, steady state conditions are preferred in continuous fermentation
processes. When considering the immobilized production strain in this respect, it usually
means that the total mass of bacterial cells inside the system remains close to constant
after the ramp up and stabilization of the continuous fermentation process. The thickness
of biofilm in steady state situation is function of growth and senescence (temperature,
nutrient supply etc) and biofilm degradation (shear forces, film stability) (Patching and
Fleming, 2003). Furthermore, several bacteria are able to produce extracellular polymeric
substances (EPS) and even use specific signalling molecules to promote the growth and
stability of the biofilms and cell-cell communication in this environment. In general, cells
in biofilms are physiologically distinct from planktonic cells and in many cases possess a
better stress tolerance than their planktonic relatives (Davies, 2003).
1.6.3.1 Potential solid supports for industrial scale immobilized fermentations
When selecting an immobilization carrier for a specific industrial process, the next aspects
should be considered case by case as proposed by Virkajärvi and Linko (1999):
x
x
x
x
type of immobilization
price of material
ease of immobilization
cell load
x
x
x
x
x
mass transfer
x
x
channeling or blocking of reactors
x
x
stability
x
rigidity
regeneration
sterilization
binding characteristics of
contaminating microorganisms
freedom to use various reactor
designs
possibility of fluidization (also for
regeneration)
approval for food use
As reviewed by End and Schöning (2004), typical solid support used for adsorption based
immobilization include those of:
Organic supports:
x cellulose, chitine (chitosan), wood,
lignin
x nylon, polypropylene fibres, polymer
nets, membranes
x polyurethane foam, polyacrylamide
Inorganic supports:
x (porous) glass, clay, bentonite,
x
x
ion-exchange resins
x
zeolithes, ceramics, mesoporous silica
x
metal oxides (Fe, Ti, Mg), metal
phosphates
mineral powder
51
1.6.3.2 The potential of new raw materials and whole cell immobilization in industrial
scale LAB fermentations
The first industrial large scale application using immobilized cells was the E. coli based
continuous L-aspartic acid manufacturing process that was successfully started in 1973 by
Tanabe Seiyaku Co. Ltd. (Chibata et al., 1979; Linko and Linko,1983). Currently,
probably the most significant commercial application that is based on the use of whole cell
immobilization technology is the beer manufacturing process with immobilized yeast
strains. Within this industry, commercialization of immobilization technology has been in
focus already for several decades and it has been utilized with various degrees of success
for a number of purposes, like those of continuous primary fermentation, low-alcohol beer
production and secondary maturation (Nedoviü et al., 2015). In wine industry, the
potential of immobilized cells has not only been studied for the alcoholic fermentation
(typical yeast fermentation), but also for the subsequent malolactic fermentation (MLF).
In the wine manufacturing process, malolactic fermentation (MLF) is a secondary
fermentation, which is needed in some wine grades to decrease the acidity and for subtle
modification of the aroma. During the malolactic fermentation, L-malic acid is
transformed into L-lactic acid and carbon dioxide. So, MLF starts when alcoholic
fermentation has finished and it is typically a result of the fermentation by lactic acid
bacteria that most typically are member of genus Lactobacillus, Pediococcus, Leuconostoc
or Oenococcus (Genisheva et al., 2014). Oenococcus oeni (formerly known as
Leuconostoc oenos) is one of the most typical strains found in the wine during
spontaneous MLF (Lafon-Lafourcade, 1983; Solieri et al., 2010). Indeed, it is also
available as commercial freeze-dried bacterial culture, readily used by the wineries, to
gain more control over this challenging process (Nielsen and Richelieu, 1999). As
reviewed by Nedoviü et al. (2015), several potential industrial whole cell immobilization
processes for wine making have been proposed, including also those based on LAB - like
the use of O. oeni for MLF. However, significant breakthrough in commercialization of
these processes in large scale is still pending.
During the last decades, several articles have been published in which the use of whole
cell immobilization of lactic acid bacteria in lactic acid (see reviews of), dairy and starter
industries have been reviewed (Abdel-Rahman et al., 2013; Kosseva et al., 2009; Kosseva
et al., 2011). In spite of the high scientific interest in this field, it seems that the final
breakthrough in the commercial scale has not occurred, yet. As reviewed by Thongchul
(2013), the reduction of product formation and instability of the immobilized cells during
long-term fermentation are the main challenges to be resolved in order to make the
immobilized (whole cell) lactic acid process commercially more attractive.
Another key element for sustainable and economical attractive lactic acid fermentation is
the source, quality and cost of the substrates. As presented in the objectives of the major
lactic acid producers in the world, technology to use the 2nd generation feedstock (nonfood cellulosic raw materials) in industrial scale has been estimated by both Corbion
(Purac) and NatureWorks to reach the maturity between 2015-2016. Indeed, according to
Lin and Tanaka (2006) cellulose materials represent the most abundant global source of
biomass. Over 90% of this biomass is in the form of lignocellulose. Clearly, there is a lot
of potential in these raw materials, but the commercial use of lignocellulose in lactic acid
production is still challenging because physicochemical pretreatments and multienzymatic reactions are required to yield monomeric hexoses and pentoses for the lactic
52
acid fermentation (Abdel-Rahman et al., 2011). Starchy non-edible raw materials, like
agro-industrial waste and food waste, provide an easier solution for the carbon source for
the lactic acid fermentations. However, as pointed out by Reddy et al. (2008) and Mazzoli
et al. (2014), relative few LAB have been reported to be capable to ferment starch directly
to lactic acid. As reviewed by Kosseva et al. (2009), the use of cheese whey as a substrate
for the lactic acid fermentation has been studied widely. Although a part of the cheese
whey can be utilized (e.g. by food and pharmaceutical industries), a significant amount of
whey is disposed with dairy wastewaters – increasing the nutrient loads in waste water
treatment plants. Economically, one of the most attractive scenario for utilizing whey is to
separate the whey protein and sell it as high value protein isolates products and then to
find relevant use for the remaining lactose fraction. As lactose is the natural substrate for
many milk related LAB, many scientists have focused on studying the use of this twocarbon sugar in LAB based lactic acid fermentations. Not surprisingly, major lactic acid
producers do not reveal details from their processes, but small Irish start-up company,
Cellulac, has opened the door for some process details. As claimed on their www-pages
(http://cellulac.co.uk/en/design-objectives/, accessed 15th of October 2015), company has
developed technology to flexible utilize whey and lignocellulosic biomass (wheat straw,
spent brewery grains from beer production, dried distilled grains from ethanol production)
for manufacturing of optically pure D(-)-lactic acid by Lactobacillus strains capable to
hydrolyze disachharides like lactose, but also pentoses and hexoses. Furthermore, in May
2014 they claimed to complete the world’s first ever industrial level continuous
production of lactic acid from deproteinized lactose whey.
In scientific literature, there are some good reviews of non-industrial scale lactic acid
fermentations based on the use of immobilized LAB. As an example, Kosseva et al.
(2009) reviewed the whey based fermentations, whereas Abdel-Rahman et al. (2013)
widely reviewed not only the use of pure sugars but also the use of different starchy and
lignocellulosic carbon sources for various fermentation approaches.
1.7 Genetic engineering of Gram-positive bacteria to achieve
whole cell immobilization with industrial potential
Although several researches studying the whole cell immobilization of Gram-positive
bacteria have been published, only a small fraction of the proposed solutions are suitable
for large scale industrial applications. Due to small number of LAB related publications
in this area, the scope here is widened to cover also some other Gram-positive bacteria. In
this section, examples are given from studies (Table 5) in which genetic engineering has
been applied to modify the surface properties of given host strain and in which the nature
of the materials used for immobilization is close enough to those that could be potentially
used also in the industrial scale. Primarily, the focus is on those solutions in which
bacteria are able to immobilize to a solid organic or inorganic carrier by direct noncovalent binding.
53
Table 5. Use of genetic engineering to achieve whole cell immobilization (Gram-positive organisms) with
industrially interesting carriers.
54
1.7.1 Immobilization on collagen surfaces by non-covalent binding
Antikainen et al. (2002) studied the collagen binding properties of the assembly region of
CbsA (S-layer protein) of L. crispatus. In this work, authors studied which regions in this
protein are required for binding to collagens and laminin, anchoring to the bacterial cell
wall and self-assembly of the S-layer protein. For this task, authors made several
constructs by inserting DNA fragments encoding various parts of CbsA in to pLPMSSA3
vector under an inducible Į-amylase promoter. In this vector, the cbsA fragments were
inserted in frame with the LPXTG cell wall-anchoring motif derived from the surface
protease (PrtP) of L. casei. The resulting constructs were expressed in L. casei and the
surface display of the N-terminal CbsA fragments was confirmed by ELISA. Successful
binding of cells to type IV of collagen, but also to laminin, was demonstrated for several
constructs. Most efficient binding to both collagen and laminin was observed with strain
expressing CbsA residues 28-287. As no S-layer structure was observed in these cells,
authors speculated that the above described binding was putatively achieved by the
LPXTG bound CbsA peptides displayed on the bacterial surface.
1.7.2 Immobilization on starch surfaces by non-covalent binding
Tarahomjoo et al. (2008) fused the C-terminal LysM domain of peptidoglycan hydrolase
(CPH) of L. lactis IL1403 with the linker region and the starch-binding domain (SBD) of
Į-amylase of S. bovis 148. The DNA construct expressing this fusion protein was
expressed in E. coli, and the resulted protein was isolated and purified from the cell lysate.
Purified fusion protein (designated as CPH-SBD) was first mixed with L. casei ssp. casei
NRRL B-441 to allow binding of the fusion protein with cells, followed by the
immobilization study with the corn starch granules. Measurement of the optical density
was used to calculate the adhesion percentage to corn starch. The adhesion percentage
under the optimal conditions for the cells was 32% whereas the same result for the control
(free cells) was 4 %. Authors also discovered by phase contrast microscopy observation
that CPH-SBD bound bacteria mediated the crosslinking of starch granules, resulting in
aggregate formation in the solution. As estimated by the authors, in average 6 ×104
molecules of CPH-SBD fusion protein were bound per one L. casei cell.
1.7.3 Immobilization on cellulose surfaces by non-covalent binding
Lehtiö et al. (2001) studied the immobilization of genetically modified Staphyloccus
carnosus on cotton fibres. Authors constructed an expression vector, in which the gene
region encoding the cellulose binding domain (CBD) of Trichoderma reesei cellulase
Cel6A of was fused with the promoter, signal sequence, propeptide encoding region of
lipase gene from Staphylococcus hyicus. This propetide encoding DNA fragment was
included in the construct, as it was known to promote the secretion of heterologous
proteins in S. carnosus when co-expressed with lipase signal. In order to enable cell wall
anchoring, the above described DNA construct was fused with gene fragments encoding
55
the serum albumin binding region (ABP) of protein G of Streptococcus sp. G148 and the
cell surface-anchoring regions X and M of protein A of S. aureus. The final expression
construct was transformed into S. carnosus host by protoplast transformation and the
resulting strain was used for immobilization studies with cotton fibers. Quantification of
the cellulose-bound cells was based on the addition of biotinylated human serum albumin
(binds with ABP tag) to the cotton fibers, followed by the addition of the streptavidinalkaline phosphatase conjugate. Finally, the amount of specific binding of the
streptavidin-alkaline phosphatase conjugate with ABP tag was visualized by addition of
chromogenic substrate. Authors demonstrated that the immobilization efficiency of the
strain expressing the CBD-fusion protein was more than double of the negative control.
This result was also confirmed by light miscroscopy.
Recently, there has been a lot of scientific interest to express minicellulosomes on the
surface of Gram-positive bacteria. Minicellulosomes, or designer cellulosomes, are
composed of a chimeric scaffoldin which displays an optional cellulose binding module
(CBM) and cohesins of different specifities which bind selectively to the cellulolytic
enzymes appended with the matching dockerings (Mingardon et al., 2007). By using this
approach, CEM (cellulose-enzyme-microbe) complexe are formed (You et al., 2012), in
which the cellulose binding domain thether the microbe-cellulosome complex with the
cellulosic substrate and allows it to progressively degrade cellulose by crawling along its
strands (Wieczorek et al., 2013). Cell wall attached minicellulosomes with cellulose
binding module attached within the scaffold have already been constructed for several
industrially attractive Gram-positive micro-organisms, like those of B. subtilis, L. lactis
and Corynebacterium glutamicum (You et al., 2012; Wieczorek and Martin; 2010; Kim et
al., 2014). In these studies different kind of cell wall anchors were applied for anchoring
of the scaffolding to the cell wall: You et al. (2010) used the LysM domain of cell wall
hydrolase (LytE) of B. subtilis, Wieczoreck and Martin (2010) used LPXTG anchor of M6
protein of S. pyogenes and Kim et al. (2014) utilized the transmemrane structures of the
mechanosensitive channel (Msc) of C. glutamicum ATCC. The cellulose binding domains
in these scaffolds were originated either from Clostridium thermocellum or Clostridium
cellulovorans (see Table 5 for details). As the primary target for these studies was not to
characterize the cellulose binding efficiency of the related recombinant cells but to
demonstrate the enhanced cellulolytic activity of these strains, no specific data to evaluate
the cellulose binding efficiency of the different recombinant strains was published.
1.7.4 Immobilization on chitin surfaces by non-covalent binding
ùimúek et al. (2013) constructed L. lactis strains capable to be immobilized on chitin. The
chitin binding property of these strains was achieved by surface display of the ChBD
(chitin binding domain) of chitinase A1 of B. subtilis on the surface of L. lactis nisin
producing strains N8 and LL27. In the fusion constructs, ChBD was fused with PrtP
anchors of three different lengths, and also with an AcmA anchor. All these anchors were
derived from L. lactis. The gene constructs coding these anchor-ChBD fusions were
expressed under the P45 constitutive promoter and usp45 signal sequence. Surface display
of the expressed ChBD in L. lactis was verified with whole cell ELISA and the whole cell
binding with chitin flakes was tested in an immobilization study. The nisin production
56
capability of the engineered strains was also evaluated by repeated cyclic fermentation test
with immobilized chitin flakes. According to the results obtained from all of these
experiments, the strains carrying either the longest or the second longest PrtP anchors,
800 and 344 C-terminal amino acids, respectively, outperformed the strains displaying the
LysM based immobilization construct (242 amino acids) and the shortest PrtP
immobilization construct (153 amino acids). In cyclic nisin production test, the highest
productivity was measured with the strain carrying the longest PrtP anchor, although it
was not statistically different from that obtained with the strain carrying the second
longest PrtP anchor. The PrtP-ChBd fusion construct with the longest PrtP anchor
demonstrated also the highest immobilization percentage on chitin, being 91 and 94 %,
when using N8 and LL27 hosts, respectively.
1.7.5 Immobilization to inorganic or synthetic surfaces by non-covalent
binding
Basicly, the strategies to enhance immobilization of bacteria on a surface can be divided
into specific methods and non-specific ones. The former include methods based on
specific affinity between the binding ligand and the binding substrate (as is the case with
CBD and cellulose), whereas the latter methods are based on more general interactions
and characteristics, like to the overall surface charges or hydrophobicities, expression of
protruding appendages on the bacterial surface or roughness of the immobilization
material. Most often, the adsorption type of bacterial immobilization on solid surfaces like
glass or plastic, has been based on these non-specific interactions, although during the
recent years efforts to generate more specific synthetic peptide adhesins also for these
materials have been increased (Karaca et al., 2014). Examples within this are the
development of bioadhesive peptides for myocardial tissue engineering with polymeric
materials (Rosellini et al., 2015) and construction of peptides with high affinity to bonelike apatite-based minerals (Segvich et al., 2009). However, it seems that the
technological maturity to use these very specific peptides for industrial scale whole cell
immobilization has not been reached, yet. Instead, during the recent years some promising
results have been published about the use of peptide adhesins with a wider binding
spectrum for immobilization of Gram-negative bacterial cells on various inorganic or
synthetic solid supports. As an example, Park et al. (2014) evaluated the use of mussel’s
adhesive catecholamine moiety to generate a sticky E. coli strain.
When considering Gram-positive bacteria, cell wall binding domains of lactococcal cell
surface proteinases (most typically PrtP) are one of the most widely used anchors in
Gram-positive surface display systems. Interestingly, PrtP is also known of its capability
to enhance bacterial adhesion to some solid surfaces, like glass or PTFE. The biological
role of this protein in milk related LAB is to initiate the proteolytic degradation of the
milk casein to smaller peptides (Siezen, 1999). Habimana et al. (2007) studied the affinity
of the lactococcal cells displaying various alleles of PrtP to solvents with different
physico-chemical properties, and cell adhesion of the resulted variant strains to solid glass
and tetrafluoroethylene surfaces. In these test three different strains of L. lactis ssp.
cremoris were used: MG1363: PRTP+ (PrtP anchored and active, pGKV2-based
construct), PRTP* (PrtP anchored and inactive, pGKV2-based construct) and PRTP(MG1363 carrying vector plasmid pGKV2 without the prtP gene) as control strain.
57
Researchers found that the exposure of an anchored cell wall proteinase PrtP, undependent
of its biological activity, was responsible for greater cell hydrophobicity and adhesion.
When compared to the negative control (PRTP-), adhesion to both solid supports was
enhanced with cells displaying either active or inactive form of PrtP on their surface. As
the properties of these solid supports were very different (PTFE: apolar and hydrophobic,
glass: polar and hydrophilic) and affinity to all various solvents (apolar, Lewis-acid or
Lewis-base) increased, Habimana et al. (2007) proposed that the increased immobilization
to glass or PTFE was due to surfacial presence of PrtP, which results in cell’s increased
capacity to exchange attractive van der Waals interactions. As a results of this phenomena,
authors suggested that bioadhesion of PrtP displaying lactococci is increased to various
types of surfaces (e.g., inert, polar or apolar, or organic). Interestingly, the highest
immobilization on solid surfaces was achieved with the surface display of the inactive
PrtP. Inactivation of the PrtP was achieved by two point mutations (Asp-30 to Asn-30) in
a catalytic site (Haandrikman et al., 1991). Habimana et al. (2007) speculated that the
weaker immobilization capacity of the strain dipalying the active PrtP was due to
degradation of AcmA (AcmA activity increases bacterial adhesion as demonstrated by
Mercier et al. (2002)) and/or by self-cleavage of the exposed surface proteins by the active
PrtP. The 100 x change in the salt concentration (NaCl: 1.5 mM to 150 mM) did not
considerably affect to the binding with solid substrates and within the tested salt
concentrations the PRTP* strain showed about six to eight times more efficient binding to
PTFE and about eight to ten times more efficient binding to glass when compared to that
of the control strain (PRTP-).
1.8 Genetic engineering of LAB to improve optical purity of
the produced lactic acid
1.8.1 Various biosynthetic routes leading to D(-)- and L(+)-lactic acid
formation in LAB
In LAB, lactic acid is most typically generated by lactate dehydrogenase which reduces
the end product of glycolysis, pyruvate, with the concominant regeneration of NAD+
(NADH oxidation). As described earlier, lactic acid is a chiral molecule and depending on
the LAB strain, D(-), L(+) or both enantiomers are produced by D(-)- and/or L(+)stereospecific lactate dehydrogenases (LDHs). In addition to lactate dehydrogenases, some
LAB produce these lactate isomers also by stereospecific 2-hydroxyisocaproate
dehydrogenases. Production of L-lactate may also take place by malolactic enzyme, which
is capable to produce this lactate isomer by decarboxylation of L-malate. Finally, in some
LAB lactic acid may putatively be produced through methylglyoxal detoxification
pathways. All these pathways are described in Fig. 11.
In lactic acid bacteria, lactate dehydrogenases that catalyze the reduction of the pyruvate
to lactic acid are typically NAD-linked (or NAD dependent) enzymes (nLDHs), which
makes sense as the reduced nicotinamide adenine dinucleotide molecules (NADHs)
generated during glycolysis need to be regenerated to maintain the fermentative life style
58
(Garvie, 1980). Furthermore, some of the L-nLHDs are allosterically activated by fructose
1,6-diphosphate (FDP dependent L-nLDHs) – by one of the key intermediates of the
glycolysis. More recently, Feldtman-Salit et al. (2013) have studied the regulation of some
L-LDHs of lactic acid bacteria more closely. According to Garvie (1980) allosteric
regulation is especially typical for many streptococcal L-nLDHs, although the most
extensively studied allosteric L-nLDH is that of L. casei.
Figure 11. Various potential metabolic pathways for lactic acid production and utilization
among LAB. Reations shown with bold white arrows are not described in detail.
*Cofactors vary for different iLDHs (NAD-independent lactate dehydrogenases).
According to Garvie (1980), the other major group among LDHs is NAD-independent
lactate dehydrogenases, i.e. iLDHs. These enzymes do not need NAD/NADH as a
coenzyme, and typically catalyze only lactate oxidation to pyruvate or acetate (lactate
utilization). Recently, Jiang et al. (2014) have reviewed the different types of bacterial
iLDHs. These enzymes are also stereospecific, and that property is beneficial for
enantiomeric purification of mixtures of D(-)- and L(+)-lactates – other optical enantiomer
of lactic acid is oxidized while the other enantiomer remains intact. Also, many D-nLDHs
and some FDP-independent L-nLDHs are capable to catalyze reaction between pyruvate
and lactate in both directions (Garvie, 1980; Goffin et al., 2004). Capability to catalyze
both oxidation and reduction reactions have also been confirmed for 2-hydroxyisocaproate
59
dehydrogenases (Hummel and Kula, 1989). The main functional difference between
lactate dehydrogenases and 2-hydroxyisocaproate dehydrogenases is the much wider
substrate specificity of the latter group. Most lactate dehydrogenases accepts only
pyruvate and in minor degree 2-oxobutyrate, the closest longer structural homologue for
pyruvate, as substrates (Hummel and Kula, 1989). However, 2-hydroxyisocaproate
dehydrogenases (HicDHs) are usually able to convert also 2-keto acids with branched or
aromatic side chains to corresponding 2-hydroxy acids (Hummel, 1999). Evolutionarily,
D-LDHs and D-HicDHs are related (Kochhar et al., 1992), as is the case with L-LDHs and
L-HicDHs (Lerch et al., 1989), too.
Not much is known about the putative methylglyoxal detoxification pathways in LAB.
Most of the cellular methylglyoxal is synthesized by methylglyoxal synthase (MGS) from
dihydroxyacetone phosphate (DHAP) by the removal of the phosphate group under
conditions of carbon excess or phosphate limitation (Booth et al., 2003). However, the
resulting metabolic intermediate, methylglyoxal (MG), is toxic for the cell because it is
structurally a reactive Į-ketoaldehyde (Murata-Kamiya and Kamiya, 2001). In bacteria,
the main role of this alternative non-ATP generating glycolytic branch is to restore
inorganic phosphate levels in stress conditions. As bacteria can also encounter MG in the
environment, too, the detoxification capability of MG is an advantage in evolutionary
competition (Chakraborty et al., 2015).
Currently, there are several different routes described in the literature for methylglyoxal
(MG) detoxification (Fig. 11). Out of these, the glyoxalase I & II or III activity dependent
routes and the route based on glycerol dehydrogenase and lactaldehyde dehydrogenase
lead to generation of the D(-)-lactate, while the route catalyzed by methylglyoxal
reductase and lactaldehyde dehydrogenase leads to production of L(+)-lactate. Out of
these enzymes, glycerol dehydrogenase and lactaldehyde dehydrogenase are most
common among LAB, whereas glyoxylase I and II –like enzymes have been reported so
far only in few LAB. Furthermore, the presence of the glyoxylase III-like enzymes in LAB
has not been reported, yet. In the following paragraphs, there are few examples of LAB
research papers, which include data of the presence and functions of the catabolic
glyoxalase enzymes among LAB.
In general, MG metabolism seems to be most abundantly detected, or at least best known,
among enteric bacteria. In accordance with this, an enteric LAB strain, E. faecalis V583 is
one of those few LABs in which the genes related to MG metabolism has been putatively
detected and expression level studied (Yan et al., 2015). In this strain, nine putative genes
have been indentified to encode proteins belonging to the glyoxalase family. As detected
by the authors, the abundancy of transcripts from several transport systems in this strain
was incresead under oxidative stress conditions. However, transcription of many other
genes, like that of the putative MG synthase and those five sharing the sequence identity
with genes of the glyoxalase I–II system, was decreased under the same stress conditions.
As suggested by the authors, the reduction in the abundance of transcripts encoding
cellular glyoxalases demonstrates the importance of the fine tuning of the intracellular MG
content for the oxidative stress adaptation for E. faecalis. As reviewed by Kant et al.
(2010), Glyoxylase I –like enzyme has been putatively identified in silico among some
LAB strains. One of these is Streptococcus mutans, in which the glyoxylase I–like enzyme
has been functionally characterized by Korithoski et al. (2007). As observed by these
authors, the lactoylglutathione lyase (LGL) in this strains (a glyoxalase I enzyme)
60
functions to detoxificy methylglyoxal, resulting in increased aciduricity. However, the
authors did not further identify the nature of the acid produced by this strain. These
authors also observed some glyoxalase II activity in S. mutans, although they could not
identify glyoxalase II genes in silico.
Finally, production of lactate may take place through lactaldehyde intermediates.
Oxidation of the L-lactaldehyde or D-lactaldehyde by NAD-dependent lactaldehyde
dehydrogenase leads to generation of L(+)-lactate or D(-)-lactate, respectively. In the case
of L(+)-lactate, the reduction of methylglyoxal to L-lactaldehyde is catalyzed by
oxidoreductase enzyme. Chaillou et al. (2005) detected genes in L. sakei 23K for
methylglyoxal bypass: These genes encoded putative enzymes capable to synthesize MG
from dihydroxyacetone phosphate (by methylglyoxal synthase) and to convert it to L(+)lactate (by oxidoreductase and aldehyde dehydrogenase). In a later study, McLeod et al.
(2011) observed up-regulation of the first two genes in this pathway (methylglyoxal
synthase and putative oxidoreductase) after the change of carbon source from glucose to
ribose. However, no induction was observed for the putative iron-containing aldehyde
dehydrogenase, suggesting that reduction of L(+)-lactaldehyde to lactate was not catalyzed
by this putative enzyme. Among LAB, lactaldehyde dehydrogenase activity has also been
proposed to function in the other direction (catalyzing lactate reduction), especially in
some silage related LAB. These strains are able to reduce lactate first to lactaldehyde and
then further to 1,2-propanediol with the concominant production of acetic acid and ethanol
from lactate (Elferink et al., 2001; Saxena et al., 2010).
It is known that the NAD-dependent lactaldehyde dehydrogenase is also able to catalyze
the oxidation of D-lactaldehyde to D(-)-lactate, although reactions with this optical
enantiomer are usually slower when compared to that of L-lactaldehyde (Purich and
Allison, 2002). In case of E. coli, the production of D-lactaldehyde from methylglyoxal by
glyoxalase I & II or III independent route has been proposed to happen by glycerol
dehydrogenase activity (Altaras and Cameron, 1999). As stated by authors, this kind of
reaction seems to be possible because the wide substrate specificity displayed by many
known glycerol dehydrogenases. In the lactate production by this route, the ketone group
of methylglyoxal is first stereospecifically reduced to give D-lactaldehyde, followed by
oxidation by lactaldehyde dehydrogenase to yield D(-)lactate (Altaras and Cameron,
1999).
Racemases catalyze the inversion of stereochemistry in biological molecules (Desguin et
al., 2014). In case of lactate, lactate racemase activity has so far been detected only in few
species among LAB, including of those of L. plantarum (Ferain et al., 1996), L. sakei
(Hiyama et al., 1968), L. curvatus (Stetter and Kandler, 1973) and L. paracasei ssp.
paracasei (formerly L. casei ssp. pseudoplantarum) (Stetter and Kandler, 1973). The best
characterized lactate racemase system is that of L. plantarum WCFS1 (Goffin et al., 2005;
Bienert et al., 2013; Desguin et al., 2014; Desguin et al., 2015). This strain is able to
produce both D(-)- and L(+)- lactate from sugars without lactate racemase, but it is also
capable of interconversion of lactate isomers by lactate racemase (Lar) that is induced by
the L(+)-lactate and transcriptionally controlled by the L(+)/D(-) lactate ratio. It has been
suggested that the physiological role of the Lar in L. plantarum is to secure the availability
of D(-)-lactate for the cells, as the structure of the L. plantarum peptidoglycan precursors
ends with D(-)-lactate (instead the more frequently met D-alanine, see section 1.3.1) and
cells lacking the capability to synthesize this lactate isomer are not viable. Indeed, this
61
uncommon peptidoglycan structure leads also to vancomycin resistance of L. plantarum
cells (Ferain et al., 1996). Reports published by Bienert et al. (2013), Desguin et al. (2014)
and Desguin et al. (2015) provide also in silico prediction for the presence of lar-related
genes in other LAB, too.
Malolactic enzyme is able to convert L-malate (natural isomer of malate) directly to lactic
acid and carbon dioxide without any other enzymatic activity. Indeed, the two other routes
for malolactic fermentation, decarboxylation of malate to pyruvate or oxidation first to
oxaloacetate and then subsequent decarboxylation of the formed oxaloacetate to pyruvate,
are both dependent on the L-lactate dehydrogenase activity when considering the
production of lactate (Schümann et al., 2013; Korkes and Ochoa; 1948; Flesch, 1969). As
reviewed by Matthews et al. (2004), malolactic fermentation in wine-borne LAB is most
typically catalysed by the malolactic enzyme. As mentioned in section 1.6.3.2, O. oeni
(formerly known as L. oenos) is one of the most typical LAB strain found in the wine
during spontaneous MLF (Lafon-Lafourcade, 1983; Solieri et al., 2010) and it also sold
for that purpose as a commercial product. In 1990’s Labarre et al. (1996) were the first to
identify and sequence the gene cluster carrying the genes responsible for expression of
malate carrier and malolactic enzyme in O. oeni. During the preceding decade, Caspritz
and Radler (1983) proposed the overall reaction equation for the L(+)-lactate synthesis by
malolactic enzyme. What they observed was that this reaction is dependent on the
presence of NAD+ and manganese. However, as no reduction of NAD+ or detection of
free reaction intermediates was detected, the exact mechanism for this reaction remained
unsolved (Casprizt and Radler, 1983; Groisillier and Lonvaud-Funel, 1999). According to
Arthurs and Lloyd (1999), malolactic enzyme is present in many species belonging to the
genera Lactobacillus, Leuconostoc, Oenococcus, and Pediococcus. However, this enzyme
has been detected also in some other LAB genera. As an example of this, it has been
proposed that malolactic enzyme has important role against lethal intracellular
acidification and it also protects cells against oxidative and starvation damage in oral
streptococci (Sheng et al., 2010).
When considering other putative biosynthetic routes for lactic acid, it is also possible that
some other enzymatic side activity or totally undiscovered enzymes are responsible of
lactate production. As an example, Aarnikunnas et al. (2002) could not find explanation
for the D(-)-lactate production in L. fermentum strain in which both stereospecific ldhgenes were inactivated by deletion. Authors speculated that the marginal formation of
D(-)-lactate in this strain was due to some other enzyme, with D-lactate dehydrogenase
side activity.
1.8.2 Use of genetic engineering to produce lactic acid with enhanced
optical purity
In this section, the main emphasis is on examples from LAB studies, which involve the
use of modern methods of genetic engineering to modify the very last step in lactic acid
biosynthesis, resulting in enhanced optical purity of the (main) lactic acid isomer
produced. Furthermore, the focus is especially on studies in which the enhanced optical
purity meets the criterium recommended for the raw materials for production of high
melting point PLA-grades: optical purity at least 95 %, preferable at least 99.5 % (EP
62
2748256 A2, Noordegraaf and De Jong, 2012). Results of these studies are presented in
Table 6.
Table 6. Genetic engineering of LAB to improve the optical purity of the produced lactic
63
The optical purity in Table 6 is expressed as enantiomeric excess (Okano et al., 2009) and
yields are expressed as the conversion ratio (g/g) of the produced optical enantiomer from
the initial amount of the sugar added in the fermentation medium.
1.8.2.1 Genetic engineering of LAB to improve the optical purity of D(-)-lactate
Ferain et al. (1994) were among the first scientists to study the consequences of the
perturbation of lactate dehydrogenase activities in L. plantarum, and how this effects on
the total production and ratio of L(+)- to D(-)-lactate in this strain. In this study, one of the
strains constructed was TF101, in which the ldhL gene was inactivated by chromosomal
deletion. This was achieved by constructing an unstable integration vector carrying an
erythromycin resistance gene and flanking DNA fragments of the desired deletion site,
followed by using this vector for two-step homologous recombination. Later, Goffin et al.
(2005) used this strain in another research to study the role of ldhL expression and the role
of L(+)-lactate for the function of the lactate racemase operon (lar operon) in L.
plantarum. In this later study, it was confirmed that TF101 strain produce only D(-)lactate and the lactate racemization activity detected in the host strain is positively
regulated by L(+)-lactate (as no lactate racemase activity could be detected in the TF101
strain). There was no difference in the total lactate production between the TF101 and
wild type strain.
Okano et al. (2009) constructed also an ldhL – deletion strain (referred as ǻldhL1) of L.
plantarum by using a plasmid with a temperature sensitive operon and an antibiotic
resistance gene for homologous recombination. For the homologous recombination events,
plasmid carried also two 1 kb regions flanking the ldhL1 of L. plantarum. After the
double-crossover homologous integration (and excision of the plasmid), screening of the
LdhL1-negative strain was carried out with PCR amplification. Finally, pCUSĮA -plasmid
expressing the Į-amylase (AmyA) of S. bovis 148 was introduced into L. plantarum host
and ǻldhL1 strain, resulting in the strains referred as WT/pCUSĮA (hereafter referred as
reference strain) and ǻldhL1//pCUSĮA, respectively. Both of these strains were able to
grow in raw corn starch as the sole carbon carbon source, which was not possible for the
original wild type host. Authors demonstrated that the ǻldhL1//pCUSĮA -strain produced
comparable amount of total lactic acid with the reference strain, but the optical purity of
the produced D(-)-lactate was improved significantly (4.0 % vs 99.6 %).
Chae et al. (2013) employed a different strategy to enhance the D(-)-lactate optical purity
in Leuconostoc citreum 95. They constructed a new E. coli-Leuconostoc shuttle vector,
which was engineered to overexpress D-lactate dehydrogenase of L. citreum 95. This was
accomplished by cloning the ldhD of L. citreum 95 into this vector, following
transformation of the resulted plasmid by electroporation into the wild type host. The
resulting strain demonstrated stable growth and lactic acid production under the
erythromycin selection. However, the control strain (wild type + shuttle vector without
ldhD gene) showed impaired growth in the fermentation test. As stated by the authors, the
observed significant difference in cell growth between these two strains suggests that the
overexpression of ldhD in L. citreum 95 putatively help cells to maintain the glucose
metabolism and NADH/NAD+ recycling in balance during the growth at high
64
concentration of glucose. The optical purity of D(-)-lactate produced by the ldhD
overexpressing strain was at least 99.8 % when expressed as enantiomeric excess (authors
reported purity higher than 99.9 % when calculated as ratio of D(-)-lactate to the total
lactate). In this study, no data for the optical purity of the wild type cells was provided. In
an other study reported by Jin et al. (2009), the optical purity for the D(-)-lactate was
lower (90.5 %) when the same wild type strain was used in fermentation tests. Thus, it
seems possible that overexpression of ldhD resulted in a minor improvement in the optical
purity in the study of Chae et al. (2013). However, these numbers are not directly
comparable, because of the differences in the fermentation conditions of these two studies.
1.8.2.2 Genetic engineering of LAB to improve the optical purity of L(+)-lactate
Bhowmik and Steele (1994) cloned and characterized the ldhD of L. helveticus CNRZ32.
This strain is industrially attractive due its homofermentative metabolism and high acid
tolerance. Also, it produces lactate with good yield, but the produced lactate is mixture of
D(-)- and L(+)-isomers. To improve the industrial attraction of this strain, authors
inactivated its ldhD gene by gene disruption. This was accomplished by homologous
recombination by using a SA3-based integration vector carrying an erythromycinresistance gene for selection and being unable to replicate at high temperatures. The
resulting recombination of this plasmid in to chromosome under high temperature led to
disruption of the native ldhD gene and generation of erythromycin resistant strain that
produced only L(+)-lactate. The total amount of L(+)-lactate produced by the mutant
strain was only marginally less than that of the wild type. However, the maximum lactate
concentration was reached somewhat earlier with the mutant strain.
Lapierre et al. (1999) focused on to develop a non-D(-)-lactate producing variant for the
well known and widely used probiotic L. acidophilus La1 strain. In typical milk
fermentations, this strain produces D(-)- and L(+)-lactates in ratio of 60:40 (%). As
reviewed by the authors, in some diseases the accumalation of D(-)-lactate in blood may
lead to a manifestation of D-acidosis and encephalopathy. In addition, food ingredients
containg D(-)-lactate are not recommended to infants and young children (until 3 years
age), because their liver may not be mature to metabolize D(-)-lactate completely.
Lapierre et al. (1999) truncated the ldhD gene in vitro by PCR (8 bp deletion), resulting in
the formation of a translational stop signal in the middle of the gene. The modified DNA
sequence was then used to construct integration vector in Lactococcus. The integration
vector was transferred to Lactobacillus johnsonii via conjugation on solid agar plates.
After homologous recombination events in L. johnsonii, growth on non-selective medium
gave raise of desired mutants in which the gene replacement and plasmid excision had
taken place. In the following fermentation test, it was demonstrated that one of these
mutans (designated as La1 ldhD-2) was able to produce L(+)-lactate with optical purity at
least 98.9 %. However, although the yield of L(+)-lactate from glucose for the mutant
strain was reasonable good (86.5 %), it was still lower when compared with that reported
for the wild type strain.
In the study reported by Aarnikunnas et al. (2002), the main focus was to enhance the
production of mannitol in combination with pure L(+)-lactic acid or pyruvate in L.
65
fermentum. One of the strains constructed to meet this goal was a LdhD negative strain
(designated as GRL1030), in which the ldhD gene was inactivated by introducing a 0.4 kb
deletion by gene replacement technique in the promoter and 5’ end region of the ldhD of
L. fermentum. Fermentation tests demonstrated that the lack of the D(-)-lactate
dehydrogenase activity in L. fermentum resulted in only minor changes in primary sugar
metabolism (and mannitol production), but the growth and sugar consumption of the
mutant was slightly slower when compared to those of the wild type cells. More
surprisingly, the L(+)-lactate dehydrogenase activity was also decreased when compared
to wild type host. However, the mutant was still able to produce high levels of pure L(+)lactate without a slowdown in mannitol production.
Viana et al. (2005) studied the effects of genetic inactivation of D-hydroxyisocaproate
dehydrogenase (HicDH) to the end-product formation in L. casei. Although L. casei is
most often referred to as an L(+)-lactate producer, about 5 % of the total lactate produced
by the wild type cells of L. casei strain BL23 was detected to be D(-)-lactate in this study.
Inactivation of the gene encoding HicDH of BL23 was accomplished by single cross-over
integration with plasmid a carrying a gene region coding for a C-terminal truncated form
of the native HicDH. The resulting mutant did not produce any D(-)-lactate, but otherwise
the fermentation end-product pattern was practically identical to that of the wild type
strain. Moreover, the amount of L(+)-lactate produced by the mutant cells was comparable
with the total lactate production measured for the wild type cells.
Finally, Goffin et al. (2005) utilized the TF102 strain of L. plantarum (Ferain et al., 1996)
to study its lactate racemase. Although the ldhD in this strain is disrupted and nonfunctional, Ferain et al. (1996) demonstrated that the production of D(-)-lactate was not
blocked during growth on glucose and almost equimolar amounts of D(-)- and L(+)-lactate
were measured from the fermentation supernatant. According to these authors, this kind of
distribution of the optical enantiomers of lactic acid suggested the presence of L(+)-lactate
inducible lactate racemase in this strain. Earlier, it had been proposed that racemization is
dependent on the L(+)-lactate induction (Stetter and Kandler, 1973), so Goffin et al.
(2005) decided to utilize this information for the identication of genes responsible for the
putative lactate racemization activity in L. plantarum. Authors managed to identify a locus
(lar) involved in lactate racemization and observed that it is composed of six genes that
are organized in an operon. Deletion of this operon in the D(-)-lactate negative TF102
strain resulted in complete loss of D(-)-lactate production but also in loss of growth in
media without supplemented D(-)-lactate. As demonstrated by Ferain et al. (1996), D(-)lactate is needed for the peptidoglycan synthesis (and vancomycin resistance) of L.
plantarum. Goffin et al. (2005) observed that when the concentration of supplemented
D(-)-lactate was higher than 0.5 mM, the growth characteristics between the wild type
cells (strain NCIMB8826) and the ldhD lar double mutant were essentially the same. For
the fermentation test with ldhD lar double mutant, MRS broth containing 20 mM added
D(-)-lactate was used. As expected, no net increase of D(-)-lactate was found in the
supernatant after the fermentation test with the mutant strain and the produced amount of
L(+)-lactate was comparable with the total lactate produced by the wild type cells.
66
2.
Aims of the study
Industrial production of thermostable PLA requires lactic acid of high enantiomerical
purity. Often, industrial fermentations are carried out with thermotolerant microorganisms, like that of L. helveticus, in order to minimize contamination risk during
fermentation. During these fermentations, the cell density is one of the most important
factors affecting to the total productivity. One of the strategies used to increase cell
density is immobilization. However, many of the current whole cell immobilization
methods are either too costly or too cumbersome, or those are not robust enough to be
used in industrial fermentations of commodity chemicals with high dilution rates.
In this work, the focus was to (roman numbers refers to related publications):
1) Develop a LAB strain that produces PLA grade L(+)- lactic acid efficiently also in the
stationary phase (I).
The main hypothesis for this aim: production of L(+)-lactic acid by L. helveticus
CNRZ32 is more efficient over the whole fermentation cycle if both native ldhpromoters drive the synthesis of this isomer.
2) Develop a novel whole cell immobilization method for LAB that would be potentially
viable to be used also in industrial scale lactic acid fermentations. This goal was
divided into two phases.
2.1)
Studying of various cell surface anchors for successful surface display in
LAB. First, potential of L. brevis S-layer as a platform for surface display
was studied (II). Moreover, the potential of various cell wall anchors
(type/length) for surface display in L. lactis was compared (III, IV).
The main hypothesis for this aim: it is possible to construct a cell envelope
anchored cell surface display system in LAB that provides measurable
surface exposure for the fused target molecule.
2.2)
Indicating the potential of whole cell immobilization of L. lactis to cellulose
by cell surface displayed binding domain (III, IV).
The main hypothesis for this aim: it is possible to construct a cell envelope
anchored cell surface display system that enables the fused cellulose binding
domain to bind with chemically unmodified cellulose and enhance the
immobilization of mutant lactococci cells to a solid cellulosic carrier.
67
3.
Materials and methods
3.1 Plasmids
The plasmids used in this work are introduced in Table 7. The detailed use of the plasmids
is described in publications I-IV.
Table 7. Plasmids used in this work.
Reference/
source
Plasmid
Relevant properties
pZeRO-2
Kmr, lacZĮ-ccdB-based E. coli cloning vector
pJDC9
Emr, Tcr, Cmr, Ts replicon, Streptococcus-E. coli
shuttle vector
Emr, E. coli cloning vector
pUC19
Apr, E. coli cloning vector
pSA3
pKTH2153
pKTH2154
pKTH2155
pKTH2156
pKTH2157
pNZ9530
pLET1
Upstream region of the L. helveticus ldhD gene in
pZeRO-2
ǻldhD construction in pSA3
slpA transcription terminator and downstream
region of the ldhD gene in pSA3
Promoter region of the ldhD gene and structural
gene of ldhL in pJDC9
ldhD::ldhL construction in pSA3
Emr, nisRK cloned in pIL252, expression of
nisRK driven by rep readthrough
Cmr, Kmr, lactococcal expression vector with T7
RNA polymerase promoter
Invitrogen Co.
Carlsbad, USA
Dao and
Ferretti, 1985
Chen and
Morrison, 1988
Yanisch-Perron
et al., 1985
This work
This work
This work
This work
This work
Kleerebezem et
al., 1997
Wells et al.,
1993
Used in
I
I
I
I
I
I
I
I
I
II
II
pKTH2151
Cmr, Kmr, pLET1 derivative carrying the L.
brevis slpA gene
This work
II
pKTH2152
Cmr, Kmr, pKTH 2151 derivative; slpA fused
with a poliovirus VP1 epitope
insert in insertion site I
Cmr, pNZ8008 derivative carrying the gusA gene
translationally fused to the
nisA promoter (PnisA )
This work
II
de Ruyter et al.,
1996
II
pNZ8032
68
Table 7 continued.
pKTH5006
Cmr, pNZ8032 derivative carrying the slpA gene
under PnisA; the VP1 epitope
encoding nucleotides in slpA insertion site I
This work
II
pKTH5007
Cmr, as pKTH5006, the VP1 insert in slpA
insertion site II
Cmr, as pKTH5006, the VP1 insert in slpA
insertion site III
Cmr, as pKTH5006, the VP1 insert in slpA
insertion site IV
CBD of xynA of Cellvibrio japonicus fused with
ssusp45 and S. aureus protein A anchor region
(spax) (resulting in: ssusp45-CBD-spax)
55 kb plasmid isolated from L. lactis MQ421.
Contains the prtP gene
E. coli-L. lactis shuttle vector used for cloning
and expression. P45 promoter and ss45’ preceding
the multiple cloning site (MCS). Transcription
termination loop downstream of the MCS
ssusp45-CBD-spax -construct cloned into pLEB124
as HindIII-BamHI fragment, resulting in
expression under P45 promoter
pLEB592 KpnI-BglII digested and blunt end
ligated to enable further XbaI clonings
nisP-anchor fragment cloned into pLEB594 as
XbaI-ApaI fragment
prtP153aa –sequence (anchor) cloned into
pLEB594 as XbaI-ApaI fragment. Constructed for
expression of CBD-PrtP 153 aa fusion protein
prtP344aa –sequence (anchor) cloned into
pLEB594 as XbaI-ApaI fragment. Constructed for
expression of CBD-PrtP 344 aa fusion protein
acmA242aa –sequence (anchor) cloned into
pLEB595 as BamHI-ApaI fragment. Constructed
for expression of CBD-AcmA 242 aa fusion
protein
srtA of Staphylococcus aureus with its own
ribosome binding site fused the with the Cterminus of prtP-anchor of pLEB597
Negative control for pLEB597 (lacks CBD)
Negative control for pLEB596 (lacks CBD)
Negative control for pLEB606 (lacks CBD)
This work
II
This work
II
This work
II
Cultor Ltd.,
Finland
III
Kiwaki et
al.,1989
Ra et al., 1996
III
This study
III
This study
III
This study
III, IV
This study
III
This study
III, IV
This study
III
This study
IV
This study
This study
This study
III
III
III
pKTH5008
pKTH5063
pN5
pLP763
pLEB124
pLEB592
pLEB594
pLEB595
pLEB596
pLEB597
pLEB606
pLEB607
pLEB685
pLEB686
pLEB687
III
3.2 Bacterial strains, growth conditions and media
The bacterial strains used in this study are shown in Table 8. The E. coli strains were
grown in Luria-Bertani (LB) medium (Miller, 1972) at +37°C on agar plates or in liquid
69
with shaking. The L. lactis strains were grown in M17 (Terzaghi and Sandine, 1975) at
+30°C medium supplemented with 0.5% (wt/vol) glucose (M17G) on agar plates or in
liquid without shaking. The L. helveticus strains were grown in MRS medium (de Man et
al., 1960) at 37°C (or at 42 °C) on agar plates or in liquid without shaking. The L. brevis
strains were also grown in MRS medium at 37°C on agar plates or in liquid without
shaking. S. aureus was grown in LB medium at 37°C on agar plates or in liquid with
shaking.
When appropriate, E. coli medium was supplemented with kanamycin (50 ȝg ml-1),
chloramphenicol (100 ȝg ml-1), tetracycline (10 ȝg ml-1), ampicillin (50 or 100 ȝg ml-1),
and erythromycin (200 or 300 ȝg ml-1), L. helveticus medium was supplemented with
erythromycin (4 ȝg ml-1), L. brevis medium was supplemented with chloramphenicol (7.5
ȝg ml-1) and erythromycin (7.5 ȝg ml-1), and L. lactis medium was supplemented with
chloramphenicol (2.5 or 10 ȝg ml-1) and erythromycin (5 or 7.5 ȝg ml-1).
Table 8. Bacterial strains used in this study.
Strains
Escherichia coli
Top10F’
Escherichia coli DH5Į
Escherichia coli
DH5ĮF’
Relevant properties
Host strain of pZERO-2
Transformation host
Transformation host
Reference/source
Invitrogen, Carlsbad,
USA
Hanahan, 1983
Hanahan, 1983
Escherichia coli
ERF573
Escherichia coli
ERF574
Escherichia coli
ERF575
Escherichia coli
ERF576
Escherichia coli
ERF577
Lactobacillus
helveticus CNRZ32
E. coli Top10F’ with pKTH2153
This work
I
E. coli DH5ĮF’ with pKTH2154
This work
I
E. coli DH5ĮF’ with pKTH2155
This work
I
E. coli DH5ĮF’ with pKTH2156
This work
I
E. coli DH5ĮF’ with pKTH2157
This work
I
Wild-type strain,
L-LDH+, D-LDH+
I
Lactobacillus
helveticus GRL86
Lactobacillus
helveticus GRL89
Lactobacillus brevis
GRL1
Lactobacillus brevis
GRL1001
(strain identification
code unpublished)
Escherichia coli TG1
ǻldhD mutant of CNRZ32
Centre National de
Recherches
Zootechniques, Jouyen-Josas, France.
This work
ldhD::ldhL mutant of CNRZ32
This work
I
L. brevis ATCC 8287
II
L. brevis GRL1 with pNZ9530
American Type
Culture Collection
This work
II
Transformation host
Gibson, 1984
III
70
Used in
I
I
I
I
Table 8 continued.
Lactococcus lactis
MG1363
Plasmid-free and prophage-cured derivative of
NCDO712. Transformation host. Source of
chromosomal DNA for acmA anchor
amplification. PrtP-
Gasson, 1983
Lactococcus lactis N8
Nisin producer. Source of chromosomal DNA
for nisP anchor amplification
Carries pLP763 (55 kb plasmid). PrtP+
Graeffe et al., 1991;
Immonen et al., 1995
Kiwaki et al.,1989
MG1363 derivative carrying a deletion in acmA
Buist et al., 1995
L. lactis IL1403 derivate containing disrupted
htrALl gene; htrALl::pVE8039, CmR
Poquet et al., 2000
L. lactis MG1363 strain carrying pLEB595.
Used for whole-cell ELISA. EmR
This work
IV
L. lactis MG1363 strain carrying pLEB606.
Used for whole-cell ELISA. EmR
L. lactis MG1363 strain carrying pLEB607.
Used for immobilization test. EmR
L. lactis MG1363 strain carrying pLEB596.
Used for whole-cell ELISA. EmR
L. lactis MG1363 strain carrying pLEB597.
Used for whole-cell ELISA. EmR
L. lactis MG1363acmAǻ1 strain carrying
pLEB597. Used for immobilization test. EmR
L. lactis IL1403 htrA strain carrying pLEB597.
Used for immobilization test. EmR
L. lactis MG1363acmAǻ1 strain carrying
pLEB685. Used for immobilization test. EmR
L. lactis MG1363acmAǻ1 strain carrying
pLEB596. Used for immobilization test. EmR
L. lactis MG1363acmAǻ1 strain carrying
pLEB606. Used for immobilization test. EmR
L. lactis MG1363acmAǻ1 strain carrying
pLEB686. Used for immobilization test. EmR
L. lactis MG1363acmAǻ1 strain carrying
pLEB687. Used for immobilization test. EmR
Source of chromosomal DNA for srtA
amplification.
This work
III
This work
IV
This work
III
This work
III,IV
This work
III
This work
IV
This work
III
This work
III
This work
III
This work
III
This work
III
Steidler et al., 1998
IV
Lactococcus lactis
MQ421
Lactococcus
MG1363acmAǻ1
Lactococcus lactis
IL1403 htrA
Lactococcus lactis
LAC236 (strain
identification code
unpublished)
Lactococcus lactis
LAC242
Lactococcus lactis
LAC243
Lactococcus lactis
LAC247
Lactococcus lactis
LAC248
Lactococcus lactis
LAC252
Lactococcus lactis
LAC257
Lactococcus lactis
LAC351
Lactococcus lactis
LAC352
Lactococcus lactis
LAC353
Lactococcus lactis
LAC355
Lactococcus lactis
LAC357
Staphylococcus aureus
Cowan I (NCTC 8530)
III
III, IV
III
III
III, IV
3.3 Oligonucleotides
Oligonucleotide primers used for the PCR amplifications done in this work are listed in
Table 9. The detailed use of the primers is described in publications I-IV.
71
Table 9. Oligonucleotides used in this study.
Oligonucleotides
O1
O2
Nucleotide sequence (5’ => 3’)
CTTGAAGAGTATCTGCAGTGTAGTC
ATTACGAATTCCAGTTAATGCCGCATTC
O3
CTTAGGATCCGCTGGAATGATTATCGTG
O4
O5
O6
O7
GATAGGATCCTTACGCTATTCGAAAAGACG
TTGTCTGGGAATTCTTTACCTTC
CCCGCGGATCCTAATAATTATTATTTAGGTG
A
GCATATCGATGTTTTTCCTAACAAAGGC
O8
O9
CGTATCTAGAATCTGGCTTTTCGCCG
TGCACGCAACTTAGTCTCTTG
O10
p3
GATAATTTTAACCATTCCTCATTATACGCTT
C
TGAGGAATGGTTAAAATTATCACTAATAAA
AAG
GAACGGATCCTTATTGACGAACCTTAACGC
CGATCCTGCTTTAACTGCTGTTGAAACTGGT
GCTACTAT
CGATAGTAGCACCAGTTTCAACAGCAGTTA
AAGCAGGAT
CTGAGAATTCAGTTACAGCAACCAACG
p4
TTTAAAGCTTGTTTTTCCTAACAAAGGCC
p5
p6
CCATGGTACAATCAAGTTTAAAGAAATCTC
CCTGCTTTAACTGCTGTTGAAACTGGTGCTA
CTGCCGCCGATCAAACTGCTC
AGTAGCACCAGTTTCAACAGCAGTTAAAGC
AGGCTTATCGCTTACCTTAGAACCTG
CCTGCTTTAACTGCTGTTGAAACTGGTGCTA
CTAATGATAAGGTTGCAGCTAACG
AGTAGCACCAGTTTCAACAGCAGTTAAAGC
AGGAGCATCAGCTGTAGTCAATGC
TACCGAATTCGGGACAGGTGCTAGAGAC
AGTAGCACCAGTTTCAACAGCAGTTAAAGC
AGGAGCCATAGTAGCCTTAGAAG
CCTGCTTTAACTGCTGTTGAAACTGGTGCTA
CTAAGTTAGCTTCTTCAAAGAGTC
GTATGAATTCGAAATGACTTCAGAAAAGG
GACAGGATCCATATAGAAGAAAAGGGC
CACAAAGCTTTGAAGAAGCAGCACTGTCC
O11
O12
p1
p2
p7
p8
p9
p10
p11
p12
p13
p14
p15
72
Use/site of hybridization
ldhD mRNA 5’-end mapping
Amplification of ldhD upstream
region
Amplification of ldhD upstream
region
Amplification of ldhD region
Amplification of ldhD region
Amplification of transcription
termination region of slpA
Amplification of transcription
termination region of slpA
Amplification of ldhD region
Amplification of ldhD upstream
region
R-PCR for ldhD-ldhL mRNA
joint
R-PCR for ldhD-ldhL mRNA
joint
Amplification of ldhL region
VP1 epitope sequence
Used
in
I
I
I
I
I
I
I
I
I
I
I
I
II
VP1 epitope sequence
II
Amplification of transcription
termination region of slpA
Amplification of transcription
termination region of slpA
Amplification of slpA region
R-PCR, site II/VP1 in slpA
II
II
II
R-PCR, site II/VP1 in slpA
II
R-PCR, site III/VP1 in slpA
II
R-PCR, site III/VP1 in slpA
II
Amplification of slpA region
R-PCR, site IV/VP1 in slpA
II
II
R-PCR, site IV/VP1 in slpA
II
Amplification of slpA region
Amplification of slpA region
Amplification of slpA region
II
II
II
II
Table 9 continued.
NIS195
NIS196
TGCTTCTAGAGGATCCACTTGGTATGCCGA
CATTGC
ATGGGGGCCCTATTCTTCACGTTGTTTCCG
Used for constructing prtP153aa
III
Used for constructing prtP153aa
and prtP344aa
Used for constructing prtP344aa
III
III
Used for constructing acmA242aa
Used for constructing acmA242aa
III
III
III
NIS197
TGGCTCTAGAGGATCCAAGTCGACTGATTT
ATACGG
NIS216
NIS217
NIS265
CTACCGGATCCAGTTATGGCGCTTCACTG
GTAATGGGCCCTTATTTTATTCGTAGATACT
GAC
GCACGAGCTCTGATAAATATGA
NIS266
GCTGTGGATCCAGCGTAAACACCTGACAAC
NIS191
TGTTCTAGAGGATCCGGGAAAAATAAAGCTTTT
AGC
Used for constructing
immobilization control plasmid
Used for constructing
immobilization control plasmid
Amplification of nisP anchor
region
ATATGGGCCCTCAATTTTTAGTCTTCCTTTT
C
Amplification of nisP anchor
region
IV
GTGAGGGCCCTAAAAGGAGCCTTAACGTAT
G
Amplification of srtA region
IV
GCGTCCCGGGTTATTTGACTTCTGTAGCTAC
Amplification of srtA region
IV
(previously
unpublished)
NIS192
(previously
unpublished)
NIS218
(previously
unpublished)
NIS219
(previously
unpublished)
3.4 Methods used in this study
During this work well-established DNA isolation, hybridization, detection, manipulation,
characterization, amplification and transformation protocols were applied (see Table 10).
Also, RNA isolations, hybridizations and detections were based on earlier documented
methods (see Table 10). Methods used for protein quantification and detection, or for
enzymatic activity assays, are also included in this same table.
The whole cell enzyme-linked immunosorbent assay (ELISA) and cellulose binding assay
were in the key role in this work. The whole cell ELISA-assay applied in this work was
used for detection of the surface exposure of anchor-fused molecules and was partially
carried out as described earlier by Laitinen et al. (2002). More detailed description for the
whole cell ELISA-assay used in this work is available in publications II and III.
The cellulose binding assay was developed for this work (applied in publications III and
IV) and this method is described in detail in publication III but is shortly covered here,
also. In this assay, chemically unmodified Whatman No. 1 filter paper (Ø 42.5 mm) was
used as immobilization support. Before the immobilization test with the filter paper, the
Petri dishes (Ø 55 mm, made out of polystyrene) in which the filter paper assay
73
III
IV
(immobilization test) was later done were first ‘saturated’ with pre-prepared cell
suspension (in PBST: PBS + Tween 20 (0.5 %, v/v), pH7.4), in order to minimize the
unspesific binding of bacterial cells to polystyrene during the actual filter paper
immobilization test. After this saturation phase, the optical density of the cell suspension
was assayed (at 600 nm wavelength, later referred as OD600) by taking a small sample
from the Petri dish and leaving the remaining cell suspension in the dish. After this,
immobilization test was initiated by adding the filter paper into the cell suspension.
Adhesion of cells to filter paper was promoted by shaking the Petri dish for 1 h at 28°C
(100 rev min-1). After the adhesion, cell suspension was completely collected from the
Petri dish by pipette for OD600 measurement, and washing solution (PBST) was added
into Petri dish. After the washing period of 10 min (100 rev min-1 shaking at 28°C), the
optical density of the washing solution was measured and that result was used to calculate
the robustness of the cellulose binding (‘washing loss’).
Table 10. Methods used in this study.
Methods
Principle of the method
described by
Used in
General methods
DNA isolation, manipulation and amplification
methods.
Sambrook et al., 1989
I, II, III, IV
Concentrations and conditions in enzyme
catalyzed reactions were adjusted to meet the
recommendations set by the enzyme manufacturers
Specific recombinant DNA technology methods
Gene replacement method
Recombinant PCR technique
Bhowmik et al., 1993
Higuchi, 1990
I
I, II
Specific DNA isolation and purification methods
L. helveticus chromosomal DNA isolation.
E. coli plasmid DNA isolation
L. lactis and L. brevis plasmid isolation
L. lactis plasmid isolation
L. lactis chromosomal DNA isolation
DNA gel extraction with kit
DNA purification with kit
Vidgren et al., 1992
Wizard Miniprep (Promega),
FlexiPrep (Pharmacia)
QIAfilter Plasmid Midi Kit
(Qiagen)
O’Sullivan and
Klaenhammer, 1993
Marmur, 1961
E.Z.N.A.® Gel Extraction
Kit (Omega Bio-tek)
QIAquick PCR Purification
Kit (Qiagen), E.Z.N.A.®
Cycle-Pure Kit (Omega Biotek)
74
I
I
II
III, IV
III, IV
III,IV
III,IV
Table 10 continued.
Methods related to DNA hybridizations
Digoxigenin (DIG) labelling of DNA probes
DIG-DNA Labeling Kit
(Boehringer Mannheim), or
DIG-High Prime Kit
(Boehringer Mannheim)
Southern, 1975
DIG Luminescent Detection
Kit (Boehringer Mannheim)
Grunstein and Hogness,
1975
Southern transfer
Southern hybridization and DIG-detection
Colony hybridization
I
I,II
I
I
DNA transformation methods
E. coli transformation
L. helveticus / L. brevis transformation
Sambrook et al., 1989
Bhowmik and Steele, 1993
L. lactis transformation
Holo and Nes, 1989
I
I, II
II, III,IV
RNA methods
Total RNA isolation
Vesanto et al.,1994, or
RNeasy Mini Kit (Qiagen) /
RNeasy Midi Kit (Qiagen)
Vesanto et al., 1995
Kahala et al., 1997
Sambrook et al., 1989,
Hames and Higgins, 1985
mRNA start site analysis
RNA dot blot
RNA hybridization
I
I
I
I
Methods used for protein quantification or detection
Protein quantification
Immunofluorescence microscopy
Whole cell ELISA
Sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE)
Western blotting
Western detection
Bradford, 1976
This work
This work and Laitinen et
al., 2002
Laemmli, 1970
I
II
II,III,IV
Sambrook et al., 1989
ProtoBlot® kit (Promega)
III,IV
III,IV
III,IV
LDH activity and lactic acid assays
Screening of D(-)-lactic acid negative strains
D(-)-LDH and L(+)-LDH activity measurements
L(+)- and D(-)-lactic acid assay
HPLC assay for total lactic acid quantification
Cellulose binding assay
von Krusch and Lompe,
1982
This work
D-/L-Lactic acid TestCombination (Boehringer
Mannheim)
This work
Whole cell immobilization test
This work
75
I
I
I
I
III,IV
4.
Results and discussion
The first goal in this work was to construct a thermostable LAB strain capable of efficient
PLA grade L(+)-lactic acid production. L. helveticus was chosen for lactic acid producer
due to its homofermentative metabolism and thermo- and acid-tolerant nature. Also, it has
been regarded as a efficient lactate producer (Roy et al., 1986; Idler et al., 2015) and it
possess the QPS status (Qualified Presumption of Safety) granted by EFSA (European
Food Safety Authority) (Anonymous, 2013). High optical purity of lactic acid is required
to produce thermostable PLA grades (over 95 %, preferable over 99.5 %), as enantiomeric
impurities decrease the melting point of the produced PLA. However, the selected host
strain, L. helveticus CNRZ32, is not as such the preferred organism for production of PLA
grade lactate, as it produces mixture of D(-)- and L(+)-lactic acid during lactic acid
fermentation. Therefore, the first focus in this work was to genetically modify L.
helveticus CNRZ32 to produce only L(+) –lactic acid. In the resulting strains, the L-LDH
expression took place under either native promoter (in GRL86 strain) or in combination of
native promoter and ldhD promoter (in GRL89 strain). In the former case, ldhD was
inactivated, whereas in the latter case the structural gene of ldhD was replaced by that of
ldhL. Both of these strains were achieved by applying homologous recombination method.
Furthermore, the relationship between growth of biomass and lactic acid production in the
D-LDH deficient mutant strains was also studied. In many conditions, growth and lactic
acid production are coupled in lactic acid bacteria. From the fermentation point of view
this is a potential drawback; when the cells are entering to the stationary growth phase,
production of lactic acid gradually ceases. This kind of characteristic, although often
unavoidable, is not usually desirable for high cell density industrial fermentations,
especially if the cells are immobilized and fermentation is operated at a continuous mode.
Another key focus area in this work was to study the potential of various anchoring motifs
for surface display and eventually for whole cell immobilization. First, the gene slpA,
encoding the S-layer protein subunit of L. brevis, was studied to detect the best locations
within this gene to allow expression of the poliovirus VP1 epitope on the cell surface of
this bacterium. The learnings from the VP1 surface display experiments were successfully
applied to develop an expression system for c-Myc epitopes from human c-myc protooncogene. The primary purpose of the original publication II was to develop a novel
whole cell platform for displaying vaccine antigens. For this thesis, the main interest is not
focused to vaccines itself, but to evaluate possibilities of S-layer as a platform for surface
display and ultimately for whole cell immobilization. Finally, the whole cell
immobilization of L. lactis was studied with cellulosic material. In this work, different
anchoring motifs and cell wall spanning sequences were fused with cellulose binding
domain (CBD) of Cellvibrio japonicus. The surface exposure of the expressed fusion
proteins was tested with whole cell ELISA by using CBD specific antibody and the
functionality of these fusion proteins was evaluated by immobilization test with filter
paper.
76
4.1 Production of PLA grade L(+)-lactic acid by genetically
modified homofermentative Lactobacillus strain
(publication I)
4.1.1
Construction of D-LDH negative L. helveticus strain by inactivation of ldhD by
gene replacement
A L. helveticus CNRZ32 derived strain with ldhD promoter deletion was constructed by
using the gene replacement method described by Bhowmik et al. (1993). The integration
vector (pKTH2154) used in this gene replacement was first constructed in E. coli and then
transferred to L. helveticus by electroporation. This vector was pSA3-based StreptococcusE. coli shuttle vector with thermosensitive replicon, carrying fused homologous DNA
fragments from both side of the ldhD promoter region of L. helveticus CNRZ32, but
missing the actual promoter region (0.6 kb deletion). The integration of this plasmid into
chromosome took place by homologous recombination after the temperature shift from
37 °C to 45°C under erythromycin selection. The second homologous recombination and
excision of integration plasmid was achieved by growing the cells for 100 generations at
37°C without any antibiotic selection. As a result, 5 % of the resulting clones were
erythromycin-sensitive. Out of these, one clone (designated as GRL86) out of the fifteen
showed D(-)-lactic acid negative phenotype.
4.1.2
Construction of D-LDH negative L. helveticus strain by replacing the structural
gene of ldhD with that of ldhL
Another L. helveticus CNRZ32 derived strain, in which the structural gene of ldhD was
replaced by that of ldhL, was constructed also by gene replacement method. First, the
transcription initiation site of the ldhD gene was determined by primer extension. This
site, base G in position -34 upstream of the first nucleotide of the start codon, was used as
a joint between the homologous ldhD region and homologous ldhL region when
constructing the DNA fragment for homologous recombination (see Fig. 12). The
beginning of the homologous ldhL fragment was set to start from the transcription
initiation site of this gene as determined by Savijoki and Palva (1997). In practise, this
DNA fragment (ldhDupstream region -PldhD - ldhLstructural gene) was built with recombinant PCR
–technique (R-PCR, Higuchi, 1990). The amplified ldhL region did not include the
transcription termination loop of ldhL, instead the transcription terminator region from
slpA of L. brevis was fused to the 3’end of coding region of ldhL. In the final integration
vector (pKTH2157), this terminator was preceeding the other homologous ldhD region
needed for the gene replacement (see Fig. 12). This ´ldhD fragment started from the ClaI
restriction site located in the 5’-end of the corresponding structural gene.
77
Figure 12. DNA construct used in cloning to express the structural gene of ldhL under the
promoter of ldhD in L. helveticus.
The integration of pKTH2157 into L. helveticus CNRZ32 chromosome and the subsequent
gene replacement events were achieved essentially as described above. However, an extra
verification by PCR after the first recombination was needed in order to be sure that
recombination had taken place only in the region of the ldhD gene. After growing the cells
for approximately 100 generations at 37°C without any selection, about 5% of clones were
erythromycin sensitive. Out of these, one clone (designated as GRL89) out of 25 tested
showed D(-)-lactic acid negative phenotype.
4.1.3
Growth and fermentation characteristics of the wild type strain and D-LDH
negative strains
First, growth, lactic acid production and transcriptional and enzymatic levels of LDHs of
L. helveticus CNRZ32 wild type strain were studied during pH controlled (pH 5.9) small
scale (1.5 liter) bioreactor fermentation at 42°C (see Fig. 13). It was noticed that the
production of L(+)-lactic acid took place during the exponential growth phase and rapidly
ceased when cells entered to the stationary phase. However, it seemed that L-LDH activity
remained at a relatively high level even through out the stationary phase, too. The bulk of
the D(-)-lactic acid was produced between the late exponential phase and the early
stationary phase. In contrast to L-LDH activity, D-LDH activity decreased rapidly after
having its maximum between the late exponential phase and the early stationary phase.
Similar pattern was also observed for the relative intensities of ldh mRNAs – mRNA for
ldhL reached its maximum to somewhat earlier than mRNA for ldhD. Interestingly, at the
time when production of L(+)-lactic acid suddenly ceased, the amount of L-LDH activity
was very near its maximum and there were still plenty of detectable ldhL mRNA present.
Based on these results it seems that the preference for D(-)-lactate production during the
growth phase in which cells are entering to the stationary phase is not a consequence of
limited presence of L-LDH enzyme but putatively due to changes in intracellular
conditions in such a way that either the affinity of L-LDH for pyruvate is diminished or
the catalytic activity of L-LDH is inhibited, resulting in a flow of pyruvate through DLDH.
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Time (h), batch fermentation
Figure 13. Expression of the ldh-genes and production of lactic acid in L. helveticus
CNRZ32 as a function of growth (pH controlled batch incubation). Symbols:z, cell
density; ǻ, L-LDH activity; ¸, D-LDH activity; Ŷ, L(+)-lactic acid; Ƒ, D(-)-lactic acid;Ÿ,
relative ldhL-mRNA intensities with a ldhL gene probe; Ƈ, relative ldhD-mRNA
intensities with a ldhD gene probe.
When the wild type host and D-LDH negative strains were compared with the above
described small scale fermentation set-up, minor but insignificant differences were
observed in the growth rate. Also, the total amount of lactic acid produced by all these
strains was essentially the same. The optical purity (or enantiomeric excess) for L(+) –
lactic acid was 16.0 % for the wild type strain but 99.8 % for the D-LDH negative mutants
as measured by enzymatic assay (unpublished results). So, practically both D-LDH
deletion mutants created were pure L(+)-producers and well qualified for production of
L(+)-lactic acid for thermostable PLA manufacturing process. As expected, no D-LDH
activity could be detected with the mutant strains GRL86 and GRL89. The highest L-LDH
activities measured for GRL86 and GRL89 were 53% and 93%, respectively, higher when
compared with that of CNRZ32 strain. All these strain reached their maximum L-LDH
activity during the late exponential/early stationary phase (at 9 h time point). The
difference between the L-LDH activities of mutant strains indicates that ldhD promoter
was functional in GRL89. Notably, the production period of L(+)-lactic acid in the mutant
strains was prolonged when compared to that observed in the wild type strain. In mutant
strains, L(+)-lactic acid production ceased approximately at the same time point (15 h, in
the stationary phase) in which the D(-)-lactic acid production stopped with the wild type
strain. Indeed, there was no decrease in the rate of L(+)-lactate synthesis at a lactic acid
concentration at which L(+)-lactate excretion ceased completely in the wild-type strain.
This observation suggests that the rate of L-LDH catalysis is not likely dependent on the
lactic acid concentration. Thus the change of flow from pyruvate to D(-)-lactate observed
in the wild type strain may be due to changes in substrate binding between D- and L79
LDHs under these conditions. However, no other data to support this hypothesis was
generated during this work.
The activity of two different ldh-promoters were also studied by a fermentation
experiment ran at higher temperature (44°C) but at lower pH (5.4). As observed by
Armane and Prigent (1999), at low pH values growth and lactic acid production in L.
helveticus are uncoupled. Interestingly, this kind of phenomenon was putatively also
observed with GRL89, but not with GRL86, at pH 5.4. With GRL86 both growth
(biomass accumulation) and lactic acid production ceased simultaneously, whereas with
GRL89, lactic acid production continued despite the biomass accumulation was levelled
(Fig. 14). However, as the lactose seemed to be depleted at the same time when the
biomass reached its maximum, it remained unanswered what was the reaction mechanism
supporting the late lactic acid synthesis in GRL89.
Figure 14. Batch fermentations of genetically engineered L. helveticus strains GRL86
(open symbols) and GRL89 (solid symbols), data from two parallel cultivations at pH 5.4
and 44°C. Concentrations of lactose (circles) and lactic acid (squares), and changes in dry
weight as a function of time are presented in panels A and B, respectively.
Under these conditions, yield from lactose based media (initial lactose content about 80
g/l) with GRL89 strain was 91.6% with the maximum productivity rate of 3.21 g liter-1h-1,
whereas the same numbers for GRL86 were 76.2% and 3.34 g liter-1h-1, respectively.
Thus, in GRL89 in which two different promoters drove the lactic acid production, the
total lactic acid production was approximately 20% higher when compared to GRL86
strain. When compared to criteria set by Datta et al. (1995) for a potential industrial scale
80
lactic acid fermentation, both the yield (91.6 vs. >90 %) and productivity (3.21 g liter-1h-1
vs. >2 g liter-1h-1) targets were met with the GRL89 strain. As the amount of sugar added
into fermentation medium was less than the criteria set by Datta et al. (1995) for the
desired end product concentration (>90 g/l), the true performance of this strain in this
sense remained unanswered.
It seems that the ldhD promoter is capable to drive the L-LDH-synthesis longer (vs.
growth phase) than the native ldhL promoter. Other explaining reason for the difference
observed in lactic acid production, however quite unlikely, is the inactivation of putative
fructosamine-3-kinase gene product in the GRL86 strain (unpublished information). When
the ldhD in this strain was inactivated by promoter deletions, this deletion also inhibited
the potential transcription of this gene. However, the putative enzymatic activity lost due
to the gene replacement in this strain is not directly affecting the lactose catabolism in L.
helveticus, as this homofermentative species possess lactose permease and ȕ-galactosidase
activity and it metabolizes galactose via the Leloir pathway (Mollet and Pilloud, 1991;
Fortina et al., 2003).
In addition, the optimum conditions for lactic acid production for GRL89 strain were
studied by using statistical experimental design and response surface methodology. As a
result of this study, pH 5.9 and 41°C were predicted to be optimum conditions for lactic
acid production in this strain.
4.2 Development of whole cell immobilization method for
LAB by surface engineering
4.2.1
Studying the potential of S-layer protein (SlpA) of L. brevis for anchoring peptides
for cell surface display (publication II)
Four insertion sites for the poliovirus VP1 epitope within the slpA gene of L. brevis ATCC
8287 (designated as L. brevis GRL1 below), were selected by using the hydrophilicity
profile of the SlpA protein. In these regions (most hydrophilic parts of the SlpA protein),
an 11-amino-acid immunodominant region of the VP1 capsid protein of enteroviruses
(Hovi and Roivainen, 1993) was expressed to verify the surface accessibility of the chosen
sites. The construction of the DNA sequence coding the VP1 epitope was carried out by
hybridization, and the resulting fragment was used to construct the modified slpA-genes
by conventional DNA manipulation techniques or by R-PCR technique. In the resulting
constructs, the VP1 epitope in the expressed fusion proteins was displayed between
between amino acid residues Asp362 and Thr363 (insertion site I), Lys249 and Ala250
(insertion site II), Ala313 and Asn314 (insertion site III) and Ala49 and Lys50 (insertion
site IV). Each of these insertions was constructed separately and the resulting slpA
constructs were expressed under the regulated nisA promoter (PnisA) in a controlled manner
in a two-plasmid system. In this system, modified slpA genes were expressed under nisA
promoter (found in pNZ8032), and the nisR and nisK genes needed for the nisin-induced
transcription from PnisA (Kleerebezem et al., 1997) were expressed by pNZ9530 plasmid.
The plasmids expressing modified slpA genes with VP1 epitope coding sequence in
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location I, II, III and IV were designated as pKTH5006, pKTH5007, pKTH5008 and
pKTH5063, respectively. These plasmids were used to transform L. brevis ATCC 8287
strain carrying pNZ9530 plasmid (hereafter referred as GRL1001) under erythromycin and
chloramphenicol selection, resulting to strains referred as GRL1001+epi-I, GRL1001+epiII, GRL1001+epi-III and GRL1001+epi-IV, respectively. As a results, four L. brevis
strains were obtained, which expressed not only the native slpA but also VP1-modified
slpA. The GRL1001 strain was used as a control strain (possess only native slpA) when
testing the surface exposure of these above mentioned VP1-modified slpA variant strains.
The surface accessibility of the VP1 epitope in the above mentioned strains was tested by
whole-cell enzyme-linked immunosorbent assays (ELISA) with anti-VP1 antibody raised
in rabbit. According to whole-cell ELISA, the strongest color response within the strains
carrying VP1-modified S-layer was obtained with the recombinant strain in which the
insertion was in site II (GRL1001+epi-II). Also, the strain expressing VP1 in site I
(GRL1001+epi-I) gave a positive color response, whereas the response for the strains
expressing VP1 in sites III (GRL1001+epi-III) and IV (GRL1001+epi-IV) was at the same
level with that measured for the control strain (GRL1001). In addition to the whole-cell
ELISA assay, surface exposure of the VP1 in these recombinant strains was analysed by
immunofluorescence microscopy using anti-VP1 antibody and FITC-conjugated
secondary antibody. However, none of the recombinant strains tested gave visible signal in
these tests. Cells for both of these tests (whole-cell ELISA and immunofluoresce
microscopy) were induced with 10 ng/ml of nisin. With this nisin concentration, both the
induction of PnisA and cell growth after the induction was at reasonable level for
subsequent immunological detection of VP1.
In the above described mutant slpA strains there is competition between the expression of
native slpA and VP1-modified slpA. Because the expression levels of these slpA-epitope
constructs were still low when compared to the high-level expression of the native
chromosomal slpA gene, epitopes were not able to be seen by immunofluorescence
microscopy. In order to increase the expression level of the epitope-SlpA protein, different
expression strategy was employed with another epitope (c-Myc). This epitope was
incorporated at site II in slpA and the resulting recombinant slpA was used for gene
replacement in L. brevis (see publication II for details) to replace the native chromosomal
slpA gene with the modified one. As a result, a recombinant strain (GRL1046, see
publication II) was achieved that displayed uniform S-layer on its surface with the desired
antigen in all of the S-layer protein subunits and which was easily detected by the
immunofluorescence microscopy, also. Further studies indicated that the S-layer lattice
structure was not affected by the presence of the additional c-Myc epitope in the S-layer
subunits.
As a summary, two insertion sites out of total four tested gave a positive signal in wholecell ELISA, which demonstrated that these insertion sites in native slpA can be used for
various cell surface display applications in which S-layer functions as a cell wall anchor.
The VP1 epitope used in this study was short (11 amino acid residues), so the potential of
displaying longer motifs in these insertion sites remained unverified in this study. As the
S-layer is usually the most outermost layer in cells in which it exists, the peptides
anchoring to it do not need to span through the peptidoglycan layer. As a comparison, the
estimated minimum length needed for the cell wall spanning and to allow proper surface
exposure of the target protein is usually at least 90–100 residues (Fischetti et al., 1990;
Strauss and Götz, 1996).
82
The advantage of the S-layer for surface dispay is the high number of protein subunits
present on the cell surface. As estimated by Sleytr and Messner (1988), an average-sized
cell consists of approximately 5 x 105 S-layer monomers on its surface. As it was
demonstrated by c-Myc epitope (10 aa), it is possible to incorporate small peptides into
every single S-layer subunit proteins without interfering the assembly and structure of the
S-layer. Regardless of the incorporated peptide and the binding mechanism involved
(mediated by this peptide), it is possible that already a weak interaction could provide a
whole cell binding with carrier. So far scientists have not managed to construct a
genetically engineered LAB strain which could be robustly immobilized to an industrially
viable carrier material by its genetically modified S-layer. However, in several cases Slayer mediated whole cell binding with various receptors, cells and biological matrixes has
been detetcted (Hynönen and Palva, 2013; Sleytr et al., 2014). This seems to be
consequence of direct interactions between various S-layer components and target
structures.
4.2.2
Indicating the potential of whole cell immobilization of L. lactis by cell surface
displayed binding domain (publication III and IV)
The final goal of this work was to develop a L. lactis strain capable to be immobilized to a
cellulosic carrier by CBD-cellulose bio-affinity. The typical properties required from an
industrial immobilization carrier have been reviewed earlier in this work (see section
1.6.3.1). Out of these, among the most important properties are good availability, chemical
and physical robustness, nontoxicity, high immobilization capacity and low price.
Cellulosic materials, like wood chips, have been used in many traditional fermentations
processes – especially in those in which mixing is not needed or the shear forces applied
are very low. In order to increase the binding force between bacterial cell and cellulose,
chemical pretreatments have been used to modify cellulose. As the surface charge of most
microorganisms is negative, basic derivates of cellulose, such as modification with DEAEgroups (diethylaminethyl), make it a more efficicient carrier for the whole cell
immobilization (Phillips and Poon, 1988). However, these kinds of treatments require the
use of additional processing time and -chemicals and are costly. Thus, transition from
DEAE-based carriers to unmodified cellulosic carriers is advantegous in many cases.
In this work, different kind of anchoring motifs and cell wall spanning sequences were
tested for both cell surface exposure and whole cell immobilization efficiency when fused
with the cellulose-binding domain (CBD) of XylA of Cellvibrio japonicus, referred
hereafter as CBDXylA. The XylA of C. japonicus is an endo-1,4-beta-xylanase A and was
originally characterized by Hall et al. (1989) from Pseudomonas fluorescens subspecies
cellulosa (reclassified as C. japonicus by Humpry et al., 2003). This enzyme has an Nterminal carbohydrate binding module, classified under CBM_2 family in the Conserved
Domain Database (CDD) (Marchler-Bauer et al., 2015). This kind of cellulose binding
domain is typically found among bacteria.
83
4.2.2.1
Construction of different cell wall anchors for whole cell immobilization of L.
lactis
During this work, both covalent and non-covalent cell wall anchors were tested to display
CBDXylA for cellulose binding (see Table 11). Covalent cell wall anchors of LPXTG-type
(Fischetti et al., 1990) included those of PrtP of L. lactis MQ421 (Kiwaki et al., 1989) and
NisP of L. lactis N8 (Immonen et al., 1995). In addition to LPXTG-based anchors, the
non-covalent LysM anchor region (containing three repeats) from AcmA of L. lactis
MG1363 was tested to anchor CBDXylA. Also, the length of the sequence between the
CBDXylA and the conserved LPXTG recognition sequence and the effect of the
heterologues sortase A (SrtA, Mazmanian et al., 1999) overexpression for the covalent
anchoring were studied with the PrtP anchor of L. lactis MQ421. The sortase gene for this
construct was amplified from chromosomal DNA of Staphylococcus aureus Cowan I
(NCTC 8530).
Table 11. Anchors used to display CBDXylA for cellulose binding.
Plasmid
Cell envelope anchor fused
with CBDXylA
Type of
anchoring
Length
of the
anchor
344 aa
pLEB597
PrtP of L. lactis MQ421
LPXTG
(covalent)
pLEB607
PrtP of L. lactis MQ421
344 aa
pLEB196
PrtP of L. lactis MQ421
LPXTG
(covalent)
LPXTG
(covalent)
pLEB606
AcmA of L. lactis MG1363
242 aa
pLEB595
NisP of L. lactis N8
LysM
(noncovalent)
LPXTG
(covalent)
153 aa
Other relevant information
Anchor part covers AN-, W-, Hdomains and also the C-terminal
part of the B-domain of PrtP (as
described by Siezen (1999)
srtA of S. aureus transcriptionally
fused to prtP anchor-sequence
Anchor part covers AN- and Wdomains and also the C-terminal
part from the H-domain of PrtP (as
described by Siezen (1999)
Anchor part covers three repeated
regions (LysM-domains)
121 aa
The DNA region encoding the CBDXylA was provided by Cultor Ltd. in a form of plasmid
(cloned in pN5). In this plasmid, the 5’end of the region encoding the CBDXylA was fused
with the sequences coding the Usp45 secretion signal of L. lactis (van Asseldonk et al.,
1990), whereas the 3’end of the CBDXylA encoding region was fused with sequence coding
the cell wall anchor (SPA) of Protein A of S. aureus (Shuttleworth et al., 1987; Steidler et
al., 1998). This fusion construct was cut out from pN5 and transferred into pLEB124
under the constitutive ss45 promoter (P45, Sibakov et al., 1991) and partial secretion
signal. The pLEB124 vector was constructed by Ra et al. (1996), and this lactococcal
expression plasmid is also able to replicate in E. coli. To allow further clonings, one XbaI
restriction site (located in the upstream region of P45) in the resulting pLEB124 based
plasmid was removed and the resulting plasmid was designated as pLEB594. In this
plasmid, the remaining XbaI-site is located at the seam between DNA fragments coding
for CBDXylA and SPA anchor. This XbaI-site was directly used for constructing the most
84
CBD-anchor fusions during this work just by directly replacing the SPA anchor with other
C-terminal anchors (as XbaI-ApaI fragments). This approach was used to constructs
plasmids with fragments encoding NisP 121 aa, PrtP 153 aa and PrtP 344 aa anchors,
resulting in plasmids pLEB595, pLEB596 and pLEB597, respectively.
Cloning of AcmA 242 aa anchor utilized BamHI site built between the DNA fragments
encoding the CBDXylA and NisP anchor. In practise, the NisP 121 aa anchor was replaced
(as BamHI-ApaI fragment) with the region coding for the AcmA 242 aa anchor, resulting
in generation of plasmid pLEB606. Finally, the pLEB607 plasmid expressing CBDXylA PrtP 344 aa –SrtA was built in the pLEB597 based vector. The DNA fragment for
expression of SrtA in this construct is transcriptionally fused with the C-terminal end of
the fragment encoding the PrtP 344 aa, and possesses also the original riboseme binding
site of the srtA of S. aureus. In detail, the construction of this plasmid started with ApaI
linearization of pLEB597, followed by ligation with an ApaI digested srtA fragment
(generated by PCR). This ligation mixture was used as template for PCR amplification of
the region, containing the P45 promoter region and the DNA fragment coding for
CBDXylA – PrtP 344 aa –SrtA. The resulting PCR-product was digested with BamHI and
XmaI, generating fragments coding the fusion of PrtP 344 aa –SrtA for the final ligation
with similarily digested pLEB597 vector. As mentioned above, the resulting plasmid was
designated as pLEB607.
All the different pLEB124-based CBDXylA-anchor constructs were used to transform L.
lactis hosts by electroporation. L. lactis MG1363 host was used to verify the surface
exposure of CBDXylA when fused with different anchors. For cellulose binding tests
(publication III), the expression host for the CBDXylA-anchor fusion was MG1363acmAǻ1.
In this strain, the major autolysin of the host (MG1363) is inactivated. As noticed by Buist
et al. (1995), the loss of AcmA activity was seen as a change in growth characteristics. As
observed by these authors, mutants did not show growth as single cells as typically
observed with the wild type strain, but seemed to grow as long chains, indicating that
AcmA is required for cell separation in this strain. The main reason why this strain was
preferred to be the host in immobilization tests instead of the MG1363 strain, was the
lower unspecific binding with the filter paper under the conditions used for the
immobilization test. This observation, suggesting the promoting effect of active AcmA on
the adhesion of cells to solid surfaces, is in agreement with the results reported by Mercier
et al. (2002). However, in publication IV the MG1363 strain was used as the host strain
also in the cellulose binding tests, as the main target was not to quantify the exact
efficiency of the binding, but rather to verify is there any difference in cellulose binding
with or without overexpression of sortase (with the same anchor construct).
4.2.2.2
Immunological verification of the surface exposure of CBDXylA when anchored
to cell with different anchors
The surface exposure of CBDXylA in constructed L. lactis MG1363 strains expressing
CBDXylA -fusion proteins with either the NisP 121 aa-, PrtP 153 aa-, PrtP 344 aa- (with or
without SrtA) or AcmA 242 aa -anchor was tested by whole-cell ELISA with the CBDantibody raised in rabbit. As indicated by the result (Fig. 15), the highest positive
response was measured for fusion constructs with the longer PrtP anchor (344 aa), but also
85
the response measured for the AcmA 242 aa anchor construct clearly exceeded that
measured for the wild type strain (MG1363). However, the response with shorter PrtP
anchor fusion was weak.
Figure 15. Selected results from whole-cell ELISA-assays. Lactococcal cells expressing
CBD-anchor fusions were tested with CBD-specific antibody. The volume of the cell
suspension used in the detection is indicated.
With the NisP 121 aa-anchor fusion, the response from the assay was not clearly
distinguishable from that of the wild type strain (see publication IV). Also, it seemed that
there were no differences in the surface exposure of the CBDXylA between the strains in
which the CBDXylA - PrtP 344 aa was expressed with or without transcriptionally coupled
heterologous srtA (see publication IV).
Based on these results, all the above strains except the one expressing CBDXylA-NisP 121
aa fusion protein were used for the immobilization test with Whatman no.1 filter paper. In
these tests, the strain expressing CBDXylA-PrtP 344 aa – SrtA was not included in the
quantitative immobilization testing (that was performed for all the other strains), but
instead it was used in one to one comparative study against the strain expressing the same
CBDXylA-anchor construct but not the SrtA enzyme.
The most probable reason for the poor surface exposure of the CBDXylA-NisP 121 aafusion protein was the significant degradation of this protein, which was clearly seen by
the Western analyses. In contrast the CBDXylA –PrtP 153 aa fusion protein seemed to
suffer from weak anchoring, as the only clearly detectable band of right size for this fusion
protein in Western analysis was found from the supernatant fraction (results not shown).
Also, significant part of the longer PrtP fusion protein and major part of the AcmA (result
not shown) fusion protein were found in the supernatant fraction. These observations
indicate restricted binding capacity of these anchors under the conditions used for
preparing the cells for this testing.
4.2.2.3
Verification of whole cell immobilization of genetically modified L. lactis by
immobilization tests
Finally, to evaluate the immobilization capability of the selected recombinant strains, a
simple filter paper-based (with Whatman filter paper no.1) immobilization assay was
developed. In this assay, immobilization was quantified by measuring the optical density
of cell suspension at 600nm before and after the filter paper was incubated with the cells
86
(cells were suspended in PBST buffer). The actual immobilization phase with the filter
paper took 1h at 28 °C (with 100 rev min-1 shaking). Optical measurement was also
applied to estimate the robustness of the binding of the cells with filter paper by measuring
the optical density from the liquid phase after the immobilized filter paper was washed
with pure PBST buffer. In order to take into account the effect of unspecific binding
caused by the host strain and the anchor, pLEB124 based control strains were constructed.
These control strains carried plasmids otherwise similar to CBDXylA-anchor –fusion
constructs, but lacked the CBDXylA sequence completely. The unspecific binding measured
for these controls were then compared with the corresponding CBD fusion proteins to
evaluate wheter the binding measured for the CBDXylA-anchor –fusions were statistically
different from those measured for the control strains.
According to results obtained from the immobilization tests (see Fig. 16), the PrtP 153 aa
anchor was not able to secure whole cell binding with the fused CBDXylA -domain.
Although an anchor peptide with 90-100 residues in the extended loop (between the target
protein and LPXTG box) may already be long enough for surface display (Fischetti et al.,
1990; Strauss and Götz, 1996), it seemed that this 153 aa anchor (with over 100 residues
in the extended loop) was too short in this case to extend the CBD domain for proper
position and distance from the cell surface for the cellulose binding. Instead, the
immobilization efficiency with the longer PrtP anchor (PrtP 344 aa) fused with the
CBDXylA -domain was significantly improved when compared to the wild type strain.
Indeed, the nature of this binding was so strong that the washing loss measured for these
cells was very low (only about 1%). Also, the immobilization of the L. lactis cells
expressing the CBD-AcmA 242 aa fusion protein was also efficient, although not
comparable with that measured for the CBDXylA –PrtP 344 aa fusion protein expressing
strain. Interestingly, the nature of the binding between the strains expressing CBD-AcmA
242 aa and CBD-PrtP 344 seems to be somewhat different, as the proportion of the
washing loss to the final adhesion was quite different between these two strains. However,
this observation is in line with that detected in the above described Western result (see
section 4.2.2.2). It seems that the nature of binding with AcmA anchor is not as strong as
that observed with properly attached PrtP (LPXTG) –anchor. Also, it is possible that this
variation is partially dependent on the number of potential anchoring ligands presented in
the cell surface. Indeed, instead of uniform distribution on the cell surface, the occurrence
of the AcmA on the cell surface is putatively localized as suggested by Steen et al. (2003).
When the immobilization efficiency was compared with the host strain (MG1363), only
the strains expressing CBD-AcmA 242 aa and CBD-PrtP 344 aa fusion proteins showed
statistically significant difference.
87
Figure 16. Filter paper immobilization tests of L. lactis strains. Legends for the bars
indicate the fusion protein expressed in MG1363acmAǻ1 host. Blue color indicates the
final immobilization % of the strains on the filter paper and purple color indicates the
calculated washing loss value for each strain. Statistical significance between the strain
expressing the CBD-anchor fusion and the corresponding control strain (expressing the
anchor only) is marked with one asterisk (*), whereas two asterisks (**) indicates
statistical significance between the strain expressing CBD-anchor fusion and the host
strain (P < 0.05, unpaired double-tailed t-test).
During the development of the immobilization test we noticed that addition of 0.5 % (v/v)
of Tween 20 into immobilization solution prevented the nonspecific binding of the
lactococcal cells. Also, we noticed that the immobilization efficiency of the strain
displaying CBD-AcmA 242 aa fusion protein increased significantly when compared to
the control strains (results not shown). Steen et al. (2003) have proposed that LTAs
(lipoteichoic acids) may potentially hinder the binding of AcmA in lactococci. As it is
known that some nonionic surfactants are able to release cell-bound LTAs (Ohta et al.,
2000) and LTAs are potentially more concentrated in lactococci in those areas in which
autolysin-anchored proteins are not able to bind, it can be speculated whether the
improved binding with the CBD-AcmA 242 aa fusion protein with Tween 20 was a
consequence of reduced number of cell-bound LTAs.
As proposed by Habimana et al. (2007), the presence of PrtP on the bacterial surface may
increase the cell’s capacity to exchange attractive van der Waals interactions. Thus, as
suggested by the authors, bioadhesion of PrtP displaying lactococci is increased to various
types of surfaces. In this work, this kind of effect with the PrtP constructs was not
observed (see Fig. 16). Indeed, the adhesion with the PrtP-anchor- control strains were at
the same level with the host strain. Notably, in this thesis the used PrtP anchors covered
88
only C-terminal part of the whole PrtP protein, whereas Habimana et al. (2007) used in
their study full length PrtP alleles. Notably, Habimana et al. (2007) did not use cellulose in
their test, but instead used solid glass and tetrafluoroethylene surfaces with different salt
concentrations.
As extensive degradation was detected during the Western analysis of many CBD-fusion
proteins, htrA negative Lactococcus-strain was tested as expression host. As reported by
Poquet et al. (2000), HtrA is a unique extracellular housekeeping protease in L. lactis, and
it is involved in propetide processing, maturation of native proteins and degradation of
recombinant proteins. The HtrA negative lactococcus strain L. lactis IL1403 htrA (Poquet
et al., 2000) was transformed with pLEB597, and the resulting strain expressing CBDXylAPrtP 344 aa fusion protein, was designated as LAC257. The expressing of CBDXylA –PrtP
344 aa fusion protein in this strain was compared with LAC248 strain (HtrA positive
MG1363 host for pLEB597) with Western analysis. The immunological detection with
CBD antibody from rabbit indicated that in HtrA negative host strain the CBDXylA -PrtP
344 aa expressed was less degraded and more of the fusion protein could be found on the
cell wall fraction when compared with the HtrA positive host strain carrying the same
expression plasmid. Based on these results, it seems most likely that HtrA activity, at least
partially, is responsible for the degradation of the CBDXylA –PrtP 344 aa fusion protein in
MG1363 based host.
Finally, the effect of heterologous SrtA activity to the whole cell immobilization of L.
lactis with cellulosic material was studied. Simply, the immobilization test method
described earlier was used to compare the immobilization efficiency of strain expressing
CBDXylA –PrtP 344 aa fusion protein in MG1363 (LAC248) with that measured for
LAC243 strain, in which the DNA fragment coding this same fusion protein was
transcriptionally fused with DNA fragment including the RBS and structural gene of srtA
of S. aureus. The result from this one to one comparision indicated that the LAC243 did
not gain any substantial advantage from the additional heterologous SrtA activity and the
difference in the immobilization efficiency between these two strains was statistically nonsignificant. Because the production of this heterologues SrtA was hard to verify due to
lack of specific antibodies, it remains unanswered whether the potential increase of sortase
activity was too low to have an effect on the proportion of cell bound CBD-PrtP/secreted
CBD-PrtP, or was this heterologous SrtA incompatible with the machinery attaching
LPXTG type cell wall anchor proteins to the cell wall in L. lactis. Also, as the covalent
anchoring of cell wall surface exposed protein is composed of a series of steps including
protein exporting, retention, sortase cleavage and cell wall linkage by transpeptidation, the
effect resulted from boosting only one of step in this process may not lead to substantial
difference for the efficiency of the whole process. After the genomic sequencing of L.
lactis MG1363 (Wegmann et al., 2007), it was found that the amino acid sequence identity
between the sortases of S. aureus and L. lactis is low and even within the sortase domain
(PF04203) the identity is just around 30 % (unpublished results). Thus, it is quite probable
that the Staphylococcus-originated SrtA is not fully effective when anchoring LPXTG
proteins in L. lactis.
89
5.
Conclusions and future prospects
In this work, one of the main targets was to genetically engineer a pure L(+)-lactic acid
producer by inactivation of the gene responsible for the synthesis of D-lactate
dehydrogenase in L. helveticus CNRZ32. This goal was achieved and two different strains
capable to produce polylactic acid grade L(+)-lactic acid were constructed. Indeed,
polylactic acid (PLA) is one of the most abundant bioplastic grades in the market. For
PLA, it has been demonstrated that the enatiomeric purity of lactic acid monomers within
the polymeric PLA molecule is directly affecting to the thermostability of the produced
plastic.
When profiles of L(+)-lactic acid production and bacterial growth were studied in detail, it
was found that the production phase of L-(+)-lactic acid in the mutant strains was
prolonged (continued also in stationary phase) when compared to the wild type strain.
Furthermore, the first experiments studying the connection between growth and lactic acid
production putatively indicated that the L(+)-lactate production and growth are uncoupled
in GRL89 strain under the conditions studied. In this strain, the structural gene of ldhD
was replaced with that of ldhL, enabling synthesis of L-LDH enzyme by two different
promoters. This putative uncoupling is very interesting observation but needs to be
verified with new set of experiments. As a conclusion, the hypothesis set in section 2,
suggesting that the L(+)-lactic acid fermentation is more efficient if the both ldhpromoters drive the L(+)-lactic acid synthesis, can’t be overruled.
One of the most effective ways to run a continuous fermentation process, in which the
product of interest is extracellular, is to use immobilized production strain. Most typically,
chemicals and/or expensive modifications are needed to achieve an immobilization system
which is capable to provide efficient and robust production of desired metabolite. The
other main goal of this work was to develop a Lactococcus strain capable to efficient
immobilization on unmodified cellulolosic materials. In more details, we wanted to
achieve efficient immobilization on cellulose without the need of specific pre-treatment
procedures, such as the modification of cellulose with DEAE-groups (diethylaminethyl).
To reach this goal and to learn to how to achieve a proper surface display, different
genetically modified lactic acid bacteria strains with different types of cell wall anchors
were constructed and tested. The goal was to study and to be able to confirm (see
hypothesis in section 2) proper surface display with various types of cell envelope
anchors. This goal was met and the hypothesis confirmed. Clearly, the type of cell
envelope anchor, length of the anchor-fusion protein, but also the location of anchoring
point within the supporting molecule (in case of SlpA-study), affected the signal level
detected in whole-cell ELISA assays.
Although the whole cell immobilization (for industrial fermentations) was not the original
focus in the S-layer study, this kind of application for S-layers remains to be interesting
90
challenge for the future. As the number of cell surface displayed S-layer subunits is high,
already a weak interaction between immobilization carrier material and genetically
engineered S-layer could enable robust whole cell binding. Potentially, this kind of
binding could be possible with some S-layer fused binding domain or with fused synthetic
‘sticky’ peptides. Also, a linker sequence with relevant length is most probably recquired
to extend this affinity domain or peptide to the necessary distance from the S-layer lattice
surface to overcome the steric effects with the immobilization carrier material.
Finally, the whole cell immobilization of Lactococcus on cellulosic material was
successfully demonstrated with two different types of cell wall anchors (LPXTG and
LysM) fused with cellulose binding domain of C. japonicus. In these immobilization
studies, statistical significant difference to the control strain was measured for two
different anchor-fusion proteins, confirming the hypothesis set in section 2. In this work,
the purpose was to prove the feasibility of this concept in lactococci. However, more work
is needed to develop a robust industrial scale process based on the concept developed in
this study. Indeed, this kind of work has already progressed, as surface display anchor
constructs developed in this work have been used to develop L. lactis strains for continuos
nisin production with immobilized cells (ùimúek et al., 2013, ùimúek 2014). In these
constructs, chitin binding domain was utilized instead of cellulose binding domain and the
length of the LPXTG anchor proteins (PrtP of L. lactis) in the fusion construct was further
optimized. As reported by ùimúek (2014), the optimized system yielded the highest nisin
production ever reported and demonstrated its potential to be used in industrial scale.
As the whole cell immobilization systems developed in this study are plasmid based and
the P45 promoter has been tested to be functional in some other LAB, too, the next logical
step in the context of this work would be to combine the immobilization capability and the
capability to produce either PLA grade D(-)- or L(+) lactic acid within the same LAB
strain. Indeed, the current host species for the above mentioned constructs, L. lactis, is
already filling these conditions as it is known to be a L(+)-lactic acid producer. Also, it
would be interesting to see how other affinity domains or other functional domains apart
from cellulose binding domain and chitin binding domain would work when fused with
the anchors constructed in this work, and what kind of new applications these new
constructs would facilitate.
91
6.
Acknowledgements
This work was carried out at the MTT Agrifood Research Finland, Food Research,
Jokioinen, at the Division of Microbiology and Epidemiology, Department of Basic
Veterinary Sciences, Faculty of Veterinary Medicine, University of Helsinki, and at the
Division of Microbiology and Biotechnology, Department of Food and Environmental
Sciences, Faculty of Agriculture and Forestry, University of Helsinki.
I am deeply grateful to my supervisor Professor Per Saris for his patience and support
during all these years. I know I have passed many deadlines, but Per has still believed in
me and without his support I would never have finished this work. Many thanks also to
my other supervisor Professor Airi Palva for her guidance and support during my early
steps in molecular biology. I’m grateful for my supervisors for giving valuable comments
when writing and editing this manuscript. I wish also to thank the late Docent Ilkka Palva
for his valuable comments and suggestions to my studies.
I wish to express my warmest thanks to the reviewers Associate Professor Alexander Frey
and Adjunct Professor Ville Santala for critical revision and constructive comments.
In addition, I wish to express my warmest thanks to all my co-authors - it was very
pleasant to work with You and I very much appreciate You both as friends but also as
scientific collaborators. My warmest thanks also to all those people I have privileged to
work with during my studies in laboratories in Jokioinen, in Hämeentie and in Viikki.
These years are among the best years in my life and I have many unforgettable memories
from all these years.
I also want to thank for all the financial support I have obtained for this work. Special
thanks also to my former colleagues in Hanko and current colleagues in Kantvik for their
flexibility which enabled me to finish this work. Warm thanks also to those people in the
former Cultor Ltd. with whom I worked with during my TEKES project. Many thanks
also for those doctoral graduate schools in which I have attended during my studies.
Most of all, I want to thank my family. My wife Päivi and daughters Elsa and Vilma are
the joy and the meaning of my life. Thank you for Your patience and loving support. You
are so precious to me.
Karjaa, May 2016
Kari Kylä-Nikkilä
92
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