Chapter ONE
1. Introduction
Amino acids, the building block of proteins, are used as raw materials in various
cellular processes, such as energy generation, nitrogen metabolism, cell wall synthesis and
intracellular communication (Stryer et al., 2002). In addition to these, they are widely
used in both human and livestock consumption. Amino acids are generally produced by
one of four different methods: hydrolysis of natural proteins, chemical synthesis,
enzymatic synthesis and bacterial fermentation. However, the fermentative production of
amino acids has been established in industry due to its low production cost. In the case of
microbial fermentation, cheap carbon and nitrogen sources (molasses and ammonia,
respectively) are frequently used as raw materials (Ikeda, 2002). Furthermore, the
production of these building blocks by fermentation yields optically active and
biologically desired L-form of amino acids. The economic importance of amino acids is
enormous since they are used as flavouring agents, food additives, feed supplements and
raw materials for the synthesis of cosmetics, shampoos, toothpaste (Eggeling and Sahm,
1999; Leuchtenberger, 1996; Mueller and Huebner, 2002). Due to their high variety of
applications, the demand for these amino acids is constantly increasing. Hence, extensive
studies on the understanding and improving the metabolic conditions leading to amino
acids overproduction have been undertaken in order to increase the yield and productivity
(Kramer, 1996; 2004; Sahm et al., 1995).
At present, most of the essential L-amino acids are industrially produced by
Corynebacteria fermentation. Corynebacterium glutamicum, a short, aerobic, rod shaped,
Gram-positive soil bacterium, is capable of growing on a minimal medium.
Taxonomically, Corynebacteria are closely related to Mycobacteria, and they belong to
the mycolic acid containing actinomycetes (Kalinowski et al., 2003). The Japanese
scientist Kinoshita and his co-workers discovered C. glutamicum (originally named
Micrococcus glutamicus) as a potential microorganism for the production of amino acids
because of its ability to excrete L-glutamic acid into the surrounding medium under
specific growth condition (Kinoshita et al., 1957). Since then, C. glutamicum is regarded
as an efficient L-glutamate secreting microorganism. Under favourable growth conditions,
this bacterium converts 100g l-1 glucose to 50g l-1 L-glutamic acid. At present, 1,000,000
1
tons of L-glutamate and 450,000 tons of L-lysine are produced per year by Corynebacteria
fermentation. In addition, L-alanine, L-isoleucine and L-proline are also produced
industrially (Kalinowski et al., 2003; Kramer, 2004).
During amino acid fermentation by Corynebacteria, an appropriate substrate
(glucose, for example) is taken up by cells through the involvement of different uptake
systems (phosphotransferase systems, PTSs). The substrate is subsequently entered into
the central metabolic pathways (glycolysis, pentose phosphate pathways, tricarboxylic
acid cycle) of these bacteria, converted to metabolic intermediates within cells, and is
finally branched off to a particular amino acid biosynthetic pathway (Ikeda, 2002). In
recombinants of Corynebacteria, the biosynthetic pathways of a particular amino acid are
altered in such a way that results in increase of internal amino acid concentration. This is
mainly achieved by the following strategies i) by increasing the activity of anabolic
enzymes ii) by altering the regulatory enzymes or pathways (loss of feedback control) iii)
by blocking the pathways leading to by-products and iv) by blocking the pathways
responsible for product degradation in the cytosol. However, the increase of membrane
permeability of Corynebacteria is the most important feature for the efficient
secretion/production of amino acid, especially L-glutamate. The secretion/excretion of a
particular amino acid into the extracellular medium is generally accomplished either by
diffusion or by treating the Corynebacteria strains with an agent or with the aid of a
carrier system (Kramer, 1994).
It is well known that the cell wall of Corynebacteria has a complex structure since
it is formed by thick meso-diaminopimelic acids containing peptidoglycans that are
covalently linked to arabinogalactan (Brennan and Nikaido, 1995). Besides the thick
peptidoglycan layer, it also contains large amount of lipids in the form of mycolic acid
(Lichtinger et al., 2001). The multilayer arrangement of different phospholipids and
peptidoglycan contributes extremely low cell wall permeability. However, the wild strains
of Corynebacteria are not suitable for the production of amino acids under normal growth
conditions. Hence, several treatments that affect the cell membrane by limiting the
synthesis of phospholipids and membrane components have been employed in order to
induce the membrane permeability of this bacterium (Kramer, 1996; 2004). For Lglutamate efflux, C. glutamicum has been grown on biotin limitation (Shiio et al., 1962);
by the addition of surfactant (Duperray et al., 1992; Takinami et al., 1965; 1968); beta2
lactam antibiotic, penicillin (Demain and Birnbaum, 1968; Ikeda et al., 1972); ethambutol
(Radmacher et al., 2005) and oleic acid or glycerol (Kanzaki et al., 1967; Okazaki et al.,
1967). All of these perturbations directly attack to the cell wall of Corynebacteria, result
in changes in the composition of cell wall material and eventually increase the Lglutamate efflux (Eggeling and Sahm, 2001; Kramer, 1994). However, it has been
mentioned that the main reasons that result in secreting of amino acids in extracellular
environment are (i) a dramatic increase of internal amino acid concentration; (ii) a
fundamental change in the permeability properties of the cell membrane; or (iii) a defect
in the corresponding uptake system that normally counteracts the efflux of a particular
amino acid (Kramer, 1994).
Apart from these treatments, several other procedures, such as mutagenesis,
screening, specific changes in both genetic and enzymatic levels have been applied in
order to develop mutants/recombinants with desirable characteristics (Jetten and Sinskey,
1995b; Nampoothiri and Pandey, 1998; Parekh et al., 2000). In general, the improvement
of amino acid producing Corynebacteria strains is carried out by an iterative procedure of
mutagenesis and selection. Mutagenic procedures are optimised in terms of mutagen and
dose applied. Selection procedures are designed in order to identify the desirable mutants
with maximum expression (Nampoothiri and Pandey, 1998). Genetic engineering is also
applied to overexpress or repress the characteristics of a particular gene, and thereby new
strains with desired genotypes are constructed (Sahm et al., 1995). To obtain high yield
and productivity of an amino acid, however, it is necessary to investigate the detailed
information of metabolic pathways and their regulation under different environmental
conditions. Hence, metabolic engineering and metabolic flux analysis are recently applied
in order to quantify the biochemical fluxes leading to the intermediates or metabolites of
central metabolic pathways of this organism (Stephanopoulos et al., 1998). In comparison
to the modern techniques applied for the characterization and manipulation of metabolic
pathways (Sahm et al., 1995; 2000), however, relatively few studies have been conducted
in order to elucidate the efflux or secretion of these amino acids into the surrounding
medium.
Electroporation, a well-established physical process dealing with living cells
(Chang et al., 1992; Neumann et al., 1989; Teissie et al., 2002; 2005; Tsong, 1991;
Weaver and Chizmadzhev, 1996), is involved with a rapid structural rearrangement of cell
3
membrane in response to an externally applied electric field resulting in pore formation in
the lipid bilayer within a short period of time (Chernomordik et al., 1987; Haest et al.,
1997; Teissie and Tsong, 1981). The opening of transient aqueous pores provides a way
to transfer ions and water-soluble molecules across the cell membrane (Prausnitz et al.,
1995; Sixou and Teissie, 1993; Tekle et al., 1994). In addition, free diffusion is observed
even with dextrans and oligonucleotides, molecular weights up to 4kD (Teissie et al.,
1999). Electroporation has been studied over the past two decades due to its numerous
applications in cellular biology and biotechnology, especially for the purpose of gene
transfer and loading of cells with extracellular molecules (Golzio et al., 2004; Faurie et
al., 2005; Mir et al., 1999; Neumann et al., 1982; Rols, 2006; Somiari et al., 2000). In
addition, the application of this technique in gene therapy, cancer therapy and transdermal
drug delivery has given a new approach to treating complicated diseases (Heller et al.,
1999; Mir and Orlowski, 1999; Mir et al., 1991; Orlowski and Mir, 1993). Apart from
those applications mentioned above, electroporation has been found to be effective in
non-thermal food pasteurization (Angersbach et al., 2000; Wouters and Smelt, 1997),
selective release of intracellular proteins from recombinant Escherichia coli (Ohshima et
al., 1999; 2000; Ohshima and Sato, 2004) and Saccharomyces cerevisiae (Ohshima et al.,
1995) and Kluyveromyces lactis (Ganeva and Galutzov, 1999, Ganeva et al., 2001).
There are two types of electroporation that are extensively applied in biosciences
i.e., reversible electroporation and irreversible electroporation. Reversible electroporation
refers to the process of treating living cells by a moderate strength of electric field in
which transient pores are formed on the cell membrane, and thus the membrane is
reversibly permeabilized (Faurie et al., 2005; Hapala, 1997; Teissie et al., 1999).
However, electropores that are usually observed on the cell membrane within one minute
of pulsation can either be resealed within short period of time or remain open for a longer
period depending on the voltage and number of pulses applied to the cell suspension
(Chang et al., 1992; Faurie et al., 2005; Weaver, 1995). It has been demonstrated that
pulsed cells usually recover their original permeability within 30min of incubation at
room temperature (RT) (Kinosita and Tsong, 1977; Teissie et al., 1999). This
phenomenon has been extensively used in molecular biology and biotechnology,
especially for the transformation of bacteria using foreign genes (Golzio et al., 2004;
Jaroszeski et al., 1999; Neumann et al., 1982). The genetic transformation of
Corynebacteria has been successfully conducted by high voltage electroporation where an
4
occurence of reversible membrane permeabilization is observed (Bonamy et al., 1990;
Dunican and Shivnan, 1989; Liebl et al., 1989; Wolf et al., 1989). On the other hand,
irreversible electroporation, the application of high intensity electric field causing
permanent breakdown of cell membrane, is used to deactivate microorganisms
(Angersbach et al., 2000; Schoenbach et al., 2000). Barbosa-Canovas et al. (1999)
demonstrated that the application of pulsed electric field is one of the most relevant nonthermal processes for food preservation without altering their organoleptic and nutritional
properties. In addition, Ohshima and Sato (2004) carried out an effective bacterial
sterilization using high intensity electric pulses in which an induced irreversible
disruption of biological membranes occurred, eventually leading to cell death.
However, the mechanisms involved in electroporation still remain unclear.
Although it has been suggested that hydrophilic pores are formed in the lipid matrix
(Haest et al., 1997; Neumann et al., 1989; Teissie and Tsong, 1981), their existence has
never been clearly shown. The molecular processes involved during electroporation are
not fully understood due to the complex nature of the cell membrane, although many
theoretical studies have been conducted on the formation of pores under the influence of
an electric field (Chernomordik et al., 1987). Nevertheless, it is obvious that electric field
intensity, number and duration of pulses are the crucial factors for successful
electroporation. Hence, the stimulation factors have to be tightly controlled and
optimized, especially when working with an uncharacterized strain. Without adjusting
those parameters, cells may not return into their normal physiological state, and
eventually lose their viability. Although killing of cells by electropulsation is vital in the
case of irreversible permeabilization, it is mandatory to maintain cell viability as high as
possible while a reversible electropermeabilization is accomplished either for genetic
transformation or bioprocessing.
The production of heterologous proteins in bacteria and yeast using recombinant
DNA technology is already well-established in the biotech industry. However, the
isolation of recombinant proteins from hosts is not straightforward as the foreign proteins
are not usually secreted into the surrounding medium. More specifically, E. coli produces
foreign protein as an inclusion body that needs to be disrupted in order to obtain the
protein of interest. In most cases, cell disruption is performed by ultrasonication or
homogenization for the recovery of recombinant proteins. Using these techniques,
5
however, complete destruction of cells makes the purification process of desired protein
complicated, and ultimately the process turns into an expensive bioprocess. Moreover, the
recombinant proteins are generally contaminated with the other host proteins during their
isolation. As these proteins are used for medical purposes or human consumption, they
should be free from other host proteins, particularly pyrogen (Hermann, 2003). To resolve
this problem, researchers have been trying to develop an alternative strategy for the
bioprocess industries.
However, two most important facts concerned with these industrially important
bacteria have been demonstrated in the literature. Firstly, C. glutamicum is one of the
most important microorganisms in amino acids production, and L-glutamate production is
caused by cultivating this bacterium under certain growth conditions, such as biotin
limitation, surfactant addition and penicillin addition (Kramer, 1996; 2004). Secondly, the
construction of Corynebacteria recombinants, for the enhancement of yield and
productivity of amino acids, is successfully carried out by electrotransformation (Bonamy
et al., 1990; Dunican and Shivnan, 1989; Liebl et al., 1989), where a reversible
permeabilization of cell membrane is observed. Furthermore, electroporation that creates
pores on the cell membrane of microorganisms enhances the selective release of proteins
and enzymes from cells (Ohshima et al., 1999; 2000; Ohshima and Sato, 2004; Ganeva
and Galutzov, 1999, Ganeva et al., 2001). Although different treatments and extensive
investigations towards the genetic engineering of Corynebacteria have been conducted in
order to increase the yield and productivity of L-glutamate, no research has been carried
out on the electropermeabilization of these bacteria for the enhancement of L-glutamic
acid production so far. Based on the above literature, it is hypothesised that
electroporation may be a potential approach by which an appropriate strength of electric
pulse will be applied to the cell suspension or fermentation broth of Corynebacteria in
which the production of L-glutamate is secured by the above-mentioned treatments, and
hence the secretion ability of L-glutamate could be enhanced through the membrane
permeabilization.
This study is focused on the production of L-glutamate by the different strains of
Corynebacteria (Brevibacterium lactofermentum, Micrococcus glutamicus and B. flavum)
using different treatments, such as biotin limitation, surfactant addition and ethambutol
addition. While developing a suitable method for the production of this amino acid in M.
6
glutamicus, the growth studies (OD600nm), glucose consumption and L-glutamate
production in presence of a range of biotin concentrations (0-200µg l-1), under the
addition of different concentrations of surfactants [Tween 20 (4g l-1), Tween 80 (4g l-1)
and Tween 40 (1-4g l-1)] at two different growth points of fermentation (both start and
exponential), and in presence of a range of ethambutol concentrations (0-500mg l-1) will
be investigated. Furthermore, a very simple and easily accessible method based on
centrifugation will be developed in which L-glutamate is separated with a purity of more
than 90% from the fermentation broth. In this research, a study on the utilization of pulsed
electric field (transient electroporation) during L-glutamate fermentation by M.
glutamicus is considered in order to improve the yield of this amino acid, and thereby
intensify the bioprocessing.
Beside L-glutamate, the effect of electric pulses on the release of malate
dehydrogenase (MDH, cytoplasmic enzyme), glutamate dehydrogenase (GDH, ammonia
assimilating enzyme) and total protein will be investigated. In addition, the effect of
electroporation factors, such as cellular (growth phase and cell wall rigidity), electrical
(field strength, capacitance, number of pulses and pulse gap/resting time in the case of
multiple pulsing) and physiochemical (medium conductivity, ionic concentration of
electroporation buffer and temperature) on the membrane permeabilization as well as the
viability of M. glutamicus will be examined. An attempt will be made in order to assess
the permeabilization of electric field treated Corynebacterial cells by Bleomycin
(antitumor agent). Furthermore, the effect of hyperosmotic conditions (addition of 0.51.5M NaCl into Seed Medium) on the growth of M. glutamicus, L-glutamate production
and the activities of MDH, GDH and total protein will be investigated. It will also be
examined whether addition of compatible solutes (glycine betaine and proline) has any
notable influence on M. glutamicus growth.
The outline of this thesis is as follows-
Chapter 2 contains literature reviews regarding Corynebacteria (taxonomy and
cell wall composition, central metabolism, anaplerortic pathways, uptake and ammonia
assimilation, and metabolic engineering); industrial production of amino acids by
microbial fermentation and the mechanism of L-glutamate efflux under different growth
conditions; and the theory of electroporation or electropermeabilization, importance of
7
this
approach
in
biotechnology
and
factors
associated
with
the
successful
permeabilization.
Chapter 3 describes the production of L-glutamate in three different strains of
Corynebacteria (B. lactofermentum, M. glutamicus and B. flavum) under several growth
conditions i.e., biotin limited (1µg l-1), surfactant (Tween 40, 2g l-1) addition and
ethambutol (100mg l-1) addition. A range of biotin or Tween 40 or ethambutol
concentrations is added in order to determine the optimum amount of agent required for
the highest production of L-glutamate. A simple method based on centrifugation is
developed for the purification of L-glutamate (90%) from the fermentation broth.
Chapter 4 represents the application of electropermeabilization for the
enhancement of L-glutamate secretion produced under biotin limited fermentation of M.
glutamicus. The effectiveness of electric pulse for the extraction of cytoplasmic enzymes
(MDH and GDH) and total protein of both M. glutamicus and E. coli is also investigated.
Chapter 5 demonstrates the effect of different factors associated with the
electroporation on cell viability (both M. glutamicus and E. coli) and membrane
permeabilization. Whether the Bleomycin based method is applicable in accessing the
membrane permeabilization of M. glutamicus after electroporation is also investigated.
Chapter 6 depicts the osmotic stress associated during amino acid production and
demonstrates the effect of hyperosmotic stress on the growth and viability of M.
glutamicus, L-glutamate production and cytoplasmic enzymes or protein level.
Chapter 7 concludes the major factors associated with the success of
electropermeabilization, and the degree to which the above-mentioned objectives have
been met. This chapter also reveals the prerequisites that are required to consider for
introducing this approach in intensified bioprocessing, and suggests the directions for
future work.
Chapter 8 References
8
Chapter TWO
2. Literature Review
2.1. Corynebacteria
2.1.1. Taxonomy and cell wall of Corynebacteria
The genus Corynebacterium, pathogenic or at least parasitic to animals
(particularly diphtheroid bacilli), is comprised of a diverse collection of microorganisms
(Liebl, 1992). The use of chemotaxonomic markers (mainly cell wall chemistry, lipid
composition and DNA base composition) and phylogenetic approaches (mainly 16S
rDNA sequence analysis) revealed that the actual hierarchic classification of genus
Corynebacterium is class Actinobacteria - subclass Actinobacteridae - order
Actinomycetales
-
suborder
Corynebacterineae
-
family
Corynebacteriaceae
(Stackebrandt et al., 1997). However, the lipid profile analysis revealed that the genera
Corynebacterium, Mycobacterium, Nocardia and Rhodococcus are closely related,
therefore these genera are considered as CMN subgroup (Barksdale, 1970). According to
Collins and Cummins (1986), the genus Corynebacterium is Gram-positive, non-sporing,
non-motile, straight or slightly curved rods, ovals or clubs, often exhibiting typical Vshaped arrangement (Figure 2.1) due to their “snapping” mode of cell division. They are
facultatively anaerobic to aerobic, catalase-positive and chemoorganotrophic. The genome
of C. glutamicum, a single circular chromosome comprising 3282708 to 3309401 base
pairs, is smaller than that of taxonomically related bacterium, M. tuberculosis (4.2 Mb),
but larger than that of its close relative, C. diphtheriae (2.5 Mb). The G+C content of the
genome of C. glutamicum is 53.8%, which is close to that of E. coli (50%) (Ikeda and
Nakagawa, 2003; Kalinowski et al., 2003).
The classification of aerobic actinomycetes based on their cell wall composition
demonstrated that the cell wall of Corynebacterium is formed by thick mesodiaminopimelic acids (meso-A2pm), a polysaccharide fraction that is rich in arabinose
and galactose. Meso-diaminopimelic acids also contain peptidoglycans that are covalently
linked to arabinogalactan, a complex branched polysaccharide (Lechevalier and
Lechevalier, 1970). The arabinogalactan that is composed mainly of D-arabinofuranosyl
and D-galactosyl residues may contain significant amounts of mannose and glucose
9
(Puech et al., 2001). Additionally, high and low molecular mass glucan, arabinomannan,
lipoglycans and a protein surface layer are present in the cell wall of Corynebacterium
(Puech et al., 2000). The cytoplasmic membrane of Corynebacterium is arranged with
long hydroxylated fatty acid chains, known as mycolic acids (Brennan and Nikaido,
1995). It has been demonstrated that mycolic acids and protein layers not only result in a
barrier against larger compounds like proteins but also reduce the membrane permeability
for small water-soluble substances i.e., amino acids or sugars (Puech et al., 2001).
Furthermore, the cytoplasmic membrane of Corynebacterium is surrounded by a rigid
murein sacculus (contains up to 100 layers of peptidoglycan) that is highly resistant
against freeze damage and pressure or osmotic stress (Komatsu, 1979). It is now obvious
that all of these complex carbohydrates available in the cell wall of Corynebacteria
decrease the membrane permeability of this bacterium.
Figure 2.1 The electron micrograph image of C. glutamicum cell (Dr. Bustard, private
communication, 2004).
Marienfeld et al. (1997) demonstrated that the cell wall of C. glutamicum is
approximately 32nm thick; and is made up with an outer layer (8.5nm), an electron
translucent region (6.5nm) and peptidoglycans (17nm). The electron microscopic
examination of ultrathin section of C. glutamicum also revealed that the cell wall of this
bacterium is comprised of (i) a plasma membrane (PM) of 6-7nm composed of two
leaflets, (ii) a thick electron-dense layer (EDL) of 15-20nm, (iii) an electron-transparent
layer (ETL) of 7-8nm and (iv) a thin outer layer (OL) of 2-3nm (Puech et al., 2001). The
plasma membrane, the innermost part of cell wall, is a typical bilayer of proteins and
phospholipids. The PM is tightly associated with the EDL due to the presence of excess
lipopolysaccharides in its outer leaflet (Marienfeld et al., 1997). The EDL, containing
10
peptidoglycans, is surrounded by a thin ETL that generally consists of mycolic acid
residues i.e., eumycoloyl, nocardomycoloyl or corynomycoloyl. These mycolic acids
together with other non-covalently linked lipids i.e., trehalose and mono- and
dicorynomycolates form a rigid bilayer. Different noncovalently linked lipids and proteins
are mainly present in OL (Puech et al., 2001).
The PM of Corynebacterium is assembled with polar lipids, mainly phospholipids
and other proteins. The main phospholipid in C. glutamicum is phosphatidylglycerol (80%
of
the
total
lipids)
although
a
trace
amount
of
diphosphatidylglycerol,
phosphatidylinositol and phosphatidylinositol dimannosides are observed (Puech et al.,
2001). The major fatty acids available in this microorganism are palmitic (C16:0) and
octadecenoic (C18:1) (Collins et al., 1982), however, 10-methyloctadecanoic acid is found
in minor quantities (Puech et al., 2001). The glycan moiety of peptidoglycans is made up
of alternating -1, 4 linked N-acetylglucosamine and N-acetylmuramic acid residues.
Peptides attached to muramic acid residues of different glycan chains usually form
interpeptide linkages, and thus result in a rigid insoluble network surrounding the plasma
membrane (Schleifer and Kandler, 1972). Mycolic acids [R1-CH(OH)-CH(R2)-COOH],
high molecular weight, α-alkyl, β-hydroxy fatty acids (Figure 2.2), are present in the cell
walls of Corynebacteriaceae, Mycobacteriaceae, Rhodococci, Nocardiae (Collins et al.,
1982). However, Corynebacterium exhibits the shortest chain mycolic acid (22-38 carbon
Figure 2.2 Structures of a representative mycolic acid from M. tuberculosis and corynomycolic
acid from C. matruchotti (Lee et al., 1997).
11
atoms) which is only one-third of that from Mycobacterium (Chun et al., 1996). These
mycolic acids are esterified to the terminal penta-arabinofuranosyl units of
arabinogalactan, and represent a second permeability barrier besides the cytoplasmic
membrane (Brennan and Nikaido, 1995). In Corynebacteria and mycobacteria, the cell
wall linked corynomycolates and mycolates certainly make the cell membrane
impermeable to other substances since the disruption of the respective genes
(mycoloyltransferases) result in decrease of corynomycolates and mycolates, and
eventually increase the membrane permeability (Jackson et al., 1999; Puech et al., 2000).
The German group analyzed more than 100 individual polypeptides in the cell
wall of C. glutamicum by two dimensional electrophoretic methods (Hermann et al.,
2001). Among them, two major extracytoplasmic proteins, namely PS2 that forms the Slayer (cell surface crystalline array of proteins) of C. glutamicum (Chami et al., 1997),
and PS1 that transfers corynomycoloyl residues into the cell wall arabinogalactan and
trehalose monocorynomycolates (Puech et al., 2000) were observed. These proteins
represent an additional barrier for the cell wall permeability of C. glutamicum since the
inactivation of PS1 gene resulted in 50% decrease of cell wall corynomycolates,
decreased the trehalose dicorynomycolates and finally altered the membrane permeability
(Puech et al., 2000). A pore-forming protein (porin) has recently been identified in the
cell wall of C. glutamicum that mediates the transport of small hydrophilic solutes across
the hydrophobic mycolic acid barrier (Niederweis et al., 1995). PorA, encoded by gene
porA, is a channel forming polypeptide that contains 45 amino acid acidic polypeptides
with an excess of four negatively charged amino acids (Costa-Riu et al., 2003). The
immunological detection of porin revealed that the channels are water-filled pores, wide
and localized in the mycolic acids layer of C. glutamicum, but not in the cytoplasmic
membrane (Lichtinger et al., 1998). The porA mutants of C. glutamicum showed
reduced growth at every stage of growth cycle as compared to the wild-type grown on
minimal medium although there is no difference observed in glutamate production
between these two strains. The result confirmed that deletion of porA does not change the
membrane permeability of C. glutamicum although this perturbation changes the cell wall
structure of this bacterium (Costa-Riu et al., 2003). However, it is now apparent that the
cell wall of C. glutamicum is rigid due to the presence of high percentage of
peptidoglycans, polysaccharides and mycolic acids that make the cell wall impermeable to
both the extracellular molecules and intracellular protein or enzymes.
12
2.1.2. Central metabolism of Corynebacteria (sugar uptake, glycolysis and TCA
cycle)
Metabolism of an organism refers to the biochemical assimilation (anabolic
pathways) and dissimilation (catabolic pathways) of nutrients by a cell. Anabolic
pathways include the reductive processes that lead to the production of new cellular
material, whereas catabolic pathways are the oxidative processes that generate energy
from the substrates or intermediates. In general, cell metabolism is involved with many
reactions in which substrates (carbon, nitrogen and phosphate) are taken up by the cell,
and finally converted into new cell or products by various catabolic and anabolic
pathways. Metabolic intermediates i.e., adenosine tris phosphate (ATP), nicotinamide
adenine dinucleotide phosphate (NADP) and reduced NADPH, produced via the
catabolism of nutrient medium, are the essential elements for microorganism’s growth as
these intermediates play a vital role in biosynthetic reactions, nutrient transport and
product excretion (Stryer et al., 2002).
Although C. glutamicum metabolises a variety of carbon and energy sources i.e.,
carbohydrates, organic acids and alcohols (Liebl, 1992), most of the research with this
organism used in amino acids production have been conducted with carbohydrates.
Several studies also showed that this bacterium co-metabolizes glucose and fructose
(Dominguez et al., 1997), glucose and lactate, glucose and pyruvate, and glucose and
acetate (Cocaign-Bousquet et al., 1993; Wendisch et al., 2000). C. glutamicum uses the
phosphotransferase systems (PTSs) for the uptake of glucose, fructose, mannose and
sucrose (Dominguez and Lindley, 1996); and all of these PTSs are responsible for
metabolising sugars across the bacterial membrane and its concomitant phosphorylation
(Barabote and Saier, 2005). Figure 2.3 shows the transport process of sugar in C.
glutamicum. There are three essential catalytic entities i.e., enzyme I, enzyme II, and HPr
(heat-stable, histidine-phosphorylatable protein) found in a PTS system in C. glutamicum.
The enzyme I (EI, encoded by ptsI) becomes autophosphorylated by phosphoenolpyruvate
(PEP) and transfers its phosphoryl group to the HPr proteins, encoded by ptsH. HPr then
phosphorylates a number of sugar specific permeases and forms enzyme II-sugar
complexes that transport their substrates by concomitant phosphorylation (Saier and
Reizer, 1992).
13
Figure 2.3 Sugar transport systems of C. glutamicum. PTSGlc, glucose PTS; PTSFru, fructose PTS;
PTSSuc, sucrose PTS; ?, unidentified transport system. Fru, fructose; Suc, sucrose; Glc, glucose;
G6P, glucose 6-phosphate; F6P, fructose 6-phosphate; FBP, fructose-1,6-bisphasphate; Suc6P,
sucrose-6-phosphate. The inset shows the phosphoryltransfer derived from PEP via the general
PTS phosphortransferases I (EI) and HPr proteins shared by the three substrate specific EII PTS
components (Eggeling and Bott, 2005).
Kiefer et al. (2002) measured approximately 8% and 30% lower L-lysine yield
during C. glutamicum growth on sucrose and fructose, respectively, as compared to the
yield obtained from glucose. Similarly, the final biomass obtained in fructose and sucrose
grown C. glutamicum was approximately 20% lower than that of glucose grown cells.
Furthermore, the NADPH generation via the pentose phosphate pathway has been shown
to be lower on fructose and glucose/fructose mixtures as compared to glucose
(Dominguez et al., 1998; Kiefer et al., 2004). These results confirm that the activity of
PTSs is dependent upon the substrates, and the growth of C. glutamicum as well as the
production of amino acids is influenced by PTSs. On the other hand, glutamate production
is not affected by the PTSs since NADPH requirement in all cases was observed to be
fulfilled by the isocitrate dehydrogenase (Georgi et al., 2005). The PTS activity in C.
glutamicum has also been shown to be dependent on the specific growth rates since the
activity at 0.1h-1 was higher (6mmol g-1 h-1) as compared to the activity (3.9-4.1mmol g-1
14
h-1) measured at 0.1h-1. However, this feature of C. glutamicum is favourable since the
amino acid overproduction generally occurred at low growth rates or during the stationary
growth phase (Cocaign-Bousquet et al., 1996). Therefore, detailed understanding of the
molecular mechanism of PTS and carbon regulation are the prerequisites for making a
more rational design for the improvement of amino acid-producing strains.
After sugar uptake and phosphorylation, further metabolism of sugar phosphate
occurred via both the central metabolic pathway (glycolysis) and pentose phosphate (PP)
pathway. Kinoshita (1985) demonstrated that the carbohydrate metabolism of C.
glutamicum performs glycolysis, PP pathway, Tricarboxylic acid (TCA) cycle and
glyoxalate cycle. Glycolysis is responsible for the conversion of one molecule of glucose
into two molecules of pyruvate that are transferred to the TCA cycle for biosynthesis of
amino acids and macromolecules. In addition, glycolysis generates several intermediates
and high-energy molecules i.e., ATP (transports chemical energy within cells) and NADH
(is used for anabolic metabolism). On the other hand, PP pathway, a bypass of glycolysis,
is branched off at glucose-6-phosphate and refuels glycolysis at the levels of F6P and
glyceraldehyde-3-phosphate. The general role of PP pathway is to supply anabolic
reducing power and precursor metabolites i.e., NADPH, ribose-5-phosphate (R5P) and
erythrose 4-phosphate for the synthesis of essential macromolecules and certain amino
acids (Stryer et al., 2002). The growth of C. glutamicum has been reported to be
dependent upon its capacity to generate adequate NADPH for the anabolic pathways, and
the requirement of this cofactor is generally met by the high flux through the PP pathway
during growth on glucose (Cocaign-Bousquet et al., 1996).
The carbon flux distribution between the glycolysis and PP pathways of C.
glutamicum grown on glucose has been shown to be slightly in favour of the PP pathway
although the phenomenon is not common in other organisms. 13C-labeling studies coupled
with nuclear magnetic resonance (NMR) analysis showed that the flux through the PP
pathway decreases with the production of L-glutamate throughout the fermentation of C.
glutamicum (Marx et al., 1996). This result indicated that the demand of NADPH is
decreased during L-glutamate production where the requirement of this cofactor is
fulfilled by the isocitrate dehydrogenase (Georgi et al., 2005). However, the situation is
entirely different in the case of L-lysine production since a high level of NADPH is
required for the biosynthetic pathways. Ishino et al. (1991) measured the contribution of
15
those two pathways in both L-glutamate and L-lysine production. Their result showed that
the ratio of glycolysis/PP during L-glutamate fermentation is 80/20, whereas it is 3040/60-70 during L-lysine production. The result demonstrated that the flux distribution
between those two pathways is strictly dependent on the type of amino acids produced in
C. glutamicum and the conditions supplied. Therefore, a better understanding and
manipulation of those two major sugar catabolism pathways is one of the most important
targets for the improvement of amino acids production by C. glutamicum fermentation.
The TCA cycle is responsible for the complete oxidation of acetyl-CoA that is
derived from pyruvate. It provides precursor metabolites for the biosynthesis of 2oxoglutarate (the precursor of L-glutamate) and oxaloacetate (the precursor of aspartate
and its derivatives). In C. glutamicum, pyruvate dehydrogenase complex (PDHC)
catalyzes the oxidative decarboxylation of pyruvate; and produces acetyl-CoA, CO2 and
reduced NAD. Acetyl-CoA is then condensed with oxaloacetate to form citrate that is
further catalyzed with a series of reactions. Figure 2.4 shows a schematic diagram of the
central metabolism of C. glutamicum during growth on glucose and acetate. During
growth on substrates (acetate, fatty acids, ethanol) that enter into the central metabolism at
the level of acetyl-CoA, TCA cycle processes glyoxylate pathway which avoids the
oxidative decarboxylation steps of isocitrate dehydrogenase (ICDH) and the 2oxoglutarate dehydrogenase complex (OGDHC), and finally produces one molecule of
malate from two molecules of acetyl-CoA (Eggeling and Bott, 2005).
However, some of the important enzymes in TCA cycle have been studied in order
to characterise their activity and to investigate their role both in C. glutamicum growth
and amino acids production. The initial reaction of the TCA cycle is catalysed by citrate
synthase (CS) that condenses acetyl-CoA and oxaloacetate in order to form citrate. CS is
considered to be the rate controlling enzyme for the entry of substrates into the TCA
cycle. However, the specific activity of CS in this organism was found to be independent
of the level of substrate and the phase of growth (Wendisch et al., 2000). Eikmanns et al.
(1994) confirmed that L-glutamate enhancement in C. glutamicum is not possible by
simply increasing the CS activity since both the gltA (encodes for CS)-negative and gltAoverexpressed strains showed an identical L-glutamate secretion [17µmol min-1 (g Dw) 1
]. Isocitrate dehydrogenase (ICDH) is an important enzyme since it provides 2-
oxoglutarate as a precursor of glutamate and reducing power (NADPH) by oxidative
16
decarboxylation of isocitrate. In C. glutamicum, the requirement of 1mol NADPH per mol
of glutamate production is fulfilled by ICDH. Like CS, the specific activity of ICDH is
also independent of the level of substrate and phase of growth. L-glutamate enhancement
in C. glutamicum is not possible by simply increasing the ICDH activity since both the icd
(encodes for ICDH)-negative and icd-overexpressed strains showed an identical glutamate
secretion rate of about 19µmol min-1 (g Dw)-1 (Eikmanns et al., 1995).
Figure 2.4 Diagram of the central metabolism of C. glutamicum during growth on glucose and
acetate. Dotted arrows represent pathways consisting of several reactions, uninterrupted arrows
represent single reactions. AK, acetate kinase; PTA, phosphotransacetylase; ICDH, isocitrate
dehydrogenase; ICL, isocitrate lyase; MS, malate synthase; PEPck, phosphoenolpyruvate
carboxykinase; Pc, pyruvate carboxylase; PEPc, phosphoenolpyruvate carboxylase (Eggeling and
Bott, 2005).
17
2-oxoglutarate dehydrogenase complex (OGDHC) catalyses the oxidative
decarboxylation of 2-oxoglutarate to succinyl-CoA. In the TCA cycle, OGDHC competes
with the glutamate dehydrogenase (GDH) for the common substrate 2-oxoglutarate.
However, the specific activity of GDH in C. glutamicum has been shown to be 20-fold
higher than OGDHC (Bormann et al., 1992). Kawahara et al. (1997) demonstrated that
OGDHC acts at the branch point of metabolic flux distribution between L-glutamate
synthesis and energy production via the TCA cycle. Although their results showed an
increase of L-glutamate production during biotin limited cultivation of C. glutamicum, a
reduction in the activity of OGDHC was observed, whereas the activity of GDH was
hardly affected. Since the TCA cycle provides precursors for the synthesis of glutamate
and aspartate family amino acids, it is obvious that the carbon fluxes through this cycle
and their control are of major importance for the fermentative production of amino acids
in C. glutamicum. In addition, the activities of some of the TCA cycle enzymes
especially, PDHC and OGDHC have a severe impact on amino acids production (Asakura
et al., 2007; Kawahara et al., 1997; Kimura et al., 1999; Shimizu et al., 2003). However,
the knowledge of C. glutamicum genome and metabolic engineering approaches will
allow us to change the specific enzymatic properties in the central metabolism, and to
increase the carbon fluxes towards the metabolic pathways leading to the synthesis of a
particular amino acid.
2.1.3. Anaplerotic pathways of Corynebacteria
Anaplerotic pathways are usually responsible for the replenishment of TCA cycle
with C4-dicarboxylic acids. Several anaplerotic enzymes have been found in C.
glutamicum, and among them phosphoenolpyruvate carboxylase (PEPc) and pyruvate
carboxylase (Pc) are the two most important enzymes (Peters-Wendisch et al., 1997) that
participate in growth and amino acid production (Delaunay et al., 1999a; Peters-Wendisch
et al., 2001). In addition to the C3-carboxylating enzymes, C. glutamicum also possesses
a few C4-decarboxylating enzymes that convert oxaloacetate or malate to PEP or
pyruvate. These are phosphoenolpyruvate carboxykinase (PEPck), malic enzymes (ME),
oxaloacetate decarboxylase and the glyoxylic cycle enzymes i.e., isocitrate lyase and
malate synthase (Gourdon et al., 2000; Jetten and Sinskey, 1993; 1995a). Figure 2.5
shows the major anaplerotic reactions occur in the central metabolism of C. glutamicum.
It has been investigated that the yield and productivity of aspartate-derived amino acids
and L-glutamate synthesis are dependent on the carbon flux through the anaplerotic
18
pathways. In addition, the anaplerotic reactions present at the junction between glycolysis
and the TCA cycle are of particular importance for the synthesis of L-glutamate (Eggeling
and Bott, 2005; Vallino and Stephanopoulos, 1993).
Figure 2.5 Anaplerotic reactions occurring in the central metabolism of C. glutamicum.
Abbreviations used: CoA, coenzyme A; PEP, phosphoenolpyruvate; PTS phosphotransferase
system for glucose uptake (Eggeling and Bott, 2005).
In order to investigate the importance of phosphoenolpyruvate carboxylase (PEPc)
activity on C. glutamicum growth and L-lysine production, Peters-Wendisch et al. (1993)
cultivated wild type of C. glutamicum, MH20-22B (a strain of C. glutamicum the
produces L-lysine) and their respective mutants in which ppc gene (encode for PEPc) was
disrupted by gene-directed mutagenesis. The results showed a similar growth pattern in all
the strains and no prominent influence on L-lysine production in the corresponding
mutants as compared to the parental strain, confirming that the growth of C. glutamicum
and L-lysine production in MH20-22B are independent on the activity of PEPc. To
investigate further, Peters-Wendisch and his co-workers (1996) constructed the
recombinants of parental strains in which both the PEPc and isocitrate-lyase (ICL) were
deleted. Their results demonstrated that both the PEPc and glyoxylate cycle are
dispensable not only for growth but also for lysine production in C. glutamicum.
Moreover, Cocaign-Bousquet et al. (1996) confirmed that PEPc does not play a
19
significant role in fuelling the TCA cycle during growth of C. glutamicum but may play
an important role during amino acid production at relatively low growth rates.
In order to explore the role of pyruvate carboxylase (Pc) in anaplerotic reactions,
Peters-Wendisch et al. (1998) constructed pyc (encodes for Pc)-inactive mutant of C.
glutamicum that showed negligible growth on lactate. In addition, a C. glutamicum mutant
in which both the pyc and ppc genes were inactivated did not show any growth even in the
presence of glucose. These results confirmed that Pc is the main anaplerotic enzyme in C.
glutamicum, and there is no further anaplerotic enzyme active during growth on
carbohydrates. The same group constructed pyc-amplified and pyc-inactive recombinants
of C. glutamicum with their respective host strains to investigate the significance of Pc in
amino acids production. The pyc-amplified strains showed more than 7-fold higher
glutamate, approximately 50% higher lysine and 10-20% higher threonine production as
compared to the original host strains. In contrast, pyc-inactive mutants showed about 2fold lower glutamate and approximately 60% lower lysine production in comparison to
the host strains. These results demonstrated that Pc of C. glutamicum has a potential role
in amino acids production, and will be an important target for metabolic engineering of C.
glutamicum (Peters-Wendisch et al., 2001). However, Delaunay et al. (1999a)
demonstrated that PEPc carries up to 70% of glutamate flux, whereas Pc is responsible for
the remaining 30% during a temperature-triggered L-glutamate production with C.
glutamicum.
Like PEPc and Pc, the role of phosphoenolpyruvate carboxykinase (PEPck) in C.
glutamicum growth and amino acids production has been investigated (Riedel et al.,
2001). Although the pck (encodes for PEPck)-inactive strain has been shown to grow on
glucose, no growth was observed in presence of acetate or lactate. This result confirmed
that PEPck has an important function during growth on substrates other than glucose.
Besides this finding, the glutamate production in pck-inactive and pck-amplified strains
was approximately 4-fold higher and 2 to 3-fold lower, respectively, as compared to the
parental strain. Similarly, the inactivation and amplification of pck gene in C. glutamicum
MH20-22B resulted in only 20% higher and lower lysine accumulation, respectively, as
compared to the original strain (Riedel et al., 2001). The results demonstrated that the
production of TCA cycle derived amino acids on carbohydrates can be increased by
attenuating the activity of this enzyme. Petersen et al. (2001) investigated PEPck activity
20
in L-lysine-producing C. glutamicum MH20-22B by deleting the respective pck gene that
resulted in an increase of intracellular concentrations of oxaloacetate, L-aspartate, ketoglutarate, pyruvate and L-lysine, whereas increasing the PEPck activity by pck
overexpression showed opposite effects.
Malic Enzyme (ME) plays an important role in NADPH generation during growth
on substrates other than glucose (Cocaign-Bousquet et al., 1995; 1996; Dominguez et al.,
1998). To investigate the physiological role of ME in C. glutamicum, Gourdon et al.
(2000) constructed malE (encodes for ME)-inactive and malE-amplified strains. Both the
recombinants grown either on acetate or glucose showed identical specific growth rates as
compared to the wild-type strain. In contrast, the malE-inactive strain grown on lactate
showed a reduced growth rate after 8h of cultivation, while the amplified strain
maintained the exponential growth throughout the fermentation. These results revealed
that the extent of exponential growth period when grown on lactate is dependent upon the
level of ME activity (Gourdon et al., 2000). From the above findings, it is apparent that
most of the anapleorotic enzymes in C. glutamicum mentioned in literature have a certain
role in cellular metabolism although pyruvate carboxylase (Pc) is considered as the most
important enzyme for carboxylation reaction, and PEP carboxykinase (PEPck) is
responsible for decarboxylation reaction.
2.1.4. Uptake and assimilation of ammonium in Corynebacteria
Nitrogen is one of the most essential components for all macromolecules i.e.,
protein, nucleic acid and cell wall components of a bacterial cell. The fermentative
cultivation of C. glutamicum is usually supplied with external nitrogen sources that are
taken up by cells, and thereafter assimilated to accomplish their metabolism (Burkovski,
2003a; b). Hence, the uptake and assimilation of nitrogen sources and their regulatory
mechanisms in C. glutamicum have a great importance in amino acids production. It is
well-known that the uptake of nitrogen sources is mediated either by passive diffusion
(ammonium and urea) or active transport. In C. glutamicum, a considerable number of
transporters involved in taking up of different nitrogen sources i.e., ammonium (MeierWagner et al., 2001; Siewe et al., 1996), urea (Siewe et al., 1998), L-glutamine (Siewe et
al., 1995) and L-glutamate (Burkovski et al., 1996; Kronemeyer et al., 1995) have been
investigated biochemically and genetically. In the presence of high concentration of
ammonium, the diffusion of uncharged ammonia (NH3) occurs through the cytoplasmic
21
membrane in order to promote the growth of cells. When the diffusion into cells becomes
limited for metabolism, a special ammonium transporter (AmtB) is activated to cope with
the nitrogen starvation (Jakoby et al., 2000; Meier-Wagner et al., 2001).
C. glutamicum hydrolyzes urea to ammonium and CO2. In presence of high
concentration of urea, it passes through the cytoplasmic membrane by passive diffusion,
whereas an energy dependent urea uptake system is active during nitrogen starvation
(Siewe et al., 1998). This bacterium can utilize L-glutamine as a nitrogen source that is
converted to glutamate by glutaminase or glutamate synthase. Siewe et al. (1995)
demonstrated that L-glutamine uptake is mediated by a sodium-dependent secondary
transport system where both the membrane potential and sodium gradient are the main
driving forces. C. glutamicum is also able to grow on L-glutamate as sole nitrogen and
carbon source where glutamate is taken up via a binding protein-dependent transport
system (Kramer and Lambert, 1990), encoded by the gluABCD gene cluster (Kronemeyer
et al., 1995) and a secondary sodium couple carrier (Burkovski et al., 1996). The DNA
microarray experiments revealed that the transcription of gluABCD gene is repressed
under high nitrogen supply (Burkovski, 2003b).
Assimilation of nitrogen has a significant influence in glutamate production by C.
glutamicum. Three major enzymes are involved in this process i.e., glutamine synthetase
(GS), glutamate synthase (GOGAT) and glutamate dehydrogenase (GDH). GS
incorporates ammonium into glutamine, GOGAT converts the glutamine to glutamate,
and GDH catalyses the reversible reductive amination of -ketoglutarate to glutamate
(Schulz et al., 2001). Figure 2.6 shows the ammonia assimilation or L-glutamate synthesis
by GDH and GS/GOGAT. C. glutamicum generally follows two different pathways for
ammonium assimilation i.e., in presence of excess nitrogen supply (>1mmol l-1);
ammonium is assimilated with one mole of NADPH to form glutamate by GDH. In
contrast, ammonium is assimilated to glutamine by GS in presence of low ammonia
concentration (<1mmol l-1), and subsequently glutamine is metabolized to glutamate by
GOGAT (Burkovski, 2003a; b). In the GS/GOGAT pathway, glutamate is amidated with
the consumption of ATP to form glutamine by GS. The amide group is then transferred
reductively to 2-oxoglutarate by GOGAT, resulting in the net conversion of 2oxoglutarate to glutamate. Both the GDH and GS-GOGAT pathways produce 1 mole of
glutamate from 1 mole of NH3, 2-oxoglutarate and NADPH. However, the GS-GOGAT
22
pathway is energetically more expensive than GDH pathway since it consumes 1 ATP
during the process. The following equations (1-3) represent the main reactions occurring
during ammonia assimilation of C. glutamicum.
Figure 2.6 Ammonia assimilation or L-glutamate synthesis by the GDH and GS/GOGAT system.
Abbreviations used GDH, glutamate dehydrogenase; GS, glutamine synthetase; and GOGAT,
glutamate synthase (Kimura, 2002a).
GDH
NH3 + 2-oxoglutarate + NADPH + H+
glutamate + NADP+ ---------- (1)
GS-GOGAT
NH3 + glutamate + ATP
glutamine + ADP + Pi ---------- (2)
glutamine + 2-oxoglutarate + NADPH + H+
2 glutamate + NADP+-------- (3)
GDH, GS and GOGAT have already been detected and characterized in C.
glutamicum and its subspecies B. flavum by several research groups since 1970’s. It is
verified that (i) GDH-negative mutants are unable to synthesize glutamate (Shiio and
Ujigawa, 1978), (ii) the activity of GDH in this organism is higher than that of GOGAT
(Sung et al., 1984) and (iii) the GS/GOGAT system is repressed under high NH4+
concentrations (Sung et al., 1985). In order to investigate the consequences of GDH
deletion and amplification on glutamate production in C. glutamicum, Bormann-El Kholy
et al. (1993) cultivated wild type, GDH-negative and GDH-amplified mutants. The results
revealed that GDH is not essential for glutamate synthesis since the glutamate production
in GDH-negative (21 ± 2µmol g Dw-1) was almost similar to the wild type (20 ± 2µmol g
23
Dw-1). Moreover, they also demonstrated that the glutamate secretion in C. glutamicum
cannot be enhanced by only elevating its activity although the GDH activity in GDHamplified mutant was 11-fold higher as compared to the wild type. The specific activities
of GS and GOGAT were also determined in all the above-mentioned strains. Surprisingly,
the GDH-negative strain showed 2 and 10-fold higher specific activities of GS and
GOGAT, respectively, as compared to the wild type. These results indicated that
GS/GOGAT pathway in C. glutamicum is regulated in response to the availability of
GDH. In addition, Beckers et al. (2001) demonstrated that glutamate synthase
(GOGAT) of C. glutamicum is not essential for glutamate synthesis, and the activity
of this enzyme is regulated by the nitrogen status.
Tesch et al. (1998) investigated the influence of different NH4+ concentrations on
cell growth, GDH, GS and GOGAT activities both in wild-type and GDH-negative
mutant of C. glutamicum. Although the specific growth rates of both strains were similar
at NH4+ concentrations ≤ 5mM, GDH-negative mutant grew significantly slower than the
wild type at NH4+ concentrations  5mM. This result indicated that GDH is dispensable
for the growth of C. glutamicum in presence of low concentrations of NH4+, and the
GS/GOGAT pathway is able to maintain the optimal growth of C. glutamicum at NH4+
concentrations ≤ 5mM. Moreover, the wild strain showed a constant activity of GDH
(1.3U mg protein-1) at NH4+ concentrations of 1 to 90mM, whereas the GS and GOGAT
activities were observed to decrease at NH4+ concentrations  10mM. In the case of GDHdeficient mutant, both the GS and GOGAT activities were distinctly higher at NH4+
concentrations 10mM than the activities measured in the wild type. These results
suggested that the GS/GOGAT pathway in ammonium assimilation is not required for the
growth of wild type C. glutamicum grown at NH4+ concentrations  10mM, and the
regulation of GS and GOGAT activities in C. glutamicum is also dependent on the
availability of NH4+.
However, the literature showed that the activities of these enzymes may be
regulated under certain growth conditions. Jakoby et al. (1999) demonstrated that the
regulation of glnA (gene encodes for GS) transcription is usually caused by
adenylylation/deadenylylation via bifunctional enzyme adenylyltransferase (ATase,
encoded by glnE gene). In order to verify the above interpretation, Nolden and his co-
24
workers (2001a) investigated the GS activity in glnE-negative mutant grown under both
nitrogen surplus and nitrogen starvation conditions. The result showed no regulation of
GS activity in the glnE deletion mutant, as compared to the wild type, indicating that
GlnE-encoded product is responsible for regulating the GS activity in C. glutamicum. The
bacterial GOGAT enzyme consists of a large and a small subunit, encoded by the gltB and
gltD genes, respectively. The GlnK-type protein (glnK, formerly named as gltB) and an
uridylytrasferase (glnD) of C. glutamicum are involved in nitrogen sensing and signal
transfer (Jakoby et al., 1999). It is also demonstrated that GlnK and uridylytrasferase i.e.,
an intact nitrogen regulation cascade are essential for the derepression of AmtR-controlled
genes in C. glutamicum under nitrogen starvation (Nolden et al., 2001b). AmtR, a TetRtype repressor protein, represses the transcription of more than 20 genes i.e., amt, amtB,
glnK, glnD, gltB, gltD, ocd, soxA and glnA in presence of nitrogen-rich medium (Jakoby
et al., 2000).
Although several studies have already been dealt with the effects of nitrogen
sources and their concentration on the regulation of nitrogen assimilatory genes, Schulz et
al. (2001) first investigated the effect of carbon status on their regulation. C. glutamicum
was cultivated in minimal medium (MM) containing 2% glucose where 2 and 100mM
NH4Cl were supplied in order to establish nitrogen-limited or nitrogen-rich conditions,
respectively. The GS and GOGAT activities in N-limited condition were approximately 5
and 7-fold higher, respectively, than the activities obtained in N-rich condition. In order to
investigate the role of carbon status on both GS and GOGAT activities, the glucose
concentration in MM was reduced to 0.05%. The results showed that GS activity is
approximately 3-fold lower in C-limited condition (MM with 0.05% glucose) than the
activity measured in N-limited MM, whereas there was no induction of GOGAT activity
observed under C-limited condition. Moreover, the transcription of glnA and gltBD genes
is approximately 5 and 7-fold higher, respectively, in N-limited than in N-excess
condition (Schulz et al., 2001). The results confirmed that both the GS and GOGAT
activities and the transcription of their respective genes are affected by both nitrogen and
glucose status. The above information has given us a clear picture about the importance of
GDH, GS and GOGAT enzymes in ammonium assimilation of C. glutamicum. It is
expected that this understanding will facilitate in implementing of metabolic engineering
techniques in order to improve the yield of amino acids in industrial scale.
25
2.1.5. Metabolic engineering and metabolic flux analysis (MFA) of
Corynebacteria
Metabolic engineering, a modern technique applied for the improvement of
properties (for example, productivity) of microorganism, is generally accomplished by
manipulating the characteristics of specific enzymes involved in the central metabolic
pathway of a microorganism. According to the definition of Stephanopoulos ‘‘metabolic
engineering is the directed improvement of cellular properties through the modification of
specific biochemical reactions or the introduction of new ones, with the use of
recombinant DNA technology’’ (Stephanopoulos et al., 1998). However, metabolic
engineering mainly deals with the synthesis of pathways leading to new products, the
elimination of pathways leading to by-product formation and the determination of both
intracellular and extracellular fluxes and their control under in vivo conditions. When the
recombinant technology is applied to a given strain, metabolic responses resulting from
the changes mostly cause an effect in metabolic fluxes as well as in the metabolic
network. In addition, the investigation of a specific pathway leading to the product of
interest is not sufficient to provide the entire information of metabolism. Therefore, a
complete biochemical reaction network is generally considered for metabolic engineering
(Christensen and Nielsen, 1999). Several applications of metabolic engineering have
already been mentioned in literature, such as improvement of yield and productivity,
extension of substrate range, production of heterologous proteins and improvement of
overall cellular physiology (Stephanopoulos et al., 1998; Nielsen, 2001).
The methodology of metabolic engineering, an iterative process, is mainly
consisted of two parts i.e., synthetic and analytical. The analytical part that suggests the
possible genetic alterations is focussed on the characterization of cell metabolism,
whereas the synthetic part deals with the molecular biological aspects i.e., construction of
new recombinants (Stephanopoulos et al., 1998). This approach allows us to determine
how metabolic fluxes are controlled by a particular pathway, and how the fluxes are
changed due to the environmental and genetic changes. A metabolic pathway is defined as
‘‘any sequence of feasible and observable biochemical reactions steps connecting a
specialized set of input and output metabolites’’. On the other hand the metabolic flux can
be defined as ‘‘the rate at which material is processed through a metabolic pathway’’.
The measurement of fluxes provides essential information regarding the cell physiology
and metabolism, and suggests the probable genetic or environmental modifications.
26
Fluxes determine the degree of participation of various enzymes in different reactions
occurring within the cell (Stephanopoulos, 1999).
The method for determining the metabolic fluxes has been named as metabolic
flux analysis (MFA). MFA is usually applied to calculate the theoretical yields, to
determine the unmeasured metabolite rates and to investigate the function of metabolic
pathways in vivo. MFA provides information about the bottleneck reactions and rigid
branch points involved in microbial growth as well as metabolite production. In the case
of MFA, a set of linear equations is constructed by studying the biochemical
stoichiometry in order to measure the unknown metabolic fluxes from the measured
fluxes i.e., substrate consumption and biomass or product formation rates. MFA has
successfully been applied in order to study Penicillium chrysogenum, Saccharomyces
cerevisiae, Bacillus subtilis, E. coli, C. glutamicum and many others (Stephanopoulos,
1999). Metabolic balancing alone or a combination of both metabolite and isotope balance
are the main approaches used for determining the metabolic fluxes of a certain pathway.
Metabolic balancing is the classical and easily accessible approach used for estimating net
fluxes in a pre-defined metabolic network. It is based on the mass balances over each
metabolite i.e., the sum of fluxes into an intermediate has to be the same as the sum of
fluxes leaving the intermediate at a steady state condition (Christensen and Nielsen, 1999;
Stephanopoulos, 1999).
Vallino and Stephanopoulos (1993) applied the concept of metabolic balancing in
order to estimate the intracellular fluxes at different growth stages of a batch cultivation of
a lysine producing C. glutamicum. The results showed that both the PP pathway and the
anaplerotic enzyme (PEPc) support significant fluxes during growth and L-lysine
overproduction. The same authors also applied this approach in order to determine the
branch points of metabolic or network rigidity during lysine production in C. glutamicum.
The flux data during the transition from glucose to lysine overproduction showed that the
glucose-6-phosphate (G6P) and pyruvate (Pry) nodes are flexible, indicating that lysine
yield is not limited by the G6P and Pry branch points and the yield must be limited due to
the rigidity at the PEP node (Vallino and Stephanopoulos, 1994a; b). This approach is
also applied in order to quantify the intracellular fluxes in the metabolism of
pantothenate-overproducing C. glutamicum (Chassagnole et al., 2002; 2003).
27
In the case of metabolic balancing, however, the metabolic network needs to be
defined properly in order to estimate all the intracellular fluxes. Furthermore, it is not
always known which pathways in the metabolism are actually active since the biological
networks are often very large and complex. Therefore, this approach is not suitable to
differentiate between two pathways when both the pathways in a given metabolic network
are leading to the same metabolite. In metabolite balancing, the cofactor balances
(NADH, NADPH and ATP) are therefore considered based on the assumption that all the
cofactors generated during the metabolism of a particular substrate are consumed in the
subsequent reactions. These balances give a number of limitations besides the flux
constraints that arise from the metabolite balances over the intermediates. To alleviate
these problems in determining the fluxes, a combination of both metabolite and isotope
balancing has recently been used (Wiechert et al., 2001). This approach is considered for
both the identification of active pathways and the estimation of relative fluxes through
two pathways. In such studies, isotope-labelled compounds (13C or 15N) are used as tracer
substrates. When the label substrate is metabolised by cells, labelled carbon atoms are
distributed all over the metabolic network according to the carbon-carbon transition of the
involved reactions. The resulting metabolic labelling patterns are then measured by mass
(MS) or nuclear magnetic resonance (NMR) spectroscopy, and thereafter the data are used
to calculate the intracellular metabolic fluxes (Christensen and Nielsen, 1999).
13
C-labeling technique has successfully been implemented in order to calculate the
in vivo fluxes through the oxidative part of PP pathway based on their intracellular
metabolite concentrations and by determining the kinetic constants of the enzymes in vitro
(Sahm et al., 2000). Their results demonstrated that the oxidative part of the PP pathway
in C. glutamicum is mainly regulated by the ratio of NADPH/NADP concentrations and
the specific enzyme activities of both glucose-6-phosphate and 6 phosphogluconate
dehydrogenases. The investigation of different anaplerotic enzymes in C. glutamicum
showed that both the carboxylation and decarboxylation reactions in the anaplerotic node
occur simultaneously. Moreover, the results demonstrated that C. glutamicum possesses
two biosynthetic pathways for the synthesis of DL-diaminopimelate and L-lysine, and the
relative use of both pathways in vivo is dependent on the ammonium concentration in the
culture medium (Sahm et al., 2000). The flux quantification in the central metabolism of
C. glutamicum MH20-22B by NMR spectroscopy combined with metabolite balancing
showed that the entry of glucose 6-phosphate flux into the PP pathway is 66.4%, whereas
28
32.3% flux enters into the glycolysis (Marx et al., 1996). Using this principle, the same
authors also determined the activity of PP pathway in a number of different strains of C.
glutamicum under the production of two essential amino acids i.e., L-glutamate and Llysine. The PP pathway was observed to be the major route for supplying the biosynthetic
reducing power since approximately 70% of the total NADPH generated via this pathway,
whereas isocitrate dehydrogenase supplied the remaining 30% of NADPH (Marx et al.,
1996). The results showed that the PP pathway contributes more flux towards L-lysine
than L-glutamate fermentation. In addition, the activity of PP pathway reduced during
glutamate production since only 1mol of NADPH is required per mol of glutamate
formation which is fulfilled by the isocitrate dehydrogenase.
Sonntag et al. (1995) investigated the carbon flux distribution in the central
metabolism of C. glutamicum during exponential growth as well as during overproduction
of L-lysine and L-glutamate. Using
13
C NMR data in conjunction with stoichiometric
metabolite balances, they observed that the molar fluxes via the PP pathway were 40 and
17 during exponential growth and L-glutamate production, respectively. On the other
hand, the flux through the PP pathway during L-lysine production was 47, indicating that
a high requirement of NADPH for C. glutamicum growth and during L-lysine
overproduction is taking place via the PP pathway (Sonntag et al., 1995).
13
C-NMR
coupled with gas chromatography-mass spectrometry (GC-MS) has been applied in order
to investigate the importance of both PEPc and Pc to the carbon flux distribution in
anaplerotic pathways of C. glutamicum. The results showed that Pc is responsible for up
to 90% of the carbon flux through the anaplerotic pathways, whereas PEPc contributes
approximately 10% of the total oxaloacetate synthesis during L-lysine production in C.
glutamicum (Park et al., 1997). Using this same strategy, Petersen et al. (2000) also
quantified the individual fluxes at the anaplerotic node where Pc is found to contribute
91 ± 7% fluxes to C3 carboxylation and PEPck is responsible to recycle the excess
oxaloacetate to PEP.
13
C-MFA has also been applied to investigate the growth and carbon-flux
distribution in the central metabolic pathways of C. glutamicum grown on different
carbon sources. Dominguez et al. (1998) observed that the growth of this organism on
fructose is significantly less than that on glucose although the substrate uptake rates were
the same in both conditions. The NMR analysis of carbon-isotope distribution to the
29
glutamate pool during C. glutamicum growth on 1-13C or 6-13C-enriched fructose revealed
that 80% of total fructose consumption occurs via glycolysis, whereas more than 50% of
total glucose consumption takes place via the PP pathway during growth on glucose. MFA
by
13
C labelling in combination with GC-MS, metabolite balancing and isotopomer
modelling revealed that intracellular flux distribution is dependent on the carbon sources
applied during fermentation (Kiefer et al., 2004). The result showed that the flux through
the PP pathway is only 14.4% of the total substrate uptake fluxes during growth on
fructose, whereas the flux through this pathway is 62.0% during growth on glucose.
The ammonia-assimilatory pathways of C. glutamicum wild type and GDHnegative mutant have been investigated by in vivo flux analysis during carbon limited but
ammonium abundant (30 to 40mM) chemostat cultivations (Tesch et al., 1999). The 15N
NMR spectroscopy analysis of ammonium flux through the wild type revealed that 28%
of NH4+ is assimilated by the GS reaction involving glutamine, whereas 72% of NH4+ is
assimilated by the GDH via L-glutamate. However, there is no GOGAT activity observed
in the wild type of C. glutamicum, indicating that the GOGAT pathway is fully inactive at
the above-mentioned condition. In contrast to the wild type, glutamate is completely
synthesized through the GS/GOGAT pathway in GDH-deficient mutant although the
GOGAT has a weak dependency on the availability of NH4+. From the above examples, it
is apparent that MFA has been successfully applied in order to understand a detailed
knowledge about the metabolic pathways, the activity (in vivo) of enzymes and their
regulation associated with the central metabolism of C. glutamicum. Since the demand for
these essential amino acids is increasing rapidly, it is necessary to apply functional
genomics and metabolic engineering in a more effective way in order to improve the yield
and productivity of this bacterium (Wendisch et al., 2006).
30
2.2. Industrial production of amino acids
2.2.1. Introduction
Amino acids are the basic structural building units of proteins. The economic
importance of L-amino acids is noteworthy due to their use for a variety of purposes,
mainly food additives and feed supplements (Eggeling and Sahm, 1999; Leuchtenberger,
1996; Mueller and Huebner, 2002). Until the 1950’s, no suitable commercial process for
the production of sodium glutamate existed except the isolation of glutamic acid from
vegetable proteins. This method, involving the hydrolysis of wheat gluten or soybean by
acid (HCl), is reasonably expensive and produces D-isomer of amino acids (Hirose et al.,
1985). Therefore, continuous efforts had been given in order to develop an alternative
process for the production of L-amino acids. At present, however, most L-amino acids
(i.e., L-lysine, L-glutamate, L-threonine and L-isoleucine) are produced by microbial
(especially Corynebacteria) fermentation (Hermann, 2003; Ikeda, 2002). In Table 2.1, the
characterized features of amino acids production by microbial fermentation are
summarized. There are two organisms mainly used for the fermentative production of
amino acids i.e., C. glutamicum is cultivated for the production of L-glutamate, L-lysine
and L-phenylalanine (Eggeling and Sahm, 1999; Kramer, 1996; 2004), whereas E. coli is
successfully used for L-threonine and L-tryptophan production (Aiba et al., 1980). A
number of other organisms (for instance, mutants of Serratia marcescens) have also been
used for L-threonine production (Komatsubara et al., 1983) although the production
processes in those organisms are not economically suitable (Leuchtenberger, 1996).
Nevertheless, C. glutamicum (M. glutamicus), a potent microorganism discovered by
Kinoshita and his co-workers (1957), occupies a central role in L-amino acids production
among the bacteria mentioned above. A few amino acids i.e., D, L-methionine, glycine,
Table 2.1 Features of amino acids production by microbial fermentation (Kinoshita, 1985).
1. The amino acids produced by microbial fermentation are active and biologically
desired L-form.
2. Both the fermentation and recovery processes are very simple.
3. Carbohydrates and NH3 are used as the main raw materials
4. Both the investment and production costs are low
5. The production facilities can be used for multi purposes
6. The risk to health and safety is low during production
31
L-cysteine and L-aspartate are still produced either by chemical or enzymatic processes
because of low yield achieved by fermentation (Eggeling and Sahm, 1999; Ikeda, 2002).
The application of these amino acids is enormous. Monosodium glutamate (MSG)
and glycine are used as flavour enhancing agents in food, whereas lysine, threonine,
tryptophan and methionine are supplemented in animal feed. Phenylalanine and aspartate
are widely used in the production of artificial sweeteners. In addition, some derivatives of
L-glutamate i.e., N-acylglutamate and oxopyrrolidinecarboxylic acid are also used as
therapeutic agents in nutritional and metabolic disorders (Hirose et al., 1985). In Table
2.2, the current production of major amino acids, their production processes and uses are
summarised. Approximately 1.5 million tons of L-glutamic acid is produced per year by
Corynebacteria, and the demand for this amino acid is growing by about 6% per year.
Alike the importance of MSG, L-lysine is widely used as a feed additive to increase the
efficiency of feed. In 2001, the demand for lysine throughout the world was 550,000 tons
with a growth rate of 7% per year (Hermann, 2003). The major companies involved in
producing these amino acids through microbial fermentation are Ajinomoto, Miwon,
Kyowa-Hakko, Cheil-Jedang, BASF, ADM and Degussa.
Table 2.2 Current production and application of amino acids (data are the average values for the
years 2002 and 2003) (Kramer, 2004).
Amino acids
Amount (t/y)
Production methods
Major uses
L-glutamate
1,000,000
Fermentation
Flavour enhancer
L-Lysine HCL
600,000
Fermentation
Feed additive
D, L. Methionine
400,000
Chemical synthesis
Feed additive
Glycine
22,000
Chemical synthesis
Food additive
L-Threonine
20,000
Fermentation
Feed additive
L-Aspartate
10,000
Enzymatic method
Sweeteners (aspartame)
L-Phenylalanine
10,000
Fermentation
Sweeteners (aspartame)
L-Cysteine
3,000
Enzymatic method
Food additive
L-Arginine
1,000
Fermentation
Pharmaceuticals
L- Tryptophan
500
Fermentation
Pharmaceuticals
L – Valine
500
Fermentation
Pharmaceuticals
L –Leucine
500
Fermentation
Pharmaceuticals
32
Due to the huge consumption of amino acids in both human and livestock sectors
(Leuchtenberger, 1996; Mueller and Huebner, 2002), the world market for these building
blocks is increasing by 10-15% per year (Hermann, 2003). Although the production of
amino acids is already established by the companies of Japan and Germany, many
academic institutions and other companies are still trying to enhance the production
capabilities of microorganisms by identifying new target genes and quantifying metabolic
activities through the system biology (Eggeling and Sahm, 1999; Sahm et al., 1995;
2000). It addition, the production processes have been improved by bioprocess and
downstream technology (Hermann, 2003). The implementation of fermentation for the
industrial production of amino acids was first introduced by Kinoshita and his colleagues
of Kyowa Hakko Koygo Co. in 1957 (Kinoshita et al., 1957). They discovered a Grampositive soil bacterium, C. glutamicum (initially reported as Micrococcus glutamicus that
is auxotrophic for biotin) which has the ability to produce significant amounts of Lglutamate under biotin limited condition. After this discovery, the same company isolated
a homoserine-auxotrophic mutant of C. glutamicum that produced a significant amount of
L-lysine (Nakayama et al., 1961). These successive achievements led biotech industries to
establish the production of these essential amino acids by C. glutamicum fermentation.
However, the biosynthesis of amino acids in this bacterium is strictly controlled by
several regulating mechanisms including feedback inhibition and repression (Nampoothiri
and Pandey, 1998). At present, most of the L-amino acids are produced by the use of
mutants that contain combinations of both auxotrophic and regulatory mutations (Parekh
et al., 2000). Auxotrophic mutants that provide the opportunity to eliminate feedback
inhibition by limiting the intracellular accumulation of feedback inhibitors and repressors,
and regulatory mutants that are insensitive to end product inhibition and repression have
been developed by the industries in order to increase the production of amino acids
(Hirose et al., 1985). In order to increase the yield and productivity of this bacterium,
however, recombinant strains have been constructed by genetic and metabolic engineering
(Sahm et al., 1995). Recently, Kramer (2004) classified amino acid producing strains into
three different groups i.e., (i) wild type strains that are capable of excreting a particular
amino acid under specific culture conditions, (ii) regulatory mutants in which feedback
control of amino acids biosynthesis is removed, and (iii) genetically modified strains in
which the biosynthetic capacity of a particular amino acid is amplified.
33
2.2.2. Fermentative production of amino acids by Corynebacteria
In the case of microbial fermentation, C. glutamicum is grown aerobically in a
liquid nutrient medium containing a carbon source (preferably glucose), a nitrogen source
(ammonium or urea), mineral salts and growth factors. However, a cost effective
production process is largely dependent on the price of carbon sources and the overall
yield and productivity of amino acids (Hermann, 2003). For laboratory purposes, glucose
is frequently used as a carbon source although sucrose, fructose, acetate and glycerol have
also been used to investigate the central metabolism of C. glutamicum (Kramer, 2004).
Fermentation is generally carried out with the presence of some control variables
(aeration, agitation, pH and temperature) that affect the overall yield and productivity of
amino acids. Once the fermentation starts, C. glutamicum passes through a series of
metabolic pathways in which the desired amino acid is produced (Ikeda, 2002). However,
all the pathways in a metabolic network are controlled by complex patterns of feedback
inhibition, induction and repression mechanisms (Kramer, 1996). Figure 2.7 represents a
schematic drawing of important steps in amino acids production by bacteria.
Figure 2.7 Schematic drawing of important steps in amino acids production by bacteria (Kramer,
1996).
34
However, the utilization of glucose as a carbon source in industrial fermentation is
very limited due to its high price. Hence, cane molasses, beet molasses and starch
hydrolysate from corn and cassava are generally used for the industrial production of
amino acids (Hermann, 2003; Ikeda, 2002). Although molasses is inexpensive, it causes
difficulty in excreting L-glutamate into extracellular medium due to its high content of
biotin. Consequently, a surfactant or lactam antibiotic is supplied as a biotin suppressing
agent at an initial or intermediate stage of cultivation (Eggeling and Sahm, 2001; Kramer,
1996; 2004). Methanol is also used as an alternative carbon source because of its low cost,
availability, high purity and water solubility. It has been observed that methylotrophic
bacteria are able to convert methanol into L-lysine (Bacillus sp.; Methylobacillus
glycogenes) or L-threonine (M. glycogenes) (Motoyama et al., 1994; 2001). Ammonia or
ammonium sulphate is commonly used as a nitrogen source during fermentation. It is
demonstrated that a continuous supply of assimilable nitrogen is favoured for L-glutamate
production (Ikeda, 2002). Nevertheless, the selection of carbon and nitrogen sources is
achieved by trial-and-error or based on the knowledge of metabolic pathways of organism
considered for fermentation (Parekh et al., 2000). In addition to these, appropriate
concentrations of phosphate, sulphate, magnesium, potassium and other minerals are
supplied during fermentation of C. glutamicum.
Beside the above-mentioned factors, the industrial production of amino acids or
any heterologous protein is dependent on the mode of fermentation used. The productivity
of L-glutamate by fermentation is represented simply by the following equation:
Pglu= glu × X × V
Where,
Pglu = L-glutamate productivity by fermentation, L-glutamate (g)/time (h)
glu = L-glutamate productivity by unit cells, L-glutamate (g)/cell (g)/time (h)
X = cell density, cell (g)/volume (l)
V= volume (l)
This equation reveals that the volume of fermentor, cell density and the specific
productivity of cells affect the productivity of a fermentation process. Although glu
depends mainly on the capability of production strains, it is strongly influenced by the
35
process conditions, such as cultivation temperature, pH, medium composition and
aeration (Eggeling and Bott, 2005). Three different modes of cultivation are usually used
in microbial fermentation. In batch cultivation, a bulk amount of carbon and nitrogen
sources are supplied in order to obtain a high titre of amino acids. However, the addition
of high concentration of carbon sources at the start of fermentation often hampers the
growth properties and the production capabilities of microorganism, and eventually
reduces the productivity of desired amino acids. Even though a batch process is
straightforward, it gives low productivity due to a long lag phase (Stanbury et al., 1998).
Therefore, the production of amino acids in industry is generally carried out by fed-batch
fermentation in order to achieve high yield and productivity (Hermann, 2003).
In a fed-batch process, the growth of microorganism can be controlled in a precise
manner. Fed-batch cultivation is initially carried out via batch fermentation. At the end of
batch cultivation, the fed-batch fermentation is started with the supply of fresh medium
containing high concentration carbon source until an optimal yield of product is obtained
(Stanbury et al., 1998). Sassi et al. (1998) obtained 110.6g l-1 L-lysine through the
cultivation of lysine producing C. glutamicum by fed-batch fermentation, whereas only
34g l-1 L-lysine was measured in the case of batch cultivation. Similarly, a fed-batch
process of C. glutamicum enabled production of 85g l-1 of L-glutamate in a biotin rich
medium where the efflux of this amino acid was obtained by increasing the culture
temperature from 33 to 39C (Delaunay et al., 1999b). Due to the high concentration of
carbon source used in fed-batch cultivation, however, the growth of microorganisms is
often inhibited and the yield of product is reduced due to the formation of by-products,
such as acetate and lactate (Ikeda, 2002).
In the case of chemostat fermentation, microorganisms are allowed to grow
exponentially by the continuous addition of fresh medium where the excess fermentation
broth is removed from the bioreactor simultaneously. This mode of operation is extremely
valuable for studying the microbial physiology, such as biochemistry, genetics and
specific enzymatic activity, and provides important information for designing a feeding
strategy of fed batch process. During this mode of cultivation, a steady state condition is
obtained where no accumulation of substrate, product or biomass in the fermentor is
observed. However, the steady state can only be obtained at a dilution rate (D) below the
maximum dilution rate (Dmax) (Stanbury et al., 1998). The continuous fermentation of a
36
L-lysine producing mutant of C. glutamicum produced 105g l-1 lysine with a volumetric
productivity of 5.6g l-1 h-1. Moreover, the productivity measured in continuous
fermentation was 2.5-fold higher than that of the fed-batch culture (Hirao et al., 1989).
Despite having higher productivity in continuous culture as compared to batch or fedbatch cultures, its application in industrial fermentation is very limited because of having
a great susceptibility to contamination, degeneration and spontaneous mutations (Ikeda,
2002).
2.2.3. Factors affecting during amino acids production in Corynebacteria
It is well-known that the growth of microorganisms is dependent on the type of
cells, and is varied in response to the physical, chemical and environmental conditions
during fermentation. However, there are some important factors i.e., dissolved oxygen
concentration and its effective transfer, intracellular pH and osmotic pressure affect the
growth of C. glutamicum as well as amino acids production. Dissolved oxygen (DO)
concentration is one of the most important parameters for any aerobic microbial
fermentation since the glucose uptake rate, yield and productivity are dependent on the
availability of DO. Kwong and Rao (1991) demonstrated that a higher oxygen supply is
mandatory during the amino acids production by C. glutamicum although the exponential
growth is favoured in presence of low oxygen. This finding has also been reported in
another study where L-lysine production by batch fermentation of B. lactofermentum was
performed with 3% DO for the first 24h, 10% for 24-48h and 5% for the rest of
fermentation. The final L-lysine concentration was measured 51.4g l-1, whereas it was 45g
l-1 in the case of cultivation carried out at a constant DO (3%) throughout the study (Yao
et al., 2001).
On the other hand, Hilliger and Hanel (1981) indicated that limitation of oxygen
caused a decrease in biomass yield, substrate consumption and L-lysine production,
concomitant with formation of the by-products L -lactate, L-alanine and L-valine. Their
results showed that the physiology of cell might be affected due to the substrate and
dissolved oxygen uptake rates, facilitating the production of by products, such as carbon
dioxide, acids and biomass. In addition, the number and concentration of by-products
produced during fermentation were observed to be dependent on the amount of O2
supplied and its effective transfer throughout the cultivation. Calik et al. (2001) observed
that pyruvic, lactic and acetic acids production are favoured at lower oxygen transfer rate
37
[at agitation rates of 150-250 rotations per minute (rpm) and an air feed rate of 0.1-0.2 air
volume per reactor volume per minute (vvm)], while the -ketoglutaric and succinic acids
production are favoured at higher oxygen transfer rates (at agitation rate of 500rpm and an
air feed rate of 0.2vvm). However, the maximum L-glutamate was measured at an
agitation rate of 250rpm and air feed rate of 0.1vvm. The influence of aeration flow
during L-lysine production in C. glutamicum has been investigated by batch cultivations.
The results showed that the substrate consumption rate, productivity and yield are
significantly higher at 0.75vvm, and approximately 50% increase in L-lysine production
was observed at 0.75vvm as compared to the L-lysine obtained at 0.5vvm, however, the
production decreased by 15% at 1vvm (Sassi et al., 1996). It is obvious from the above
findings that cell growth, product and by-product formation are also dependent on the
availability of DO, aeration flow and agitation speed during an aerobic cultivation.
The osmotic pressure of fermentation media is one of the major operational
parameters since an increase of external osmolarity causes an efflux of water from cells,
decreases the cell turgor pressure and eventually leads cells to die. The medium
osmolarity is often increased due to the accumulation of amino or organic acids during the
industrial amino acids production. Guillouet and Engasser (1995b) demonstrated that
increase in medium osmolality (0.4 to 2.0Osmol kg-1) interrupts the cell growth, decreases
both the specific growth rate (0.7 to 0.2h-1) and biomass yield (0.6g Dw g-1 to 0.3g Dw g1
) during the batch cultivation of C. glutamicum. Although the level of glutamic acid
(55mg g Dw-1) was not shown to be affected by this perturbation, a decreased level of
biomass (7.5 to 5.5g Dw l-1) was observed. In addition, the glucose uptake rate was found
to be increased in proportion to the specific growth rates at a given medium osmolality
(Guillouet and Engasser, 1995b). Gourdon et al. (2003) confirmed that increase in
medium osmolarity inhibits the PTS sugar uptake capacity, and eventually decreases the
metabolic activity of C. glutamicum.
The intracellular pH (pHi) has a major influence on the metabolic activity of cells
since it determines the in vivo activity of enzymes and modulates the transport kinetics of
nutrients and metabolites. The pHi has been shown to vary from 7.7 to 8.3 during
glutamate production in C. glutamicum although a constant extracellular pH (7.3) was
maintained throughout the batch fermentation. Although the pHi was relatively stable at
7.7 during the lag period, it increased to 8.1 at the growth phase and finally decreased to
38
7.3 at the decline phase. Furthermore, pH gradient was also observed to be related to the
specific glucose consumption rate during both the initial growth and glutamate production
phase of fed-batch cultivation (Leyval et al., 1997). However, it is often observed that the
production yield obtained in the laboratory is not reproduced in industrial scale. This
considerable variation occurs within the bioreactor due to low mixing efficiency during
the scale of operation (Hermann, 2003). Therefore, a better understanding of fluid
dynamics in bioreactor as well as the interaction among the cultural conditions,
environment and microbial physiology is required for scaling up a process successfully.
2.2.4. Mechanisms of L-glutamate efflux in Corynebacteria
The mechanism of L-glutamate secretion has initially been described by the ‘Leak
Model’. According to this hypothesis, L-glutamate is passively leaked out through the cell
membranes that are damaged due to biotin limitation and other treatments. Lack of biotin
decreases the activity of fatty acid synthase that results in the reduction of fatty acids and
phospholipids synthesis, and subsequently alters the physiological properties of plasma
membrane (Takinami et al., 1968). In addition, the feedback inhibition has been shown to
be decreased during biotin limited condition that apparently accelerates the internal
synthesis of L-glutamate and favours its secretion into the external medium (Shiio et al.,
1962; 1963). However, the permeabilization of some ions and organic acids through the
cell membrane during L-glutamate secretion revealed that the secretion of L-glutamate is
not fully influenced by biotin limitation (Kramer, 1994). Clement et al. (1984) proposed
an additional hypothesis, known as ‘Inversion Model’, where the secretion of L-glutamate
is mediated by the glutamate permease that functions as an uptake system. It is observed
that biotin limitation or surfactant treatment, altering the cell membrane composition of C.
glutamicum, selectively uncouples the glutamate uptake system from its energy (proton
motive force) source without affecting the ability of permease to interact with its
substrate. Once uncoupled, the permease causes secretion or diffusion of L-glutamate into
the extracellular medium due to the electrochemical potential (Clement et al., 1984).
However, the investigation of the transport kinetics of L-glutamate and its
regulation demonstrated that both the ‘Leak’ and ‘Inversion’ Models are very uncertain
and not reliable. Hoischen and Kramer (1989) confirmed that a special efflux carrier
system ‘Secretion Carrier’ is involved in the secretion of L-glutamate by C. glutamicum.
According to this hypothesis, L-glutamate secretion in biotin limited cells of C.
39
glutamicum occurs against an existing chemical gradient where the secretion carrier
system is not driven by the membrane potential, pH or other ion gradients. Nevertheless,
it has been reported that the alteration in the lipid state of membrane is necessary, but is
not sufficient to induce glutamate secretion by this bacterium (Hoischen and Kramer,
1989; 1990). From a biochemical point of view, two factors are mentioned as the principal
prerequisites for the production of L-glutamate (Kramer, 1994). First, the central
metabolism must be in an imbalanced situation that results in secretion of an intermediate
(L-glutamate). L-glutamate secretion is caused by an overflow metabolism whenever (1)
the carbon and energy source is present in excess, (2) growth is limited by the lack of an
essential nutrient or another component, and (3) substrate uptake is not effectively
regulated. In addition, effective glutamate efflux is only observed if the plasma membrane
and cell wall are somehow altered resulting in changes in the activity of glutamate
secretion carrier system (Kramer, 1994).
Secondly, L-glutamate production has been observed to co-exist with several
glutamate uptake systems. C. glutamicum possesses a highly active binding-proteindependent glutamate uptake system with a maximal uptake rate (Vmax) of 16nmol min-1
(mg dry weight) -1 (Kronemeyer et al., 1995), and a secondary transporter with a Vmax of
about 15nmol min-1 (mg dry weight)
-1
(Burkovski et al., 1996). The primary system,
under the control of glucose catabolite repression, was shown to be down-regulated in
typical fermentation media (Kronemeyer et al., 1995). On the other hand, the secondary
transporter of C. glutamicum (glutamate permease) that excretes L-glutamate to the
extracellular environment is exclusively active in complex media (Burkovski et al., 1996).
In addition, the L-glutamate secretion is regulated by the metabolic and/or energetic state
of cells since the secretion activity is decreased drastically in presence of different
experimental conditions i.e., limitation of carbon source, change in oxygen availability
and inhibition of the respiratory chain (Hoischen and Kramer, 1989). However, the actual
mechanism of L-glutamate efflux by this bacterium is still uncertain. Therefore, proper
investigations are required in order to discover the mechanisms responsible for Lglutamate production.
Due to these inconsistencies, it has also been assumed that L-glutamate secretion
is induced due to the changes in cytoplasm rather than cell wall of C. glutamicum. To
investigate this interpretation, several studies have already been carried out with the
40
different enzymes or genes associated with L-glutamate biosynthesis, glycolysis, TCA
cycle, anaplerotic pathways and nitrogen assimilation. The role of those pathways or
enzymes or genes in amino acids production was already discussed in Sections 2.1.22.1.5. However, it is well investigated that the activity of 2-oxoglutarate dehydrogenase
complex (OGDHC) is a crucial factor for L-glutamate production by C. glutamicum. In
TCA cycle, the OGDHC competes with glutamate dehydrogenase (GDH) for the common
substrate (2-oxoglutarate). Shingu and Terui (1971) observed approximately 40-60%
decrease of OGDHC activity in B. flavum grown on biotin limited condition as compared
to a biotin rich condition. Similarly, Kawahara et al. (1997) showed a significant
reduction (approximately 80%) of OGDHC activity in glutamate production conditions
(biotin limitation, penicillin addition and surfactant addition) as compared to the nonglutamate production condition, whereas the activity of GDH remained almost constant.
Shimizu et al. (2003) investigated the effect of changes in ICDH, GDH and
OGDHC activities on metabolic flux distribution at the 2-oxoglutarate branch point of C.
glutamicum. Even though ICDH and GDH activities in both icd and gdh-overexpressed
strains increased 2.96 and 3.21-fold, respectively, the flux distribution at the 2oxoglutarate node was not affected. In contrast, the carbon flux towards the 2oxoglutarate branch point (glutamate production) has been shown to be more than 75%
when the OGDHC activity decreased by 50% due to biotin limitation. The results
conclude that the attenuation of OGDHC activity has the greatest impact on glutamate
production in C. glutamicum. In order to investigate the relationship between the decrease
in OGDHC activity and L-glutamate production by C. glutamicum, Asakura et al. ( 2007)
constructed a mutant of this organism in which odhA (encodes for OGDHC) gene was
disrupted. The odhA-disrupted mutant has been shown to accumulate L-glutamate
(265mM) with a final cellular dry weight of 1.3g l-1 in the presence of excess biotin,
whereas the wild type accumulated 1.3mM of L-glutamate and produced a biomass of
3.0g l-1. These results confirmed that the mechanism of L-glutamate production by
Corynebacteria is not primarily related to the membrane structure, and the production of
this amino acid is caused by a change in metabolic flux towards L-glutamate synthesis.
dtsR gene, encodes a component of a biotin-containing enzyme complex involved
in fatty acid synthesis, has been identified in C. glutamicum (Kimura et al., 1996). To
elucidate the role of DtsRl in vivo, Kimura et al. (1997) constructed dtsR1-disrupted
41
mutant of C. glutamicum that requires fatty acids for growth. The growth and L-glutamate
production by both the wild type and mutant in the presence of excess of biotin
supplemented with 1mg ml-1 Tween 40 showed a slower growth as compared to the
mutant. In addition, the dtsR1-disrupted mutant was observed to produce L-glutamate
efficiently even in the presence of excess biotin, whereas the production of this amino
acid in the wild strain was zero. In order to investigate further, the same group revealed
that the amplification of dtsR1 inhibits the induction of L-glutamate overproduction under
biotin limitation, Tween 40 addition and penicillin addition. In addition, they showed that
the activity of OGDHC reduced due to Tween 40 addition, biotin limitation and dtsR
disruption. These results indicate that decrease in the level of DtsR or a complex
containing DtsR triggers the increased synthesis of glutamate from 2-oxoglutarate by
lowering the OGDHC activity (Kimura et al., 1999). Moreover, Kimura (2002a; b)
identified a dtsR1-regulator protein (DRP) that represses the expression level of dtsR1 by
binding to the promoter region of dtsR1. It was assumed that DRP may be a global
metabolic regulator since it induces a drastic metabolic flux change from energy
production via the tricarboxylic acid cycle to L-glutamate overproduction by controlling
the expression level of dtsR1. Table 2.3 represents the mechanisms of L-glutamate
secretion described in literature so far.
Table 2.3 Leak Model and Metabolic Flux Change (MFC) Model (Eggeling and Bott, 2005)
Trigger factor
Leak Model
MFC Model
Biotin limitation
Biotin limitation
Surfactant addition
Surfactant addition
Penicillin addition
Penicillin addition
Oleate auxotrophy
Oleate auxotrophy
Glycerol auxotrophy
Targets
Cell surface
Biotin-containing complex
Cytoplasmic membrane
DtsR1
Fatty acid biosynthesis
dtsR1-regulator protein (DRP)
Cell wall
Hypothetical effect
Membrane permeability
DtsR1 containing complex activity
Leakage through membrane Metabolic flux change
42
2.3. Electroporation
2.3.1. Introduction
Electroporation is regarded as a molecular biology technique in which cell
membrane is reversibly permeabilized due to the application of moderate strength electric
pulses. It is one of the non-viral methods successfully used to transfer genes into living
cells in vitro as well as in vivo (Golzio et al., 2004; Jaroszeski et al., 1999). In 1982,
Neumann and his colleagues first performed in vitro electroporation of mouse lyoma cells
in which the plasmid DNA containing the herpes simplex thymidine kinase (TK) gene is
transferred into cells by imposing electric pulses (8kV cm-1, 5µs) (Neumann et al., 1982).
Since then, it has been applied for gene transformation i.e., the introduction of foreign
DNA into mammalian cells (Mir et al., 1999; Neumann et al., 1982; Nickoloff, 1995),
bacteria (Bonamy et al., 1990; Calvin and Hanawalt, 1988; Dower et al., 1988; Dunican
and Shivnan, 1989; Miller et al., 1988), plant protoplast (Fromm et al., 1985; Riggs and
Bates, 1986) and yeast cells (Karube et al., 1985; Meilhoc et al., 1990). Moreover, this
approach has been widely used in a range of medical applications, such as
electrochemotherapy and transdermal drug delivery (Belehradek et al., 1993; Dev et al.,
2000; Heller et al., 1999; Jaroszeski et al., 1997; Mir and Orlowski, 1999; Orlowski and
Mir, 1993). It has several advantages over the conventional techniques of gene
transformation. These are
 technical simplicity, ease of operation, rapidity and reproducibility;
 greater transformation efficiency (TE) as compared to the chemical methods like
calcium
chloride
(CaCl2)
and
polyethylene
glycol
(PEG)
mediated
transformations. In Escherichia coli, the TE by electroporation is 108-109
transformants/µg of DNA which is much higher than the efficiencies (105-106
transformants/µg of DNA) obtained by chemical methods;
 provides a way to avoid the deleterious toxic effects of chemicals (PEG);
 better control of the position and size of electropores, minimizing leakage of
cytosolic components;
 determination of electrical parameters, their control and optimization result in
greater performance;
 no need for the pre-incubation of cells and DNA, since DNA penetration into the
cytoplasm is associated with the electric pulses;
 in the case of CaCl2 mediated transformation, the TE is inversely related to the size
43
and form of plasmid DNA; whereas the electroefficiency is directly proportional
to the concentration of input DNA and is independent on the size and form of
DNA (Prasanna and Panda, 1997).
Heat treatments (pasteurization or ultra-high temperature are the predominantly
used techniques) are applied in order to extend the durability of foodstuffs in food
industry. However, the application of these techniques often causes undesirable side
effects, such as denaturation of proteins, destruction of vitamins and deterioration of the
taste of foods (Adams, 1991). To resolve this problem several researchers have already
developed a non-thermal process in which pulsed electric fields (PEF) with a range of 25100kV cm-1 are applied to kill spoilage microorganisms (Ohshima and Sato, 2004;
Schoenbach et al., 2000). In addition, PEF is effective for the preservation of foods
because of its potential to inactivate microorganisms without altering organoleptic and
nutritional properties of foods (Barbosa-Canovas et al., 1999). The phenomenon of
sterilization or inactivation by high intensity electric pulses is resulted in a certain
condition in which the total area of electropores appeared the cell membrane becomes
extremely large, causes irreversible permeabilization, and eventually kills the
microorganisms (Husheger et al., 1981; Wouters and Smelt, 1997).
However, the electrosterilization efficiency is dependent on the electrical field
strengths (a quantitative expression of the intensity of an electric field at a particular
location) and treatment time (Husheger et al., 1981), shape of the treatment chambers
(plate and neddle) and temperature of cell suspension (Ohshima et al., 1997) and growth
temperature of bacteria (Ohshima et al., 2002). Jayaram et al. (1992) reported that an
electric field strength of 25kV cm-1 and treatment time of 10ms at 60C resulted in a high
level of destruction (survival ratio of 10-9) of Lactobacillus brevis. The effects of PEF on
the inactivation of four different organisms (E. coli, Listeria innocua, Leuconostoc
mesenteroides and Saccharomyces cerevisiae) suspended in same medium showed that
the electroefficiency is dependent on the cell size, shape and cell wall composition
(Aronsson et al., 2001). The results showed that S. cerevisiae is the most sensitive
organism with a 6-log reduction, followed by E. coli with a 5.4-log reduction, whereas
only a 3-log reduction was obtained in both L. innocua and L. mesenteroides although all
the bacteria were exposed to a constant electrical condition (30kV cm-1 electric voltage,
4μs pulse duration and 20 pulses). In addition, while investigating the inactivation kinetics
44
of L. innocua, Wouters et al. (1999) observed that heat inactivation is less effective than
PEF although the treatment time and temperature were kept constant in both conditions.
Electrochemotherapy (ECT) has been established as an emerging drug delivery
method for the treatment of cancer in which a cytotoxic nonpermeant drug (for example,
Bleomycin) having high intrinsic cytotoxicity is delivered to the infected tumors by
applying of electric pulses (Belehradek et al., 1993; Mir, 2000; Mir et al., 1991). After
permeabilization of membrane, the nonpermeant molecules have a direct access into the
cytosol, and thereby infected tumors are killed (Gothelf et al., 2003; Mir et al., 1988; Mir
and Orlowski, 1999). Electroporation with micro- to millisecond duration and field
strengths of 100-1500V cm-1 generally enhance the delivery of certain chemotherapeutic
drugs by three to four orders of magnitude and DNA by several 100-fold (Rabussay et al.,
2003). The combination of Bleomycin (BLM) and pulse electric field increase the toxicity
of this anticancer drug by hundreds of thousands folds in vitro. The most significant
increase of toxicity has been observed with molecules, such as netropsin (200-fold) and
Bleomycin (700-fold) that do not strongly diffuse through the plasma membrane under
normal conditions (Orlowski et al., 1988). Okino and Mohri (1987) observed that 17%
decrease of initial mass of infected tumors after 4 days of electroporation using of a highvoltage electrical pulse (5kV cm-1, 2ms) with the administration of BLM. They also
observed that the tumor growth cannot be inhibited either by imposing of high-voltage
electric field or by administering of BLM alone. In general, the electric field strengths,
pulse widths and multiple pulses ranging from 1 to 5kV cm-1, µs to ms and 1 to 8,
respectively, are applied for ECT. However, the appropriate electric field needed for ECT
is dependent on the specific cells or tissues applied (Jaroszeski et al., 1997).
Apart from the application mentioned above, there are few reports available in the
literature that demonstrated the possibility of using this approach in bioprocess
intensification. Ohshima et al. (1995) observed an increase of invertase and alcohol
dehydrogenase (ADH) secretion in the supernatant of samples while pulsed at 6 and 12kV
cm-1, respectively. The same group also extracted nucleic acid molecules within 1min of
pulsation (Ohshima et al., 1999), and recovered the recombinant proteins (-glucosidase,
-amylase and cellobiohydrolase) from the recombinant strains of E. coli (Ohshima et al.,
2000). In addition, the specific activities of -amylase and cellobiohydrolase were
approximately 9 and 1.9-fold higher than that of ultrasonic treatment. Like bacteria, a
45
considerable release of cytoplasmic proteins of S. cerevisiae, such as glutathione
reductase (GLR), 3-phosphoglycerate kinase (PGK) and ADH was observed after 3-8h of
pulsation (Ganeva and Galutzov, 1999). They also extracted approximately 80-90% of
intracellular enzymes of S. cerevisiae i.e., hexokinase, PGK and glyceraldehyde-3phosphate dehydrogenase (GAPDH) by applying a series of electric field pulses (Ganeva
et al., 2003). Form the above findings, it is apparent that pulse electric field easily disrupts
the outer membrane of cells, and the extraction or secretion of specific protein or enzymes
from an organism is possible by imposing of high intensity electric pulses.
2.3.2. Theory and kinetic studies of electropermeabilization
Although electroporation or electropermeabilization has successfully been applied
in a range of purposes, the mechanisms or the molecular processes by which the
membrane permeability of cells increased are not fully understood. Electroporation is
established with the transient reversible permeabilization of cell membrane by imposing
an external electric field to living cells (Faurie et al., 2005; Hapala, 1997; Sowers, 1992;
Teissie et al., 1999; 2002; 2005; Tryfona and Bustard, 2006; Tsong, 1991; Weaver, 1995;
Weaver and Chizmadzhev, 1996). Permeabilization involves two distinct processes (i) the
introduction of transient inhomogenities in membrane structure and (ii) the generation of
passage in order to transfer or exchange molecules across the membrane through these
defects or electropores (Hapala, 1997). Due to the low conductivity of phospholipid
bilayers, application of external electric field (voltage) generates a potential difference
across the membrane, termed as transmembrane potential (TMP). TMP is the electrical
potential (the potential energy per unit of charge associated with an electric field,
expressed by voltage) difference across the plasma membrane of the cell.
Membrane permeabilization only occurs on the part of cell surface where the TMP
difference is equal to or higher than its critical rupturing value i.e., the intensity of electric
pulse applied to cell suspension must be higher than the rupturing value (Teissie et al.,
1999; Weaver and Chizmadzhev, 1996). The degree of permeabilization is dependent on
the TMP (Teissie et al., 1999). In the case of a lipid bilayer, the TMP value is about
200mV (Teissie and Rols, 1993), whereas the threshold TMP for dielectric breakdown of
NG108-15 cells (a hybridomal cell line of a rat neuroblastoma and a mouse glioma) is
250mV (Ryttsen et al., 2000). TMP is not evenly distributed over the cell surface since
the highest value is obtained at the sites of plasma membrane closest to the electrode,
46
whereas the lowest value of TMP is observed at the sites that are far from the electrode
(Ho and Mittal, 1996; Pavlin et al., 2002; Valic et al., 2003). It has been reported that the
induced TMP difference brings major changes on the phospholipid bilayer, such as
rupture of cell membrane, alteration of membrane proteins and increase of membrane
permeability (Zimmermann et al., 1974; Zimmermann, 1982). Hence, it is obvious that
the most important factor responsible for electroporation is the induced transmembrane
potential difference (), describing by the following equation
() = F g rcell Ee cos
Where, F is the shape of the cell (in case of spherical cell, F =1.5);
g is the conductivity (cell membrane is considered as a pure dielectric, g =1);
rcell is the radius of cell;
Ee is the applied field strength; and
 is the angle between the site on the cell membrane where  is measured
and the direction of Ee
Electropermeabilization assay by propidium iodide (PI) penetration into Chinese
hamster ovary (CHO) cells showed that permeabilization mainly occurs at the sites of cell
membrane facing the two electrodes (Golzio et al., 2002). In addition, Gabriel and Teissie
(1997) demonstrated that permeabilization in CHO cells is first observed on the side of
cell facing the anode that is more permeabilized than the other. Tekle et al. (1994)
demonstrated that pores with smaller size (but greater in number) are created on the
membrane facing the anode, whereas larger pores (with a lower population) appear on the
membrane facing the cathode. Since the induced TMP difference is also proportional to
the cell radius, it is evident that the threshold value of electric field varies with cell size.
i.e., larger cells are more sensitive to lower electric field strength than smaller ones
(Teissie et al., 1999). The induced TMP difference also depends on the density of cells,
arrangement and position of cells in electroporation chamber (Canatella et al., 2001,
Pavlin et al., 2002). However, electroporation is a multi-step process in which formation
of transient pores, their expansion, exchange of molecules across the cell membrane and
resealing of pores are observed. The first three events occur during the pulse, whereas
pore resealing takes place after the pulsation (Kinosita and Tsong, 1977; Rols and Teissie,
1990b; 1998). However, the mentioned steps are prone to vary depending on the physical
and electrical conditions supplied during pulsation (Ho and Mittal, 1996). Teissie et al.
47
(1999;
2005)
and
Faurie
et
al.
(2005)
described
the
kinetic
studies
of
electropermeabilization in four steps (Table 2.4).
Table 2.4 Steps of cell electropermeabilization due to electroporation
Induction step
the applied electric field induces the membrane potential difference
that leads to local defects when transmembrane potential difference
exceeds the threshold value (about 200mV). However, in the case
of mammalian cells electropores appear at a critical rupturing value
of 0.2-1.5V;
Expansion step
the size or diameter of pores that are created due to the electric
pulses starts to expand until a certain voltage is placed on. The
expansion of pores (20 to 120nm in diameter) takes place within
20ms after the pulse is given;
Stabilization step
when the electric field intensity is decreased below the threshold
value, a stabilization state is observed within a few ms where the
cell membrane allows small molecules to be permeated or
intracellular products to be secreted, and finally
Resealing step
a slow resealing process of cell membrane is usually observed after
removing the electric pulse. The membrane conductance is
significantly decreased within a few ms, and the radii of
electropores start to reduce. The number of pores is reduced
significantly, and a complete disappearance of pores occurs soon
after.
Rols and Teissie (1990a) demonstrated that the application of electric pulses to a
cell membrane reorganizes lipid molecules, induces leakage and forms electropores
within a few ms of pulsation. The
31
P NMR analysis of pulsed membrane (CHO cells)
revealed that a reorientation of polar heads leads to altering the organization of
phospholipids (Lopez et al., 1988). The freeze-fracture electron microscopy showed that
electropores appeared within 3 milliseconds of pulsation (Chang et al., 1992). Although
electropores in smaller size are formed just after the imposing of pulses, only a few of
them expand locally to create passages for macromolecules to pass through the membrane
(Tsong, 1991). The expansion of pores is only controlled by the mechanical parameters of
cell membrane, such as surface tension or viscosity. Wilhelm et al. (1993) demonstrated
48
that increase in field strength increases the number of electropores on the lipid bilayer
membrane but does not influence the kinetics of pore opening. Furthermore, Ryttsen et al.
(2000) investigated the kinetics of pores formation, opening and closing due to pulses by
patch-clamp and fluorescence microscopy. Their results showed that electropores do not
allow a passage for the entry of fluorescein into NG108-15 cells until the electric field
reaches its threshold value.
However, the electron microscopic analysis of an electropermeabilized cell
membrane showed prominent structural changes with the formation of many transient
pores (Teissie et al., 1999). These pores provide a way of transferring target molecules,
ions and water from one side of cell membrane to the other (Figure 2.8). It has also been
demonstrated that the cell-impermeant solutes added in the extracellular medium can
easily enter into the cell interior by diffusion during the pore-opening time (Weaver,
1995). However, if the total area of electropores is small in relation to the total surface
area of membrane, the electropores resealed automatically after removing the external
electric field to ensure the survival of electrically stimulated recipient cells (Sowers, 1992;
Tsong, 1991; Weaver, 1995). This phenomenon is observed in the case of reversible
membrane permeabilization. The electron microscopy of electroinduced permeabilized
membrane of CHO showed that numerous microvilli and blebs are formed just after the
Figure 2.8 A schematic diagram of theoretical cell membrane before and after electroporation
(Dr. Bustard, private communication, 2004).
49
application of electric pulses, however, these effects are reversible and disappeared within
30min at 37°C (Escande-Geraud et al., 1988).
It has been reported that the field-treated cells recover their original membrane
impermeability within few minutes or hours depending on the electroporation and postelectroporation conditions (Chang et al., 1992; Prasanna and Panda, 1997; Teissie et al.,
1999). The reversible behaviour of biological membrane alteration is strongly controlled
by the post pulse temperature (Golzio et al., 1998). The induced permeabilization state of
CHO cells has been shown to remain in a permeable state without loss of viability for
several hours at 4C (Lopez et al., 1988), whereas cells were observed to be resealed in
less than 5min at 37°C (Golzio et al., 2004; Rols et al., 1994). However, a fast resealing
process increases the viability of cells and transfection yield (Rols et al., 1994). The
stability of the permeabilized state is also influenced by the nature of the membrane. Pure
phospholipids bilayer membrane has been shown to have very short-lived permeability
(Teissie and Tsong, 1981), whereas biological cell membranes can be maintained
permeable for longer periods i.e., from seconds to hours depending on the post pulse
conditions (Kinosita and Tsong, 1977).
Irreversible membrane breakdown often occurs due to the application of nonoptimal electrical conditions to the cell suspension. If huge numbers of electropores are
formed or the diameters of individual pores are enlarged enough due to the application of
high field strength, the membrane is no longer able to repair these perturbations (Teissie
et al., 1999; Weaver and Chizmadzhev, 1996). Two common phenomena are mainly
observed during irreversible electroporation i.e., creation of permanent holes on the
membrane due to the expansion of electropores, and cell lysis as a result of chemical
imbalances caused by molecular transport through the transient pores. During irreversible
electropermeabilization, essential cytoplasmic components have been observed to leak out
from the cells. Nevertheless, no dramatic rupture is monitored if the electrical parameters
applied during electroporation are inappropriate (Sowers, 1992; Weaver and
Chizmadzhev, 1996). Moreover, membrane fusion among the pulsed cells is often
observed after electropermeabilization. During electropermeabilization, cells loose the
repulsive forces that prevent membranes of two cells fusing spontaneously (Rols and
Teissie, 1989; Sowers, 1986; Teissie and Rols, 1986). Figure 2.9 shows a schematic
diagram of the effects that occurred during and after electroporation.
50
Figure 2.9 Exposure of a cell to an electric field may result either in permeabilization of cell
membrane or its destruction (Puc et al., 2004).
2.3.3. Factors affecting cell electropermeabilization
The efficiency of a successful electroporation is dependent on many critical
factors and parameters that influence the transmembrane transport of molecules into cells
as well as the yield of transformants or electroporants. Table 2.5 presents a number of
factors that are required to investigate during establishing an electroporation or
electropermeabilization process.
Table 2.5 Critical factors and parameters that influence electroporation (Faurie et al., 2005;
Prasanna and Panda, (1997)
Type
Cellular factors
Factors
growth phase of cell, cell density, cell diameter and cell
wall rigidity;
Physiochemical factors
medium
conductivity,
ionic
concentration
of
the
electroporation buffer, osmolarity, post-electroporation
incubation
conditions,
temperature,
pH
and
DNA
concentration;
Electrical parameters
optimal field strength, critical voltage, pulse length, number
of repetitive pulses and different electrodes geometry.
51
2.3.3.1. Cellular factors
The efficiency of transformation is dependent on the growth stage of a microbial
culture used for electroporation. Several experiments showed that electro-TE is
significantly higher with cells grown at early or mid exponential or log phase than that of
cells harvested for electroporation (Bonamy et al., 1990; Calvin and Hanawalt, 1988;
Dunican and Shivnan, 1989; Miller et al., 1988). When E. coli cells harvested from early
exponential growth phase were pulsed, approximately 3-fold increase in TE was reported
as compared with the cells harvested from late exponential phase (Calvin and Hanawalt,
1988). The highest yield of transformants (107) was obtained with Corynebacterium cells
grown at the mid-exponential phase, and the efficiency of transformation may be varied
from 100 to 1000-fold due to the difference (2-3h) of culture age of C. glutamicum
(Bonamy et al., 1990). In another study, PEF treatment with an electric field strength of
1.5V mm-1 and an energy input of 95J ml-1 resulted in 23% permeabilized cells when the
cells were in the stationary growth phase, whereas an energy input of only 24J ml-1 was
required to obtain a similar level of membrane permeabilization with cells in the
exponential growth phase (Wouters et al., 2001). Like the growth stage, the yield of
transformants is also dependent on the density of cells. The TE of C. glutamicum has been
shown to increase up to cell density of 1010 cells ml-1 (Bonamy et al., 1990; Liebl et al.,
1989), and reduce linearly with a concentration of below 1010 cells ml-1 (Dunican and
Shivnan, 1989). Pucihar et al. (2007) determined a decrease in permeabilized CHO cells
(approximately 50%) due to increase of cell density from 10 × 106 to 400 × 106 cells ml-1
although the electroporation conditions were kept constant. Furthermore, Delorme (1989)
demonstrated that the electro-TE of S. cerevisiae with an OD600 of 11.0 is approximately 4
to 5-fold lower than the OD600 of 0.3-1.0.
The success of electroporation is also dependent on the type of organism used for
transformation. Since the induced TMP difference is proportional to the radius of cell, it
has been reported that the threshold value of electric field for membrane permeabilization
is varied with the size of cell (Teissie et al., 1999). In addition, the magnitude of external
electric field required to generate a TMP sufficient for pore formation is inversely
proportional to the size and shape of cells (Miller et al., 1988). This means that the field
strengths required for small and spherical shaped cells are considerably higher than those
for larger and rod shaped cells (Prasanna and Panda, 1997; Wouters et al., 2001). It has
also been reported that the electropermeabilization of eukaryotic cells requires lower
52
electric field strength than prokaryotic cells (Miller et al., 1988; Prasanna and Panda,
1997). Bonamy et al. (1990) reported that the electrical and other factors are more
stringent for efficient transformation of Corynebacterium species than for other strains
i.e., E. coli and C. jejuni. Prasanna and Panda (1997) reported that the Gram-positive
bacteria as a group are somewhat less efficient in transformation by electroporation than
are Gram-negative species. However, increasing the electric field intensity may overcome
this barrier to a certain degree, and hence improve electro-TE of Gram-positive bacteria.
Several researchers observed that the voltage required for pore formation and the
efficiency of transformation are varied from species to species. The intact cells of B.
japonicum were transformed with a TE of 106 to 107 transformants/g of DNA in
presence of field strengths of 10.5 to 12.5kV cm-1. Wouters et al. (2001) showed that L.
fermentum is more resistant and less permeabilized than L. plantarum although the
electroporation conditions were kept constant throughout the study. These results
indicated that the effect of electroporation is not only dependent on the species but also on
the genus levels. The level of leakage of UV-absorbing substances from Salmonella
typhimurium after pulsation is approximately 4-fold higher than that of Listeria
monocytogenes although the electrical conditions were kept constant (Simpson et al.,
1999). Moreover, it has been shown that induced TMP difference also depends on cell
angle in relation to the direction of electric field applied (Pavlin et al., 2002). However,
cellular factors (especially growth phase, cell density and type of organism) in
comparison to the electrical factors have relatively minor influence on the efficiency of
transformation (Miller et al., 1988).
2.3.3.2. Physiochemical factors
The electroporation buffer in which the targeted cells are suspended has the major
influence on both TE and cell viability (Muller et al., 2001). The conductivity of
electroporation medium that is often increased due to electric pulses have an effect on
pulse duration and expansion of transient pores (Rols and Teissie, 1989), which play a
vital role in transferring the molecules across the cell membrane. The ionic composition
of the pulsing buffer determines its specific resistivity, and hence the RC time constant
(resistance of the cell suspension and the capacitance of the capacitor) of electric pulse.
V = V0exp(-t/RC)
53
=RC
Where,
V is the voltage across the pulsing chamber;
V0 is the initial voltage;
T is the time after the pulse starts;
R is the resistance of the suspension;
C is the capacitance of the capacitor; and
 is the time constant
McIntyre and Harlander (1989b) showed that L. lactis cells suspended in
deionised distilled water (lowest conductivity) were more rapidly electroporated than the
cells suspended in different pulsing medium, such as EPB (0.5M sucrose, 1mM MgCl2,
7mM K2HPO4-KH2PO4 at pH 7.4), EPM (0.3M raffinose, 1mM MgCl2.6H2O, 5mM
K2HPO4-KH2PO4 at pH 7.4) or EB (222mM sucrose, 1mM MgCl2, 7mM N-2hydroxyethylpiperazine-N-2-ethanesulfonic acid at pH 7.4), respectively. The medium
conductivities of EPB, EPM and EB were 2.04, 1.43 and 0.5mS cm-1, respectively,
whereas it was only 0.07mS cm-1 in the case of deionised distilled H2O. Moreover, the
highest TE was obtained when the least conductive suspending solution (deionised
distilled H20) was used. Furthermore, the pulse duration applied during their study was
shown to vary with the medium conductivities although cells were treated for a constant
time period (5ms). Use of the highest ionic strength and most conductive solutions (EPB
and EPM) resulted in reduced pulse duration (0.60ms for EPB-suspended cells and
4.02ms for EPM-suspended cells) and transformation efficiencies (McIntyre and
Harlander, 1989b).
Similarly, Djuzenova et al. (1996) observed that the incorporation of PI into the
reversibly permeabilized cells (murine myeloma) increased significantly with the
decreasing of medium conductivity. The electropermeabilization of mouse myeloma cells
by single square-wave electric pulse (150kV cm-1 and 10-100ns) also showed that a
significant increase of PI uptake occurred with the decreasing of medium conductivity
(Muller et al., 2001). Like mammalian cells, the number of permeabilized cells (L.
plantarum) was observed to be increased due to the decrease of medium conductivity
from 1.5 to 0.4S m-1 although a constant energy input (2.5V µm-1) was supplied
throughout the study (Wouters et al., 2001). Furthermore, the electropermeabilization of
54
S. cerevisiae has been investigated by suspending cells in media containing different salts,
such as NaCl, KCl, MgCl2, CaCl2 or MgSO4 (Muraji et al., 1998). Although the
conductivities of pulsing media were adjusted to 15.4mS cm-1 by varying concentration of
salts and a constant energy input was supplied throughout the study, the yield of
electropermeabilized cells measured in the medium containing NaCl was significantly
higher than that of media containing other salt solutions, especially CaCl2. The results
demonstrated that extracellular ions affect the functioning of membrane electropores by
interacting with the cell membrane, and confirmed that the ionic content of the
extracellular media has an important influence on the yield of electropermeated cells
(Muraji et al., 1998).
Several studies have already established that osmotic pressure has a major impact
on electroporation. Rols and Teissie (1990a) investigated the effects of osmotic pressure
(due to pulsing buffer) on the electropermeabilization of CHO cells. The results showed
that osmotic pressure has no effect on the induction step of permeabilization, whereas the
expansion and resealing steps were inhibited due to the increase of osmotic pressure of
pulsing buffer. During voltage-induced membrane permeabilization and gene transfer in
CHO cells, Golzio et al. (1998) showed that the size of CHO cells (12.2µm) increased
with the decrease in medium osmolarity, whereas the size of cells decreased with an
increase of medium osmolarity. The membrane permeabilization of osmotically shocked
CHO cells has been shown to occur at low TMP as compared to the cells electroporated
under normal condition (Barrau et al., 2004). Suga et al. (2003) showed approximately 5fold increase in electro-TE (6.8 ± 1.7 × 106) while S. pombe cells pre-incubated with
hyperosmotic solution (2M sorbiol/1.5M NaCl at 30C for 60min) were pulsed at 11kV
cm-1, 25μF and 200Ω.
It has been demonstrated that the temperature generated during electroporation of
samples has an important influence on the efficiency of transformation (Miller et al.,
1988). While keeping the electric field strength constant at 5.25kV cm-1, samples (cell
suspension of Campylobacter jejuni) pre-incubated at 22C for 10min resulted in a TE of
5.0  0.9 × 103 transformants/g DNA, whereas it was 3.3  1.9 × 105 transformants/g
DNA in the case of samples pre-incubated at 4C for 10min. Rols et al. (1994) also
showed that the electroporated samples with a pre-incubation at 4C and post-incubation
at 37C enhanced the transfection efficiency of mammalian cells. They also mentioned
55
that pre-incubation of cells at 4C with plasmid DNA increased the interfacial
concentration of plasmid at the cell membrane and inhibited DNA degradation by
extracellular nucleases. The TE of slow-growing species (M. tuberculosis, M. bovis and
M. intracellulure) increased by several orders of magnitude due to the increase of
temperature to 37C. In contrast, the TE of M. smegmatis (a fast-growing species) was
higher at 0°C, and decreased with the increase of temperature (Wards and Collins, 1996).
Furthermore, Gallo et al. (2002) investigated the effects of temperature on the
electropermeabilization of stratum corneum (the outermost layer of porcine skin). The
results showed that the stratum corneum is susceptible to permeabilization at high
temperature, and cells pulsed at high temperature needed a prolonged time to recover their
original structure. However, this detrimental effect due to Joule heating could be
minimized by conducting the pulse experiments at low temperature (Rols et al., 1994;
Teissie et al., 1999). Moreover, if the cells suspended in deionized water are pulsed and
the parameters, such as cell concentration, field intensity, number and duration of pulses
are properly optimized, this injurious effect can be overcome (Ganeva et al., 2003).
Although several authors demonstrated that the pH of the pulsing medium does
not affect the inactivation of microbial cells by PEF (Husheger et al., 1981), it was
recently found that pH has also a major influence on electroporation efficiency. VegaMercado et al. (1996) reported that E. coli is more PEF-sensitive under acidic conditions,
whereas Alvarez et al. (2000) demonstrated that Salmonella senftenberg is more resistant
at acidic pH. The electric field and ionic strength are more likely related to the poration
rate and physical damage of the cell membranes, while pH is more likely related to
changes in the cytoplasmic conditions due to the osmotic imbalance caused by the
poration (Vega-Mercado et al., 1996). DNA concentration is also reported to be a limiting
factor for obtaining an optimal electro-TE. Dunican and Shivnan (1989) demonstrated that
there is a saturation level above which increasing DNA level decreases the efficiency of
transformation. In their experiments, the optimum DNA concentration was observed to be
1ng while transforming plasmid DNA into C. glutamicum. A similar observation is also
mentioned by Delorme (1989) during the electrotransformation of S. cerevisiae. The yield
of transformants (107 transformants per µg DNA) obtained through the use up to 1-2g of
DNA was shown to be higher than that achieved by means of high concentration of DNA
(more than 4g) although the electric parameters were identical in both conditions
56
(Bonamy et al., 1990). The result suggests that the TE could be improved by lowering the
amount of plasmid DNA during electroporation.
To increase the efficiency of electroporation, several compounds (penicillin G,
ampicillin, isonicotinic acid or Tween 80) were also added into the growth medium in
order to make the cells more permeable for the exogenous DNA (Bonnassie et al., 1990;
Haynes and Britz, 1990). Haynes and Britz (1989) showed an increase of TE (up to 4 ×
107 trasformants/μg of DNA) due to the addition of glycine plus Tween 80 into the
electroporation medium of Brevibacterium lactofermentum and C. glutamicum. The same
group later obtained the TE of 5 × 105 transformants/μg of homologous DNA and 2 × 103
transformants/μg of E. coli DNA due to the supplementation of glycine and isonicotinic
acid hydrazide (INH), respectively, into the culture medium (Haynes and Britz, 1990).
These results might have occurred due to the fact that glycine incorporates into the cell
wall and leads to a less tightly cross-linked structure, while INH changes the mycolic acid
composition. Similarly, it has been demonstrated that addition of Tween 80 altered the
degree of unsaturation of the side chains and the overall composition of corynemycolic
acids (Chevalier et al., 1988).
It has been reported that penicillin and other -lactam antibiotics alter the
membrane structure of Corynebacteria, and cause the cellular excretion of Nacetylglucosamine derivatives and phospholipids, regardless of the carbon sources used
for cultivation (Kikuchi and Nakao, 1986). Kurusu and his co-workers showed
approximately 2-fold increase in TE by supplying C. glutamicum cells with 1U ml-1
penicillin G into the pulsing medium prior to electroporation (Kurusu et al., 1990). Pretreatment of cells with ampicillin (a -lactam antibiotic) has also been reported to increase
the electro-TE of B. lactofermentum (Bonnassie et al., 1990). The result showed that
ampicillin (0.5-1.5µg ml-1) partially disorganised the cell wall structure of this bacterium
and increased the TE, however, no transformant was observed without imposing the
electric pulse. In addition, the TE of C. glutamicum has been increased by allowing an
additional freeze/thaw cycle and treating the cells with lysozyme prior to electroporation
(25-30kV cm-1). The optimal yields of transformants per µg of plasmid DNA were 3.0 ×
103 for intact, 2 × 104 for freeze/thawed and 7 × 104 for lysozyme treated cells,
respectively (Wolf et al., 1989). However, careful choice of permeabilizing agents and
adjusting the experimental conditions are the most important factors while increasing the
57
membrane permeability is conducted by the use of these chemicals into the
electroporation medium in which cells are suspended.
2.3.3.3. Electrical parameters
Among all the factors discussed above, electrical parameters (mainly electric field
strength) have a major influence on the efficiency of electroporation. The electric field
strength, pulse duration, pulse repetition frequency, number of pulses and pulse shape
determine whether electropermeabilization of cell membrane is reversible or irreversible.
It is often observed that cells are irreversibly permeabilized and lose their viability if these
parameters exceed a certain limit i.e., amplitude of pulses is too high or duration of pulses
is too long. Several groups have already investigated the effects of electric field strength/
pulse amplitudes on the TE of different stains. Table 2.6 presents a list of transformation
experiments that have been effectively carried out by controlling the electrical parameters.
The TE of C. jejuni was observed to be increased with an exponential manner in
the range of field strengths 5 to 13kV cm-1, and the frequency of transformants was
approximately doubled in every 1kV cm-1. Furthermore, an increase (2-fold) in voltage
results in more than 1000-fold increase of TE (Miller et al., 1988). However, no
transformants were obtained at 3kV cm-1 while electrotransfomation was carried out with
B. japonicum (Hattermann and Stacey, 1990). The number of transformants per µg DNA
has been shown to be increased from 2300 to 4500 due to the increase of voltage from
800V to 900V, however, pulsing at 1000V sharply reduced the number of transformants
to 1630 during the electrotransformation of DNA into S. cerevisiae (Delorme, 1989).
Dunican and Shivnan (1989) demonstrated that high voltage is required in order to
generate electropores on the cell wall of C. glutamicum since an electric field strength of
6.25kV cm-1 was not shown to be sufficient to improve the uptake of DNA into
protoplast. Bonamy et al. (1990) showed that the uptake of plasmid DNA in
Corynebacteria only occurs at field strengths 5kV cm-1 and pulse duration of 0-20ms.
The number of transformants also increased with the pulse duration that has been
successfully investigated with different type of strains i.e., bacteria (Eynard et al., 1992),
yeast (Meilhoc et al., 1990) and mammalian cells (Wolf et al., 1994). In the case of E.
coli, no transformation was observed with pulse duration shorter than the millisecond
(ms) range although the membrane permeabilization occurred with microsecond (µs)
58
Table 2.6 A list of transformation experiments carried out with the different types of strains by varying the electrical conditions of electroporation
Strains
Cell density
Field strength
Pulse duration
Yield of
(cells/ml)
(kV cm-1)
(ms)
transformants
N/M
12.5
7ms
1.0 × 107
Hattermann and Stacey, 1990
Brevibacterium flavum
1.0 × 1010
12.5
N/M
5.0 × 104
Satoh et al., 1990
C. jejuni
5.0 × 109
13.0
2ms
1.2  0.9 × 106
Miller et al., 1988
Clostridium cellulolyticum
1.0 × 1011
7.5
5ms
1.0 × 107
Tardif et al., 2001
C. glutamicum
3.0 × 108
35.0
500µs
3.0 × 103
Wolf et al., 1989
C. glutamicum
7.0 × 1010
12.5
N/M
4.0 × 107
Liebl et al., 1989
C. glutamicum
1.0 × 1010
12.5
5ms
1.0 × 107
Dunican and Shivnan, 1989;
Bradyrhizobium japonicum
References
Bonamy et al., 1990
E. coli K-12
2.2 × 109
13.0
N/M
4.5 × 109
E. coli
2.5 × 1010
13.0
5ms
1.6  0.1 × 109
L. casei
2.0 × 108
5.0
N/M
8.5 × 104
Chassy and Flickinger, 1987
L. lactis
5.0 × 1010
17.0
5ms
1.0 × 103
McIntyre and Harlander, 1989a
Staphylococcus aureus
3.0 × 1010
23.0
2.5ms
4.0 × 103
Schenk and Laddaga, 1992
N/M: Not mentioned
59
Calvin and Hanawalt, 1988
Dower et al., 1988
pulses (Dower et al., 1988). However, pulse duration with high electric voltage often kills
the microorganisms, and eventually decreases the TE. Miller et al. (1988) observed that
an increase of pulse duration from 10 to 20ms decreased the TE by a factor of 100.
Hattermann and Stacey (1990) also observed that increasing the pulse duration from 6.6
to 30.5ms with a constant field strength of 12.5kV cm-1 resulted in a decrease (200-fold)
in TE. Miller et al. (1988) observed that pulse amplitude and duration have compensatory
effects on the efficiency of transformation. An efficiency of 5.0 x 105 transformants of
C. jejuni/g of DNA was obtained with a field strength of 5.25kV cm-1 and pulse
duration of 21ms or a field strength of 10.3kV cm-1 and a duration of 2.4ms. A similar
phenomenon was also observed where the exponential decay pulses of either 7kV cm-1
and 20ms or 12.5kV cm-1 and 5ms yielded the same level of transformants (109 to 1010)
while plasmid DNA was transformed into E. coli by high-voltage electroporation (Dower
et al., 1988). Nevertheless, Jayaram et al. (1992) demonstrated that the application of
high electric voltage is more effective for the destruction of L. brevis cells than is
obtained due to the increase of treatment time or pulse duration.
Pulse type has also a significant influence on the efficiency of electroporation.
Two types of electric pulses i.e., exponential decay pulses, EDPs (Neumann et al., 1982)
and square wave pulses, SWPs (Rols and Tessie, 1990b) are frequently used in order to
permeabilize cells. The main difference between these two types of pulses is that the
voltage applied to cell suspension remains constant during pulsation (in the case of
SWPs), whereas voltage that discharged into the cell suspension decreases over time (in
the case of EDPs). In general, exponential decay pulses are used for the electroporation of
bacteria, yeast and other microorganisms, where a high voltage is applied to a cell sample
suspended in a small volume of high resistance media (Neumann et al., 1982). On the
other hand, SWPs, applied for the electropermeabilization mammalian cells, resulted in
high survivability of permeabilized cells that could not be obtained by EDPs (Takahashi
et al., 1991). It is now apparent that several factors or parameters are required to consider
for the the success of electroporation, and hence it is necessary to investigate all the
factors while conducting electrotrasformation or applying this approach in chemotherapy
or improving the bioprocesses through electropermeabilization.
60
Chapter THREE
3. Production of L-glutamate by Fermentative
Cultivation of Corynebacteria under Different
Growth Conditions
3.1. Introduction
In 1957, Kinoshita and his co-workers first observed that C. glutamicum (initially
reported as Micrococcus glutamicus) is able to produce L-glutamate under biotin limited
condition (Kinoshita et al., 1957). The same group also constructed a homoserineauxotrophic mutant of C. glutamicum that was observed to produce a large amount of Llysine by fermentation (Nakayama et al., 1961). Since then, a number of amino acids i.e.,
L-lysine, L-glutamate, L-threonine and L-isoleucine have been produced by microbial
fermentation (Hermann, 2003; Ikeda, 2002). Under specific culture conditions, however,
Corynebacteria convert almost 50% of the supplied carbohydrate to L-glutamate with the
formation of insignificant amount of by-products, and the maximal production rate can
reach up to 100g l-1 (Kinoshita et al., 1985). The maximal specific growth rate of C.
glutamicum in CGXII Minimal Medium was reported to 0.33h-1. After 30h of
fermentation on CGXII Medium supplemented with 220mM glucose, this fast-growing
bacterium was observed to reach an OD600 of 53.0 (equivalent to approximately 14g Dw
l-1) (Keilhauer et al., 1993).
However, it is well investigated that the cell wall of C. glutamicum consists of
three different layers i.e., cytoplasmic membrane, peptidoglycan layer and mycolate layer
that result in low cell wall permeability (Hirose et al., 1985; Kramer, 1994). Jarlier and
Nikaido (1990; 1994) demonstrated that the cell wall permeability of M. chelonae to
cephalosporins is about three orders of magnitude lower than that of E. coli, and ten times
lower than the permeability of P. aeruginosa. Because of having low membrane
61
permeability, wild strains of C. glutamicum excrete a limited amount of L-glutamate
under normal growth condition, and hence the product yield is often inadequate (Hirose
et al., 1985). Furthermore, a major rate-limiting step is observed not only in the amino
acid secretion but also in glucose and nutrient uptake due to the presence of rigid cell
wall (Kramer, 1994).
Glutamic acid is produced by C. glutamcum grown under sufficient amounts of
glucose and nitrogen supply where the growth of cells is inhibited due to the lack of an
essential compound responsible for fatty acid synthesis (Gutmann et al., 1992). The
secretion of L-glutamate produced in C. glutamicum has been occurred by various
treatments i.e., biotin limitation (Shiio et al., 1962); surfactant addition (Duperray et al.,
1992; Takinami et al., 1965; 1968); addition of beta-lactam antibiotic, penicillin (Demain
and Birnbaum, 1968; Ikeda et al., 1972); ethambutol addition (Radmacher et al., 2005)
and addition of oleic acid or glycerol (Kanzaki et al., 1967; Okazaki et al., 1967). It has
been well investigated that all of these treatments affect the cell wall by either limiting
the synthesis of phospholipids compounds (auxotrophic for biotin or glycerol or fatty
acid) and membrane components (penicillin or ethambutol) or directly influencing the
membrane state (addition of amine surfactants) (Eggeling and Sahm, 2001; Kramer,
2004). Hoischen and Kramer (1989) mentioned that L-glutatmate secretion is an energy
dependent process, and is regulated by the metabolic status of cells. Gebhardt et al.
(2007) demonstrated that the lack of mycolate is sufficient to induce L-glutamate
secretion since Corynebacterial cells with a reduction in mycolate content were observed
to show higher permeability for the uptake of substrates or the secretion of intracellular
products into the extracellular medium. However, the appropriate concentration of those
agents and the time (growth point) at which they are added into the fermentation medium
have a major influence on L-glutamate production (Takinami et al., 1965; 1968).
Biotin (vitamin H) is one of the most fascinating cofactors involved in central
metabolic pathways of pro-and eukaryotic organisms (Streit and Entcheva, 2003). It is the
most representative growth promoting substance generally used as a vitamin supplement
in fermentation medium. In addition, biotin, the cofactor of acyl-CoA carboxylase, plays
62
a major role in fatty acid biosynthesis (Kramer, 1994). The L-glutamate production is
remarkably increased when C. glutamicum is cultivated in presence of an optimum
amount of biotin. The idea of cultivating this organism under biotin limitation is to
provide biotin as a limiting factor in which the rate of fatty acid biosynthesis is decreased,
and the composition of plasma membrane is altered (Eggeling and Sahm, 2001; Kramer,
1996). Hoischen and Kramer (1990) demonstrated that the deficiency of biotin results in
the changes in phospholipids and fatty acids composition, decreases of phospholipids
content, and ultimately increases the membrane permeability to L-glutamate. It is
therefore always a challenge to determine the optimum concentration of biotin required
for the highest production of L-glutamate. However, the literature showed that the
optimum range of biotin (0.5 to 2.5g l-1) is dependent on the concentration of
carbohydrate (10 to 15%) (Kramer, 1996; 2004). The production of L-glutamate
measured in B. flavum at a biotin concentration of 3µg l-1 was 122µM ml-1, whereas it
was 105, 22 and 0µM ml-1 at 6, 15 and 30µg l-1 of biotin, respectively (Shiio et al., 1962).
The biotin concentration in laboratory based fermentation media where glucose is
used as a carbon source can easily be controlled, however, it is very difficult to optimize
the biotin level during industrial fermentation since molasses (feedstock that is generally
used in biotech industry) often contains high levels of biotin (Kramer, 1996; 2004). In
order to make the use of these feedstocks, addition of surfactants is generally applied for
the production of L-glutamate (Hermann, 2003; Ikeda, 2002). It has been demonstrated
that polyoxyethylene sorbitane monopalmitate (Tween 40) and polyoxyethylene sorbitane
monostearate (Tween 60) are the most effective fatty acid derivatives for L-glutamate
production, however, monolaurate and monoleate esters (Tween 20 and Tween 80,
respectively) were observed to be unsuccessful (Shiio et al., 1963). In practical exercises,
about 100mM of L-glutamate can be achieved from 220mM glucose by adding 1.5%
Tween 60 into the growing culture of C. glutamicum (Eggeling et al., 2001). In addition,
L-glutamate production in a biotin rich condition is dependent on the concentration of
surfactants and the growth stage at which they are introduced into the culture medium
(Takinami et al., 1965; 1968). It has been observed that the early stages of exponential
63
growth is the best suitable time for the addition of surfactant into the fermentation
medium in order to obtain the highest production of L-glutamate (Naji et al., 2000).
Marquet et al. (1986) demonstrated that addition of surfactants at the exponential
phase of C. glutamicum growth results in decrease of cells volume and accumulates Lglutamate (100g l-1) in the extracellular medium. Naji et al. (2000) showed that the
addition of surfactants, such as polyoxyethylene glycol (POEFE), polyoxypropylene
glycol (POPFE) and polyoxyethylene–polyoxypropylene glycol (POAFE) influences not
only the L-glutamate secretion but also affects the growth and respiration of C.
glutamicum. Although POAFE and POPFE showed a tiny effect on the respiration of
cells grown at the exponential growth phase, a significant decrease in respiration (28.5
and 75%, respectively) was observed in cells harvested at the stationary growth phase.
However, the sterilization of surfactants is too complicated, and hence few alternative
treatments have been conducted, such as addition of penicillin or tetracaine or ethambutol
or oleic acid or glycerol in order to induce L-glutamate secretion in a biotin rich
condition.
Ethambutol (EMB) is widely used as anti-mycobacterial agent. The addition of
EMB disorders the cell envelope and alters lipid composition of the plasma membrane of
C. glutamicum that represents a permeability barrier. The critical target of EMB is to
inhibit the pathway responsible for the biosynthesis of cell wall arabinogalactan. EMB
causes less arabinan deposition in cell wall arabinogalactan, and reduces the content of
cell wall bound mycolic acids (Radmacher et al., 2005). At a concentration of 10mg l-1,
EMB reduced the specific growth rate and biomass yield of C. glutamicum. After 32h of
fermentation, OD600 in control (without ethambutol) was 53, whereas it was
approximately 18 at 10mg l-1 of EMB. However, the growth of this bacterium over a
wide range of EMB concentrations (up to 500mg l-1) was almost identical. It has also
been demonstrated that the addition of this anti-mycobacterial agent to the growing
cultures of C. glutamicum caused L-glutamate efflux at rates of up to 15nmol min–1 (mg
Dw)–1, whereas no efflux was observed in the absence of EMB (Radmacher et al., 2005).
64
It is now obvious from the literature that L-glutamate efflux involves an
interaction of several cell wall components i.e., cytoplasmic membrane, peptidoglycan,
mycolic acid layer and exporter, and the extent L-glutamate secretion is dependent on the
treatments mentioned above. For example, the addition of penicillin led to the
accumulation of only one third of L-glutamate produced by biotin limited condition
(Kimura et al., 1999). On the other hand, Takinami et al. (1965) obtained the maximum
L-glutamate yield (0.45g g-1) both in a biotin limited (3µg l-1) condition and by adding
Tween 60 (1mg ml-1) into a biotin-sufficient medium. Moreover, C. glutamicum is found
to have a number of highly elaborated regulatory networks (nitrogen control, for
instance) (Burkovski, 2003a; b), and the cellular responses of this bacterium due to
osmotic stress (Morbach and Kramer, 2003) revealed that L-glutamate production in C.
glutamicum is not straightforward. Therefore, intensive research is still needed in order to
minimize the problems encountered during C. glutamicum fermentation, and to increase
the production of L-glutamate.
Downstream processing is one of the most important and expensive steps in any
bioprocess industry since the separation of desired product is often limited for the success
of biological processes (Hermann, 2003). Hence, the development of a cost-effective
purification process is crucial in order to reduce the investment and production costs.
Bioprocess engineers have been trying to develop efficient methods in order to produce
the product of interest in a purified form. The separation of biomass from fermentation
broth is the first step of downstream processing that is usually accomplished either by
gravitation-based techniques (centrifugation or decantation) or by filtration (Hermann,
2003). However, a significant amount of product might be lost during this biomass
removal step. Once the biomass is removed from the fermentation broth, the purification
of the product begins. In general, glutamic acid is recovered from its fermentation broth
by removing the bacterial cells or any other impurities through centrifugation or
filtration. Filtrate is then collected, evaporated and adjusted to a pH of 3.2 (iso-electric
point of glutamic acid) by the addition of acid, and thereby crystalline glutamic acid is
obtained through precipitation (Ikeda, 2002). As the evaporation of filtrate before
acidification is very expensive and the purification by this common technique gives low
65
product yield (Hermann, 2003), proper investigations are still required in order to develop
a suitable purification process of L-glutamic acid.
In this study, considerable emphasis is given on the growth of different strains of
C. glutamicum, substrate consumption as well as L-glutamate prodcution in presence of a
range of biotin concentrations (0-200µg l-1), under the addition of different
concentrations of surfactants [Tween 20 (4g l-1), Tween 80 (4g l-1) and Tween 40 (1-4g l1
)] and in presence of a range of ethambutol concentrations (0-500mg l-1). The objective
of
this
research
is
to
investigate
the
optimum
amount
of
agent
(biotin/surfactants/ethambutol) required for the highest production of L-glutamate by the
fermentative cultivation of M. glutamicus. Furthermore, the influence of the addition time
of surfactants to L-glutamate production will be examined by adding Tweens at two
different growth stages (both start and exponential) of fermentation. The main purpose of
this investigation is to make a comparative study among the different treatments
generally applied for the production of L-glutamate by M. glutamicus fermentation, and
thereby the yield and productivity of this amino acid can be increased significantly. In
addition, a simple purification method based on centrifugation and acid-base addition will
be developed by which L-glutamate is separated from fermentation broth in a purified
form.
66
3.2. Material and Methods
3.2.1. Chemicals
Peptone from pancreatically digested casein and meat extract were obtained from
VWR (Merck, UK). Yeast extract and bacteriological agar were bought from Oxoid, UK;
and D-glucose, urea, NaCl and all other chemicals were purchased from Fisher, UK
unless otherwise mentioned. Tween (20, 40 and 80), 3-[N-Morpholino] propanesulfonic
acid (MOPS) and BHI Medium were procured from Fisher, UK. Ethambutol, biotin and
3, 4-Dihydroxybenzoic acid were purchased from Sigma-Aldrich, UK.
3.2.2. Organism and cultivation
The bacterial strains used for this study were Brevibacterium lactofermentum
DSM 1412 (Liebl et al., 1991), B. flavum DSM 20411 (Collins et al., 1979; Suzuki et al.,
1981) and Micrococcus glutamicus DSM 20300 (Collins et al., 1977; Suzuki et al., 1981;
Yamada and Komagata, 1970; Yamada et al., 1976) supplied by the German Collection
of Microorganisms and Cell Culture (DSMZ-Deutsche Sammlung von Mikroorganismen
und Zellkulturen GmbH). B. lactofermentum DSM 1412 was cultivated in Nutrient
Medium (5g l-1 peptone, 3g l-1 meat extract, 1000ml distilled water, pH adjusted to 7.0),
whereas B. flavum DSM 20411 and M. glutamicus DSM 20300 were cultivated in
Medium 53 (10g l-1 peptone, 5g l-1 yeast extract, 5g l-1 glucose, 5g l-1 NaCl, 1000ml
distilled water, pH adjusted to 7.2-7.4) according to the supplier’s instructions. pH of all
the solutions used in this study was adjusted by Microprocessor pH Meter 210 (Hanna
Instruments, USA). Agar plates with respective media were prepared by adding 15g l-1
agar to the above-mentioned compositions. The resulting solution was dissolved by
heating under stirring, and thereafter autoclaved (Astell Scientific, UK). The media were
cooled to about 50°C and mixed gently before pouring approximately 30-40ml of
solution into the sterile Petri dishes. The media were allowed to solidify, the agar plates
were dried overnight, and thereafter stored at 4°C in an inverted position.
A loop of growing culture cultivated on both Nutrient Medium and Medium 53
was transferred on to the respective agar plates, streaked under the aseptic condition, and
thereafter incubated (Camlab, UK) at 30C until the colonies were visible (for 48-72h).
67
These plates (stock cultures) were preserved in refrigerator for further experiments. The
same procedure was carried out at least for five generations in order to make those
bacteria suitable for the growth study and L-glutamate fermentation. However, fresh
stock cultures were generated every month throughout the study. Inocula were prepared
by selecting a single colony from the stock agar plates and transferring into Seed Medium
(Tatsuya et al., 1997) or Brain Heart Infusion (BHI) Medium depending on the purpose
of investigation. In the case of growth study, bacteria were cultivated in Seed Medium
(30g l-1 glucose, 3g l-1 yeast extract, 4g l-1 urea, 1g l-1 potassium dihydrogen phosphate,
20mg l-1 ferrous sulphate and 20mg l-1 manganese sulphate, 1000ml distilled water, pH
adjusted to 7.0) in which Soyabean protein hydrolaste used in the original composition
was replaced by yeast extract. A colony from the stock agar plate was cultivated
overnight into 3.5% BHI Medium in order to prepare the preculture for L-glutamate
production. In both cases, the cultivation was carried at 30C and shaken at 220rpm in an
orbital incubator shaker (Weiss Gallenkamp, UK). Both the Seed and BHI Media were
sterilized at 110C for 10min, and adjusted to pH 7.0 before inoculating the colonies
from stock agar plates. All the experiments were performed in 250ml shake flasks
containing 50ml of culture media.
3.2.3. Microbial growth measurement
Microbial cells growth is generally measured by UV-visible spectrophotometer,
dry weight measurement (g Dw) and cell counting through the use of a haemocytometer
both in laboratory and biotech industry. In most experiments carried out throughout the
study, however, the growth of different strains of C. glutamicum was monitored by
measuring the absorbance at 600nm (600) using spectrophotometer [S1200, WPA (a
wholly owned subsidiary of Biochrom Ltd)]. Bacterial cultures or samples were diluted
(if concentrated) to keep the absorbance within the range of 0.1-0.3, and thereafter
multiplied with the dilution factor in order to obtain the actual optical density (OD). In
addition, the determination of biomass concentration (g Dw l-1) was started with the
filtration of a known amount of sample followed by drying the membrane filter
(Whatman plain cellulose acetate, white, 0.2µm pore size, 47mm diameter, purchased
68
from Fisher Scientific, UK) in a microwave oven. The following steps were carried out
for the measurement of biomass
The membrane filters were dried for 10min at 150W in a microwave oven.

The filter papers were cooled at RT for 15min in a desiccator and weighed.

Approximately 2-5ml of fermentation broth was filtered and washed with the
same amount of distilled water.

The filter papers were dried in the microwave oven for 15min at 150W.

The papers were cooled again at RT for 15min in a desiccator and weighed.

The amount of biomass was determined by substracting the weight of filter papers
before filtration from the weight of filter papers after filtration.
3.2.4. Calculation of maximum specific growth rate (µmax)
Specific growth (µ) rate is defined as the increase in cell mass per unit of cell
mass per unit of time (g. g-1. h-1). However, the widely used unit of growth rate is h-1. The
specific growth provides a quantitative expression of the ability of microorganisms to
grow on a particular substrate. In an environment where the unlimited cell growth is
possible, the growth rate of cells (dx/dt) is proportional to the density of population (cells
ml-1, expressed by x).
dx/dt=µmax x
Where, µmax is described as the maximum specific growth rate under which the
substrate concentration is high enough that does not impose any limitation to the reaction
process. In this study, µmax was calculated by taking the slope of a plot of cell density
(OD), corresponding to the exponential growth phase versus time.
3.2.5. L-glutamate production in M. glutamicus under biotin limited condition
L-glutamate was produced in M. glutamicus by cultivating cells in CGXII
Minimal Medium that has been developed by Keilhauer and his co-workers. The
preparation of CGXII Minimal Medium is as follows-
69
Preparation of CGXII Minimal Medium:
50% (w/v) glucose solution
50g glucose was dissolved in 60ml distilled water by heating under stirring. After
complete solubilization, distilled water was added up to 100ml and thereafter autoclaved.
CaCl2
1g CaCl2 was dissolved in 100ml of distilled water.
Biotin
20mg biotin was transferred in 100ml of distilled water, dissolved by heating, and
the resulting solution was sterilized by 0.2µm Nalgene filter.
Trace elements
1g FeSO4 × 7H2O
1g MnSO4 × H2O
0.1g ZnSO4 × 7H2O
0.02g CuSO4
0.002g NiCl2 × 6H2O
90ml distilled water was added and dissolved the above salts by adding of
concentrated HCl. The final pH of the resulting solution was kept about 1.0. The solution
was then sterilized by 0.2µm Nalgene filter.
3, 4-Dihydroxybenzoic acid
300mg of 3, 4-Dihydroxybenzoic acid was added to 8ml of distilled water. The
solution was dissolved by adding of about 1ml of 10N NaOH, sterilized by filtration and
stored at 4°C.
CGXII-salts minus biotin
20g (NH4)2SO4
5g urea
70
1g KH2PO4
1g K2HPO4
0.25g MgSO4 × 7H2O
42g MOPS (3-[N-Morpholino] propanesulfonic acid)
1ml CaCl2 solution
All the above-mentioned constituents were weighed accurately and transferred
into a bottle. 800ml of distilled water was added and dissolved the constituents with
stirring. The resulting solution was adjusted to pH 7.0 with 1N NaOH, filled up the bottle
with distilled water up to 920ml (again adjusted to pH 7.0) and thereafter autoclaved. 1ml
trace element solution was added into the sterilized solution. CGXII minus biotin was
prepared by adding 1ml of 3, 4-Dihydroxybenzoic acid and 80ml of 50% (w/v) glucose
solution.
Glutamate production in CGXII Minimal Medium:
Preculture preparation (Day 1)
One colony from a fresh Luria-Bertani (LB) agar plate was inoculated into 50ml
BHI medium and cultivated overnight in a 500ml Erlenmeyer flask on an orbital
incubator shaker at 220rpm and 30°C.
Biotin depletion (Day 2)
50ml of CGXII minus biotin medium was dispensed into a 500ml Erlenmeyer
flask and labelled the flask “5” that stands for 5μg of biotin per liter. The biotin stock
solution (200µg ml-1) was diluted 1:100 with sterile distilled water, and added 125μl of
diluted solution into the flask labelled “5”. The OD (600nm) of the overnight BHI
preculture was measured, and the appropriate amount of inoculum was transferred into a
sterile 15ml Falcon tube and centrifuged (Heraeus, Germany) at 3000g for 10min. The
cell pellets were resuspended in a few milliliters of CGXII Medium and the resulting
volume was transferred into the Erlenmeyer flask labelled “5” where the starting OD600
was 0.1. The flask was then incubated on an orbital incubator shaker at 220rpm and
30°C.
71
Glutamate production (Day 3)
50ml of the CGXII minus biotin medium was dispensed into seven 500ml
Erlenmeyer flasks labelled “0”, “0.5”, “1.0”, “1.5”, “2.0”, “2.5”and “200” which stand
for 0 (without biotin), 0.5, 1, 1.5, 2, 2.5 and 200μg of biotin per liter, respectively. The
biotin stock solution was diluted 1:10 and 1:100 with sterile distilled water and
appropriate amount of biotin was transferred into the mentioned flasks, whereas no biotin
was added to the flask labelled “0”. The OD (600nm) of the biotin depletion culture was
measured, and the appropriate amount of inoculum was transferred parallelly into seven
sterile 15ml Falcon tube and centrifuged (Heraeus, Germany) at 3000g for 10min. Cell
pellets were dissolved in few milliliters of medium, and the resulting volumes were
poured into the corresponding flasks where the starting ODs at 600nm were 1.0. The
flasks were incubated on an orbital incubator shaker at 220rpm and 30°C, and 1ml of
sample from the mentioned flasks was collected randomly over a period of 72h in order
to determine the OD600, glucose consumption and glutamate production.
3.2.6. L-glutamate production in M. glutamicus by surfactant addition
In this study, three different surfactants (Tween 20, Tween 40 and Tween 80)
were added into CGXII Minimal Medium (Section 3.2.5) in order to excrete L-glutamate
under biotin- rich (200µg l-1) cultivation of M. glutamicus. Preculture propagation (day 1)
was the same as the method used in Section 3.2.5. 50ml of the CGXII minus biotin
medium was dispensed into seven 500ml Erlenmeyer flasks labelled “0”, “TW 20 (4)”,
“TW 40 (1)”, “TW 40 (2)”, “TW 40 (3)”, “TW 40 (4)”and “TW 80 (4)” which stand for 0
(without Tween), 2g l-1 Tween 20, 1 to 4g l-1 Tween 40 and 2g l-1 Tween 80, respectively.
The OD600 of the overnight BHI preculture was measured, and the appropriate amount of
inoculum was transferred parallelly into seven sterile 15ml Falcon tube and centrifuged
(Heraeus, Germany) at 3000g for 10min. Cell pellets were dissolved in few milliliters of
medium, and the resulting volumes were poured into the corresponding flasks where the
starting ODs at 600nm were 0.5. Tween was added into the flasks both at the start of
fermentation and exponential phase. The flasks were then incubated on an orbital
incubator shaker at 220rpm and 30°C, and samples were collected randomly over a
period of 72h to determine the OD600, glucose consumption and glutamate production.
72
3.2.7. L-glutamate production in M. glutamicus by ethambutol addition
A range of ethambutol concentrations (10-500mg l-1) was added into CGXII
Minimal Medium (Section 3.2.5) in order to excrete L-glutamate under biotin rich
(200µg l-1) condition of M. glutamicus. Preculture propagation (day 1) was the same as
the procedure used in Section 3.2.5. 50ml of the CGXII minus biotin medium was
dispensed into eight 500ml Erlenmeyer flasks labelled “0”, “10”, “20”, “30”, “50”,
“100”, “200’’ and “500” which stand for 0 (without ethambutol), 10, 20, 30, 50, 100, 200
and 500mg l-1, respectively. The OD600 of the overnight BHI preculture was measured,
and the appropriate amount of inoculum was transferred parallelly into seven sterile 15ml
Falcon tube and centrifuged (Heraeus, Germany) at 3000g for 10min. Cell pellets were
dissolved in few milliliters of medium, and the resulting volumes were poured into the
corresponding flasks where the starting ODs at 600nm were 0.5. Ethambutol was only
added at the exponential phase of fermentation. The flasks were then incubated on an
orbital incubator shaker at 220rpm and 30°C, and samples were collected randomly over
a period of 72h in order to determine the OD600, glucose consumption and glutamate
production.
3.2.8. Quantification of glucose, L-glutamate and other amino acids
The quantification of substrate (glucose), L-glutamate and all the other amino
acids were achieved with the use of AAA-DirectTM Amino Acid Analysis System
(Dionex, UK). This method has been developed in such a way that glucose and a range of
amino acids are separated by gradient (Table 3.1) anion exchange chromatography, and
simultaneously determined their concentration through the Pulsed Electrochemical
Detector of HPLC (ED40, Dionex, UK) (Ding et al., 2002; Yu et al., 2002).
Preparation of eluents and standards
Eluent 1: Deionised water
Approximately 2.0L of Milli Q water (Direct Q5, Millipore, USA) with a
resistivity of 18.2 M-cm at 25°C were degassed using helium, and then transferred into
the eluent bottle. The bottle was sealed immediately in order to minimize the time to
expose with atmosphere.
73
Eluent 2: 0.250 M Sodium hydroxide
About 986ml of degassed Milli Q water and 14ml of 46-48% NaOH (Fisher, UK)
were transferred into a volumetric flask, and mixed properly by inversion. The solution
was then poured immediately into the eluent bottle supplied by the manufacturer, and
sealed properly to minimize the carbon dioxide absorption. The pressure was allowed to
build up inside the bottle, and the cap was reopened briefly several times in order to
replace the trapped air by the inert gas.
Eluent 3: 1.0 M Sodium acetate
Approximately 450ml of degassed 18.2M-cm water was added into a sodium
acetate container (Dionex, UK), and shaken until the contents were completely dissolved.
The resulting sodium acetate solution was then transferred into a volumetric flask (1.0L).
The sodium acetate container was rinsed with approximately 100ml of degassed water,
transferred into the flask and filled with the water up to the mark. The eluent was then
mixed properly, and transferred into the eluent bottle. All of these procedures were done
quickly in order to minimize the time to exposure with atmospheric carbon dioxide.
Standard
The standard amino acids obtained from Sigma (AAS18-10X1ML) contained
amino acids at 2.5µM ml-1 except cystine at 1.25µM ml-1. A stock solution of glucose and
amino acids was prepared by mixing 0.5ml standard (2.5mM) with 12ml glucose solution
(0.1mM), which contained 100µM of each amino acid, 50µM of cystine and 100µM of
glucose (3.1). This stock solution was stored in aliquots at -20C prior to use.
Gradient conditions for analysis of amino acids and carbohydrates
The gradient conditions for analysis of amino acids and carbohydrate are
summarized in Table 3.1. 25µl of standard/sample was injected for analysis, where the
eluent flow rate and column temperature were adjusted to 0.25ml min-1 and 30C.
74
Figure 3.1 Simultaneous analysis of glucose and amino acids in fermentation broth via HPLC.
Table 3.1 Gradient conditions for analysis of amino acids and carbohydrates
Time (min)
%E1
%E2
%E3
Comments
Init
84
16
0
Autosampler fills the sample loop
0.0
84
16
0
Valve from load to inject
0.0
84
16
0
Begin hydroxide gradient
12.1
68
32
0
16.0
68
32
0
24.0
36
24
40
40.0
36
24
40
40.1
20
80
0
42.1
20
80
0
42.2
84
16
0
Equilibrate to starting conditions
65.0
84
16
0
Ready for the next run
Begin acetate gradient
Column wash with hydroxide
75
3.2.9. Recovery of glutamic acid from fermentation broth
Even though satisfactory yields of glutamic acid were obtained by the above
mentioned treatments of M. glutamicus, numerous challenges have already been
mentioned in the literatures that make the recovery of this amino acid extremely difficult
and expensive. A purification method for isolating glutamic acid by adding a zinc salt
(ZnSO4.7H2O) into the culture liquid was successfully developed in which bacterial cells
or any other impurities were separated by means of centrifugation. A systematic diagram
is presented as follows:
Fermentation broth
Centrifuged at 4600g for 1hr at 4C (Heraeus, Germany), and collected the supernatant
(200ml)
ZnSO4.7H2O (1:1.32 ratio) was added into the supernatant and dissolved the salt properly
pH was adjusted to 6.3 by the addition of NH4OH solution (25-30% NH3), stirred the
resulting solution for 30min
Centrifuged the solution at 4600g for 30min at 4C, and collected the precipitated zinc
glutamate
The precipitated salt was washed with dH2O, and slurried with 10ml of dH2O
50% NaOH solution was added until the pH reached to 12.3, stirred the solution for 1h
The solution was centrifuged at 4600g for 30min at 4C, and collected the precipitated
sodium glutamate
The precipitated sodium glutamate was washed with dH2O, and slurried with 10ml of
dH2O
76
Concentrated H2SO4 was added in order to adjust the pH 3.2, stirred the solution for 1h
The solution was centrifuged at 4600g for 30min at 4C, and removed the supernatant
The precipitated L-glutamate acid was collected, and dried at room temperature (RT)
1.49g of L-glutamate was weighed that is approximately 89% recovery of the HPLC
measurement
77
3.3. Results
3.3.1. Growth properties of the different strains of Corynebacteria in a defined
medium
To examine the growth properties of C. glutamicum strains i.e., B.
lactofermentum, B. flavum and M. glutamicus, colonies from the respective bacterial
cultures grown on agar plates (stock) were aseptically inoculated in 250ml shake flasks
containing Seed Medium. The cultivation was carried out overnight at 30C and the
flasks were shaken on an orbital incubator shaker (220rpm). Appropriate amount of
growing cultures were then transferred in new flasks having Seed Medium in such way
the starting ODs in all flasks were around 1.0. Each strain of C. glutamicum was
cultivated in three shake flask (i.e., total of 9), and the number of samples analyzed in
order to determine the growth of cells (OD600) and glucose consumption on each occasion
was two. Samples were collected throughout the cultivation at certain intervals for the
measurement of cells growth and residual glucose concentrations. The growth of cells
was determined either by optical density (OD) at 600nm or dry weight (g Dw)
measurements. A correlation between OD and dry weight measurement is presented in
Figure 3.2. High Performance Liquid Chromatography (HPLC) was used in order to
determine the residual glucose concentrations in samples.
In Figure 3.3a, the growth of cells and substrate consumption of three different
strains of C. glutamicum over time can be seen; and it is clear that all the strains started
growing with approximately 4h of lag phase, grew exponentially up to 12h and finally
shifted to the stationary phase after 13-14h of inoculation. The ODs measured after 24h
of cultivation were 44, 45 and 44 (Figure 3.3a) in the case of B. lactofermentum, M.
glutamicus and B. flavum, respectively. The exponent of each curve was considered as
the specific growth rate (µ) that was almost constant to 0.38h-1 in the case of all the
strains investigated in this study (Figure 3.3b). In addition, this study showed that the
glucose (Glu) available (30g l-1) in Seed Medium was consumed within 20h of
cultivation, irrespective of the strains of C. glutamicum considered for this investigation
(Figure 3.3a). However, no prominent differences in growth properties (cell biomass,
specific growth rate and substrate consumption) were observed among these three strains.
78
50
OD at 600 nm
40
y = 4.2923x - 1.143
2
R = 0.9992
30
20
10
0
0
2
4
6
8
10
12
Biomass, g Dw/l
Figure 3.2 A correlation between OD and dry weight measurement of M. glutamicus growth.
50
35
OD at 600nm
25
30
20
20
15
10
10
5
0
0
0
5
10
15
20
25
30
Time after inoculation, hr
OD, B. lactofermentum
OD, M. glutamicus
OD, B. flavum
Glu, B. lactofermentum
Glu, M. glutamicus
Glu, B. flavum
Figure 3.3a Growth studies of different strains of Corynebacteria on Seed Medium.
79
Glucose, g/l
30
40
30
y = 0.6419e 0.3751x y = 0.5844e 0.383x y = 0.5309e 0.3788x
R2 = 0.9993
OD at 600nm
25
R2 = 0.9964
R2 = 0.9993
20
15
10
5
0
0
2
4
6
8
10
12
Time after inoculation, hr
B. lactofermentum
M. glutamicus
B. flavum
Figure 3.3b The growth (OD600) of different strains of Corynebacteria on Seed Medium.
3.3.2. L-glutamate production by the different strains of Corynebacteria under
biotin limited condition
The important feature of cultivating different strains of Corynebacteria under the
supply of limited amount biotin was to supply a biotin concentration that is low enough
to trigger L-glutamate secretion but high enough to allow sufficient growth of bacteria. A
colony from the stock agar plate of M. glutamicus was initially cultivated (overnight) in
BHI medium (Day 1). In order to optimize the appropriate concentration of biotin
essential for L-glutamate production, a precultivation step was carried out in which the
requirement of biotin for cells growth was depleted (Day 2). The overnight culture of M.
glutamicus obtained from the biotin depletion step (Day 2) was then transferred in CGXII
Minimal Medium in which a range of biotin concentrations i.e., 0, 0.5, 1.0, 1.5, 2.0, 2.5
and 200µg l-1 was added at the start of fermentation. The idea of supplying a range of
biotin concentrations was to determine the optimum biotin required for the highest
production of L-glutamate.
At each concentration of biotin, the preculture of M. glutamicus collected from
day 2 was transferred into three shake flasks (i.e., total of 21) containing CGXII Minimal
Medium, and the number of samples analyzed in order to determine the growth of cells
80
(OD600), glucose consumption and L-glutamate production on each occasion was two.
Figure 3.4 depicts the growth of M. glutamicus and glucose consumption in presence of a
range of biotin concentrations. The result showed that increasing the concentration of
biotin is fermentation medium increased both cell biomass and the rate of glucose
consumption. The ODs after 48h of cultivation were measured and found to be 5, 18, 31,
32, 34, 40 and 53 in cultures containing biotin at 0, 0.5, 1.0, 1.5, 2.0, 2.5 and 200µg l-1,
respectively. The residual glucose concentrations measured after 72h of fermentation
were 3, 0.7, 0.4, 0.13, 0.1 and 0g l-1 in cultures containing biotin at 0.5, 1.0, 1.5, 2.0, 2.5
and 200µg l-1, respectively (Figure 3.4). When biotin was not added in the CGXII
Minimal Medium, only 30% glucose was observed to be consumed even after 72h of
fermentation.
The amounts of L-glutamate produced after 48h of fermentation were 41, 56, 28,
20 and 8mM in presence of 0.5, 1.0, 1.5, 2.0 and 2.5µg l-1 biotin, respectively (Figure
3.5). Although the highest production of L-glutamate by M. glutamicus was measured as
57mM (after 52h) at a biotin concentration of 1µg l-1, the production this amino acid at
0.5µg l-1 biotin was 40mM (after 52h) which is a reasonably significant level of
production. In contrast, the amount of L-glutamate measured in control (without biotin)
and the culture cultivated at 200µg l-1 biotin were 1.5 and 0.1mM, respectively. The
results confirmed that cell growth, glucose consumption and L-glutamate production are
completely dependent on biotin concentration supplied in CGXII Minimal Medium.
Moreover, the results revealed that the optimum concentration of biotin required for Lglutamate production by M. glutamicus is approximately 1µg l-1 (Figure 3.5). Since the
previous study (Figure 3.3a) clearly demonstrated that biotin has an influence on M.
glutamicus growth, the specific growth rates of this organism under the supply of
different concentrations of biotin were investigated. Table 3.2 shows that increasing the
concentration of biotin in CGXII Minimal Medium increased the specific growth rate of
M. glutamicus, however, little cell growth occurred in the absence of biotin (Figure 3.4).
As the earlier investigation demonstrated that there is no difference in growth properties
and glucose consumption among the three strains, the production of L-glutamate by these
strains under the presence of 1µg l-1 biotin was also examined. The results showed that
81
60
45
40
50
35
30
25
30
20
20
15
10
10
5
0
0
0
10
20
30
40
50
60
70
Time after inoculation, hr
OD, 0µg/l
Glu, 0µg/l
OD, 0.5µg/l
Glu, 0.5µg/l
OD, 1µg/l
Glu, 1µg/l
OD, 1.5µg/l
Glu, 1.5µg/l
OD, 2.0µg/l
Glu, 2.0µg/l
Figure 3.4 Effect of different concentrations of biotin on M. glutamicus growth and glucose consumption.
82
OD, 2.5µg/l
Glu, 2.5ug/l
OD, 200µg/l
Glu, 200µg/l
80
Glucose, g/l
OD at 600nm
40
70
60
Glutamate, mM
50
40
30
20
10
0
0
20
22
24
26
48
Time after inoculation, hr
0µg/l
0.5µg/l
1.0µg/l
1.5µg/l
2µg/l
2.5µg/l
Figure 3.5 L-glutamate production in presence of different concentrations of biotin by M. glutamicus.
83
200µg/l
52
72
Table 3.2 Effect of different concentrations of biotin on specific growth rate of M. glutamicus
Concentration of biotin (µg l-1)
Specific growth rate (h-1)
0
0.12
0.5
0.16
1.0
0.18
1.5
0.18
2.0
0.18
2.5
0.19
200
0.21
the productions of L-glutamate measured after 48h of fermentation were 54, 53 and
56mM in the case of B. lactofermentum, B. flavum and M. glutamicus, respectively,
indicating that there is no notable difference in L-glutamate production among the three
strains of C. glutamicum considered in this study (Figure 3.6).
70
60
Glutamate, mM
50
40
30
20
10
0
0hr
24hr
48hr
72hr
Time after inoculation, hr
M. glutamicus
B. lactofermentum
B. flavum
Figure 3.6 L-glutamate production by different strains of Corynebacteria under biotin limited
(1µg l-1) condition.
84
3.3.3. Secretion of L-glutamate produced by M. glutamicus due to surfactant
addition
The overnight culture of M. glutamicus cultivated in BHI medium was transferred
into the CGXII Minimal Medium in which 200µg l-1 was already added at the start of
fermentation. Tween 20 (4g l-1), Tween 80 (4g l-1) and Tween 40 (1-4g l-1) were added
both at the start (ST) and exponential (EXP) growth phase of M. glutamicus in order to
investigate the most favourable time of their addition into the fermentation medium. At
each concentration of surfactants, the preculture of M. glutamicus grown in BHI medium
was transferred into three shake flasks (i.e., total of 21) containing CGXII Minimal
Medium, and the number of samples analysed in order to determine the growth of cells
(OD600), glucose consumption and L-glutamate production on each time was two. The
results showed that addition of Tween 40, irrespective of the concentration and growth
phases at which it was added into CGXII Medium, reduced the growth of M. glutamicus,
decreased the rate of glucose consumption, however, excreted L-glutamate into the
extracellular medium to a certain degree in spite of having sufficient amount of biotin
(200µg l-1) in the fermentation medium.
The ODs after 48h of cultivation were measured and found to be 49, 37, 35, 33,
30 and 54 in cultures containing Tween 20 (4g l-1), Tween 40 (1g l-1), Tween 40 (2g l-1),
Tween 40 (3g l-1), Tween 40 (4g l-1) and
Tween 80 (4g l-1), respectively, where
surfactants were added at the start of fermentation (Figure 3.7). On the other hand, ODs
measured after 48h of fermentation were 51, 40, 37, 35, 33 and 55, respectively, in
cultures where these agents were added after 8h of cultivation (Figure 3.8). In the case of
control (without surfactant), however, the OD was 52 (after 48h). At both conditions, the
growth of cells was observed to decrease after the addition of Tween 40 into the
fermentation medium, nevertheless, cells started to grow exponentially after a while. The
results also demonstrated that the inhibition of cell’s growth due to Tween 40 addition at
the start of fermentation was intense, and finally produced less biomass as compared to
the cultures in which Tween 40 was added at the exponential growth phase (after 8h of
cultivation). In addition, M. glutamicus cells that were grown with Tween 40 at the start
of cultivation showed slower glucose consumption than that of exponential addition. The
85
residual glucose concentrations after 72h of fermentation were measured and found to be
3.5, 4.0, 5.4 and 7.0g l-1 in cultures grown at 1, 2, 3 and 4g l-1 Tween 40 (start addition),
respectively, whereas there was no glucose left in cultures in which Tween 40 was added
exponentially. From these experiments, however, it is observed that the growth of cells
was not inhibited due to the addition of Tween 20 and 80 in CGXII Minimal Medium
Like biomass and glucose consumption, the production of L-glutamate varied
depending on the concentration and addition time of Tween 40. The amounts of Lglutamate produced after 48h of fermentation were 20, 22, 19 and 17mM (Figure 3.9) at
1, 2, 3 and 4g l-1 Tween 40 (start addition), respectively, whereas the productions were
39, 45, 40 and 36mM (Figure 3.10) in cultures where Tween 40 was added to the
exponential growth phase of M. glutamicus. However, the results clearly demonstrated
that the highest amount of L-glutamate production was observed by supplying of 2g l-1
Tween 40 to a biotin rich (200µg l-1) CGXII Minimal Medium, irrespective of the growth
phases of M. glutamicus at which this surfactant was added. After 48h of cultivation, in
contrast, the amounts of L-glutamate measured in control, cultures grown at 4g l-1 Tween
20 and 4g l-1 Tween 80 were 0.2, 0.5 and 0.4mM (start addition) and 0.4, 1.9 and 3.8mM
(exponential addition), respectively. Nevertheless, the results clearly demonstrated that
addition of Tween 20 and 80, regardless of the growth phases of M. glutamicus at which
these surfactants were added, were not effective in excreting L-glutamate by M.
glutamicus.
86
60
45
40
50
35
30
25
30
20
20
15
10
10
5
0
0
0
10
20
30
40
50
60
70
80
Time after inoculation, hr
Control (No TW), OD
TW-20 (4g/l), ST OD
TW-40(1g/l), ST OD
TW-40(2g/l), ST OD
TW-40(3g/l), ST OD
TW-40(4g/l), ST OD
TW-80(4g/l), ST OD
Control (No TW), Glu
TW-20 (4g/l), ST Glu
TW-40(1g/l), ST Glu
TW-40(2g/l), ST Glu
TW-40(3g/l), ST Glu
TW-40(4g/l), ST Glu
TW-80(4g/l), ST Glu
Figure 3.7 Effect of a range of surfactants (added at the start of cultivation) on M. glutamicus growth and glucose consumption.
87
Glucose, g/l
OD at 600nm
40
70
45
40
60
35
50
40
25
30
20
Glucose, g/l
OD at 600nm
30
15
Tween
addition
20
10
10
5
0
0
0
10
20
30
40
50
60
70
80
Time after inoculation, hr
Control (No TW), OD
TW 40 (4g/l), EXP OD
TW 40 (2g/l), EXP Glu
TW 20 (4g/l), EXP OD
TW 80 (4g/l), EXP OD
TW 40 (3g/l), EXP Glu
TW 40 (1g/l), EXP OD
Control (No TW), Glu
TW 40 (4g/l), EXP Glu
TW 40 (2g/l), EXP OD
TW 20 (4g/l), EXP Glu
TW 80 (4g/l), EXP Glu
TW 40 (3g/l), EXP OD
TW 40 (1g/l), EXP Glu
Figure 3.8 Effect of a range of surfactants (added at the exponential phase of growth) on M. glutamicus growth and glucose consumption.
88
50
Glutamate, mM
40
30
20
10
0
0hr
24hr
48hr
72hr
Time after inoculation, hr
Control, ST
TW-40(3g/l), ST
TW-20 (4g/l), ST
TW-40(4g/l), ST
TW-40(1g/l), ST
TW-80(4g/l), ST
TW-40(2g/l), ST
Figure 3.9 L-glutamate production by M. glutamicus under different concentrations of surfactants
(added at the start of cultivation).
50
Glutamate, mM
40
30
20
10
0
0hr
24hr
48hr
72hr
Time after inoculation, hr
Control, EXP
TW-40(3g/l), EXP
TW-20 (4g/l), EXP
TW-40(4g/l), EXP
TW-40(1g/l), EXP
TW-80(4g/l), EXP
TW-40(2g/l), EXP
Figure 3.10 L-glutamate production by M. glutamicus under different concentrations of
surfactants (added at the exponential phase of growth).
89
3.3.4. Secretion of L-glutamate produced by M. glutamicus due to ethambutol
addition
The overnight culture of M. glutamicus cultivated in BHI medium was transferred
into the CGXII Minimal Medium in which 200µg l-1 was added at the start of
fermentation. A range of EMB concentrations (10-500mg l-1) was added to the
exponentially (8h after inoculation) grown M. glutamicus in order to excrete L-glutamate.
The idea of supplying a range of EMB concentrations was to determine the optimum
level of ethambutol excreting the highest amount of L-glutamate. At each concentration
of ethambutol, the preculture of M. glutamicus grown in BHI medium was transferred
into three shake flasks (i.e., total of 24) containing CGXII Medium, and the number of
samples analyzed in order to determine the growth of cells (OD600), glucose consumption
and L-glutamate production on each occasion was two. The results showed that addition
of EMB reduced both the growth of M. glutamicus and glucose consumption as compared
to the control (Figure 3.11). The ODs measured at 600nm after 48h of fermentation were
39, 36, 33, 32, 29, 29, and 27 in the case of cells treated with 10, 20, 30, 50, 100, 200, and
500mg l-1 EMB, respectively, whereas the OD600 in control (without EMB) was 52. In the
case of control, however, all the available glucose (initially, 40g l-1) was observed to be
consumed after 32h of fermentation.
On the other hand, the concentrations of residual glucose in samples (culture that
was supplied with 100mg l-1 of EMB) collected after 36 and 48h of fermentation were 5.0
and 3.8g l-1, respectively. In addition, the results showed that the production of Lglutamate increased due to the increase of EMB concentration (up to100mg l-1) in the
CGXII Minimal Medium. After 48h of fermentation, the amounts of L-glutamate
measured in samples supplied with 10, 20, 30, 50, 100, 200, and 500mg l-1 of EMB
were15, 18, 24, 34, 49, 47 and 48mM (Figure 3.12). In the case of control, on the
contrary, only 0.14mM L-glutamate was measured. The result from this experiment
showed that the optimum amount of ethambutol needed for the highest secretion of Lglutamate was 100mg l-1, and the enhancement of production was not possible by
increasing the concentration of ethambutol more than 100mg l-1. In order to determine the
most suitable treatment that can be applied for glutamic acid production, the amounts of
90
60
45
40
50
35
30
25
30
20
20
Glucose, g/l
OD at 600nm
40
15
Ethambutol
addition
10
10
5
0
0
0
10
20
30
40
50
60
70
80
Time after inoculation, hr
Control (No E), OD
10mg/ml E, OD
20mg/ml E, OD
30mg/ml E, OD
50mg/ml E, OD
100mg/ml E, OD
500mg/ml E, OD
200mg/ml E, OD
Control (No E), Glu
10mg/ml E, Glu
20mg/ml E, Glu
30mg/ml E, Glu
50mg/ml E, Glu
100mg/ml E, Glu
500mg/ml E, Glu
200mg/ml E, Glu
Figure 3.11 Effect of a range of concentrations of ethambutol (E) on M. glutamicus growth and glucose consumption.
91
60
50
Glutamate, mM
40
30
20
10
0
0mg/l
10mg/l
20mg/l
30mg/l
50mg/l
100mg/l
200mg/l
Concentrations of Ethambutol
24hr
48hr
72hr
Figure 3.12 L-glutamate production by M. glutamicus due to the exponential addition of a range of ethambutol concentrations.
92
500mg/l
L-glutamate produced by the fermentative cultivations of M. glutamicus under the three
different treatments, such as biotin limitation, surfactant addition and ethambutol addition
were plotted in a graph. Figure 3.13 shows that the productions of glutamic acid after 48h
fermentation were 56, 45 and 49mM, respectively, indicating that the production of Lglutamate in biotin limited condition is significantly higher than those obtained under the
other growth conditions.
70
Glutamate, mM
60
50
40
30
20
10
0
24hr
48hr
72hr
Time after inoculation, hr
1µg/l biotin
200µg/l biotin, Tween 40 (2g/l) (EXP)
200µg/l biotin, Ethambutol (100mg/l) (EXP)
Figure 3.13 L-glutamate production by M. glutamicus fermentation under three different
conditions.
93
3.4. Discussion
In this research, L-glutamate production was investigated by the three different
strains of Corynebacteria i.e., B. lactofermentum, B. flavum and M. glutamicus cultivated
on CGXII Minimal Medium under biotin limited condition. It is clearly demonstrated that
there are no prominent differences in growth properties, substrate consumption and Lglutamate production among the three different strains of Corynebacteria (Figure 3.3a;
3.6). A study on DNA homology among the Corynebacterial strains, such as B.
lactofermentum, B. flavum, B. divaricatum, M. glutamicus and C. lilium has already been
revealed that there are no prominent differences among the various glutamic acid bacteria
(Goodfellow et al., 1976). Hence, these bacteria can all be grouped within the taxon of
Corynebacterium sensus stricto, and be regarded as a single species of the genus
Corynebacterium (Eikmanns et al., 1991). Since the discovery of glutamic acid
production by Corynebacteria fermentation, many researchers investigated the action of
biotin, surfactants, penicillin, ethambutol and oleic acid or glycerol on the membrane
permeability of these bacteria (Demain and Birnbaum, 1968; Eggeling and Sahm, 2001;
Ikeda et al., 1972; Kanzaki et al., 1967; Okazaki et al., 1967; Radmacher et al., 2005;
Shiio et al., 1962; Takinami et al., 1965; 1968). However, the influence of these agents
on the production of L-glutamate is not well-established so far.
In this study, the effect of biotin, Tween 40 and ethambutol on the production of
L-glutamate by M. glutamicus fermentation was investigated. The results clearly showed
that biotin has an important influence on the growth of M. glutamicus. In biotin rich
condition (200µg l-1), the OD at 600nm was 53, whereas it was 18 and 40 in presence of
0.5 and 2.5µg l-1 biotin, respectively, indicating that increasing the concentration of biotin
in fermentation (CGXII Minimal) medium increases biomass production. In general,
substrate taken up by this organism is converted into acetyl-CoA through glycolysis.
However, this building block (acetyl-CoA) is inactivated by the limited concentration of
biotin available in fermentation medium, and therefore cannot participate in fatty acid
synthesis (Kramer, 1994). As the fatty acid synthesis is interrupted due to biotin
limitation, M. glutamicus cells might not be able to build up their cytoplasmic membrane
and cell wall, and eventually inhibited the growth of cells.
94
However, there was not much difference observed in biomass production (ODs
after 48h of fermentation were 31, 32 and 34) among the cultures cultivated in presence
of 1.0, 1.5 and 2.0µg l-1 biotin, respectively (Figure 3.4). This result confirmed that the
minimum amount biotin required for sufficient growth of M. glutamicus is 1-2µg l-1.
Nevertheless, when biotin was not added in the CGXII Minimal Medium (control), this
investigation showed insignificant cell growth (OD600 of 5.3), extremely low glucose
consumption (only 25% even after 72h of cultivation) and trace amount of L-glutamate
production (Figure 3.4 and 3.5a). Hence, it is now concluded that biotin is directly
involved in fatty acid synthesis, and decrease in cell biomass during biotin limited
condition was possibly caused by a change in metabolic activities related to fatty acid
synthesis of M. glutamicus. Hoischen and Kramer (1990) analyzed the changes in lipid
content, cell wall composition and the kinetic and energetic properties of substrate
transport across the membrane between producer (biotin limited) and non-producer
(biotin rich) cells. Their result showed that the total amount of lipids (represented by fatty
acids) as well as the phospholipids content decreased by about 50% in cells cultivated in
biotin limited condition.
Although biomass production in presence of 1.0, 1.5 and 2.0µg l-1 biotin was
almost similar (Figure 3.4), the productions of L-glutamate at these concentrations were
56, 28 and 20mM, respectively (Figure 3.5a), indicating that the concentration of biotin
available in the CGXII Minimal Medium has also a major influence on the production of
L-glutamate. On the other hand, no L-glutamate was obtained in the presence of 200µg l-1
biotin, confirming that L-glutamate production is not possible in biotin rich condition
(Figure 3.5a). Furthermore, it has been reported that addition of excess biotin in
production medium produces lactate as a by-product although it encourages cells to grow
optimally, whereas biotin limitation directly affects fatty acid synthesis and results in the
secretion of L-glutamate (Kramer, 1994). Shiio et al. (1962) demonstrated that the
amount of intracellular L-glutamate in cells grown in a biotin limited medium was less
than that in cells grown in a biotin rich medium. In the former case, the intracellular Lglutamate was almost released from the unstructured cell wall, whereas in other condition
the release of this amino acid was only observed when the cells were grown with
95
surfactants. In biotin rich condition, the production of L-glutamate in biotin rich
condition was not achieved because of a change in the aerobic metabolism of glucose or
any other carbon source used during fermentation, but not due to the consumption of
glutamate or its precursors for the synthesis of cellular constituents and increased amount
of biomass (Shiio et al., 1962).
This study confirmed that a tiny amount of biotin is required for L-glutamate
production by M. glutamicus, and the optimization of biotin concentration in fermentation
medium is mandatory for the enhancement of L-glutamate yield. The appropriate amount
of biotin required for the efficient growth of cells as well as for the highest production of
glutamic acid is 1.0µg l-1 that led to the production of 57mM L-glutamate after 52h of
cultivation (Figure 3.5). Although the highest production of glutamic acid was obtained
by cultivating the C. glutamicum strains in presence of 2.5-3µg l-1 biotin (Shiio et al.,
1962), this study demonstrated that L-glutamate production at 1.0µg l-1 biotin was
approximately 7-fold higher than that of L-glutamate measured at 2.5µg l-1 biotin. The
limiting concentration of biotin required for L-glutamate production depends on the type
of strain, the concentration and nature of carbon sources, however, it is generally
somewhat below 5µg l-1 (Kinoshita et al., 1985). Even though 20g l-1 biotin was observed
to be sufficient to allow the growth of C. glutamicum, glutamate production was only
observed at a biotin concentration of 3g l-1 (Takinami et al., 1966). However, this study
showed that the production of this amino acid reduced extremely by the increase of biotin
concentration ( 1µg l-1) in the CGXII Minimal Medium (Figure 3.5a). These results
might have occurred due to the changes in the metabolic activity of cells caused by the
addition of excess amount of biotin resulting in a decrease in the membrane permeability
of M. glutamicus to L-glutamate.
This study confirmed that the production of L-glutamate by M. glutamicus is only
occurred during biotin limited condition. This phenomenon might have observed due to
the fact that acetyl-CoA that participates in fatty acid synthesis is inactivated because of
limited amount of biotin inhibiting the synthesis of cell wall, and hence the plasma
membrane became permeable to L-glutamate. In addition, a very low activity of -
96
ketoglutarate dehydrogenase is observed in samples cultivated under biotin limitation
directing the metabolism of this bacterium to the synthesis of L-glutamate (Kramer,
1994). In this study, glucose was used as the only carbon source in fermentation medium.
However, it has been demonstrated that the concentration of biotin required for cell’s
growth and L-glutamate production is dependent on the carbon sources supplied in
fermentation medium (Shiio et al., 1962). Their results showed that 3µg biotin per liter of
glucose based fermentation medium led to the maximum production of L-glutamate
(122µM ml-1), whereas the same amount of biotin excreted only 55µM ml-1 L-glutamate
where acetate was used as the main carbon source. However, due to time limitation, the
influence of other carbon sources to L-glutamate production by this bacterium was not
investigated in this study.
The cultivation of C. glutamicum under biotin limited condition is often restricted
in industry since the molasses based fermentation medium (contains high amount of
biotin) is generally used for L-glutamate production (Ikeda, 2002). However, this study
confirmed that the production of L-glutamate could be achieved by the addition of Tween
40 into a biotin rich fermentation (CGXII Minimal) medium. When an appropriate
amount of Tween 40 (1.0g l-1) was added, irrespective of the time (both start and
exponentially) of addition into the CGXII Minimal Medium containing 200µg l-1 biotin,
the growth of M. glutamicus and glucose consumption were observed to reduce as
compared to the control (without surfactant). After the addition of Tween 40, cell’s
growth was almost stopped or grew at a slower rate for a while, however, cells started to
grow exponentially after few hours of cultivation (Figure 3.7 and 3.8). Nevertheless, Lglutamate accumulated considerably or at least to a certain degree due to Tween 40
addition although the fermentation medium contained sufficient amount of biotin. This
study clearly demonstrated that Tween 20 and 80 are not suitable for the production of
this amino acid (Figure 3.9 and 3.10), whereas significant amount of L-glutamate (47mM
after 72h) was obtained due to the addition of Tween 40 (2.0g l-1, at the exponential
growth phase) in a biotin rich (200µg l-1) cultivation of M. glutamicus (Figure 3.10).
97
This study demonstrated that the growth of cells (Figure 3.7 and 3.8) and Lglutamate production (Figure 3.9 and 3.10) due to Tween 40 addition are dependent on
the concentration and the addition time of this surfactant, regardless of the concentration
of biotin available in the CGXII Minimal Medium. Takinami et al. (1965; 1968) also
reported that the time at which surfactants are added into a biotin rich cultivation of B.
lactofermentum caused a remarkable change in the growth of this industrially important
microorganism as well as on the yield of L-glutamic acid. In the case of Tween 40
addition at the start of fermentation, the amounts of L-glutamate measured after 48h of
cultivation were 19, 22, 19 and 17mM (Figure 3.9) in presence of 1, 2, 3 and 4g l-1 Tween
40, respectively, whereas it was 39, 45, 40 and 36mM (Figure 3.10) due to the addition of
similar concentrations of Tween 40 to the exponentially grown M. glutamicus. In both
cases, the results clearly showed that the enhancement of L-glutamate production was not
possible by increasing the concentration of Tween 40 to 2.0g l-1. Furthermore, it has
been reported that higher concentration of surfactant than the optimum amount leads to
cellular death, and results in decrease of L-glutamate production (Hashimoto et al.,
2006).
Similar to this study, the concentration and addition time of Tween 40 to a biotin
rich cultivation of B. lactofermentum have been observed as a decisively important factor
for the production of L-glutamate. The highest amount of L-glutamate (62g l-1) was
obtained at 4g l-1 Tween 40 after 51h of fermentation, where this surfactant was added
after about 10h of cultivation (Shiratsuchi et al., 1995). Like Tween 40, the enhancement
of glutamic acid production was not possible by increasing the concentration of Tween 60
more than 2.0g l-1. The highest yield of glutamic acid (50%) was obtained by adding
1.0mg ml-1 of Tween 60 to a growing culture of B. lactofermentum cultivated on biotin
rich fermentation medium (Takinami et al., 1965; 1968). Their results also demonstrated
that the effect of surfactant addition on the growth of B. lactofermentum was less due to
delay in addition, and resulted in decrease of L-glutamate accumulation. On the other
hand, adding of Tween 60 earlier than the optimum time inhibited the growth, and
consequently reduced the yield of L-glutamic acid. However, due to time constraint, Lglutamate production by Tween 60 was not possible in this study.
98
Similar to the Tween 40 addition, this study showed lower cell densities and
slower rate of glucose consumption in cultures treated with a range of EMB
concentrations as compared to the control (Figure 3.11). These results might have
occurred because addition of EMB inhibits arabinofuranosyl transferase resulting in
decrease of the amount of covalently-bonded mycolate in the outer layer of Mycobacteria
and related species (Takayama et al., 1979). Light microscopy showed that C.
glutamicum cells change their morphology in the presence of EMB. Furthermore, the
content of arabinose, mycolic acids and the specific growth rate of C. glutamicum were
observed to reduce extremely in the presence of 50mg l-1 EMB (Radmacher et al., 2005).
However, this study showed that the production of L-glutamate increased due to the
increase of EMB concentration (up to 100mg l-1) in the CGXII Minimal Medium. After
48h of fermentation, the amount of L-glutamate measured in flasks containing 100mg l-1
EMB was 49mM, whereas it was only 0.14mM in control (Figure 3.12). Since
ethambutol inhibits the biosynthesis of bacterial cell wall, an incomplete cell wall in
presence of this antibiotic might be formed leading to the secretion of L-glutamate.
Moreover, the results showed that the optimum amount of ethambutol needed for the
highest production of L-glutamate is 100mg l-1, and the enhancement of L-glutamate
production was not possible by increasing the concentration of ethambutol more than
100mg l-1 (Figure 3.12).
In this study, however, low production of L-glutamate as compared to the yield of
production obtained by the other research groups (Radmacher et al., 2005; Shiio et al.,
1962; Shiratsuchi et al., 1995) was observed. These results might have occurred because
all the fermentation experiments were carried out in shake flasks that do not allow
researchers to optimize the cultivation parameters properly, resulting in decrease of Lglutamate yield. However, process variables, such as temperature, aeration, agitation and
pH play a significant role in fermentation. It has been reported that the rate of oxygen
transfer in shake flask is dependent on the design of flasks, speed of shaking and culture
volume (Ikeda, 2002). The appropriate culture volume required during shake flasks
fermentation is generally determined by the volume of flask. In this research, the growth
studies and L-glutamate production in M. glutamicus were performed in 250ml and
99
500ml shake flasks, respectively, where the medium volume in both conditions were only
50ml. Nevertheless, Ikeda (2002) mentioned that the volume of medium must be less than
70ml in the case of a standard 250ml flask, whereas it should not be more than 200ml in
the case of 1L flask.
In this study, all the experiements were carried out at 220rpm in an orbital
incubator shaker. However, it has been demonstrated that the growth properties of
microorganisms are affected by the speed of agitation during shake flask cultivations
(Ikeda, 2002). While conducting the L-lysine fermentation by the shake flask cultivations
of C. glutamicum, substrate consumption was observed to increase with the increase of
agitation speed from 50 to 300rpm (Shah et al., 2002). Although higher oxygen transfer
rates are usually achieved by continuous shaking of flasks, the supply of oxygen is
observed to be limited while the oxygen demand exceeds the oxygen transfer capacity.
This occurrence often gives inaccurate specific growth rates of microorganisms, and
finally decreases product and biomass yields (Ikeda, 2002). Hence, the production of
amino acids is currently achieved by fed-batch and continuous fermentations
(Chassagnole et al., 2003; Sassi et al., 1998; Kiss and Stephanopoulos, 1991; 1992).
However, it was not possible to investigate these approaches further due to the limitation
of time. Furthermore, this study demonstrated that production of L-glutamate after 72h of
fermentation is a bit lower than that of L-glutamate obtained after 48-52h. This result
might have occurred due to the fact that B. flavum is capable of growing on glutamic acid
as a carbon and nitrogen source (Shiio et al., 1982).
Apart from the treatments applied in this research, penicillin has been used to
induce the production of L-glutamate in which this antibiotic is usually added into the
fermentation medium before the microbial growth reaches its maximum limit (Demain
and Birnbaum, 1968). Like surfactants, a combination of both the concentration and the
time at which penicillin is added in fermentation medium have a great influence on Lglutamate production (Shiio et al., 1963). Adding 0.5U ml-1 penicillin at the start of
fermentation produced approximately 81mM l-1 of L-glutamate after 48h of cultivation,
whereas the production increased up to 100mM l-1 in presence of 2.0U ml-1 penicillin
100
added after 9h of cultivation. However, it has been confirmed that this enhancement was
not caused by increase of penicillin concentration in fermentation medium since the
addition of 2.0U ml-1 penicillin at the start of fermentation produced only 2.9mM l-1 of Lglutamate (Shiio et al., 1963). L-glutamate secretion has also been triggered in C.
glutamicum by applying different osmotic gradients combined with the addition of
1.3mM tetracaine (Lambert et al., 1995).
It has also been demonstrated that the temperature at which C. glutamicum is
grown has an influence on the secretion of L-glutamate. Due to the temperature upshift
after a certain hours of cultivation of C. glutamicum 2262, the activities of OGDHC and
pyruvate dehydrogenase were observed to decrease redirecting the flux of 2-oxoglutarate
towards L-glutamate production (Uy et al., 2003). In another investigation, 85g l-1 Lglutamate was achieved after 24h of fed-batch cultivation of C. glutamicum (temperaturesensitive strain) by changing the culture temperature from 33C (initial growth phase) to
39C (production phase) (Delaunay et al., 1999b). A 6-fold increase in the efflux rate of
glutamate (6mM g Dw-1 h-1) is observed when the glutamate export system has been
activated by a temperature shift during the temperature triggered process (Delaunay et al.,
1999b; Lapujade et al., 1999). However, due to time limitation, L-glutamate production
by the addition of penicillin/tetracaine or by increasing the growth temperature of M.
glutamicus was not investigated in this study. Apart from the L-glutamate production
facilitated by the above-mentioned treatments through the cell wall of C. glutamicum,
genetic or metabolic engineering has been applied in order to increase the production of
this amino acid. The amplification of glutamate dehydrogenase (GDH) activity resulted
in increase of intracellular glutamate concentration (Bormann-El Kholy et al., 1993).
Kimura (2002b) showed higher production of L-glutamate in the mutant lacking of 2oxoglutarate dehydrogenase (OGDHC) activity, confirming that the glutamate efflux
might be occurred by a change at the 2-oxoglutarate branch point.
However, this study concluded that L-glutamate production is not possible
without treating the cell membrane of M. glutamicus by the addition of agents i.e., limited
amount of biotin or surfactant or ethambutol. The secretion of this amino acid might have
101
occurred due to the formation of an incomplete cell membrane by the above-mentioned
treatments increasing the membrane permeability of M. glutamicus to L-glutamate.
During biotin limitation or surfactant or ethambutol addition, it is evident that a
modification in phospholipids of plasma membrane i.e., increase in the ratio of saturated
to unsaturated fatty acids (Demain and Birnbaum, 1968; Marquet et al., 1986), or a
decrease in the content of phospholipids (Clement and Laneelle, 1986; Huchenq et al.,
1984) or a decrease in total lipids content (Hoischen and Kramer, 1990) must occur
resulting the secretion of L-glutamate. However, this study showed that the production of
L-glutamate in biotin limited condition was considerably higher than that of L-glutamate
obtained from the other two conditions since the amounts of glutamic acid measured after
48h fermentation of M. glutamicus were 56, 45 and 49mM under the condition of biotin
limitation, surfactant addition and ethambutol addition, respectively. Hashimoto et al.
(2006) demonstrated that both the cellular constituents of Corynebacterineae synthesized
from fatty acids and the content of mycolic acid decreased under the all conditions
generally used for L-glutamate overproduction. They showed a similar amount of Lglutamate production i.e, 121, 122, 101 and 113mM under the supply of limited amount
of biotin, the addition of Tween 40, penicillin G and cerulenin into a biotin rich medium,
respectively.
Nampoothiri et al. (2002) reported that the intracellular concentration of Lglutamate decreased constantly due to it’s secretion to the extracelular medium, and
thereby the feedback inhibition that regulates the internal glutamate pool is relaxed while
cultivating C. glutamicum in presence of the above-mentioned conditions. Hoischen and
Kramer (1989) demonstrated that change in the lipid state of C. glutamicum membrane is
not the only parameter for inducing glutamate secretion. For effective efflux of Lglutamate, a mechanism for crossing the permeability barrier of the plasma membrane
must be present. It has been also reported that the reduced phospholipids content of the
cytoplasmic membrane due to lack of biotin does not lead to the permeabilization of cells
although a limited supply of biotin is necessary for the induction of L-glutamate
secretion. Furthermore, it is not clear whether L-glutamate secretion is linked to an
alteration in membrane biosynthesis due to these treatments and/or any unknown
102
regulatory effects on the level of transcription and translation (Hoischen and Kramer,
1990). Over the past several years, three models have been described as the essential
steps for L-glutamate efflux, such as diffusion, functional inversion of uptake systems
and the presence of specific excretion systems (Kramer, 1994).
Eggeling and Sahm (1999) demonstrated that three components are involved in Lglutamate secretion of C. glutamicum, such as oxoglutarate dehydrogenase activity, the
presence of a specific exporter and the membrane status. Although all the treatments are
mainly involved in the alteration of bacterial membrane, Nampoothiri et al. (2002)
demonstrated that some additional parameters must be considered for increasing the Lglutamate efflux, such as the energetic state of cell, a low α-ketoglutarate dehydrogenase
activity, the carrier itself and an altered permeability of mycolic acid layer. It is now
obvious that the enhancement of membrane permeability, identification of limiting steps,
eliminating the feedback and regulatory controls, changing the cellular metabolism of C.
glutamicum and developing of a cost effective production process are the main
prerequisites to achieve high yield of amino acids. However, intensive researches are still
required in order to examine the L-glutamate production by the different strains of C.
glutamicum. It is expected that this research provided substantial knowledge regarding
the different approaches usually used for L-glutamate production by Corynebacteria, and
will assist in improving the yield and productivity of glutamic acid by metabolic
engineering and functional genomics.
The isolation, purification and concentration of many biomolecules produced in
fermentation processes are extremely important since downstream processing often
contributes a large portion of the product cost. In the case of fermentative production of
L-glutamate, the downstream processing costs are approximately 30-40% of the selling
price (Hacking, 1986). The purification process of L-glutamate developed in this study is
mainly based on centrifugation. Using this conventional method, the problems associated
during scale-up are enormous, and eventually leads the recovery processes uneconomical
and expensive unless the product is of very high market value (Hermann, 2003).
However, the application of suitable methods for purifying the desired product depends
103
on several factors, such as the physico-chemical properties of product (solubility,
isoelectric point), composition of process liquid (quality and quantity of by-products),
environmental regulations (waste liquor treatment) and the uses of product (feed or
pharmaceutical use). Furthermore, the raw materials that are used in fermentation media
have a major influence on the downstream processing since the removal of unused
components of molasses make the process expensive (Hermann, 2003). As the purity of
amino acids or any other therapeutic products is a major concern for the biotech industry,
it is mandatory to develop an economically viable recovery process with the highest level
of purity. Nevertheless, the development of sophisticated chromatographic methods and
crystallisation techniques will increase the efficiency of downstream processing.
104
Chapter FOUR
4. Enhancement of the Secretion of L-glutamate
Produced under Biotin Limited Fermentation of M.
glutamicus by Electropermeabilization
4.1. Introduction
Electroporation or electropermeabilization is a well-established approach used in
molecular biology where cell membrane is exposed to high intensity electrical pulses
with several milliseconds duration in order to increase the membrane permeability for
exogenous molecules (Chang et al., 1992; Neumann et al., 1989; Teissie et al., 1999;
2002; 2005). This approach is an elegant way to gain access into the cytoplasm of cells
(Teissie et al., 1999). The electric pulses may form transient pores on cell membrane
depending mainly on the electric field strength in which the entry of foreign molecules
(DNA, RNA and proteins) into cells could be achieved (Faurie et al., 2005; Golzio et al.,
2004; Weaver and Chizmadzhev, 1996). This technique has been used for inserting the
foreign genes into microorganisms (Potter, 1993; Somiari et al., 2000; Rols, 2006).
Moreover, it’s application in medical sectors i.e., electrochemotherapy, transdermal drug
delivery and gene transfer is noteworthy (Belehradek et al., 1993; Dev et al., 2000; Heller
et al., 1999; Mir and Orlowski, 1999; Orlowski and Mir, 1993). Pulsed electric field is
also regarded as a non-thermal process to kill spoilage microorganisms (Schoenbach et
al., 2000; Wouters and Smelt, 1997). Although this approach has been successfully
applied in gene transfer, medical sector and bacterial sterilization purposes, there are few
reports published in literature revealing it’s application in bioprocess intensification.
Electropermeabilization is based on the dielectric properties of a biological
membrane. The cell membrane is composed of phospholipids and amphipatic molecules
having a hydrophilic head group attached to a hydrophobic tail. These molecules are
105
observed to be polarized due to electric fields (Kinosita and Tsong, 1977). The
application of external electric field generates a transmembrane potential (TMP)
difference because of the dissimilarity in conductivity across the cell wall that results in
formation of transient electropores, and eventually increase the membrane permeability
of cells (Hapala, 1997). However, a prominent structural alteration i.e., asymmetry in
bilayer membrane of phospholipids with the formation of many transient electropores is
usually observed within a short time (ms) of pulsation (Chernomordik et al., 1987; Haest
et al., 1997). These pores transfer ions across the cell wall, allows introducing the small
and large molecules into the cytoplasm and permits the insertion of proteins into cells
although the membrane usually represents a considerable barrier for them in its normal
state (Hapala, 1997; Somairi et al., 2000; Tessie et al., 1999; Tsong, 1991). However, the
amount of molecules that will be transported depends on their size, extracellular or
intracellular concentration of molecules and on the degree of permeabilization (Ho and
Mittal, 1996).
The appearance of electropores on cell membrane is dependent on the pulse
intensity, whereas the dynamics of pores (expansion of electropores, their stabilization
and resealing) depends on pulse duration and membrane physical conditions, for instance
medium conductivity (Rols et al., 1990). In addition, the duration of pulses (defined by
pulse decay halftime) is also dependent on the capacitance of loaded capacitors and the
conductivity of pulsing medium (Djuzenova et al., 1996; Pucihar et al., 2001). Within a
certain limit, a longer pulse duration may compensate with lower field strength and vice
versa in order to obtain same level of permeabilization (Rols and Teissie, 1989). It has
been demonstrated that pulses with short duration (100µs) and high field strength
(>700Vcm-1) are the most favourable for the delivery of anti-cancer drugs in solid tumors
(Hofmann et al., 1999), whereas longer pulses (20-60ms) and low field strength (100200Vcm-1) are suitable for DNA delivery in skeletal muscle (Smith and Nordstrom,
2000). Furthermore, it has been shown that the temperature-dependent resealing process
as well as the amount of cells permeabilized and survived after the treatment depends on
the electric field and electroporation medium (Rols and Teissie, 1989).
106
In the case of reversible electroinduced membrane permeabilization, the
incorporation of DNA or foreign molecules into cells and the exchange of
macromolecules across the cell membrane occur for a very short period during and
immediately after the pulsation (Chang et al., 1992; Tsong, 1991; Weaver, 1995; Weaver
and Chizmadzhev, 1996). The transport of molecules across the membrane mainly occurs
via three different mechanisms, diffusion, electrophoresis and electroosmosis (Tekle et
al., 1994). In general, these processes keep going until the cell membrane is resealed
completely. The involvement of each mechanism is dependent on the pulse length and
amplitude as well as on the type of molecule being transported (Puc et al., 2003). Mir et
al. (1988) demonstrated that the direct access of nonpermeant molecules into cell’s
cytoplasm is mostly caused by passive diffusion. Although it has been reported that
diffusion is the main mechanism involved in transmembrane transport of small molecules
into cells (Sixou and Teissie, 1993; Tekle et al., 1990), Dimitrov and Sowers (1990)
proposed that electroosmosis is responsible for the molecular exchange of small
molecules into cells. On the other hand, endocytosis-like processes were considered to be
the possible mechanisms for the uptake of larger molecules (Lambert et al., 1990).
Furthermore, Gabriel and Teissie (1999) observed that the exchange of calcium ions
through the electropermeabilized membrane of CHO cells during a millisecond pulse is
mostly caused by diffusion-driven processes i.e., governed by concentration gradient.
Furthermore, electrophoresis has been found to play a major role in transporting
of macromolecules, particularly DNA (Klenchin et al., 1991; Satkauskas et al., 2002).
Somiari et al. (2000) demonstrated that the mechanism of electroporation-mediated gene
transfer is based on the formation of electropores on the cell membrane followed by DNA
electrophoresis into the cells. It has been reported that the electrophoretic transport of
charged molecules only occurs during the pulses (100s), whereas the transport of other
molecules through diffusion takes place throughout the lifetime of permeabilized state
(Mir, 2000). Similarly, the electroefflux of calcein (a fluorescent molecule) is mostly
caused by electrophoresis or electroosmosis during a pulse, whereas the transport of
calcein after pulsation is partially or completely caused by diffusion (Prausnitz et al.,
1995). It is, however, uncertain whether the transformation or transfection is achieved
107
due to permeabilization itself or rather due to the subsequent electrophoresis and
electroosmosis caused by very long pulse duration. Hence, there is still plenty of research
needed for the investigation of mechanisms involved in transporting molecules across the
membrane during electroporation
Microorganisms, both bacteria and yeast have become a suitable host for the
production of recombinant proteins with biotechnological and pharmacological
applications. However, the recovery of recombinant proteins from host cells is always a
significant challenge for bioprocess industry since it is very unusual that the foreign
proteins are simply secreted in the extracellular medium (Hermann, 2003). Therefore, the
disruption of cells is usually performed by ultrasonication or homogenization, physiochemical (autolysis by solvent, membrane disintegration, pH and osmotic shock) and
enzymatic (zymolase, lyticase) treatments for the recovery of recombinant proteins
(Hermann, 2003). Using these conventional techniques, however, a complete destruction
of cells is required that causes the purification process complicated, and eventually leads
to an expensive bioprocess (Teissie et al., 2002). Therefore, alternative methods for the
extraction of enzymes or proteins are needed in order to ensure a higher selectivity of
their release and to preserve their activity at a maximal level.
Ohshima et al. (1999) extracted nucleic acid molecules from the recombinant E.
coli within 1min of electropulsation. Like bacteria, the efflux of proteins or enzymes
during or after electroporating of yeast has been reported in several studies (Ganeva et
al., 1995; 2001; 2003; 2004; Ohshima et al., 1995). Ohshima et al. (1995) demonstrated
the release of invertase and alcohol dehydrogenase due to electropermeabilization of S.
cerevisiae at field strengths of 6 and 12kV cm-1, respectively, indicating that electric
pulses could lead to the selective release of intracellular proteins or enzymes from cells.
The application of high intensity electric pulses (2 × 9 pulses, 990ms duration, 2.75kV
cm-1) to a yeast (S. cerevisiae) suspension showed a considerable release of some
cytoplasmic proteins, glutathione reductase (GLR), 3-phosphoglycerate kinase (PGK)
and alcohol dehydrogenase (ADH) after 3-8h of pulsation (Ganeva and Galutzov, 1999).
Similarly, the electroporation of E. coli, Listeria innocua and S. cerevisiae resulted in a
108
leakage of intracellular compounds (ATP and UV-absorbing substances) into the
extracellular medium (Aronsson et al., 2005). -galactosidase was extracted from
Kluyveromyces lactis by a series of electric pulses (2ms duration, 1Hz frequency, and 44.5kV cm-1 field strength) within 8h of electropermeabilization (Ganeva et al., 2001). In
another study with S. cerevisiae, approximately 80-90% of intracellular enzymes i.e.,
hexokinase, PGK and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) are
obtained by applying a series of electric pulses (Ganeva et al., 2003).
Similarly, Ganeva et al. (2004) observed a release of intracellular enzymes
(GAPDH and PGK) by applying high intensity electric pulses to the yeast,
Schizosaccharomyces pombe. Furthermore, it has been demonstrated that the specific
activities of these electroextracted enzymes are higher than those obtained by mechanical
disintegration or enzymatic lysis (Ganeva et al., 2003). The secondary metabolites (ionic
betalains, mainly negatively-charged betanin from Beta vulgaris and ionic alkaloids,
particularly positively-charged ajmalicine and yohibine from Catharanthus roseus) from
plant cells have also been extracted by using electric pulses, whilst maintaining the
viability of cells (Yang et al., 2003). From the literature, it is obvious that electroporation
creates transient pores on the cell membrane of living organisms, and the part of the
plasma membrane that is permeabilized due to electric pulses provides a path in order to
transfer the different molecules across the cell membrane. However, the quantity of
molecules that will be introduced or extracted into or from cells, respectively, depends on
their size, concentration and the degree of permeabilization of cell membrane (Trissie et
al., 1999).
It is not usual that all the primary or secondary metabolites of commercial interest
are excreted by cells into the extracellular medium. Furthermore, they may be stored
intracellularly or in certain cases, the permeability barrier of cell wall causes low reaction
rates that result in decrease the product yield. Hence, the cell wall porosity has been
regarded as an important factor for the recovery of proteins from microorganism (Ganeva
et al., 2001; White and Marcus, 1988). When the plasma membrane loses its barrier due
to electric pulses, the efficiency of protein efflux is controlled by the cell wall porosity as
109
well as by the size and electric charge of proteins (De Nobel and Barnett, 1991). During
pulsation, various substances (chemical permeabilizing agents) that alter the structure of
cell membrane are used in order to increase the efficiency of electroporation. These
substances often casue extensive cell injury, and result in releasing of specific
intracellular constituents (De Nobel et al., 1989). Hapala (1997) demonstrated an
improved uptake of substrate and product excretion across the cell membrane without
killing cells by the addition of cell wall permeabilizing agents (for instance, detergent)
into the cell suspension before electropulsation.
During L-glutamate production by C. glutamicum, the extracellular transport of
this amino acid is the major rate limiting factor affecting both the yield and productivity
of amino acids (Kramer, 1994). Extensive efforts on the improvement of these bacterial
strains for amino acids overproduction have been directed towards deregulation of the
corresponding pathways by classical mutagenesis and screening procedures (Kinoshita
and Nakayama, 1978). Although the increase of yield and productivity of amino acids are
accomplished through the use of auxotrophic and regulatory mutants, their application in
the food and beverage industry is generally unacceptable (Parekh et al., 2000). Hence,
much attention is required in order to develop a rapid method for reducing the
permeability barrier of these bacteria, and to increase the yield of amino acids. The recent
development of electroporation in bioprocessing has given us the opportunity to apply
this technique for the increase of L-glutamate production by M. glutamicus fermentation.
The goal of this study is to enhance the membrane permeability of L-glutamate
through the electropermeabilization of M. glutamicus cultivated under the different
growth conditions (i.e., biotin limitation, surfactant addition and ethambutol addition) by
which glutamic acid is excreted into the extracellular medium (Chapter 3). In order to
achieve this, the fermentative culture of M. glutamicus cells grown to production phase
(stationary growth period) will be imposed by a range electric field strengths. The effect
of electric pulses on the release of glutamate dehydrogenase (GDH), malate
dehydrogenase (MDH) and total protein will also be examined. The effect of
physiochemical [(providing resting time between pulses, controlling the temperature of
110
pulsed samples at 4C in order to prolong the permeabilized state and treating the cells by
chemical permeabilizing agent (D-L-1, 4-Dithiothreitol, DTT)] and cellular (using a
Gram-negative strain, E. coli) factors to the electroextraction of L-glutamate, GDH,
MDH and total protein will also be investigated. In addition, the effectiveness of this
new approach for release intracellular protein or enzymes will be compared with the
established techniques i.e., sonication and French press that are routinely used in biotech
industry. The ultimate purpose of this research is to gain sufficient knowledge for
improving the secretion or extraction of intracellular product by the transient
electropermeabilization, and thus intensify the bioprocesses.
111
4.2. Material and Methods
4.2.1. Chemicals
Peptone from pancreatically digested casein and meat extract were obtained from
VWR (Merck, UK). Yeast extract and bacteriological agar were procured from Oxoid,
UK; D-glucose, urea, NaCl and all other chemicals were purchased from Fisher, UK
unless otherwise mentioned.
4.2.2. Organism and cultivation
The bacterial strains used for this study were M. glutamicus DSM 20300 (Collins
et al., 1977; Suzuki et al., 1981; Yamada and Komagata, 1970; Yamada et al., 1976) and
E. coli DSM 498 (Farnleitner et al., 2000) supplied by the German Collection of
Microorganisms and Cell Culture (DSMZ-Deutsche Sammlung von Mikroorganismen
und Zellkulturen GmbH). M. glutamicus DSM 20300 was cultivated in Medium 53,
whereas E. coli DSM 498 was cultivated in Nutrient Medium. In the case of L-glutamate
production by M. glutamicus, inoculum was prepared by selecting single colony from the
stock agar plate and transferring into 3.5% BHI Medium. In order to investigate the
efficiency of electroporation on the release of protein or enzymes, M. glutamicus was
grown on Seed Medium, whereas E. coli was grown in Nutrient Medium. The
composition and preparation of all the media and stock agar plates used in this study were
described previously in Section 3.2.2. M. glutamicus was cultivated at 30C and 220rpm,
whereas E. coli was cultivated at 37C and 150rpm.
4.2.3. L-glutamate production in M. glutamicus under biotin limited condition
L-glutamate was produced in M. glutamicus by cultivating cells in biotin limited
CGXII Minimal Medium that has been used by Keilhauer and his co-workers (Keilhauer
et al., 1993). The preparation of CGXII Minimal Medium was described previously in
Section 3.2.5.
4.2.4. Electroporator and pulse treatment
For electroporation experiments, cell suspension (fermentation broth) is usually
placed between two electrodes connected to the generator of a high-voltage electric field.
112
A pulse with an intensity of several kilovolts per centimeter (kV cm-1) leads to membrane
permeabilization at the sites where TMP is observed to be its highest level. The
exponential decay pulses were generated by a Gene PulserTM apparatus (Bio-Rad
Laboratories, USA) that allows us to apply a capacitance of 3 to 25µF and voltage of 0.2
to 2.5kV. The output of pulse generator was directed through a Pulse Controller unit
(Bio-Rad) having a selection of resistors of 100 to 1000. The effective resistance used
in parallel with the electrodes determines the time constant of pulse (for example, 200
with the capacitor of 25µF gives a 5ms time constant). Cuvettes (Bio-Rad) with an
electrode gap of 0.2cm were used to achieve the desired level of field strengths. Samples
(fermentation broth) were collected after 24 and 48h of cultivation and transferred into
Bio-Rad electroporation cuvettes, and subsequently pulsed with a range of voltages (1.5,
2.0 and 2.5kV that produce field strengths of 7.5, 10.0 and 12.5kV cm-1) although the
capacitance (25F) and parallel resistance (200) were
kept constant in all the
experiments. The number of samples pulsed for increasing the secretion of of Lglutamate and the release of total protein and enzymes on each time was three, and each
sample was pulsed by a single to multiple pulses (up to 5). Electroporation was
performed at RT (22  1°C) by transferring 0.4ml of cell suspension into 0.2cm gap
electroporation cuvettes.
4.2.5. Quantification of glucose, L-glutamate and other amino acids
The quantification of substrate, L-glutamate and all the other amino acids were
achieved with the use of AAA-DirectTM Amino Acid Analysis System (Dionex, UK). The
preparation of eluents and standard were described previously in Section 3.2.8.
4.2.6. Preparation of crude extract
The crude extract of M. glutamicus was prepared by two different methods i.e.,
sonication or French press. A schematic diagram of these procedures is as follows-
Sonication
Fermentation broth (50ml)
113
Centrifuged at 4500g for 10min at 4C
The supernatant was discarded, and cell pellets were washed twice with 0.2% KCl
The pellets (1g Dw) were resuspended in 4ml of buffer (100mM Tris-HCl +
2.5mM MgCl2, pH 7.2).
The resulting cell suspension was disrupted by sonication (Fisher, UK) for 15min
(samples were kept in ice bath while sonicating)
Sonicated samples were centrifuged at 4600g for 10min at 4C
The supernatant was then collected for determining the concentration of total
protein, GDH and MDH.
French press
Fermentation broth (50ml)
Centrifuged at 4500g for 10min at 4C
The supernatant was discarded, and cell pellets were washed twice with 0.2% KCl
The pellets (1g Dw) were resuspended in 4ml of buffer (100mM Tris-HCl +
2.5mM MgCl2, pH 7.2).
The resulting cell suspension was disrupted by French press (Aminco, USA) at
20000psi
Sonicated samples were centrifuged at 4600g for 10min at 4C
114
The supernatant was then collected for determining the concentration of total
protein, GDH and MDH.
4.2.7. Determination of total protein, GDH and MDH
Analysis of Total Protein (TP) in cell suspension
Principle
The Bradford Reagent (Sigma-Aldrich, UK) was used to determine the
concentration of total protein in cell suspension. It is the most widely employed method
for determining protein concentration. The reagent is consisted of Coomassie Brilliant
Blue G-250 solubilized in phosphoric acid and methanol. After the addition of this
reagent in samples that contained proteins, a complex between the dye and proteins is
formed in the reaction mixture. The dye in solution is in the cationic form and has an
absorption maximum at 465nm (red). When the dye binds with proteins, both the
hydrophobic and ionic interactions stabilize the anionic form of the dye, and cause a
visible color change. The maximum absorbance of Brilliant Blue G in an acidic solution
of is obtained at 595nm. The amount of absorption is proportional to the proteins
available in supernatant (Bradford, 1976). The protein assay using this reagent has great
advantages over the other assays, such as Lowry and BCA methods. There is no need to
dilute this reagent, and it is compatible with the reducing agents (i.e., dithiothreitol and 2mercaptoethanol) that are often used to stabilize the proteins in solution. The protein
assay by this method is less susceptible to interfere by various chemicals that are
available in samples.
Reagents and Equipment
 Bradford Reagent (B6916, Sigma-Aldrich, UK).
 Protein (Bovine Serum Albumin, BSA) standard solution (P0834, Sigma-Aldrich,
UK).
 Deionized H2O: Milli Q water (Direct Q5, Millipore, USA).
 Spectrophotometer (S1200, WPA).
 Test tubes.
 Macro disposable polystyrene 4.0ml cuvettes (FB55143, Fisher, UK).
115
Procedure
The Bradford Reagent in the bottle was gently mixed and kept at RT prior to use.
The assay was performed by adding 3ml of Bradford Reagent into 0.1ml of standard or
sample per test tube. A standard curve of protein was prepared by diluting the protein
standard (BSA) to 0.25, 0.5, 0.75 and 1.0mg ml-1 using deionized water (Figure 4.1). A
blank assay was performed by adding deionized water, and the OD of blank was deducted
from all the readings obtained from the standard and samples.
Blank (ml)
Standard (ml)
Unknown samples (ml)
Deionized water
0.1
--
--
Standard (0.25-1.0mg ml-1)
--
0.1
--
Unknown samples
--
--
0.1
Bradford Reagent
3.0
3.0
3.0
y = 0.8976x + 0.0122
R2 = 0.9982
1.0
OD at 595nm
0.8
0.6
0.4
0.2
0.0
0.0
0.2
0.4
0.6
0.8
1.0
1.2
Concentration of BSA, mg/ml
Figure 4.1 Standard curve of total protein (BSA).
The above-mentioned volume (3.1ml) in test tubes was vortexed gently in order to
mix properly, and incubated at RT for 5 to 45min. The resulting solution was then
transferred into polystyrene cuvettes, and the absorbance was measured at 595nm. The
absorbance was recorded within 30min since the protein dye complex is usually stable up
116
to 60min (reported by the manufacturer). In the case of samples with unknown protein
concentrations, samples were diluted in order to keep the absorbance within the linear
range of 0.25-1.0mg ml-1. The total protein concentrations in unknown samples were
determined from the standard curve.
Analysis of malate dehydrogenase (MDH) in cell suspension
Principle
Malate dehydrogenase (MDH) is an enzyme in the citric acid cycle that catalyzes
the inter-conversion of L-malate and oxaloacetate using nicotinamide adenine
dinucleotide (NAD+) as a coenzyme. In this study, MDH was used as a marker enzyme in
cytoplasmic fractions.
MDH
L-malate + NAD+
Oxaloacetate + NADH + H+
Reagents and Equipment
 0.1M Phosphate buffer, pH 7.5: 10ml of 1M phosphate buffer (P3619, SigmaAldrich, UK) was diluted with 90ml of deionized water, and adjusted to pH 7.5.
 15mM Oxaloacetate: 19.81mg of oxaloacetic acid (O4126, Sigma-Aldrich, UK,
Mol. Wt. 132.07) was dissolved with 10ml of deionized water.
 14.11mM NADH: 10mg NADH (N1161, Sigma-Aldrich, UK, Mol. Wt. 709.4)
was dissolved in 1ml of deionized water.
 MDH (as a standard): Malic dehydrogenase from Thermus flavus (M7032, SigmaAldrich, UK).
 Deionized H2O: Milli Q water (Direct Q5, Millipore, USA).
 UV-Visible spectrophotometer (DR/2000, HACH, Germany).
 1.5ml disposable semi-micro PMMA UV grade cuvettes (CXA-110-040H, Fisher,
UK).
Procedure
This assay was performed directly in 1.5ml disposable UV visible cuvettes. All
the reagents (phosphate buffer, oxaloacetate and NADH) were added into the cuvettes
117
and equilibrated for 5min at RT. In the case of standard, a known concentration of MDH
was added, and the disappearance of NADH or the formation of NAD+ was measured at
340nm every 30sec for 2-3min by the UV-Visible spectrophotometer. Figure 4.2 shows a
standard curve of MDH that was prepared by plotting the ∆A340nm 30sec-1 at a range of
malate dehydrogenase concentrations (1-10U ml-1). A blank (deionized water) assay
without adding standard MDH or sample that contained MDH was performed in order to
investigate whether these reagents had any effect on the reduction of NADH to NAD+.
Blank (l)
Standard (l)
Unknown samples (l)
0.1M phosphate buffer
940
940
940
Oxaloacetate
30
30
30
NADH
10
30
30
Deionized water
20
--
--
Standard (1-10U ml-1)
--
20
--
Unknown samples
--
--
20
y = 0.0118x - 0.0021
R2 = 0.9985
Reduction of NADH to NAD+ (OD
difference at 340nm)
0.14
0.12
0.10
0.08
0.06
0.04
0.02
0.00
0
2
4
6
8
Concentration of MDH, U/ml
Figure 4.2 Standard curve of malate dehydrogenase (MDH).
118
10
12
The resulting solution was mixed immediately by inversion, and the decrease was
recorded in A340nm for approximately 2-3min. In the case of samples, appropriate dilution
was made in order to keep the ∆A340nm 30sec-1 within the values of standard curve. The
concentrations of MDH in unknown samples were calculated from the standard curve.
Analysis of L-glutamate dehydrogenase (GDH) in cell suspension
Principle
Glutamate dehydrogenase (GDH), located in the mitochondria, is an important
branch-point enzyme between carbon and nitrogen metabolism (Stillman et al., 1993).
GDH catalyzes the oxidative deamination of L-glutamate into -ketoglutarate and
ammonia. GDH is able to utilize both NADP+ and NAD+. NADP+ is utilized in the
forward reaction of -ketogluterate and free ammonia, which are converted to Lglutamate via a hydride transfer from NADPH to glutamate. On the other hand, NAD+ is
utilized in the reverse reaction, in which L-glutamate is converted to -ketoglutarate and
free ammonia via an oxidative deamination reaction (Stryer et al., 2002).
GDH
-KG + -NADH + NH4+
L-glutamate + -NAD + H2O
-KG = -Ketoglutarate
NH4+ = Ammonia
-NADH = -Nicotinamide adenine dinucleotide, reduced form
-NAD = -Nicotinamide adenine dinucleotide, oxidized form
Reagents and Equipment
 100mM
Triethanolamine
buffer,
pH
7.3
at
25C:
Weighed
1.8565g
triethanolamine hydrochloride (T1502, Sigma-Aldrich, UK, Mol. Wt. 185.65) in a
volumetric flask (100ml), and dissolved with deionized water up to the mark of
the flask. Adjusted to pH 7.3 at 25C with 1M NaOH (BPE358-212, Fisher, UK).
 200mM α-KG solution: Weighed 292.2mg -ketoglutaric acid (K1750, Sigma-
119
Aldrich, UK, Mol. Wt. 146.10) in a volumetric flask (10ml), and dissolved with
deionized water up to the mark of the flask. Adjusted to pH 6.5-7.5 using solid
sodium bicarbonate (S8875, Sigma-Aldrich, UK).
 3.2M Ammonium acetate solution (NH4OAc): 1.233g ammonium acetate
(A7262, Sigma-Aldrich, UK, Mol. Wt. 77.08) was dissolved in 5ml of deionized
water.
 10mM -NADH solution: 35.47mg -NADH, disodium salt (N8129, SigmaAldrich, UK, Mol. Wt. 709.4) was dissolved in 5ml of deionized water. A freshly
prepared solution was always used in this study.
 25mM Ethylenediaminetetraacetic acid (EDTA) solution: 104.05mg EDTA,
Tetrasodium Salt (ED4SS, Sigma-Aldrich, UK, Mol. Wt. 416.20) was dissolved
in 10ml of deionized water.
 L-Glutamic dehydrogenase enzyme solution: A standard curve having of 0.2-0.6U
ml-1 of L-glutamic dehydrogenase (G2501) was freshly prepared in cold Reagent
A.
 Deionized H2O: Milli Q water (Direct Q5, Millipore, USA).
 UV-Visible spectrophotometer (DR/2000, HACH, Germany).
 2.5ml disposable semi-micro PMMA UV grade cuvettes (CXA-110-025D, Fisher,
UK).
Procedure
A reaction cocktail was prepared by pipetting the following reagents into a 50ml
centrifuge tube.
Reagents
Amount (ml)
Reagent A (Buffer)
26.0
Reagent B ( -KG)
2.0
Reagent C (NH4OAc)
0.5
Reagent D (-NADH)
0.3
Reagent E (EDTA)
0.3
120
The resulting solution was mixed by stirring, equilibrated to 25C, and adjusted to
pH 7.3 at 25C with 1M NaOH or 1M HCl. In the case of standard, a known
concentration of GDH was added, and the disappearance of NADH or the formation of
NAD+ was measured at 340nm in every 1min for 3-5min by the UV-visible
spectrophotometer. Figure 4.3 shows a standard curve of GDH which was prepared by
plotting the ∆A340nm min-1 at a range of glutamate dehydrogenase (0.2-0.6U ml-1). A
blank (deoinzed water) assay without adding standard GDH or sample that contained
GDH was performed in order to investigate whether these reagents had any effect on the
reduction of NADH to NAD+.
Blank (ml)
Reaction cocktail
Standard (ml)
2.90
Unknown Samples (ml)
2.90
2.90
0.10
--
--
Reagent F (standard)
--
0.10
--
Unknown samples
--
--
0.10
Equilibrated to 25C and added
Deionized water
Reduction of NADH to NAD+ (OD
difference at 340nm)
0.06
y = 0.0796x + 0.0001
R2 = 0.9973
0.05
0.04
0.03
0.02
0.01
0.00
0.0
0.1
0.2
0.3
0.4
0.5
Concentration of GDH, U/ml
Figure 4.3 Standard curve of glutamate dehydrogenase (GDH).
121
0.6
0.7
The resulting solution was mixed immediately by inversion, and the decrease in
A340nm was recorded for approximately 3-5min. In the case of samples, appropriate
dilution was made in order to keep the ∆A340nm 1min-1 within the values of standard
curve. The concentrations of GDH in unknown samples were calculated from the
standard curve. One unit of GDH reduced 1.0μM of α -ketoglutarate to L-glutamate per
minute at pH 7.3 at 25C in presence of ammonium ions.
122
4.3. Results
4.3.1. Effect of electric pulses on the secretion of L-glutamate produced in M.
glutamicus
In order to investigate the efficiency of electropermeabilization for increasing the
production of L-glutamate, cell suspension of M. glutamicus (fermentation broth)
cultivated in CGXII Medium under biotin limited (1µg l-1) condition that induced Lglutamate secretion to the extracellular medium (Chapter 3.2.2) was collected after 24
and 48h of cultivation where sufficient amounts of L-glutamate were already determined
by HPLC. Appropriate amounts (400µl) of samples were transferred in 0.2cm
electroporation cuvettes, pulsed at 7.5, 10.0 and 12.5kV cm-1 where both the resistance
and capacitance were kept constant at 200 and 25µF, respectively. The number of
samples pulsed for increasing the secretion of L-glutamate on each time was three. The
results from the following experiements clearly showed that increasing the field strength
(voltage) and pulse number (up to 5 pulses) applied to the fermentation broth of M.
glutamicus, irrespective of the age of culture (both 24 and 48h), have no prominent
influence in enhancing the L-glutamate secretion (Figure 4.4 and 4.5). The maximum
increase (only 2.6% as compared to the control) of L-glutamate secretion was obtained in
samples treated at 12.5kV cm-1 by 4 pulses.
123
Glutamate, mM
31
29
27
25
Control
1
2
3
4
5
No. of pulses treated
Control
7.5kV/cm
10.0kV/cm
12.5kV/cm
Figure 4.4 Effect of electric field strengths on L-glutamate secretion (fermentation broth taken
after 24h of cultivation).
Glutamate, mM
61
59
57
55
Control
1
2
3
4
5
No. of pulses treated
Control
7.5kV/cm
10.0kV/cm
12.5kV/cm
Figure 4.5 Effect of electric field strengths on L-glutamate secretion (fermentation broth taken
after 48h of cultivation).
124
4.3.2. Effect of providing resting time and controlling the temperature of
samples between two pulses while pulsing repetitively on the secretion of Lglutamate produced in M. glutamicus
In the case of experiments mentioned above (Fig 4.4 and 4.5), the samples were
repetitively pulsed without giving any resting time or gap between two consecutive
pulses. It has been reported that if the pulse gap between pulses (in the case of multiple
pulses) is high enough, pulsed cells get enough time to reseal or rearrange their
membrane that eventually increase the viability of cells. In addition, the viable cells are
observed to be at permeabilized state during the given pulse gap (Teissie et al., 1999).
Furthermore, the literature showed that the temperature at which cells are electroporated
or the temperature that is raised into the electroporation cuvettes due to multiple pulses
has a prominent influence on the secretion of proteins or enzymes as well as on the
viability of cells (Teissie et al., 1999). It is also reported that maintaining the temperature
at low level helps to avoid rapid resealing of electropores, and facilitates the maximum
uptake of DNA or macromolecules into cells (Prasanna and Panda, 1997). For that
reason, it is assumed that allowing resting time or the combination of both pulse gap and
temperature control between two successive pulses may increase the secretion of Lglutamate produced in M. glutamicus. Hence, an experiment was conducted in which
30min pulse gap was given between two consecutive pulses (up to 5 pulses) at 12.5kV
cm-1, 200 and 25µF, and the electroporation cuvettes containing pulsed samples were
kept in an ice bath (4C) while allowing 30min resting time between pulses in order to
enhance the L-glutamate production. However, the results showed that L-glutamate
secretion was not increased significantly (less than 1% as compared to the control) by the
effort (Figure 4.6 and 4.7).
125
Glutamate, mM
31
29
27
25
Control
2
3
4
5
No. of pulses treated
Control
30min pulse gap
Cuvettes kept in ice bath while allowing pulse gap
Figure 4.6 Effect of providing pulse gap (30min) and controlling the temperature of pulsed
samples at 4C between two consecutive pulses on the secretion of L-glutamate (fermentation
broth cultivated for 24h was pulsed at 12.5kV cm-1, 200 and 25µF).
Glutamate, mM
61
59
57
55
Control
2
3
4
5
No. of pulses treated
Control
30min pulse gap
Cuvettes kept in ice bath while allowing pulse gap
Figure 4.7 Effect of providing pulse gap (30min) and controlling the temperature of pulsed
samples at 4C between two consecutive pulses on the secretion of L-glutamate (fermentation
broth cultivated for 48h was pulsed at 12.5kV cm-1, 200 and 25µF).
126
4.3.3. Effect of electric field strengths, providing resting time and controlling the
temperature of samples while pulsing repetitively on the release of total protein
and enzymes (GDH and MDH) by M. glutamicus
Besides L-glutamate, the effect of electric pulses on the secretion of total protein
and enzymes (GDH and MDH) of M. glutamicus was investigated. In this attempt, M.
glutamicus cells grown on Seed Medium were collected after 16h of cultivation, and
approximately 400µl of samples was transferred into the electroporation cuvettes.
Cuvettes were then repetitively pulsed (up to 5 pulses) at 7.5, 10.0 and 12.5kV cm -1
where cells were treated without giving any resting time between two consecutive pulses,
with giving 30min pulse gap and by keeping the cuvettes in ice bath (4C) while allowing
pulse gap in order to increase the secretion of total protein and enzymes (GDH and
MDH) of M. glutamicus. The number of samples pulsed for increasing the release of total
protein and enzymes (GDH and MDH) on each occasion was three. However, the results
demonstrated that electric pulse, irrespective of the field strengths and number of pulses
applied, has no prominent influence on the secretion of total protein (Figure 4.8). In
addition, it was observed that the secretion of total protein cannot be increased
significantly by providing resting time between pulses (up to 5 pulses) or by maintaining
the temperature at 4C while allowing pulse gap (Figure 4.9 and 4.10, respectively). The
maximum increase (only 1.9% higher as compared to the control) of total protein
secretion was determined in samples treated at 12.5kV cm-1 by 2 pulses (Figure 4.9). In
the case of samples kept at 4C in between two successive pulses, only 0.8% increase of
total protein secretion due to 12.5kV cm-1 by 5 pulses was observed as compared to the
samples treated without maintaining the temperature (Figure 4.10). However, the
activities of MDH and GDH in both control and pulsed samples were below the detection
level.
127
225
Total protein, µg/ml
220
215
210
205
200
Control
1
2
3
4
5
No. of pulses treated
Control
7.5kV/cm
10.0kV/cm
12.5kV/cm
Figure 4.8 Effect of electric field strengths on the total protein release by M. glutamicus.
225
Total protein, µg/ml
220
215
210
205
200
Control
2
3
4
5
No. of pulses treated
Control
7.5kV/cm
10.0kV/cm
12.5kV/cm
Figure 4.9 Effect of providing pulse gap (30min) between two pulses (in the case of multiple
pulsing) on the total protein release by M. glutamicus.
128
225
Total protein, µg/ml
220
215
210
205
200
Control
2
3
4
5
No. of pulses treated
Control
30min pulse gap
Cuvettes kept in ice bath while allowing pulse gap
Figure 4.10 Effect of controlling the temperature of samples at 4C while allowing 30min pulse
gap on the total protein release by M. glutamicus (pulsed at 12.5kV cm-1, 200 and 25µF).
4.3.4. Effect of the type of strains, providing resting time and controlling the
temperature of samples while pulsing repetitively on the release of total protein
and enzymes (GDH and MDH) by E. coli
It was assumed that the enhancement of L-glutamate and total protein secretion by
electropermeabilization might not have occurred because of the presence of a rigid cell
wall of M. glutamicus. Hence, the effect of similar strength of electric pulses to the cell
suspension of E. coli, a Gram-negative microorganism that contains less peptidoglycan in
the cell wall, was investigated whether electroporation can improve the secretion of total
protein and enzymes (GDH and MDH) of E. coli. As with M. glutamicus, E. coli cells
grown on Nutrient Medium were collected after 16h of cultivation, and 400µl of samples
was transferred into the electroporation cuvettes. Samples were repetitively pulsed (up to
5 pulses) at field strengths of 7.5, 10 and 12.5kV cm-1 where cells were treated without
giving any resting time between two consecutive pulses, with giving 30min gaps and by
keeping the cuvettes in ice bath (4C) while allowing pulse gap in order to increase the
129
secretion of total protein and enzymes (GDH and MDH) of E. coli. The number of
samples pulsed for increasing the release of total protein and enzymes (GDH and MDH)
on each occasion was three. As with M. glutamicus, only 1.2% increase of total protein of
E. coli as compared to the control was observed in samples treated at 12.5kV cm-1 by 5
pulses. However, the results demonstrated that electric pulse, irrespective of the field
strengths and number of pulses applied, has no major influence on total protein secretion
of E. coli (Figure 4.11). In the case of samples kept at 4C in between pulses, only 0.3%
increase of total protein due to 12.5kV cm-1 by 5 pulses was observed as compared to the
samples treated without maintaining temperature (Figure 4.12). However, the activities of
MDH and GDH in both control and pulsed samples were below the detection level.
275
Total protein, µg/ml
270
265
260
255
250
Control
1
2
3
4
No. of pulses treated
Control
7.5kV/cm
10.0kV/cm
12.5kV/cm
Figure 4.11 Effect of electric field strengths on the total protein release by E. coli.
130
5
275
Total protein, µg/ml
270
265
260
255
250
Control
2
3
4
5
No. of pulses treated
Control
30min pulse gap
Cuvettes kept in ice bath while allowing pulse gap
Figure 4.12 Effect of controlling the temperature of samples at 4C while pulsed on the total
protein release by E. coli (field strength, 12.5kV cm-1 with 30min gap between pulses).
4.3.5. Effect of the pre-treatment of cell permeabilizing agent (DTT) before
pulsation on the release of total protein and enzymes (GDH and MDH)
Since C. glutamicum is has an unusual cell wall structure with a lipid-rich mycolic
acid layer (Brennan and Nikaido, 1995; Puech et al., 2001), it is assumed that the cell
wall of this bacterium treated with any chemical agent followed by high voltage electric
pulses may reduce the cell wall resistance, and eventually increase the secretion of total
protein and enzymes (MDH and GDH). In order to investigate this hypothesis, DTT was
added into the cultivation broth of both M. glutamicus (grown on Seed Medium) and E.
coli (grown on Nutrient Medium) in which the concentration of this reagent was 20mM,
incubated for 30min, and thereafter pulsed at 12.5kV cm-1, 200 and 25µF. The number
of samples pulsed for increasing the release of total protein and enzymes (GDH and
MDH) on each occasion was three. Samples were repetitively treated with multiple
numbers of pulses (up to 5) without given any resting time between pulses. The results
showed that addition of DTT into cell suspension before pulsation does not increase the
131
total protein secretion significantly, irrespective of the field strengths, number of pulses
and the bacterial strains used in this study (Figure 4.13). However, the activities of MDH
and GDH in both control and pulsed samples were below the detection level.
Total protein, µg/ml
280
260
240
220
200
Control DTT Control DTT
(-), pulses(-) (+), pulses(-)
1
2
3
4
No. of pulses treated
M. glutamicus
E. coli
Figure 4.13 Effect of DTT addition followed by pulsation at 12.5kV cm-1, 200 and 25µF on the
release of total protein.
4.3.6. Effect of the pulsing media in which cells are suspended on the release of
total protein and enzymes (GDH and MDH) by M. glutamicus
It has been demonstrated that the efficiency of membrane electropermeabilization
is significantly affected by the pulsing media (especially by its conductivity and
osmolarity) that are generally used for washing and suspending cells before and during
electroporation (McIntyre and Harlander, 1989b). Since all the investigations conducted
in this study so far failed to increase the production of L-glutamate and total protein, it
was assumed that the transfer of electricity in the cultivation broth of both M. glutamicus
(grown on Seed Medium) and E. coli (grown of Nutrient Medium) might be affected
although a definite electric field strength was applied during pulsation. It was then
decided to investigate how the medium conductivity affects the pulse electric fields as
well as the molecular transport of protein and enzymes from cells. Hence, M. glutamicus
132
5
cells were harvested at 4500g 4C for 10min, and pellets (1g Dw) were dissolved with
4ml of sterilized distilled water (least conductivity medium). Approximately 400µl of
resulting cell suspension was transferred in electroporation cuvettes and treated with up
to 4 pulses at 12.5kV cm-1, 200 and 25µF, where pulsed cells were kept at 4C while
giving 30min pulse gap in between two consecutive pulses. The number of samples
pulsed for increasing the release of total protein and enzymes (GDH and MDH) on each
occasion was three. Surprisingly, Figure 4.14 demonstrated that electric pulse has a
prominent effect in intracellular protein secretion since the concentrations of protein were
measured 132, 145 and 139µg ml-1 in the case of cells treated with 2, 3 and 4 pulses,
whereas it was 122µg ml-1 in control (without pulse treatment).
Intracellular protein secretion µg/ml
200
150
100
50
0
Control
2
3
4
No. of pulses treated
Figure 4.14 Release of intracellular proteins of M. glutamicus due to pulse at 12.5kV cm-1, 200
and 25µF (cell pellet, 1g DW was dissolved in 4ml of sterilized distilled water).
As with total protein, the activities of MDH and GDH (U ml-1) in these pulsed
samples were analyzed. Nevertheless, the activities of those enzymes in both control and
pulsed samples were below the detection level, irrespective of the electric field strengths,
number of pulses, and the bacterial strains used in this study. In order to investigate the
133
efficiency of electropermeabilization as compared to the methods usually used for
intracellular protein and enzymes recovery, crude extract of M. glutamicus was prepared
by harvesting cells at 4500g 4C for 10min. Cell pellet (1g Dw) were dissolved in 4ml
of Tris-HCl 100mM plus 2.5mM MgCl2 buffer (pH 7.2), and thereafter disrupted the
resulting cell suspension by both sonication and French press (Section 4.2.6). The results
showed that the concentrations of intracellular protein measured in sonicated and French
pressed samples were 315 and 1011µg ml-1 (Figure 4.15). Like the pulsed samples, no
activities of GDH and MDH were detected in sonicated samples, whereas the activities in
French pressed samples (M. glutamicus) were 0.4U ml-1 and 29U ml-1, respectively.
Intracellular protein secretion, µg/ml
1200
1000
800
600
400
200
0
Control
Sonicated
French
pressed
2 pulses
3 pulses
4 pulses
Figure 4.15 Release of intracellular proteins of M. glutamicus due to sonication, French press and
electroporation (pulsed at 12.5kV cm-1, 200 and 25µF).
134
4.4. Discussion
The scope of this study was to investigate whether the electropermeabilization of
M. glutamicus can offer a potential advantage over the conventional methods of amino
acids production, and to establish this approach for the intensification of industrial
bioprocesses. It was expected that high intensity of electric pulses create electropores on
the cell membrane of M. glutamicus, increase the membrane permeability to L-glutamate,
and thereby enhance the glutamic acid production. A series of experiments was carried
out where the electroporation parameters were optimized carefully in order to increase
the secretion of L-glutamate that is produced by cultivating of M. glutamicus under biotin
limited condition. In order to investigate the most favourable production phase that needs
to be considered for increasing the L-glutamate production by electropermeabilization,
M. glutamicus cells collected after 24 and 48h of fermentation (production phase of Lglutamate) were pulsed with a range of electric field strengths (i.e., 7.5 to 12.5kV cm-1).
The results clearly demonstrated that increasing the intensity of electric pulses (7.5 to
12.5kV cm-1) and numbers of treatment (up to 5) have no significant influence in
increasing the L-glutamate secretion, irrespective of the production (L-glutamate) phases
applied for electropulsation (Figure 4.4 and 4.5). The maximum increase of L-glutamate
production was measured only 2.6% as compared to the control while samples were
treated at 12.5kV cm-1 by 4 pulses. Nevertheless, it has been confirmed that the
electroinduced secretion of proteins is dependent on the age of cultures used for
electroporation (Ganeva and Galutzov, 1999). Ganeva et al. (2003) reported that cells
growing in the stationary phase are suitable for the electroextraction of proteins or
enzymes.
The
release
of
-galactosidase
from
Kluyveromyces
lactis
by
electropermeabilization was shown to be dependent on the growth phase of bacteria since
a significant electroextraction of -galactosidase (75-80%) was obtained from cells
grown in the stationary phase without any further treatment (Ganeva et al., 2001).
In this study, the influence of similar strength of electric field and pulse number
on the release of cytoplasmic enzymes (MDH and GDH) and total protein of M.
glutamicus was also investigated. However, no remarkable influence of electric pulses,
irrespective of the field strengths and number of pulses applied, on the release of total
135
protein was observed (Figure 4.8). Hence, it was supposed that the electroenhancement of
L-glutamate, protein and enzyme secretion might be restricted because of having a rigid
cell wall of M. glutamicus. Nevertheless, this assumption was ruled out while the
application of similar intensity of pulses did not exhibit any major influence on the
secretion of total protein of E. coli (Figure 4.11). Although this study showed that electric
pulses have no prominent influence on L-glutamate, total protein and enzymes (MDH and
GDH) secretion, it is well-established that the application of high-intensity, short, electric
field pulses to living cells permeabilizes the plasma membrane, and allows a free
exchange of ions and molecules between the cytoplasm and surrounding media (Tsong,
1991). Ohshima and his colleagues (2000) recovered the foreign proteins (-glucosidase,
-amylase and cellobiohydrolase) from the recombinant strains of E. coli (E. coli/pNC1,
E. coli/pHI301A and E. coli/pNB6, respectively) by pulsing at 10kV cm-1 and 200J ml-1.
Furthermore, Rols and Teissie (1998) demonstrated that an increase in the number
of pulses enhances the rate of permeabilization. The uptake of Cr51 - EDTA has been
shown to be enhanced greatly due to an increase of pulse number above 8 (Gehl and Mir,
1999). Ganeva et al. (1995) demonstrated the leakage of intracellular proteins from S.
cerevisiae after the application of a single rectangular pulse (25kV cm-1) with a duration
of 10-25ms. Their results also showed that the number of permeabilized cells and the
flow of macromolecules (fluoresceinated dextran, FD70) across the cell membrane
increased with an increase in electric field strength, pulse duration and number of pulses
(Ganeva et al., 1995). Deng et al. (2003) demonstrated that the pulse with longer duration
results in increase of membrane permeability. Pulse duration has been shown to be a
crucial factor for the penetration of macromolecules [fluorescein isothiocyanate (FITC)dextran, -galactosidase and plasmid DNA)] into CHO cells since there was no
permeabilized cells detected at pulse duration shorter than 1ms (Rols and Teissie, 1998).
The uptake of Cr51 - EDTA (non-radioactive marker) into skeletal muscle fibers was
approximately 1nM at a pulse duration of 5ms, whereas it increased 6-fold higher in
presence of pulse duration of 25ms (Gehl and Mir, 1999). It has been demonstrated that a
higher electric field strength and longer pulse duration resulted in a greater number of
permeated cells, irrespective of the type of microorganisms, and excreted higher
136
concentrations of ATP in the extracellular environment (Aronsson et al., 2005).
Moreover, the expansion of transient electropores and the resealing rate of
electroporation depend on both the pulse number and duration (Rols and Teissie, 1989).
However, the influence of pulse duration on the secretion of L-glutamate and total protein
and enzymes of M. glutamicus was not investigated since the electroporator that was used
in this study does not offer to control this parameter.
Teissie et al. (1999) demonstrated that the kinetics of protein efflux due to
electropermeabilization is a long lasting process although a very fast leakage is detected
just after the pulse. It was presumed that the secretion of L-glutamate and total protein of
M. glutamicus might be limited due to the rapid releasing of electropores. Therefore, it is
vital to increase the opening or resealing time of electropores in order to enhance the
efficiency of extraction by electropermeabilization. Although pulsed cells of M.
glutamicus and E. coli were given 30min pulse gap between two consecutive pulses at
12.5kV cm-1 with up to 5 pulses, no promising influence of electric pulses on the
secretion of L-glutamate (Figure 4.6 and 4.7) and total protein (Figure 4.10 and 4.12 in
the case of M. glutamicus and E. coli, respectively) was observed. However, it has been
reported that pulsed cells that are given with certain resting time between two consecutive
pulses have a prolonged permeabilized state facilitating the transfer of DNA or foreign
molecules or intracellular proteins or enzymes across the membrane (Prasanna and
Panda, 1997; Teissie et al., 1999).
It has also been reported that increase of medium temperature during pulsation
has a harmful effect on the properties of protein or enzymes that are inserted or excreted
into or from the cells, respectively (Gallo et al., 2002; Rols et al., 1994). For instance, an
increase of intensity from 2.7 to 3kV cm-1 resulted in about 90% decrease of GAPDH
activity in the supernatant of pulsed cells (Ganeva et al., 2003). Electrotransformation
protocols are, therefore, carried out at low temperatures (0-4°C) with the use of cold
buffers or by chilling the electroporation cuvettes contained the targeted cells (Teissie et
al., 1999). In this study, while pulsed cells of M. glutamicus and E. coli were kept in ice
bath (4C) between consecutive pulses, no promising influence of electric pulses on the
137
secretion of L-glutamate (Figure 4.6 and 4.7) and total protein (Figure 4.10 and 4.12 in
the case of M. glutamicus and E. coli, respectively) was observed. However, Nanda and
Mishra (1994) demonstrated that the permeabilization of erythrocyte membranes by
electric pulses is substantially improved by controlling the incubation temperature of cells
before, during and after pulsation. It was supposed that all the above-mentioned efforts
conducted in this study in order to improve the secretion of L-glutamate and total protein
might have failed because of having high conductivity of fermentation and culture media
(Seed and Nutrient Media for M. glutamicus and E. coli, respectively) used for
electroporation. Since the electroporation medium with high conductivity decreases the
intensity of electric pulse, cells might not be exposed by the actual voltage generated
from the Bio-Rad Gene Pulser.
In literature, several experiments have been demonstrated that addition of
different chemicals or reagents in cell suspension affects the structure of membrane, and
apparently increases the number of pores on the cell wall after electropulsation (Ganeva
et al., 1995; 2003; Ganeva and Galutzov, 1999). However, this study revealed that pretreatment of M .glutamicus and E. coli cells with DTT (20mM, final concentration)
before electroporation has no prominent influence on the secretion of total protein, MDH
and GDH, irrespective of the field strengths, number of pulses and the bacterial strains
used in this study (Figure 4.13). Nevertheless, it has been reported that thiol compounds
reduce the disulphide bridges in the mannoprotein layer and increase the cell wall
porosity. Macromolecule transfer through the yeast cell wall was successfully
accomplished by pre-incubating cells with thiol compounds (De Nobel et al., 1989).
Approximately 7-fold increase of protein and a dramatic improvement in transfer of
FD70 have also been observed while incubating yeast cells with DTT prior to pulsation
(Ganeva et al., 1995). The electroinduced effluxes of protein and enzymes have also been
achieved by the pre-treatment of S. cerevisiae with DTT (Ganeva and Galutzov, 1999).
In the case of all studies mentioned above, however, pre-treatment of cells with
DTT before electropulsation has been carried out for the secretion or extraction of protein
or enzyme of yeast. It is obvious that the cell wall structure of bacteria is more rigid than
138
that of yeast, and hence the cell wall of M. glutamicus might not be affected by the
addition of DTT. Furthermore, a buffer (Tris-Tricine buffer, pH = 7.5) having low
conductivity as compared to the growth medium was used for washing and suspending
the yeast cells that were pre-treated with DTT before pulsing with a series of high
intensity electric pulses (Ganeva et al., 1995; 2003; Ganeva and Galutzov, 1999). In this
study, on the other hand, DTT was added directly into the Seed Medium in which M.
glutamicus was grown, and thereafter pulsed at 12.5kV cm-1, 200 and 25µF. It is,
therefore, assumed that the secretion of protein and enzymes by electropermeabilization,
irrespective of the treatment of cells with DTT before pulsation, might be restricted due
to the high conductivity of culture media used for pulsation.
Like DTT, the effect of glycine on the release of enzymes from the recombinant
of E. coli has been investigated by pulsing cells at 10kV cm-1 and 200J ml-1. The
secretion of -glucosidase increased due to increase of glycine concentration into the
pulse medium, and the maximum amount of enzyme was achieved when the glycine
concentration was 5%, whereas an opposite effect was observed in the case of
cellobiohydrolase and -amylase. In addition, the highest amount of -amylase was
secreted at 5% PEG although the concentration of total protein gradually decreased due to
increase of PEG in electroporation medium (Ohshima et al., 2000). These results
confirmed that the efficiency of protein or enzyme secretion due to these chemicals or
reagents depends on the type of product excreted by electropermeabilization. Similar to
the preincubation with a detergent, Ganeva et al. (2001) demonstrated that the efficiency
of electroinduced protein release is dependent on the composition of post-pulse medium.
The addition of glycerol and DTT in the post-pulse medium showed an increase of
GAPDH and PGK activities in the supernatant of electrically treated cells (Ganeva et al.,
2003). Ganeva and Galutzov (1999) also demonstrated that addition of potassium or
sodium chloride (50mM) in the post-pulse medium (50mM Tris-Tricine, pH 7.5)
provoked an accelerated release of proteins and enzymes i.e, PGK, GLR and ADH. Due
to time restriction, however, the influence of using post-pulse medium on the secretion of
protein and enzymes of M. glutamicus was not investigated in this study.
139
However, when M. glutamicus cells (1g Dw) dissolved in 4ml of sterilized dH2O
were treated with up to 4 pulses at 12.5kV cm-1 in conjunction with the 30min resting
time at ice bucket (4C), approximately 19% increase of total protein (M. glutamicus) as
compared to the control was measured in sample treated by 3 pulses. This result indicated
that the conductivity of medium in which cells are suspended has a major influence on
the efficiency of electropermeabilization. However, treatment of this cell suspension with
more that 4 pulses was not possible due to high concentration of cells in dH2O (Figure
4.14). Several experiments suggested that the ionic content of the extracellular medium is
an important factor that can influence the yield of electropermeated cells, as well as initial
field strength, pulse time constant and salt concentration (Muraji et al., 1993; Tatebe et
al., 1995). Ohshima et al. (1995) demonstrated that the medium conductivity that is
increased (over 2mS cm-1) due to pulse electric field reduces the electric voltage and
pulse width, and eventually results in lower concentration of released proteins. This
phenomenon will be severe while cells are treated with multiple numbers of pulses for the
electroextraction of a particular intracellular product (Ohshima and Sato, 2004).
The conductivity of an electroporation buffer greatly affects the electroporation
efficiency (Pucihar et al., 2001). Muller et al. (2001) showed that a reduction in ionic
conductivity increases the uptake or release of various molecules. The uptake of
membrane-impermeable dye PI in low-conductivity media (1mS cm-1) was considerably
higher than that in high-conductivity media (4-5mS cm-1) (Muller et al., 2001). Like
mammalian cells, the amount of energy required in order to obtain a certain percentage of
permeabilized cells is dependent of the conductivity of electroporation buffers.
Approximately 70% permeabilized cells of L. plantarum was obtained by delivering an
energy of 14J ml-1 to a phosphate buffer having a conductivity of 0.4S m-1, whereas it
was 100J ml-1 in the case of a conductivity of 1.5S m-1 in order to achieve the same
permeabilization (Wouters et al., 2001). Although this study demonstrated that medium
conductivity is the major barrier to increasing the secretion of amino acids and
intracellular protein and enzymes, no conductometer was used to determine the
conductivities of media (Seed and CGXII) and pulsing buffers. However, the time
constants obtained during pulsation at a certain voltage were used as an indication of the
140
conductivities of media or buffers used in this study. The time constant depends on both
the resistance of suspension and the capacitance of the capacitor. Furthermore, the
resistance of suspension is inversely related to its conductivity. When M. glutamicus cells
suspended in H2O were pulsed, higher time constants (2.87msec for 24h culture) as
compared to those achieved by electropulsing the cells suspended in Seed Medium
(1.08msec for 24h culture).
This study clearly demonstrated that the efficiency of electropermeabilization in
secreting of total protein was lower than that of conventional methods since the amount
of protein measured in sonicated and French pressed samples were 2.2 and 7.0-fold,
respectively, higher than that of samples pulsed at 12.5kV cm-1 by 3 pulses (Figure 4.15).
In addition, the activities of GDH and MDH in French pressed samples were 0.4U ml-1
and 29U ml-1, respectively, whereas no activities were determined both in pulsed and
sonicated samples. Similar to these results, the maximal protein concentration (40µg ml-1)
obtained due to electric pulses was approximately 30% of glass beads homogenization
(Ohshima et al., 1995). The same group also observed that the amount of released βglucosidase from the recombinants of E. coli by pulsed electric fields (PEF) is only 26%
of that by ultrasonic treatment, whereas the specific activities of -amylase and
cellobiohydrolase were 9 and 1.9-fold, respectively, higher than that of the ultrasonic
treatment (Ohshima et al., 2000). Their results demonstrated that the applied PEF easily
disrupted the outer membrane of cells, but was not enough to disrupt the cytoplasmic
membrane simultaneously, indicating that this technique will be suitable especially for
the release of periplasmic proteins.
Although the concentrations of released proteins by pulse treatment are relatively
lower than that by the usual methods, electropermeabilization permits the selective
release of the products of interest from microorganism (Ganeva and Galutzov, 1999;
Ohshima et al., 1995; 2000). In addition, electropermeabilization is simpler and less
destructive to proteins or enzymes as compared to the routinely applied methods
(mechanical breakage and chemical extraction) used in protein or enzyme extraction
(Ganeva and Galutzov, 1999). The isolation of desired proteins with a high percentage of
141
purity by use of traditional methods often encounters with many technical difficulties
since the protein or enzyme of interest is often contaminated with host proteins and other
unknown contaminants (Ohshima et al., 1995). Electroporation has low operating cost,
and
the time required for the maximum recovery of intracellular products by this
approach is considerably shorter than that of chemical extraction where a long incubation
(24h to several days) with a particular agent is mandatory (Breddam and Beenfeldt,
1991). It has been reported that electroporation does not cause cell fragmentation
(Ganeva et al., 1995), and this approach will be effective for the isolation of recombinant
proteins that are highly sensitive to proteolytic degradation (Ganeva et al., 2003).
Electropermeabilization might have the greatest potential in industrial
bioconversion processes where biological systems with the continuous production of
metabolites and their subsequent extraction by high voltage electric pulses will increase
the speed and efficiency of production (Ganeva et al., 2003). However, the
electroporation units that are currently available commercially cannot generate high
electric field strength in order to achieve maximal permeabilization efficiency in many
bacterial strains (Mercenier and Chassy, 1988). The volume of sample that can be pulsed
by a single treatment using the available electroporator is very small. Nevertheless, the
recent improvements in the pulsing instruments may allow us to apply electric pulses
with controlled parameters. A large volume flow electroporator has been designed by
MaxCyte Inc. (USA) in order to overcome the limitation of sample size. In this system,
cells are first suspended in pulsing buffer together with the target molecules. The
resulting cell suspension is pumped through the disposable flow chamber, and pulsed
with an appropriate electric field strength while flowing in the chamber (Li et al., 2002).
High transfection efficiency was achieved by this approach using cell volumes up to 50ml
with a cell densities ranging from 1 to 8  107 cells ml-1.
Finally, the results obtained from this study confirmed that the extraction or
isolation of proteins or enzymes (that are not usually secreted or excreted into the
fermentation medium) through the electropermeabilization would be an alternative
approach in the field of biotechnology. In this case, cells growing at a certain growth
142
stage are required to suspend and pre-incubate in a low conductivity buffer before
applying the electric pulses. After pulsation, cells are needed to keep in a specific
medium in which the secretion or excretion of the product of interest will take place.
However, the electroenhancement of L-glutamate or any proteins and enzymes secretion
is strongly dependent on the conductivity of fermentation medium, electric field strength
and number of pulses. The intracellular L-glutamate (produced in M. glutamicus) that is
not usually excreted into the extracellular medium by the well-established treatments
(biotin limitation, surfactant addition and penicillin/ethambutol addition) could easily be
secreted by electroporation increasing the overall yield of L-glutamate. It is expected that
the findings obtained in this study may assist researchers to design the
electropermeabilization experiment in which the yield of a certain product will be
increased. Hence, careful optimization of electroporation factors, such as cellular,
physiological and electrical is mandatory in order to excrete intracellular products
through the membrane electropermeabilization.
143
Chapter FIVE
5.
Transient
Bioprocessing:
Electroporation
Cell
Viability
in
Intensified
and
Membrane
Permeabilization
5.1. Introduction
Developing an eletropermeabilization procedure for either molecular biology or
bioprocessing purposes is always a challenge since the efficiency of this approach is
directly related to the membrane permeability and viability of cells (Teissie et al., 1999).
However, these two important features are generally influenced by the different factors,
such as electrical (pulse amplitude, pulse duration, number of pulses and pulse type),
cellular (cell size, shape, growth stage and structural configuration of cell wall) and
physical (temperature, osmotic pressure and electroporation medium) associated with
electroporation. Among these, however, membrane permeabilization is mostly controlled
by the electrical factors (Prasanna and Panda, 1997). Canatella et al. (2001) mentioned
that the typical electrical conditions for electroporation are the field strengths of 1 to
20kV cm-1 and pulse duration of 10µs to 10ms, which may vary depending on the type of
cells and its particular application in biotechnology. Without adjusting the abovementioned parameters, nevertheless, cells may not return into their normal physiological
state and eventually lose their viability although a high level of membrane
permeabilization is usually observed after pulsation (Teissie et al., 1999).
Haest and his co-workers (1997) observed an asymmetry in lipid bilayer and a
loss of phospholipids in the membrane of human erythrocyte after electroporation. It has
also been reported that membrane permeabilization results in an entrance of water into
cells, and consequently increases the volume of cells that may lead to the rupture of cell
membrane (Golzio et al., 1998). Under mild pulse conditions, permeabilization appears
144
as a reversible process that weakly affects cell viability, while drastic electrical conditions
lead to cell death (Vernhes et al., 1999). However, pulsed cells recover their original
permeability within 30min of incubation at room temperature (RT), although the
resealing process may be varied depending on the conditions applied. Teissie and Rols
(1988) demonstrated that the viability of CHO cells reduced due to increase of electric
field strength although the permeability of cell membrane was observed to be increased
significantly. Furthermore, Vernhes et al. (1999) demonstrated that the viability of CHO
cells was affected more severely in the presence of a strong electric field with short pulse
duration than in a weak electric field with longer pulse duration.
There is close correlation between the cell viability and the efficiency of
electrotransformation since the decrease in transformation efficiency (TE) occurs due to
the reduction of cell viability (Rols et al., 1992). Apart from the effect of high intensity
electric pulses, several factors have been mentioned that show a strong influence on the
viability of cells while conducting an electroporation experiment. Miller et al. (1988)
demonstrated that the efficiency of transformation (C. jejuni) is affected by the increased
temperature generated during electroporation. Hence, pre and post incubation at certain
temperature before and after electroporation have been suggested in order to increase the
viability of cells as well as the efficiency of transformation. For instance, Rols et al.
(1994) observed that electroporated samples (mammalian cells) with a pre-incubation at
4C and post-incubation at 37C increased cell viability. The survivability of cells during
electroporation is also dependent on the type of cells and their membrane configuration.
Hattermann and Stacey (1990) demonstrated that B. japonicum cells are more resistant to
high-voltage electric pulses than mammalian cells, carrot protoplasts, yeast and bacterial
cells (E. coli). Garcia et al. (2003) investigated the occurrence of sub-lethal injury of E.
coli by PEF at different pH. Although E. coli cells were observed to be injured slightly
due to pulse (400µs at 19kV) at pH 7, cells mortality increased up to 99.5% at pH 4.
These results clearly demonstrated that pH of pulsing buffer has also an important
influence on cell viability. Since the yield of electrotransformants is limited due to the
decrease in cell viability, it is important to maintain the number of viable cells as high as
possible while conducting an electroporation experiment.
145
Although the different factors of electroporation (especially, electric pulse) affect
the viability of cells, appropriate strength of electric field induces transient electropores
on the cell membrane and causes reversible permeabilization that allows the foreign
molecules or DNA to be introduced into cells or intracellular products to be secreted to
the extracellular medium (Teissie et al., 1999). Apart from the electrotransformation, in
recent years, this approach has been successfully applied for the isolation of a certain
product (protein or enzyme that is not usually secreted into the extracelllular medium)
from both bacteria and yeast (Aronsson et al., 2005; Ganeva et al., 1995; 2001; 2003;
2004; Ohshima et al., 1995). Due to the recent advancement of electroporation in
bioprocessing, extensive efforts have also been given in order to establish this technique
for the continuous production of metabolites by fermentation and their subsequent
extraction with the pulse electric field treatment, and thereby increase the yield of
production. Nevertheless, it is confirmed that the permeabilization of cell membrane by
high intensity electric pulses is not straightforward especially for the enhancement of
excretion or secretion of a certain product (for example, L-glutamate) during
fermentation. There are several factors associated with the electroporation, which have a
major impact for increasing the production of L-glutamate through the membrane
permeabilization (Chapter 4).
In the case of previous study, M. glutamicus cells growing at the production phase
(stationary growth period, 24-48hr of fermentation) were pulsed with a range of electric
field strengths in order to increase the secretion of L-glutamate. However, it is obvious
that a certain percentage of cells might have killed due to electric pulse, irrespective of
the electric field strength applied, and this phenomenon will interrupt the bioprocess
while electropermeabilization is considered for the enhancement of product secretion
during fermentation. If the electric pulse is severe, a large population of cells will be
killed resulting in decreased the overall yield of product. Therefore, it is mandatory to
maintain cell viability as high as possible with having the highest level of membrane
permeabilization by optimizing the electroporation parameters since pulsed cells need to
complete the rest of fermentation. Furthermore, Chen and Lee (1994) demonstrated that
the application of high electric voltage causes electroconformational changes of proteins
146
affecting the properties of protein. Therefore, intensive research towards the
electroporation factors, method development and optimization are crucial for either
developing an electrotransformation protocol or establishing this approach in
bioprocessing.
It is well defined that viable cells are capable of performing all the cellular
functions necessary for survival or growth or reproduction under the given conditions.
The number of viable cells in a population is traditionally measured by the agar plate
assay in which samples are serially diluted and spread on the agar plates (Hattori, 1988).
Cells are then incubated at 25-37C (depending on organism) for 2-3 days that allow cells
to form countable colonies on the agar plates. While conducting the electroporation
experiments, however, it is necessary to determine not only the cells that survived the
treatment but also the number or percentage of cells that are permeabilized (the efficiency
of
electropermeabilization)
by
the
electric
pulses.
The
quantification
of
electropermeabilized cells in a population is generally achieved by analyzing the uptake
of marker molecules (fluorescent or nonpermeant dyes) through the fluorometry and
fluorescence microscopy (Gabriel and Teissie, 1997; 1999). In the case of quantitative
analysis of fluorescent [propidium iodide (PI) and calcein] or nonpermeant [trypan blue
(TB)] dyes uptake, samples are first labelled with a certain concentration of dye,
incubated at 30C and thereafter electropulsed. Pulsed cells are then incubated for 530min at RT and visualized under the fluorescence microscope. The colour of the
cytoplasm of permeabilized cells is altered i.e., the cytoplasm turns blue in the case of TB
or becomes highly fluorescent in the case of PI (Teissie et al., 1999). Moreover, the
release of intracellular metabolite (glucose-6-phosphate) or leakage of ATP has been
measured to determine the percentage of permeabilized cells (Rols and Teissie, 1990b).
However, these widely established methods have several drawbacks: (i) a random
choice of dyes with minimum fluorescence intensity for the characterization of
permeabilized cells; (ii) failure to detect cells due to severe electropermeabilization; and
(iii) false detection of cellular ghosts lacking of fluorescence due to DNA leakage caused
by electropermeabilization (Kotnik et al., 2000). Furthermore, the electropermeabilized
147
CHO cells have been observed to be fused each other during the resealing of pores
(Teissie and Rols, 1986). The membrane generally contains some interfacial water
molecules that form a well structured network. The energy provided by the external field
induces structural change of polar head groups and disorganizes the interfacial water
lattice (Sowers, 1986). It has been observed that cells loose the repulsive forces that
prevent membranes of two cells fusing spontaneously during electropermeabilization
(Rols and Teissie, 1989). It is, therefore, always challenging to detect the permeabilized
cells among the total cells population.
Bleomycin (BLM), is used as an antitumor agent, causes nucleotide sequencespecific DNA cleavage (Quada et al., 1998) and inhibits the growth of both bacteria and
mammalian cells (Kross et al., 1982). It also has the ability to cleave RNA to a lesser
extent (Carter et al., 1990). However, BLM is impermeable to an intact cell membrane
under normal physiological conditions (Mir et al., 1996). Orlowski et al. (1988)
demonstrated that electropermeabilization allows a defined number of BLM molecules to
enter directly into the cytoplasm of cells, and increases the cytotoxicity of this molecule
by 100-1000 times in vitro (Dev et al., 2000; Mir and Orlowski, 1999; Mir, 2000; Rols,
2006). This compound is currently used in anticancer therapy (Gothelf et al., 2003),
where the cytotoxicity of BLM is induced by delivering the electric pulses to the infected
tumors. BLM molecules enter into intact cells not by diffusion through the plasma
membrane but by a mechanism of receptor-mediated endocytosis (Pron et al., 1999).
However, it has been reported that the concentration of BLM (5nM) has no effect on nonpermeabilized cells, whereas it causes the death of electropermeabilized cells (Kotnik et
al., 2000). The permeabilization of mammalian cells (Chinese hamster lung, DC3F) has
been quantified by using BLM that was added in the electroporation medium before the
pulsation. This method is highly selective, accurate, affordable, and has been used for
evaluating the membrane permeabilization (Kotnik et al., 2000; Puc et al., 2003).
The purpose of this study is to examine how the different factors of
electroporation (voltage, capacitance, pulse number, pulse gap/resting time, growth stages
of cell, buffers in which cells are suspended and the type of strains) influence the viability
148
and membrane permeability of M. glutamicus. Efforts will be given in order to establish a
method for determining the number of permeabilized cells in a given pulsed population
with the aid of Bleomycin added (before pulsation) into the cell suspension of M.
glutamicus followed by measuring the cell viability through the agar plate assay. The
main objective of this investigation is to achieve an indication how these two important
features influence the yield of electroextraction while the continuous production of any
intracellular protein or enzyme and its secretion during fermentation will be carried out
by imposing of high voltage electric pulses.
149
5.2. Material and Methods
5.2.1. Chemicals
Peptone from pancreatically digested casein and meat extract were obtained from
VWR (Merck, UK). Yeast extract and bacteriological agar were procured from Oxoid,
UK; and D-glucose, urea, NaCl and all other chemicals were purchased from Fisher, UK
unless otherwise mentioned.
5.2.2. Organisms and cultivation
The bacterial strains used for this study were M. glutamicus DSM 20300 (Collins
et al., 1977; Suzuki et al., 1981; Yamada and Komagata, 1970; Yamada et al., 1976) and
E. coli DSM 498 (Farnleitner et al., 2000) supplied by the German Collection of
Microorganisms and Cell Culture (DSMZ-Deutsche Sammlung von Mikroorganismen
und Zellkulturen GmbH). M. glutamicus DSM 20300 was cultivated in Medium 53 (10g
l-1 peptone, 5g l-1 yeast extract, 5g l-1 glucose, 5g l-1 NaCl, 1000ml distilled water, pH
adjusted to 7.2-7.4), whereas E. coli DSM 498 was cultivated in Nutrient Medium (5g l-1
peptone, 3g l-1 meat extract, 1000ml distilled water, pH adjusted to 7.0). The stock agar
plates were prepared according to the method described previously in Section 3.2.2. M.
glutamicus was grown on Seed Medium, whereas E. coli was grown in Nutrient Medium.
The composition and preparation of both media were described previously in Section
3.2.2. M. glutamicus was cultivated at 30C and 220rpm, whereas E. coli was cultivated
at 37C and 150rpm. All the experiments were performed in 250ml shake flasks
containing 50ml of culture media.
5.2.3. Electroporator and pulse treatment
Electropermeabilization was carried out using a Gene PulserTM apparatus (BioRad Laboratories, Richmond, CA). The description of this apparatus was mentioned in
Section 4.2.4. Bacterial cultures were collected from both the exponential and stationary
growth phases, transferred into Bio-Rad electroporation cuvettes (0.2cm), and
subsequently pulsed at RT with a range of voltages (1.5, 2.0 and 2.5kV) although the
capacitance (25F) and parallel resistance (200) were kept constant in all the
experiments.
150
5.2.4. Viability assay on agar plates
The viability of cells was measured by the Luria Bertani (LB) agar plates, which
contain 10g l-1 peptone, 5g l-1 yeast extract, 10g l-1 sodium chloride and 15g l-1 agar
(Bertani, 1951). LB agar plates were prepared according to the method described
previously in Section 3.2.2. Pulsed cells were serially diluted with sterile distilled H2O
and spread on to the agar plates in an aseptic condition. Plates were then incubated at
30C and 37C for M. glutamicus and E. coli, respectively, and colonies were counted
after 48-72h of incubation. Since the viability assay by the agar plate is not precise
enough, all the experiments conducted in this study were carried out at least two times,
and the number of pulsed samples spread on agar plates was three. The viability of cells
was calculated by normalizing the non-electroporated samples (control) to have 100%
viability. The ratio of recovered viable colonies in pulsed samples to that of control was
then used to determine the relative cell viability at a given electric field strength and pulse
number.
5.2.5. Permeabilization assay by Bleomycin (BLM) treatment
BLM sulphate, a mixture of glycopeptide antibiotics isolated from a strain of
Streptomyces verticillus, was purchased from Sigma-Aldrich, UK. The activity range of
this product is 1.2-1.7U mg-1, and the vial contains total of 15U in 10mg of powder. The
appropriate concentration required for tissue culture application is dependent on the type
of cells. The entire crystalline solid in vial was dissolved with sterile distilled water to
make a stock solution (1mg ml-1). This solution was reported to be stable for several days
at 0-5°C in a glass container. In the case of electropermeabilization assay, a range of
concentrations of BLM was added to the cell suspension of both M. glutamicus and E.
coli. Approximately 400µl of BLM added samples were transferred into the
electroporation cuvettes (0.2cm electrode gap), and subsequently pulsed with a range of
voltages (1.5, 2.0 and 2.5kV) although the capacitance (25F) and parallel resistance
(200) were constant. The pulsed samples were kept at RT for approximately 30min
after the treatment. Samples were serially diluted with sterile pre-cooled distilled water,
and thereafter 100µl samples were spread on agar plates in order to determine the viable
colonies.
151
5.3. Results
5.3.1. Effect of voltages, capacitance, number of pulses and pulses gap on the
viability of M. glutamicus
Exponentially growing cultures of M. glutamicus were directly transferred into the
Bio-Rad electroporation cuvettes, and subsequently pulsed with the field strengths of 7.5,
10.0 and 12.5kV cm-1 where the capacitance and parallel resistance were kept constant at
25µF and 200, respectively. In addition, the samples were treated with up to 5 pulses
without giving any resting time (gap) between pulses. The results showed a gradual loss
of cell viability due to increase of field strength. After applying a single pulse, the
percentages of killed cells were measured as 5, 8 and 13 (%) at field strengths of 7.5, 10.0
and 12.5kV cm-1, respectively. A similar trend was obtained when bacterial culture was
pulsed with multiple numbers of pulses (up to 5). The relative cell viabilities were
measured as 95, 91, 85, 80 and 74 (%) while cell suspension was treated by 1 to 5 pulses,
respectively, although the field strength was kept constant at 7.5kV cm-1 (Figure 5.1). A
similar phenomenon was also observed in presence of 10.0 and 12.5kV cm-1. The
percentages of viable cells measured after treating with 5 pulses were 74, 71 and 64 (%)
at field strengths of 7.5, 10.0 and 12.5kV cm-1, respectively.
Moreover, the results showed that the viability of cells decreased gradually due to
increase in the number of pulses; and the higher the number of pulses, the high cells
mortality (Figure 5.1). The percentage of killed cells measured at 12.5kV cm-1 by single
pulse was 13%, whereas it was 36% in presence of 5 pulses. Furthermore, the effect of
capacitance on the viability of M. glutamicus was investigated where a range of
capacitances (5 to 50µF) was applied in presence of field strengths 7.5 and 12.5kV cm -1
(Figure 5.2). The result showed a gradual decrease of cell viability due to increase of
capacitance. When the capacitance was set to 50µF, the relative cell viabilities were
measured 89 and 78 (%) at 7.5 and 12.5kV cm-1, respectively. Furthermore, pulses at low
capacitance (<10µF) did not greatly affect the viability of cells since more than 94% of
cells were viable even after pulsed at 12.5kV cm-1 (Figure 5.2). This result demonstrated
that capacitance higher than 10µF is required to create large number of electropores on
the cell membrane of M. glutamicus.
152
Relative cell viability (%)
120%
100%
80%
60%
40%
20%
0%
Control
1
2
3
4
5
No. of pulses treated
7.5 kV/cm
10.0kV/cm
12.5kV/cm
Figure 5.1 Effect of voltages and multiple pulses on M. glutamicus viability.
Relative cell viability (%)
120%
100%
80%
60%
40%
20%
0%
5
10
15
20
25
50
Capacitance (µF)
7.5kV/cm
12.5kV/cm
Figure 5.2 Effect of capacitance on the viability of M. glutamicus (cells were treated by a single
pulse).
153
In the case of repetitive pulsing (no pulse gap between two consecutive pulses),
the viability of M. glutamicus cells was affected greatly. Since the viability of cells is
related to the efficiency of transformation, it is necessary to reduce cells mortality while
developing an electroporation protocol. It is assumed that given a certain interval or gap
between two successive pulses (in the case of multiple pulsing) may facilitate the
recovery of original membrane structure of cells. In order to investigate this assumption,
samples were treated with a resting time of 20min between pulses. After treating with a
single pulse at certain field strength, electroporation cuvettes were kept at RT for 20min
before applying the second one. In both conditions i.e., without gap and 20min gap, it
was observed that cells mortality increased with increasing the field strengths and number
of pulses. However, the results showed that the viability of M. glutamicus cells increased
(approximately 3-10%) due to 20min pulse gap given between two consecutive pulses,
regardless of the field strengths and number of pulses applied (Figure 5.3).
5.3.2. Effect of growth stages on the viability of M. glutamicus
In order to investigate the effect of growth stage of the cell cycle (used for
electroporation) on the viability of M. glutamicus, cell suspensions collected from two
growth points were pulsed by imposing a range of field strengths (7.5 to 12.5kV cm-1)
and a number of pulses (1 to 5). The results demonstrated that M. glutamicus cells
growing at the stationary phase (24h of cultivation) are more sensitive to killing by
electric pulses than that of cells growing at the early or mid-exponential phase (12h of
cultivation). In the presence of a single pulse at 7.5kV cm-1, the relative cell viabilities
measured in exponentially growing cells and cells growing at the stationary phase were
95 and 88 (%), respectively, whereas the viabilities reached to 87 and 78 (%) at 12.5kV
cm-1 by treating with a single pulse. Furthermore, the difference in cell viabilities was
profound when the cell suspension was treated with more than one pulse (Figure 5.4).
However, the viability of M. glutamicus cells varied (approximately 5-20%) due to the
difference in growth stages of the cell cycle, irrespective of the field strengths and
number of pulses applied in this study. These results confirmed that the viability of cells
due to electric pulses is influenced by the growth stages of bacterial culture.
154
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
2
3
4
5
No. of pulses treated
7.5kV/cm
7.5kV/cm + 20min gap
10.0kV/cm
Figure 5.3 Effect of pulse gap or resting time on the viability of M. glutamicus.
155
10.0kV/cm + 20min gap
12.5kV/cm
12.5kV/cm + 20min gap
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
1
2
3
4
5
No. of pulses treated
7.5kV/cm (12h)
7.5kV/cm (24h)
10.0kV/cm (12h)
10.0kV/cm (24h)
12.5kV/cm (12h)
12.5kV/cm (24h)
Figure 5.4 Relative cell viability by treating with a range of electric pulses to the M. glutamicus cells grown at two different growth stages.
156
5.3.3. Effects of different suspending fluids on the viability of M. glutamicus
To examine the effect of using electroporation media or buffers with different
conductivities on the viability of M. glutamicus, 5ml of exponentially growing cultures
were centrifuged at 4000g and 4C for 5min. Cell pellets were washed twice with the
following buffers i.e., distilled H2O, 1mM Tris plus 270mM sucrose (pH 7.2) and 50mM
Tris HCl (pH 7.4), and thereafter dissolved in 5ml of mentioned buffers. All the media
were sterilized at 121C for 15min, and cooled to 4C before use. Approximately 400µl
cell suspension was then transferred into the electroporation cuvettes and treated with up
to 5 pulses at 12.5kV cm-1. Pulsed samples were serially diluted, spread on to the agar
plates and kept in incubator for further evaluation. The results showed that cells
suspended in Seed Medium maintained their viability at a higher level (64%) even after
treating with 5 pulses at 12.5kV cm-1, however, the relative cell viabilities measured in
distilled H2O, 1mM Tris plus 270mM sucrose and 50mM Tris HCl were 3, 10 and 7 (%),
respectively (Figure 5.5). This study also showed that multiple pulses affected the
viability of cells, regardless of the media or buffers applied to suspend M. glutamicus
cells. In the presence of single pulse at 12.5kV cm-1, the highest percentage of cells
reduction was 23% observed in distilled H2O, whereas the relative cell viabilities were
87, 66 and 54 (%) in the case of Seed Medium, 1mM Tris plus 270mM sucrose and
50mM Tris HCl, respectively. The results obtained from this study confirmed that cell
viability due to electroporation is also dependent on the composition of buffers in which
cells are suspended.
157
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
1
2
3
4
No. of pulses treated
-20%
Medium
Distilled Water
Tris 1mM + 270mM Sucrose
Tris HCl 50mM
Figure 5.5 Effect of suspending fluids on the viability of M. glutamicus (pulsed at 12.5kV cm-1, 200 and 25µF).
158
5
5.3.4. Effects of electric pulses on viability of different strains
In order to investigate the effect of electric pulses on the viability of other
bacteria, cell suspension of a Gram-negative bacterium (E. coli) were treated with up to 5
pulses at 12.5kV cm-1. The results showed that E. coli is more sensitive to electric pulses
than M. glutamicus. Furthermore, the relative cell viability of E. coli was 74 (%), whereas
it was 87 (%) in the case of M. glutamicus although the electric field strength (12.5kV
cm-1) and pulse number (1) were kept constant. However, a strong influence of electric
pulses on the viability of E. coli cells was observed when the number of pulses was 5.
Figure 5.6 clearly shows that multiple pulses (5) affect the viability (almost 98% cells
were killed) of E. coli tremendously. On the other hand, M. glutamicus cells maintained
their viability to 64% even in the presence of similar field strength and pulse number. In
addition, the cell viability of M. glutamicus was approximately 13-fold higher than that of
E. coli. The results also confirmed that the viability of cells decreased with increasing the
pulse number, regardless of the strains applied for electroporation.
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
1
2
3
4
5
-20%
-40%
No. of pulses treated
M. glutamicus
E. coli
Figure 5.6 Effect of electric pulses on bacterial cell viability (pulsed at 12.5kV cm-1, 200 and
25µF).
159
5.3.5. Assessment of cells electropermeabilization with the aid of Bleomycin
M. glutamicus cells growing on Seed Medium were subjected to pulse at 12.5kV
cm-1, 200 and 25µF in presence of 5nM Bleomycin (BLM) in order to determine the
number of permeabilized cells by agar plate assay. It is observed that cell mortality
reached 17% in the case of samples treated with 5nM BLM, whereas approximately 10%
cells were killed in samples (without added BLM) although both cell suspensions were
pulsed at 12.5kV cm-1, 200 and 25µF (Figure 5.7), indicates that only 7% cells were
permeabilized due to this treatment. However, this study showed that approximately 5%
cells were killed due to the addition of BLM even though cell suspensions were not
treated by the electric pulses. It was assumed that higher concentrations of BLM might be
required for determining the membrane permeabilization of M. glutamicus since the size
and cell wall composition of this bacterium are completely different from the mammalian
cells. Therefore, cell suspensions were pulsed in presence of a range of BLM
concentrations (5 to 50nM). Nevertheless, the result did not show any trend of decreasing
the number of viable cells due to increase of BLM concentrations in the cell suspensions
of M. glutamicus (Figure 5.8).
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Pulse (-) bleo (-)
Pulse (-) bleo (+)
Pulse (+) bleo (-)
Pulse (+) bleo (+)
Figure 5.7 Effect of Bleomycin (5nM) on the viability of M. glutamicus subjected to a single
pulse at 12.5kV cm-1, 200 and 25µF.
160
100%
Relative cell viability (%)
80%
60%
40%
20%
0%
Control
5
10
15
20
25
50
Concentration of Bleomycin (nM)
Figure 5.8 Effect of a range of blemoycin concentrations on the viability of M. glutamicus
exposed to a single pulse at 12.5kV cm-1, 200 and 25µF.
It was also assumed that the electrical parameters (field strength, pulse duration
and number of pulses) imposed during pulsation in presence of BLM might not be
sufficient to create electropores on the cell membrane as well as to transfer BLM
molecules into the pulsed cells. Since the Bio-Rad Gene Pulser does not allow us to
increase the field strength 12.5kV cm-1 and regulate pulse duration, M. glutamicus cell
suspensions were repetitively pulsed (up to 5, without given any resting time between
two consecutive pulses) in presence of 5nM BLM. However, no prominent effect of BLM
addition on the viability of cells was observed due to increase of pulse numbers (Figure
5.9). It was hypothesized that M. glutamicus cells grown in the Seed Medium presented a
considerable barrier in transferring the electric current into the cell suspensions.
Therefore, cell pellets were suspended in a buffer (Tris buffer 1mM plus 270mM sucrose,
pH 7.2), and subsequently pulsed at 12.5kV cm-1. Although the number of killed cells due
to electric pulses was higher in the mentioned buffer as compared to the Seed Medium,
there was no indication that the use of a particular buffer instead of culture medium for
increasing the BLM cytotoxicity since higher cells mortality might
161
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
1
2
3
4
5
No. of pulses treated
Without Bleomycin
With Bleomycin
Figure 5.9 Effect of Bleomycin (5nM) on the viability of M. glutamicus grown on Seed Medium
subjected to multiple pulses at 12.5kV cm-1, 200 and 25µF.
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
1
2
3
4
5
-20%
No. of pulses treated
Without Bleomycin
With Bleomycin
Figure 5.10 Effect of Bleomycin (5nM) on M. glutamicus viability suspended in buffer (Tris
Buffer 1mM + 270 mM Sucrose) subjected to multiple pulses at 12.5kV cm-1, 200 and 25µF.
162
120%
100%
Relative cell viability (%)
80%
60%
40%
20%
0%
Control
1
2
3
4
5
-20%
-40%
-60%
No. of pulses treated
Without Bleomycin
With Bleomycin
Figure 5.11 Effect of Bleomycin (5nM) on the viability of E. coli grown on Nutrient Medium
subjected to multiple pulses at 12.5kV cm-1, 200 and 25µF.
have occurred due to the low due to the low conductivity of buffer (Figure 5.10). It was
then assumed that BLM might not be effective in the case of M. glutamicus cells because
of the presence a rigid cell wall. Hence, the electropermeabilization of E. coli grown on
Nutrient medium was investigated by adding 5nM BLM, however, there was no cytotoxic
effect observed in the pulsed cells of E. coli (Figure 5.11).
163
5.4. Discussion
The results obtained from this study clearly showed that increasing the electric
voltage and pulse number decreased the viability of M. glutamicus. After imposing a
single pulse to the cell suspension of M. glutamicus grown on Seed Medium, the
percentages of viable cells reached to 95, 92 and 87 (%) at 7.5, 10.0 and 12.5kV cm-1,
respectively (Figure 5.1). A similar trend was observed while the cell suspension was
treated with up to 5 pulses i.e., the relative cell viabilities were measured 74, 71 and 64
(%), respectively (Figure 5.1). It has been reported that high intensity of electric field
strength leads to the permeation of a wider area or to the formation of a large number of
pores that may exceed the resealing limit, and eventually results in large reduction in
number of cells (Hui, 1996). Dower et al. (1988) showed that increasing the field
strengths and pulse number decreased the viability of E. coli whilst developing a
transformation procedure by electroporation. The highest yield of transformants was
obtained while only 30 to 40% cells were survived due to the electric shock. Moreover,
Fiedler and Wirth (1988) demonstrated that application of multiple pulses reduced the TE
due to loss in cell viability while electroporating the plasmid DNA into Enterococcus
faecalis, E. coli and Pseudomonas putida. In literature, most of the investigators
determined the yield of transformant rather than cell viability while examining the effect
of electroporation factors to a particular microorganism. However, the efficiency of
electrotransformation is linked to cell viability since the transformation efficiency (TE)
has been observed to be limited by the viability of cells (Rols et al., 1992). For this
reason, maintaining the cell viability as high possible during pulsation is the most
important factor for the success of electroporation.
Similar to bacterial cells, Canatella et al. (2001) demonstrated that the viability of
prostate cancer cells (DU 145) decreased due to increase of field strengths although the
uptake of calcien, a model cell-impermeant, was observed to be increased with increasing
field strengths. The loss in cells (CHO) viability is dependent on the electrical
parameters, although the permeability of CHO cells increased because of increasing
electric field strengths. At 800V cm-1, the level of permeabilization reached 75%, whereas
the viability of cells decreased to 60% (Golzio et al., 1998). However, this study
164
confirmed that pulse with high voltage has a severe effect on the viability of cells
although a high membrane permeabilization might be observed after electroporation.
Hence, it is obvious that the pulsed electric field must be high enough to create
electropores on the cell membrane, nevertheless, it should not be exceeded to a certain
level that leads to excessive cell death. The viability of cells could be preserved by
decreasing the electric field intensity with longer pulse duration (Gabriel and Teissie,
1995; Wolf et al., 1994). Gabriel and Teissie (1997) reported that increasing the field
strength tends to increase the area of cell membrane, whereas increasing the pulse
duration enhances the degree of perturbation of the affected membrane area.
Nevertheless, the influence of pulse duration on the viability of M. glutamicus was not
investigated in this study since the Bio-Rad Gene Pulser does not allow the user to
regulate this parameter.
This study clearly showed that the viability of cells affected greatly whilst M.
glutamicus cells were repetitively pulsed without given any resting time (pulse gap)
between two consecutive pulses. This phenomenon was observed because the resealing
process of electroinduced cells might be hampered by not getting enough time to return
back to their normal physiological state. While allowing 20min resting time between two
successive pulses, the results showed that the relative cell viability of M. glutamicus was
increased approximately 3-10% depending on the field strengths and number of pulses
(Figure 5.3). Although cell viability was affected due to multiple pulses, pulsed cells with
given resting time might have enough time to rearrange their structure, whereas the
viability in the case of cells pulsed without providing any resting time was affected
enormously. Teissie et al. (1999) also demonstrated that the number of resealed cells after
pulsation was increased considerably by providing resting time between two pulses.
Apart from the consequence of multiple pulses on cell viability, there might be an
additional effect due to increase of temperature during multiple pulses. Fologea et al.
(2004) demonstrated that a prolonged exposure of cells at high temperature decreased cell
viability. In the case of in vitro experiments, this injurious effect might be controlled by
using a low ionic content pulsing buffer or treating cells with low electric voltage (Teissie
165
et al., 1999). Fiedler and Wirth (1988) obtained higher transformation efficiencies in
presence of three successive pulses with two intermittent cooling steps as compared to the
control (without maintaining the temperature). Furthermore, Teissie et al. (1999) also
demonstrated that the cytoplasmic concentration after pulsation is reduced because of
draining out of intracellular compounds from the cells. If this is the case, cells must face
hyperosmotic stress even after resealing of electropores, and consequently the viability of
cells is hampered due to the difference in turgor pressure. On the other hand, membrane
permeabilization also results in an entrance of water into cells, and consequently
increases cells volume that leads to the rupture of membrane (Golzio et al., 1998).
While investigating the effect of culture age/growth stage of cells that are used for
electroporation on the viability of M. glutamicus, the results showed that the relative cell
viabilities in mid-exponentially growing culture were significantly higher (5 to 20%,
depending on the electric field strengths and number of pulses) than that in cells grown at
the stationary phase. Although the deviation in cell viabilities between two different
growth stages of pulsed samples was not so prominent in presence of a single pulse, the
results demonstrated approximately 1.1 to 1.5-fold higher cell viabilities (due to 5 pulses)
in mid-exponentially growing cells as compared to the cells grown at the stationary phase
(Figure 5.4). Similar to this study, the electroporation of Corynebacteria growing at the
mid-exponential phase were resistant to killing by electric pulses and yielded the highest
level of transformants (107 transformants per µg DNA), whereas the number of
transformants was observed to be reduced in the case of cells grown at the stationary
phase (Bonamy et al., 1990). As with Corynebacteria cells, Miller et al. (1988) obtained
approximately 2 to 5-fold higher TE of C. jejuni when the mid-exponentially growing
cells were electroporated as compared to the cells growing at the early and late stationary
phases. However, this study confirmed that cells growing at the stationary phase are
sensitive to killing by electric pulses, and the intensity of electric field and energy
required for the membrane permeabilization of cells depend on the growth stage of cells.
However, this statement does not indicate that cells growing in exponential phase will be
suitable for pulsed electric field treatment while increasing the secretion of an
intracellular product during microbial fermentation is enhanced by transient membrane
166
permeabilization. In general, the production of metabolites during fermentation is
observed at the end of exponential or the start of stationary growth phase of
microorganism. If the cells growing at the production or stationary phases are treated by
the high intensity electric pulses, a major percentage of cells might be killed, cells are not
able to complete the rest period of fermentation and eventually result in decreasing the
overall yield of product.
Previous investigation already confirmed that the conductivity of medium in
which cells are suspended has a major influence on the electroexcretion of proteins or
enzymes from a particular host (chapter 4). Since the continuous enhancement of any
intracellular product secretion by electropermeabilization during microbial fermentation
is limited by cell viability (Rols et al., 1992), the effect of medium conductivity on the
viability of M. glutamicus was examined in this study. The results showed that cells
suspended in distilled H2O and buffers (1mM Tris plus 270mM sucrose and 50mM Tris
HCl) are more easily killed than cells suspended in Seed Medium; and the highest
percentage of cells reduction was observed in the case of cells suspended in distilled H2O
although the electrical parameters (field strengths and number of pulses) were kept
constant throughout the study. Furthermore, the effect of multiple pulses on the viability
of M. glutamicus was severe in the case of cells suspended in distilled H2O and
electroporation buffers as compared to the Seed Medium. After pulsing the cell
suspensions at 12.5kV cm-1 with 5 pulses, the percentages of viable cells suspended in
electroporation buffers and distilled water reduced to less than 7%, whereas cell mortality
measured in cells suspended in Seed Medium was only 36% (Figure 5.3).
The phenomenon observed in this study could be explained on the basis of
electrical conductivity of media/buffers in which M. glutamicus cells were suspended. It
is obvious that the medium conductivities of distilled H2O, 1mM Tris plus 270mM
sucrose and 50mM Tris HCl are lower than that of Seed Medium, and H2O has the least
conductivity among the all media used in this study. It has been demonstrated that
electroporation media with high conductivities reduced the electric voltage and pulse
duration (McIntyre and Harlander, 1989b). If this is the case, M. glutamicus cells might
167
not be imposed by the actual electric field strength applied in this study, and eventually
lower cell mortality was observed. On the other hand, medium with low conductivity
permits more pulses to be applied, results in higher membrane permeabilization, and
eventually more inactivation (Sixou et al., 1991). Djuzenova et al. (1996) also observed
that decreasing the extracellular medium conductivity resulted in lower viability of
murine myeloma cells. Furthermore, the rate of resealing of electropores is significantly
enhanced in high ionic strength medium resulting in high cell viability (Tekle et al.,
1994). Apart from the consequences of electric pulses, an additional effect on the
reduction of cell viability might have occurred due to the hypoosmotic stress (Csonka,
1989) caused by suspending cells in H2O and electroporation buffers. However, the
results obtained in this study confirmed that the composition of buffers used during
electroporation has an important influence on the viability of cells as well as on the yield
of product that will be excreted through the membrane permeabilization.
In Table 2.6, it is clearly showed that the number of electrotransformants is
dependent on the type of strains. The results obtained from this study confirmed that the
cell wall of M. glutamicus is more rigid to electric pulses than that of E. coli. Even after
imposing of 5 pulses at 12.5kV cm-1, approximately two-third (64%) of M. glutamicus
cells maintained their viability, whereas it was only 2% in the case of E. coli. This finding
can be explained in regards to the membrane structures of two different strains used in
this investigation. C. glutamicum (in this study, M. glutamicus), a Gram-positive
bacterium, has a cell wall of approximately 32nm that is composed of thick mesodiaminopimelic acid containing peptidoglycans and arabinogalactan (Marienfeld et al.,
1997). The inner cell membrane of C. glutamicum is composed of a lipid bilayer and the
outer membrane is arranged with long hydroxylated fatty acid chains called mycolic acids
(Puech et al., 2000). On the other hand, E. coli, a Gram-negative bacterium, has a thin
(generally 2-3nm thick) inner wall composed of peptidoglycan. Moreover, E. coli
contains an outer membrane with a lipid bilayer of 7nm thick composed of phospholipids,
lipoproteins, lipopolysaccharides and proteins (Madigan and Martinko, 2005). Weaver
and Chizmadzhev (1996) demonstrated that the mechanical stability of cell membrane
has an influence on the viability of cells as well as the efficiency of electroporation since
168
the reversible behaviour of membrane is dependant on its configuration. Hattermann and
Stacey (1990) also showed that the survival rate of cells due to electric pulses is
dependent on the type of cells while conducting electrotransformation of various cells.
Therefore, it is necessary to gain information regarding the structure or composition of
cell membrane of microorganism in which foreign gene or molecules will be transferred
or from which intracellular enzymes or protein are extracted by electropermeabilization.
In this study, the relative cell viability after pulsation was measured by the agar
plate assay although the determination of viable cells by this approach is not accurate.
This study showed that this process takes long incubation times (48-72hr) to form
countable colonies, and colonies were often observed to clump together and formed a
chain like structure. In addition, this technique was very sensitive to cross contamination
with airborne microorganisms. The measurement of cell viability by this method often
underestimates the number of viable cells in a given population, and only expresses the
cells that are actively multiplied or divided in selected culture conditions (Barer and
Harwood, 1999). Hence, the reduction in cells number does not necessarily correlate with
the cell mortality. However, a range of fluorescence techniques in combination with
florescence microscopy or fluorometry or flow cytometry have been carried out in order
to determine cell viability (Breeuwer and Abee, 2000; Nebe-von Caron et al., 1998;
Shapiro, 1995). All of these methods involve the use of different dyes that are either
excluded from or decolourised the viable cells, and thereby viable or non-viable cells are
easily detected under the fluorescence microscope.
DNA or nucleic acid probes, such as propidium iodide (PI) and ethidium bromide
(EtBr) are used for the assessment of cell viability where these probes are excluded from
the viable cells. It has been demonstrated that both the PI and EtBr are not capable of
crossing intact membrane of living microorganisms, but are able to pass through the
damaged cell membranes. Once PI enters inside the cells, it intercalates with DNA or
RNA and forms a bright red fluorescent complex. When EtBr binds with DNA, the
intensity of its colour increases approximately 20-fold, and it fluoresces with a red-orange
colour in presence of ultraviolet light (Bank, 1987). In general, PI/EtBr-stained cells are
169
assumed to be non-viable. Although PI has been used for the measurement of cell
viability, the major limitation of using this probe is its low extinction coefficient giving
relatively low fluorescence. Therefore, new DNA probes have been used as an alternative
that are highly fluorescent when bound to DNA. SYTOX Green, a nucleic acid stain, is
completely excluded from live eukaryotic and prokaryotic cells. When this stain binds to
the nucleic acids, the fluorescence emission is increased by more than 500-fold, and the
dead cells fluoresce green (Roth et al., 1997). Recently LIVE/DEAD BacLight Bacterial
Viability Kits, produced by Molecular Probes (USA) have been widely used to determine
the viability of bacteria in a large population. The kit contains two fluorescent nucleic
acid stains: the permeant SYTO 9 (green) and the non-permeant PI (red). When cells are
incubated with these two stains, the intact cells are labelled green, whereas cells with
damaged membranes are labelled red (Haugland, 1992). This kit has been used for
various bacterial species, such as Pseudomonas aeruginosa, E. coli, S. aureus and
Lactococcus lactis (Suller and Lloyd, 1999).
Flow cytometry in conjunction with different fluorescent dyes has recently been
established as a valuable tool to evaluate cell viability. It is more effective than agar plate
or microscopy or fluorometry technique (Diaper et al., 1992). In the case of flow
cytometry, samples are delivered at low flow rate and passed through a light beam that
permits the rapid measurement of scattered light and fluorescence emission from the
individual cells. This method is reliable since it determines the total cells number as well
as the number of fluorescence-labeled cells simultaneously (Breeuwer and Abee, 2000).
The use of fluorescent probes with flow cytometry also allows the simultaneous
multiparametric analysis of physical and chemical characteristics of a single cell flowing
through an optical or electronic detection apparatus (Davey and Kell, 1996). Flow
cytometric analysis offers the assessment of cell viability at a rate up to 50,000 cells per
second (Shapiro, 1995). A rapid flow cytometric method in conjunction with rhodamine
123 (below 0.5µM) has been developed in order to assess the viability of Micrococcus
luteus, a Gram-positive bacterium (Davey et al., 1993). Jepras et al. (1995) also
determined the viabilities of E. coli, Staphylococcus aureus and P. aeruginosa by flow
170
cytometry in combination with several fluorescent probes. Due to the limitation of time,
however, it was not possible to determine cell viability by this approach.
In this study, the effectiveness of BLM (5 to 50nM) for determining the
electropermeabilized cells in a pulsed population was investigated. However, none of the
attempts (i.e., increasing the concentration of BLM and number of pulses, the addition of
BLM to M. glutamicus cells suspended in a low conductivity medium and the
determination of electropermeabilized cells of E. coli by BLM addition) conducted in this
study did not show any prominent cytotoxic effect of BLM in pulsed cells. The entry of
BLM into the pulsed cells of M. glutamicus, even after treating with the high intensity
electric pulses, might be limited because of the resistivity of C. glutamcium cell wall to
antibiotics (Jarlier and Nikaido, 1994; Nikaido, 1994). However, the determination of
electropermeabilized mammalian cells in a pulsed population has been successfully
accomplished by the addition of 5nM BLM (Kotnik et al., 2000). Although BLM has
been regarded as a non-permeant cytotoxic drug (Orlowski et al., 1988), this study
showed that a tiny percentage of cells (5%) were killed by the addition of BLM in cell
suspensions. This phenomenon might have occurred due to the inaccuracy of agar plate
measurement since the viability of CHO cells was not shown to be affected without
imposing of electric pulses even after 18h of incubation in a culture medium contained
7000nM BLM (Akiyama et al., 1979). However, this study revealed that the evaluation of
electropermeabilized cells of M. glutamicus or E. coli in a pulsed population by means of
BLM is not such an efficient method as fluorescent nonpermeant dye measurement.
It is now obvious from this study that permeabilization is not similar to the
poration or induction of structural defects on the cell membrane due to electric pulses.
Electropermeabilization should be selective to a specific target membrane without
affecting the rest of the cell severely. Electroporated membranes may be permeabilized
for a protein or enzyme but may represent as an efficient barrier for others. Although this
study demonstrated that increasing the electric field strengths and number of pulses
enhanced the secretion of protein or enzymes (MDH and GDH) of both M. glutamicus
and E. coli by suspending the respective cells in low conductivity medium (distilled
171
H2O), this approach for increasing the yield of L-glutamate is complicated whilst the
fermentation is carried on (Chapter 4). In the case of product or enzyme (for example,
heterologous protein production in E. coli) that is not usually excreted into the
extracellular medium during fermentation, it is obvious that the resealing of
permeabilized membranes or maintaining cell viability after electropulsation is not
essential. However, a high degree of membrane reversibility or a long-term viability of
pulse-treated cells is mandatory while the continuous release of a product during
fermentation (for instance, the increase of L-glutamate yield) will be accomplished by
electropermeabilization. If the electric pulses kill a major percentage of cells, the rest of
viable cells will not be sufficient to complete the fermentation, and eventually result in
reducing the product yield and make the process expensive. Furthermore, the application
of electric fields often causes the electroconformational change of proteins (Chen and
Lee, 1994), which is a potential drawback in considering this approach for bioprocessing
i.e., effective secretion or separation of intracellular proteins or enzyme. Hence, further
studies are still needed in order to confirm the applicability of electroinduced extraction
of amino acids at an industrial level.
In conclusion, the results obtained from this research brought direct experimental
evidences that electroporation is a stress for biological cells. It is now apparent that
electropermeabilization of membrane depends on the electric factors (field strength, pulse
number and duration), physiochemical factors (medium conductivity and ionic
concentration of buffers), the properties of membrane (cell wall composition, structural
homogeneity) and the permeant molecule itself (polarity and size). It is, therefore,
necessary to optimize all the factors in order to maintain cell viability as high as possible
whether it will be designed for either gene transformation or the secretion of intracellular
products or introducing foreign molecules into the cell. If the application of intense
electric field affects cell viability, the efficiency of transformation or intracellular product
secretion, where biological systems with the continuous production of metabolites and
their subsequent extraction will be carried by the electric pulses, is reduced. It is expected
that the important facts observed in this study will assist in optimizing, developing and
validating the electropermeabilization protocols, and hence intensify the bioprocessing.
172
Chapter SIX
6. Osmoregulation of M. glutamicus: Effect of
Hyperosmotic Stress on Its Growth, Viability, Lglutamate Production and Cytoplasmic Enzymes or
Protein Level
6.1. Introduction
Although the cytoplasmic membrane of a microorganism is permeable to water, it
forms an effective barrier for solutes suspended in extracellular medium or metabolites
present in the cytoplasm. In general, the total concentration of osmotically active solutes
within cells is generally higher than that in extracellular environment, and therefore the
flow of water into cells is often observed due to its chemical potential (Csonka, 1989;
Csonka and Hanson, 1991). Since the permeability of water through the cytoplasmic
membrane is high, imposed imbalances between the turgor pressure exerted by cell
membrane and the osmolality gradient across the bacterial cell wall are short in duration,
and thus the cell division and growth of a microorganism are occurred (Koch, 1983). The
osmotic strength of an environment is one of the most important physical parameters that
determine the ability of organisms to grow in a given condition. However, cells are often
observed to face osmotic stresses that are defined as an increase or decrease in osmotic
strength or osmolality of the cultural medium in which an organism is usually grown.
Osmolality describes the osmotic pressure of a solution in osmoles (Osm) of osmolytes
per kg of solvent, whereas osmolarity is a measure of Osm of osmolytes per litre of
solution.
When fermentation is started with high concentration of nutrients, cells often
experience with the hyperosmotic stress. Moreover, the medium osmolarity is often
increased due to the efflux of amino or organic acids from cells (Guillouet and Engasser,
173
1995b). Hyperosmotic shock causes considerable shrinkage of cytoplasmic volume, may
result in cell mortality (Csonka, 1989), and eventually reduces the yield and productivity
of a bioprocess. On the other hand, a hypoosmotic stress is observed when the
intracellular products and by-products are accumulated within cells during fermentation.
Since the bacterial cell walls are rigid and can withstand pressures up to 100 atm
(Carpita, 1985), hypoosmotic shock generally results in minor increase of cells volume.
Figure 6.1 shows a schematic overview of the effects of osmotic stresses on water flux
and turgor pressure. In general, microorganisms have to cope with both conditions due to
the fluctuation in medium osmolarity during bioprocess, and are forced to develop
efficient adaptation mechanisms (Galinski, 1995). The active process by which
microorganisms survive with the osmotic stresses is defined as osmoregulation (i.e.,
accumulation or efflux of compatible solutes) that is dependent on the type of osmotic
stresses faced by cells (Morbach and Kramer, 2003). It has been demonstrated that
bacterial compatible solutes (Figure 6.2), cytoplasmic low-molecular-mass solutes, are
accumulated either by de novo biosynthesis (endogenous osmolytes, such as glutamate,
proline, ectoine, trehalose and sucrose) or by taking up (exogenous osmolytes, such as
glycine betaine) from the environment during osmoregulation (Csonka and Hanson,
1991; Poolman and Glaasker, 1998). These solutes cause rehydration of cytoplasm by
increasing the internal osmolarity, stabilize and protect enzymes, and support cells to
grow at their normal rate (Record et al., 1998).
In the case of hyperosmotic condition, an increase in external osmolarity or a
decrease in external water activity causes a rapid efflux of water that leads to the
dehydration of cells. Simultaneously, a reduction in cytoplasmic volume and the increase
in the concentrations of intracellular metabolites are occurred due to this perturbation,
and thus the growth of cells is diminished (Csonka, 1989). Three overlapping phases are
usually observed when bacterial cells are stressed by hyperosmotic conditions: 1)
dehydration of cytoplasm due to water efflux, 2) rehydration of cytoplasm by
accumulating ions or compatible solutes, and 3) cellular remodelling that results in
exchange of ionic osmolytes against compatible solutes, and leads cells to grow again
(Wood, 1999). However, the extent of cell volume reduction due to hyperosmotic stress
174
Hyperosmotic stress
Normal condition
Hypoosmotic stress
Figure 6.1 The effects of osmotic stresses on water flux and the turgor pressure (Morbach and
Kramer, 2002).
Figure 6.2 List of compatibles solutes involved in osmoregulation (Eggeling and Bott, 2005).
175
is dependent on the osmolality of media and the type of species. The volume reduction of
C. glutamicum cells measured at 1.5 and 3.3Osm kg-1 was 30% and 65% higher,
respectively, than the reduction observed while cells were cultivated at 0.4Osm kg-1
(Guillouet and Engasser, 1996). In the case of Bacillus subtilis, an increase in osmotic
upshift to 0.8Osm kg-1 resulted in decrease of 30% cells volume (Whatmore et al., 1990),
whereas the volume of cells (Halomonas elongata) decreased only 20% due to an
osmotic upshift to 2.2Osm kg-1 (Miguelez and Gilmour, 1994).
Under hyperosmotic stress, however, compatible solutes cause rehydration of
cytoplasm by increasing the internal osmolarity, and can be accumulated (up to molar
concentrations) in cytoplasm without disturbing the cellular functions (Record et al.,
1998). Furthermore, compatible solutes stabilize and protect enzymes mainly by being
excluded from the protein surface, thus leading to the preferential hydration of protein
(Arakawa and Timasheff, 1985). To overcome this hyperosmotic stress, bacteria rapidly
accumulate K+ ions into cells via specific transporters, and synthesize glutamate (the
counterion of K+) (Booth and Higgins, 1990). At high osmolarity, however, K+-glutamate
is not sufficient to ensure the growth of cells, and therefore bacteria replace the
accumulated K+ ions with the compatible solutes (Lucht and Bremer, 1994). Figure 6.3
represents a schematic diagram of the mechanism of compatible solutes involved during
osmoregulation. The mechanism of osmoregulation due to the hyperosmotic stress has
been investigated in both Gram-negative bacteria, such as E. coli (Grothe et al., 1986),
Salmonella typhimurium (Cairney et al., 1985) and Gram-positive bacteria, for instance
C. glutamicum (Farwick et al., 1995) and B. subtilis (Whatmore and Reed, 1990;
Whatmore et al., 1990). In C. glutamicum, a rapid but transient influx of Na+ during the
first 30min of osmotic upshocks and a strong accumulation of proline (increased from 5
to 110mg g Dw-1, at the end of the growth phase) were observed due to increase of
medium osmolality, from 0.4 to 2Osm kg-1 (Guillouet and Engasser, 1995a).
Peter et al. (1998) investigated the mechanisms involved during osmoadaptation
of C. glutamicum revealing that this organism is equipped with four uptake systems for
compatible solutes (Figure 6.4). Among them, three systems are osmoregulated. BetP,
176
Figure 6.3 A schematic diagram of the influx and efflux of compatible solutes during
osmoregulation (Morbach and Kramer, 2002).
Figure 6.4 Systems involved during the adaptation of Corynebacteria to osmotic stresses
(Morbach and Kramer, 2003).
177
specific for glycine betaine, is the highest affinity system; ProP, a medium affinity system
that is involved for taking up proline and ectoine; and EctP, a low affinity system that
could be activated for all the three compounds (Peter et al., 1998). Farwick et al. (1995)
demonstrated that glycine betaine is accumulated by a secondary transport system (BetP)
coupled to the transport of two Na+ ions, which was observed to be induced up to 8-fold
at hyperosmotic stress. However, the above-mentioned three systems are effectively
regulated at the level of activity, and the former two are controlled at the level of gene
expression (Peter et al., 1998). The expression of betP gene was observed to be
dependent on the medium osmolality, and the maximum rate of betaine uptake in cells
grown in media with low osmolality was 20-fold higher than that of high osmolality
(Peter et al., 1996). An additional proline uptake system (PutP) has been found in C.
glutamicum, however, it is used for anabolic purposes, and its physiological function is
not related to osmoprotection (Peter et al., 1997). However, all of these studies revealed
that bacteria are able to uptake or synthesize compatible solutes (glycine betaine, proline,
trehalose, glutamate and glutamine) during growth in high osmolality media. The former
two solutes are accumulated in higher intracellular concentration as compared to the
others, and play an important osmoprotective function in bacteria (Peter et al., 1998).
Nevertheless, it is evident that the synthesis of these compatible solutes through
the activation of their respective pathways is dependent on the type of bacterial species
and the composition of medium in which a microorganism is grown. Frings and his coworkers (1993) demonstrated that ectoines (for example, tetratrahydropyrimidines) are
the main compatible solutes in the genus Brevibacterium, whereas the accumulation or
synthesis of glycine betaine is mainly occurred during the osmoadaptation of
Corynebacterium. In addition, the intracellular proline concentration was observed to
increase (up to 100µmol g Dw-1) in C. ammoniagenes grown on media containing 1-5%
NaCl (Tomita et al., 1992), whereas the proline pool of C. ammoniagenes grown on
glucose-yeast-salt medium (8% NaCl) was not shown to exceed 50µmol g Dw-1 (Frings et
al., 1993). Whatmore et al. (1990) reported that a sudden osmotic upshock with 0.4M
NaCl triggered potassium uptake, raising the potassium pool from 350nM to 650nM.
178
In the case of hypoosmotic condition, on the other hand, an increase in external
water activity or a sudden decrease in external solute concentration causes massive influx
of water into cells, increases the turgor pressure and membrane tension, and eventually
disrupts the plasma membrane (Csonka, 1989). Wood (1999) mentioned that the
responses due to osmotic downshifts occur in three phases: 1) water uptake 2) extrusion
of water and cosolvents and 3) cytoplasmic cosolvent reaccumulation and cellular
remodelling. To survive in this dangerous situation, bacteria release compatible solutes in
extracellular medium through the emergency release valves, so-called mechanosensitive
(MS) channels, and consequently the flow of water into cells is reduced by lowering the
internal osmolarity (Figure 6.4). The hypoosmotic induced efflux of compatible solutes
has been investigated both in Gram-negative (Koo et al., 1991; Lamark et al., 1992) and
Gram-positive bacteria (Glaasker et al., 1996; Ruffert et al., 1997). In all cases, the
effluxes of glycine betaine, proline and choline are dependent on the osmotic conditions.
In C. glutarnicum, the release of glycine betaine and proline were observed to occur
through the osmoregulated channel (similar to the MS channels of E. coli) with an efflux
rate of 6000µmol min-1 g dm-1 or higher. However, the release of glutamate or lysine was
restricted, and ATP was completely retained in cells even after severe hypoosmotic
stress. The efflux of compatible solutes through the osmoregulated channel is tightly
regulated at the level of activity, but is independent on the growth conditions (Ruffert et
al., 1997). Although the efflux of compatible solutes occurred due to the action of MS
channels, it can also be mediated by carriers. In Lactobacillus plantarurn, a rapid efflux
is most probably mediated by channel activity followed by a slow process, carrier
mechanism (Glaasker et al., 1996).
During the industrial fermentation of C. glutamicum, a large amount of carbon
and nitrogen source is required in order to produce bulk quantity of L-amino acids (Ikeda,
2002). Since the osmotic pressure of industrial media is generally higher than that of
media used in laboratory experiments, the growth of this bacterium is often stressed
leading to the reduction of yield and productivity of amino acids (Kawahara et al., 1989).
It has been demonstrated that increase of medium osmolality from 0.4 to 2Osm kg-1
during the batch cultivation of C. glutamicum resulted in a decrease of specific growth
179
rate, from 0.7 to 0.2h-1, and biomass yield, from 0.6 to 0.3g g-1 (Guillouet and Engasser,
1995a). While conducting a glucose-limited continuous cultivation of C. glutamicum for
the production of L-glutamate, Guillouet and Engasser (1995b) observed a decrease of
biomass production, from 7.5 to 5.5g Dw l-1, due to increase of medium osmolality, from
0.4 to 2Osm kg-1. Even though the mechanisms involved during osmoregulation i.e.,
accumulation or efflux of compatible solutes (Guillouet and Engasser, 1995a; Kawahara
et al., 1989; Peter et al., 1998) and the effects of hyperosmotic stress on the growth and
viability of C. glutamicum (Guillouet and Engasser, 1995b; Skjerdal et al., 1995) have
been investigated, the consequence of this stress on the production of amino acids is not
intensively examined.
So far, only a few studies have been conducted in order to investigate the effects
of osmolality on amino acids production in C. glutamicum and its related strains
(Delaunay et al., 1999b; Gourdon et al., 2003; Kawahara et al., 1990; Ronsch et al.,
2003). Delaunay et al. (1999b) demonstrated that C. glutamicum produced 100g l-1 Lglutamate with a resulting osmotic potential of approximately 2Osm kg−1 decreasing the
productivity of this amino acid throughout the fermentation due to its accumulation.
Furthermore, the metabolism of C. glutamicum is subjected to strong and rapid changes
by the alteration of its surrounding osmolality although this soil bacterium is equipped
with powerful mechanisms to adapt to the hyperosmotic and hypoosmotic conditions
(Morbach and Kramer, 2003). Therefore, intensive researches are still required in order to
understand the osmotic stresses associated during the industrial production of amino acids
by C. glutamicum (in this study, M. glutamicus). The objective of this study is to
investigate the effects of hyperosmotic condition on cell viability, L-glutamate
production,
cytoplasmic
enzymes
(i.e.,
malate
dehydrogenase
and
glutamate
dehydrogenase), and total protein concentration of M. glutamicus. In addition, the
consequence of externally added compatible solutes (i.e., glycine betaine and proline) on
the growth of osmotically stressed cells will be examined.
180
6.2. Material and Methods
6.2.1. Chemicals
Peptone from pancreatically digested casein and meat extract were obtained from
VWR (Merck, UK). Yeast extract and bacteriological agar were procured from Oxoid,
UK; and D-glucose, urea, NaCl and all other chemicals were purchased from Fisher, UK
unless otherwise mentioned. Glycine betaine (N, N-Dimethylglycine) and L-proline were
ordered from Sigma-Aldrich, UK.
6.2.2. Organism and cultivation
M. glutamicus DSM 20300 (Collins et al., 1977; Suzuki et al., 1981; Yamada and
Komagata, 1970; Yamada et al., 1976) supplied by the German Collection of
Microorganisms and Cell Culture (DSMZ-Deutsche Sammlung von Mikroorganismen
und Zellkulturen GmbH) was cultivated in Medium 53 (10g l-1 peptone, 5g l-1 yeast
extract, 5g l-1 glucose, 5g l-1 NaCl, 1000ml distilled water, pH adjusted to 7.2-7.4)
according to the supplier’s instructions. The stock agar plates were prepared according to
the method described previously in Section 3.2.2. This bacterium was grown on Seed
Medium, and the composition and preparation of medium was described previously in
Section 3.2.2. M. glutamicus was cultivated at 30C and 220rpm, and all the experiments
were performed in 250ml shake flasks containing 50ml of culture medium.
6.2.3. Microbial growth measurement
The growth of M. glutamicus cells was periodically determined by measuring the
absorbance at 600nm (600) using spectrophotometer (S1200, WPA). Samples were
diluted (if concentrated) to keep the absorbance within the range of 0.1-0.3, and thereafter
multiplied with the dilution factor in order to obtain the actual optical density (OD). The
steps for dry weight measurement were described previously in Section 3.2.3.
6.2.4. Calculation of maximum specific growth rate (µmax)
The maximum growth rate (µmax) of M. glutamicus under different growth
conditions was determined according to the method described in Section 3.2.4. In this
181
study, µmax was calculated by taking the slope of a plot of cell density (OD),
corresponding to the exponential growth phase versus time.
6.2.5. Hyperosmotic shock to the cell suspension of M. glutamicus
Growing culture of M. glutamicus was transferred into the Seed Medium (50ml in
250ml flask) in such a way that the starting OD of culture medium was 1. Cells were
hyperosmotically shocked by the addition of NaCl where the concentrations of salt in
medium reached 0.5M, 1.0M and 1.5M. Three parallel shake flasks at each osmotic
strength were cultivated throughout the study, and the number of samples analyzed on
each occasion was two. In order to investigate the effect of hyperosmotic stress on the
different stages of growth cycle, NaCl was added both at the start and exponential phase
of cultivation. A control experiment was carried out simultaneously in which no salt was
added into the Seed Medium. The effect of NaCl addition on the growth of M. glutamicus
was determined by measuring the ODs at different growth points of cultivation. In the
case of biotin limited CGXII Minimal Medium, however, NaCl was added only at the
start of fermentation in order to investigate the effect of this stress on L-glutamate
production.
6.2.6. Viability assay
The viability of osmotically stressed M. glutamicus cells was determined by the
traditional agar plate (LB) assay. The composition and preparation of plates were
mentioned in Section 5.2.4. The osmotically stressed cells were collected, serially diluted
and 100µl samples were spread on agar plates aseptically. Plates were then kept in an
incubator at 30C, and colonies were counted after 24-72h of incubation. Since the
viability assay by the agar plate is not precise enough, all the experiments conducted in
this study were carried out at least two times, and the number of hyperosmotically
stressed samples spread on agar plates was three. The viability of cells was calculated by
normalizing the control (without NaCl addition) to have 100% viability. The ratio of
viable colonies in hyperosmotic stressed samples to that of control was then used to
determine the relative cell viability at a given osmotic strength.
182
6.2.7. Osmoregulation by the addition of compatible solutes
In order to investigate the effect of compatible solutes on the growth of this
industrially important bacterium, a range of concentrations (10 to 100mM) of both
glycine betaine and proline were added to the hyperosmotically stressed M. glutamicus
grown on Seed Medium. A control experiment was carried out at the same time where
none of those solutes were added to the hyperosmotically stressed culture of M.
glutamicus.
6.2.8. L-Glutamate production in M. glutamicus under biotin limited condition
L-glutamate was produced in M. glutamicus by cultivating cells in biotin limited
CGXII Minimal Medium (Keilhauer et al., 1993). The preparation of this medium was
described earlier in Section 3.2.5.
6.2.9. Quantification of glucose, L-glutamate and other amino acids
The quantification of substrate (glucose), L-glutamate and all the other amino
acids were achieved with the use of AAA-Direct TM Amino Acid Analysis System
(Dionex, UK). The preparation of eluents and standard were described previously in
Section 3.2.8.
6.2.10. Determination of total protein, GDH and MDH
The concentrations of total protein, glutamate dehydrogenase (GDH) and malate
dehydrogenase (MDH) were determined according to the methods described in Section
4.2.7.
183
6.3. Results
6.3.1. Effects of hyperosmotic stress on M. glutamicus growth and viability
In order to investigate the effect of hyperosmotic stress on M. glutamicus growth,
this bacterium was grown on Seed Medium having a range of NaCl concentrations (0.5 to
1.5M) added at the start of cultivation. The results demonstrated that the growth of cells
was inhibited due to the addition of salt in Seed Medium, and the amounts of biomass
produced in an osmotically stressed cultivation are lower than that of control (without
NaCl). The biomasses (OD600) measured after 24h of cultivation were 42.80, 35.0, 28.80
and 18.10 in the case of control, medium containing of 0.5M, 1.0M and 1.5M NaCl,
respectively (Figure 6.5). The results indicated that the degree of biomass reduction is
dependent on the osmotic strength of medium in which cells are grown. In addition, the
effect of NaCl on the specific growth rate (µ) of M. glutamicus cells was investigated i.e.,
0.30h-1, 0.23h-1, 0.19-1 and 0.19h-1 in the case of control, medium containing of 0.5M,
1.0M and 1.5M NaCl, respectively. These results confirmed that the growth rate of
microorganism is decreased with the increase of medium osmolarity.
50
OD at 600nm
40
30
20
10
0
0
5
10
15
20
25
Time after inoculation, hr
No NaCl
0.5M NaCl
1.0M NaCl
1.5M NaCl
Figure 6.5 The growth of M. glutamicus in the presence of different concentrations of NaCl,
added at the beginning of cultivation.
184
30
50
OD at 600nm
40
30
NaCl
addition
20
10
0
0
5
10
15
20
25
Time after inoculation, hr
No NaCl
0.5M NaCl
1.0M NaCl
1.5M NaCl
Figure 6.6 The growth of M. glutamicus in the presence of different concentrations of NaCl,
added at the exponential phase (7h) of cultivation.
In the case of the experiment mentioned above (Figure 6.5), NaCl was added at
the start of cultivation. Since the number of cells (inoculum) at the beginning of
cultivation is usually less, it is obvious that the number of hyperosmotically stressed
viable cells after the initial osmotic upshock will be apparently low. Because of this, a
prolonged lag phase and decreased biomass production at the end of cultivation were
observed. Therefore, the effect of NaCl addition to the exponentially growing (7h)
cultures of M. glutamicus cultivated in Seed Medium was investigated. The biomass
(OD600) measured after 24h of cultivation were 42.80, 36.75, 31.60 and 26.10 in the case
of control, medium containing of 0.5M, 1.0M and 1.5M NaCl, respectively (Figure 6.6).
These results demonstrated that hyperosmotic stress results in decrease of cell’s growth
and biomass production, irrespective of the growth stages of cell at which NaCl is added.
It has already been investigated that microbial cells often could not maintain their
viability due to this sudden osmotic up and down-shock (Csonka, 1989; Wood, 1999).
During industrial fermentation, if a certain percentage of cells are killed due to the
185
30
osmotic stresses, the yield and productivity of a bioprocess are affected enormously.
Hence, the effect of hyperosmotic condition on the viability of M. glutamicus was
investigated. Both the osmotically stressed (by adding NaCl at the start and exponential
growth phase of cultivation), and control cells grown on Seed Medium were collected
after 2 and 4h of salt addition, serially diluted with sterile H2O and spread on agar plates.
The results demonstrated that addition of NaCl in growth medium increased the mortality
of cells, and the degree of cell mortality is dependent on the osmotic strength of medium
(Figure 6.7 and 6.8). However, the extent of cell viability reduction in cultures in which
salt was added at the start of cultivation is considerably higher than that observed in
cultures where cells grown at the exponential growth phase were hyperosmotically
stressed. In the case of the former experiment (Figure 6.7), the percentages of killed cells
reached 79 and 81 (%) after 2 and 4h of cultivation, respectively, whereas it was 55 and
57 (%) for the latter case (Figure 6.8) although NaCl concentration in both conditions was
kept constant to 5M.
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
0.5M
1.0M
1.5M
2.5M
5.0M
Concentration of NaCl
2hr
4hr
Figure 6.7 The reduction in cell viability in the presence of different strengths of hyperosmotic
stress (addition of NaCl at the start of cultivation).
186
120%
Relative cell viability (%)
100%
80%
60%
40%
20%
0%
Control
0.5M
1.0M
1.5M
2.5M
5.0M
Concentration of NaCl
2hr
4hr
Figure 6.8 The reduction in cell viability in the presence of different strengths of hyperosmotic
stress (addition of NaCl at the exponential phase, after 7h of inoculation).
6.3.2. Effects of glycine betaine and proline addition on the growth of
hyperosmotically stressed M. glutamicus
In order to investigate the effect of compatible solutes (for instance, glycine
betaine and proline) on the growth of hyperosmotically stressed M. glutamicus, a range of
glycine betaine concentrations (10 to 100mM) was added (at the start of cultivation) into
the Seed Medium containing 0.5M NaCl. However, there was no significant effect
observed on the growth of M. glutamicus due to the addition of this compatible solute in
culture medium. Moreover, the result showed that increase of glycine betaine
concentration more than 30mM decreased biomass production at the end of cultivation
(Figure 6.9). Ronsch et al. (2003) demonstrated that the addition of glycine betaine does
not accelerate the growth of cells or change the time at which the maximum growth rate
is reached under the condition of low or medium osmotic stress, although the cytoplasmic
volume was observed to be higher in presence of glycine betaine. It was thus assumed
that the effect of externally added glycine betaine to the growth of hyperosmotically
stressed M. glutamicus might only be observed in the presence of osmotic strengths
187
40
OD at 600nm
30
20
10
0
0
5
10
15
20
25
30
Time after inoculation, hr
Control
10mM
20mM
30mM
50mM
100mM
Figure 6.9 Effect of a range of concentrations of glycine betaine (mM) on the growth of M.
glutamicus grown on hyperosmotic condition (Seed Medium containing 0.5M NaCl).
50
OD at 600nm
40
30
20
10
0
0
5
10
15
20
25
30
Time after inoculation, hr
Control
Control + Betaine
0.5M
0.5M + Betaine
1.0M
1.0M + Betaine
1.5M
1.5M + Betaine
Figure 6.10 Effect of glycine betaine (20mM, added at start of cultivation) on the growth of M.
glutamicus grown on a range of hyperosmotic strengths (Seed Medium containing 0.5-1.5M
NaCl, added at start of cultivation).
188
50
OD at 600nm
40
30
Addition of both NaCl
and Glycine betaine
20
10
0
0
5
10
15
20
25
30
Time after inoculation, hr
Control
Control + Betaine
0.5M
0.5M + Betaine
1.0M
1.0M + Betaine
1.5M
1.5M + Betaine
Figure 6.11 Effect of glycine betaine (20mM, exponential phase) on the growth of M. glutamicus
grown on a range of hyperosmotic conditions (Seed Medium containing 0.5-1.5M NaCl, added to
the exponential growth phase).
40
35
OD at 600nm
30
25
20
15
10
5
0
0
5
10
15
20
25
30
Time after inoculation, hr
Control
10mM
20mM
30mM
50mM
100mM
Figure 6.12 Effect of a range of concentrations of proline on the growth of M. glutamicus grown
on hyperosmotic condition (Seed Medium containing 0.5M NaCl).
189
higher than 0.5M NaCl. Hence, 20mM glycine betaine was added both at the start and to
the exponential growth phase of cells grown on Seed Medium having a range of NaCl
concentrations (0.5 to 1.5M). However, the results showed that the inhibition of cells
growth due to NaCl addition could not be minimized by this compatible solute, regardless
the growth phases at which it was added into the Seed Medium (Figures 6.10 and 6.11).
Like glycine betaine, proline with a range of concentrations (10 to 100mM) was added (at
the start of cultivation) into Seed Medium containing 0.5M NaCl, however, no prominent
effect on the growth of M. glutamicus was observed (Figure 6.12).
6.3.3. Effects of hyperosmotic stress on L glutamate production by M.
glutamicus
M. glutamicus was grown on biotin limited (1µg l-1) CGXII Minimal Medium
with a range of osmotic strengths (0.5 to 1.5M NaCl, salt was added at the start of
fermentation in order to investigate the effect of NaCl addition on cells growth, glucose
consumption and L-glutamate production in M. glutamicus. Three parallel shake flasks at
each osmotic strength were cultivated throughout the study, and the number of samples
analyzed on each occasion was two. The results demonstrated that the growth of cells and
glucose consumption decreased due to the addition of salt in CGXII Medium, and the
degree of biomass reduction and substrate consumption is dependent on the osmotic
strength of the medium (Figure 6.13). The amount of biomass (OD600) after 72h of
cultivation measured in CGXII Medium containing 1.5M NaCl was approximately 50%
lower than that of the control. In addition, the residual glucose concentration in glutamate
production medium containing 1.5M NaCl was 13.05g l-1 (at the end of fermentation),
whereas it was only 3.73g l-1 in the case of the control (Figure 6.13). As with cells growth
and glucose consumption, it was observed that L- glutamate production was reduced
noticeably by the addition of salt in biotin limited CGXII Minimal Medium. Figure 6.14
shows that the extent of L-glutamate production is dependent on the osmotic strength of
the production medium. After 72h of fermentation, the amount of L-glutamate in control
was measured as 52.95mM, whereas it was only 16.13mM in the case of medium with an
osmotic strength of 1.5M NaCl (Figure 6.14).
190
35
45
40
30
20
25
15
20
15
10
10
5
5
0
0
0
10
20
30
40
50
60
70
80
Time after inoculation, hr
OD, Control (1.0µg/l biotin)
OD, 1.0µg/l biotin + 1.5M NaCl
Glu, 1.0µg/l biotin + 1.0M NaCl
OD, 1.0µg/l biotin + 0.5M NaCl
OD, 1.0µg/l biotin + 1.0M NaCl
Glu, Control (1.0µg/l biotin)
Glu, 1.0µg/l biotin + 0.5M NaCl
Glu, 1.0µg/l biotin + 1.5M NaCl
Figure 6.13 Effect of hyperosmotic stress on the growth and glucose consumption of M.
glutamicus grown on biotin limited (1µg l-1) CGXII Minimal Medium.
60
50
Glutamate, mM
OD at 600nm
30
Glucose, g/l
35
25
40
30
20
10
0
0hr
24hr
48hr
72hr
Time after inoculation, hr
Control (1.0µg/l biotin)
1.0µg/l biotin + 0.5M NaCl
1.0µg/l biotin + 1.0M NaCl
1.0µg/l biotin + 1.5M NaCl
Figure 6.14 Effect of hyperosmotic stress on the production of L-glutamate in M. glutamicus.
191
6.3.4. Effects of hyperosmotic stress on the activity of enzymes (MDH and GDH)
and total protein of M. glutamicus
In this study, the effect of hyperosmotic stress on the activity of total protein and
enzymes (GDH and MDH) of M. glutamicus was examined. M. glutamicus was
cultivated in Seed Medium with a range of NaCl concentrations (0.5 to 1.5M). The results
showed that addition of NaCl into medium increased the amount of total protein, GDH
and MDH of M. glutamicus (Figure 6.15 to 6.17). The total protein concentration (mg ml1
) in control was 1.94, whereas it increased to 2.47, 2.47 and 2.95 in Seed Medium with
osmotic strength of 0.5, 1.0 and 1.5M NaCl, respectively. The activity of GDH (U ml -1)
in control was 0.34 while hyperosmotic stress increased it to 0.43, 0.43 and 0.45 in
presence of 0.5, 1.0 and 1.5M NaCl, respectively. Similarly, the activity of MDH (U ml-1)
in control was 12.22, whereas it increased to 17.30, 16.03 and 19.84 in Seed Medium
with osmotic strength of 0.5, 1.0 and 1.5M NaCl, respectively. These results confirm that
hyperosmotic stress also influences the activity of cytoplasmic enzymes and proteins.
Total protein, mg/ml
4
3
2
1
0
Control
0.5M
1M
1.5M
Figure 6.15 Effect of hyperosmotic stresses on total protein (TP) of M. glutamicus grown on
Seed Medium.
192
0.6
GDH, U/ml
0.4
0.2
0
Control
0.5M
1M
1.5M
Figure 6.16 Effect of hyperosmotic stresses on glutamate dehydrogenase (GDH) of M.
glutamicus grown on Seed Medium.
30
MDH, U/ml
20
10
0
Control
0.5M
1M
1.5M
Figure 6.17 Effect of hyperosmotic stresses on malate dehydrogenase (MDH) of M. glutamicus
grown on Seed Medium.
193
6.4. Discussion
In this study, M. glutamicus was cultivated in Seed Medium containing different
concentrations of NaC1 in order to investigate the effect of this hyperosmotic stress on
the growth and viability of cells, substrate consumption and L-glutamate production. The
results obtained from this study clearly demonstrated that addition of NaCl in Seed
Medium decreased the specific growth rate of M. glutamicus and reduced the biomass
production, irrespective of the growth phases at which salt was added. However, the
effect of this stress on cell’s growth was more severe in cultures that were
hyperosmotically stressed from the beginning of cultivation than in cultures in which salt
was added to the cells grown at the exponential phase. In presence of high medium
osmolarity (medium containing 1.5M NaCl), the amount of biomass (OD600) measured
after 24h of cultivation was 26.10 (in latter condition, Figure 6.6), whereas it was only
18.10 (in former condition, Figure 6.5). It is obvious that microbial fermentation is started
with inoculum in which a low number of cells are usually inoculated. If cells are
hyperosmotically stressed at the beginning of fermentation, a large percentage of cells
must be killed within short period of NaCl addition affecting the growth properties
severely. On the other hand, the application of similar strength of osmotic stress to the
exponentially grown cells will not affect the growth of cells as like as the former
treatment.
The results also showed that the degree of cell growth inhibition is dependent on
the osmotic strength of medium in which M. glutamicus was grown. During the first few
hours (2-6h) of salt addition, the growth of cells was inhibited severely by the high
osmolarity (medium containing 1.5M NaCl). On other hand, when the osmotic stress was
not rigorous (medium containing 0.5M NaCl), cells were shown to regain their ability to
grow exponentially even after an extended lag phase. Similar to this observation, Csonka
(1989) mentioned that the cytoplasmic volume of cells stressed by low medium
osmolarity is adjusted as a result of osmotic balances even after a slow growth period.
Skjerdal et al. (1995) observed that the specific growth rates of these bacteria (B.
lactofermentum and C. glutamicum) decreased in a linear fashion by the increase of
medium osmolality. Moreover, the total volume of cells was observed to decrease
194
immediately upon exposure to 0.6M NaCl, and approximately 25-30% of cell volume
was reduced within 2-15min after hyperosmotic stress. Ronsch et al. (2003) showed that
the growth rate and cytoplasmic volume of C. glutamicum MH20-22B (a L-lysine
producing strain) decreased linearly with increasing osmotic stress.
This study demonstrated that the percentage of viable cell (M. glutamicus)
reduction is increased with the increase of medium osmolality. In the case of the former
condition (where NaCl was added at the start of cultivation), only 21 and 19 (%) cells
were viable after 2 and 4h of cultivation, respectively, whereas it was 44 and 43 (%) in
cultures where salt was added at the exponential growth phase although the concentration
of NaCl in both conditions was 5M (Figure 6.7 and 6.8). This result confirmed that cells
grown in the lag phase are more susceptible to hyperosmotic stress than cells grown in
the exponential growth phase. This phenomenon might have observed due to the fact that
hyperosmotic stress causes diffusion of water from cells, resulting in cell shrinkage,
arrests growth of cells and eventually causes cell death indicating that medium osmolarity
is one of the major operational parameters during industrial fermentation of
microorganism. Similar to this study, Skjerdal et al. (1995) reported that increase or
decrease of turgor pressure due to differences in solute concentration across the
cytoplasmic membrane causes cell inactivation. Furthermore, the reduction in cell
viability of Gram-negative bacteria due to osmotic stress is higher than that of Grampositive strains (Poolman and Glaasker, 1998). The reason behind this phenomenon
might be the strong adhesion between the cytoplasmic membrane and peptidoglycan in
Gram-positive bacteria.
Although a range of concentrations (10 to 100mM) of glycine betaine and proline
was added into the cells stressed by the addition of 0.5M to 1.5M NaCl in the Seed
Medium, no prominent influence of compatible solutes on the growth of M. glutamicus,
irrespective of the growth phases at which salt was added (Figure 6.9, 6.10, 6.11 and
6.12). In addition, a decreased amount of biomass was observed at the end of cultivation
due to the addition of compatible solutes ( 50mM). However, the wild type of C.
glutamicum (ATCC 13032) was shown to uptake glycine betaine and its precursor
195
dimethyl-glycine, ectoine and proline when these compatible solutes were added into the
culture medium having an osmotic strength of 1.5M NaCl (Farwick et al., 1995). Their
results also demonstrated that the accumulation and uptake rates of compatible solutes are
dependent on the osmotic strengths of medium, and addition of these solutes enhances the
growth rate of osmotically stressed C. glutamicum. Ronsch and his co-workers (2003)
reported that the addition of betaine reduces the time required to reach the maximal cell
growth at high osmolality, and results in increase of cell viability.
Apart from the influences of compatible on C. glutamicum growth, the addition of
20mM glycine betaine to the fermentation medium with an osmotic pressure of 2.12Osm
kg-1 increased the sugar consumption rate, from 8.8 to 14nmol min-1 mg Dw-1 and Llysine production rate, from 5.3 to 8.7 8.8 to 14nmol min-1 mg Dw-1 (Kawahara et al.,
1990). In Corynebacteria, however, potassium glutamate was observed to be the main
osmoregulator at low to moderate stress, while proline was accumulated at higher stresses
(Kawahara et al., 1989). It has also been demonstrated that this bacterium is able to
synthesize glutamine, proline and trehalose after an osmotic upshift even in the absence
of externally added compatible solutes (Frings et al., 1993). Ronsch et al. (2003)
observed an accumulation of proline (increased 8-fold) and betaine (reached up to
250nM) under severe osmotic stress (increase of osmolality from 1.0 to 2.5Osm kg-1).
Guillouet and Engasser (1995b) also demonstrated a rapid increase of intracellular
proline, from 5 to 125mg g Dw-1 and trehalose, from 20 to 60mg g Dw-1 due to increase
of medium osmolality, from 0.4 to 2Osm kg-1. Nevertheless, it has been reported that this
bacterium prefers the uptake compatible solutes rather than synthesis within cells because
of lower energy cost (Morbach and Kramer, 2002). Hence, much research are still
required in order to investigate the effect of externally added compatible solutes (mainly
glycine betaine and proline) to the high osmolality fermentation medium in which amino
acids (L-glutamate and L-lysine) are produced industrially.
In the case of industrial bioprocesses, particularly in fed-batch cultivation, cells
are often exposed to changes in osmolality due to the composition of medium and the
accumulation of product (Delaunay et al., 1999b; Guillouet and Engasser, 1995b).
196
Moreover, the excreted product in extracellular medium increases the osmotic strength of
the environment, modifies the biochemical activity of the substrate transport system,
affects the sugar consumption capacity, and finally leads to slow down in the metabolic
rates of cells (Gourdon et al., 2003). This study showed that addition of NaCl (at the start
of fermentation) in CGXII medium markedly reduced the L-glutamate production by M.
glutamicus. Also, the extent of L-glutamate production is dependent on the osmotic
strengths of fermentation media since the amounts of L-glutamate measured after 72h of
fermentation were 52.95mM, 32.66mM, 24.37mM and 16.13mM in the case of CGXII
medium without NaCl (control), 0.5M, 1.0M and 1.5M NaCl, respectively. These results
might have occurred due to the fact that hyperosmotic stress causes a decrease in specific
growth rate of M. glutamicus resulting in an increase proportion of sugar catabolized for
maintenance requirements, and eventually decreased the yield of L-glutamate.
Nevertheless, this study confirmed that the osmotic pressure of industrial media is one of
the limiting factors during amino acid fermentation.
Similar to this study, Guillouet and Engasser (1995b) demonstrated that the
medium osmolality represents one of the major operational parameters for an efficient
production of glutamate by C. glutamicum. Gourdon et al. (2003) showed that medium
osmolarity is directly related to the product accumulation during L-glutamate
fermentation, and increasing the medium osmolarity inhibits the L-glutamate
overproduction in C. glutamicum. However, this inhibitory effect is reversible since
washed cells retained their ability to transport sugar while removing the osmotic stress.
Furthermore, the specific rate of glutamate production was observed to decrease
throughout the fermentation as a function of glutamate accumulation (Delaunay et al.,
1999b). However, this severe effect of osmotic stress could be minimized by the addition
of glycine betaine during amino acid fermentation. In the case of L-lysine production in
C. glutamicum MH20-22B, addition of this compatible solute into fermentation medium
having high osmolality reduced the time necessary to reach the stationary phase, resulted
in synthesis of L-lysine at earlier fermentation times and increased both product
concentration and yield (Ronsch et al., 2003). Nevertheless, the effect of compatible
solutes on the production of L-glutamate was not investigated due to time limitation.
197
Furthermore, this study showed increased activities of total protein, GDH and
MDH of M. glutamicus because of NaCl addition into Seed Medium. At an osmotic
strength of 1.5M NaCl, the concentrations of total protein, GDH and MDH were
approximately 1.7, 1.38 and 1.55-fold, respectively, higher than that of control. It is now
obvious that hyperosmotic stress has also a major influence on the expression of proteins
or enzymes, and these results indicated that cells may need to synthesis some proteins in
order to cope with the hyperosmotic stress. Like this investigation, Clark and Parker
(1984) showed an induction of a group of proteins (E. coli) because of increasing the
osmolality in culture medium, whereas an osmotic downshift resulted in the repression of
those proteins. Furthermore, increase of medium osmolality due to salt or sucrose
addition resulted in changes in the protein content of periplasm (E. coli) and reduced the
rate of -galactosidase synthesis (Barron et al., 1986). 2-DE protein analysis coupled to
MALDI-TOF MS showed that the protein patterns of Enterobacter sakazakii grown on
NaCl reflect more or less a general down-regulation of central metabolic pathways
(Riedel and Lehne, 2007). Like bacteria, 2-D gel analysis of epithelial cells under
osmotic stress showed a total number of 40 proteins in which 25 proteins were
overexpressed, whereas 15 proteins showed a down-regulation (Dihazi et al., 2005).
The previous study (Chapter 3) confirmed that the L-glutamate produced in M.
glutamicus is starting to be excreted at the end of exponential or the start of stationary
phase of fermentation. In addition, the rate of glutamic acid production was observed to
decrease because of accumulating this amino acid in extracellular medium. This
phenomenon is caused by the process constraints, observed at the late stage of Lglutamate fermentation, which are directly related to the product accumulation and the
stresses due to an osmotic imbalance resulting in the decrease of sugar uptake rate, and
eventually reduce the productivity of glutamic acid (Gourdon et al., 2003). However, this
study demonstrated that the growth of M. glutamicus and the yield of L-glutamate are
affected significantly because of increasing the medium osmolality by NaCl addition. It is
expected that the outcome of this study will assist in developing a suitable production
process of L-glutamate in which the osmotic stresses could be minimized. Nevertheless,
the optimization of medium osmolality in conjunction with other physical parameters
198
associated with the fermentative cultivation of microorganism is crucial for the
enhancement of intracellular proteins or enzymes production. Amino acid production by
chemostat fermentation may be an alternative approach for avoiding the hyperosmotic
stress caused by the medium composition or product accumulation during industrial
fermentation (Kawahara et al., 1990). Furthermore, it would be beneficial to develop
mutant strains with high resistance to osmotic stresses that are capable of maintaining
high metabolic activity towards the synthesis of product while a high product titre is
required (Gourdon et al., 2003).
199
Chapter SEVEN
7. Conclusions and Future Works
The following points can be concluded from this research-
1. This study revealed that there are no remarkable dissimilarities in growth
properties, substrate consumption and L-glutamate production among the three
different strains of Corynebacteria i.e., B. lactofermentum, B. flavum and M.
glutamicus (Figure 3.3a; 3.6).
2. This research concluded that the secretion of L-glutamate into the extracellular
medium is not occurred without cultivating these bacteria in the presence of a
limited amount of biotin or by the addition of surfactant (Tween 40) or
ethambutol into the CGXII Minimal Medium.
3. This investigation confirmed that biotin concentration in fermentation medium
has also a major influence on the yield of biomass as well as on the production
of L-glutamate. The highest L-glutamate production (57mM after 52h of
cultivation) was measured in the presence of 1.0µg l-1 biotin (Figure 3.5).
4. Although surfactants have been used for the industrial production of Lglutamate, this study confirmed that Tween 20 and 80 are not suitable for the
production of this amino acid, whereas the addition of Tween 40 into the
CGXII Minimal Medium resulted in L-glutamate secretion effectively.
5. The production of glutamic acid is significantly influenced by the
concentration and the period of growth cycle at which Tween 40 was supplied.
The addition of Tween 40 (2.0g l-1, after 8h of fermentation) into a biotin rich
cultivation of M. glutamicus produced 45mM L-glutamate after 48h of
fermentation (Figure 3.10), whereas it was only 22mM (Figure 3.9) due to the
addition of similar concentration of Tween 40 at the start of cultivation.
200
6. This study confirmed that L-glutamate can also be produced by the addition of
ethambutol into a biotin rich culture of M. glutamicus, and the production of
glutamic acid increased by enhancing the concentration of EMB (up to 100mg
l-1) in the CGXII Minimal Medium. The highest amount of L-glutamate, 49mM
after 48h of fermentation, was measured at 100mg l-1 EMB (Figure 3.12).
7. This investigation clearly demonstrated that there is a significant difference
among the different treatments applied for the production of L-glutamate. The
amounts of L-glutamate measured after 48h fermentation of M. glutamicus
under the condition of biotin limitation, surfactant addition and ethambutol
addition were 56, 45 and 49mM, respectively (Figure 3.13), indicating that the
most suitable method for L-glutamate production is to cultivate these bacteria
in presence of limited amount of biotin.
8. Although electroporation has successfully been used for the transformation of
Corynebacterial cells, this study revealed that the electroinduced secretion of
L-glutamate produced in M. glutamicus and the release of intracellular
products (MDH, GDH and total protein) from both the M. glutamicus and E.
coli are limited by the conductivity of medium in which cells are suspended.
9. This examination showed that there was prominent influence of electric pulses
on the enhancement of L-glutamate production (Figure 4.4-4.7) or the release
of intracellular protein (Figure 4.8-4.10) or enzymes (GDH and MDH) until
cells suspended in H2O were electroporated. The intracellular protein release
increased approximately 14% while cell pellets (1g Dw) dissolved in a least
conductivity medium (distilled H2O) were electroporated by 4 pulses at 12.5kV
cm-1, 200 and 25µF (Figure 4.14).
10. This study demonstrated that the viability of cells decreased enormously due to
the increase of electric voltage and pulse number. Although the greater number
of pulses increases the yield of electroporation due to the formation of a large
number of pores on the cell membrane, over-stimulation may lead to apoptosis.
201
11. This investigation concluded that cell viability could be preserved by providing
resting time between pulses. The relative cell viability of M. glutamicus
increased approximately 3-10% (depending on the field strengths and number
of pulses) while giving 20min resting time between two pulses (Figure 5.3).
12. This examination also confirmed that the relative cell viability after
electropulsation is strongly dependent on the physiological state of cells since
higher cell mortality (around 5-20%) was observed in cells grown on late
exponential or stationary phase than in cells grown on early to mid-exponential
phase, regardless of the electric field strengths, pulse number and the length of
intervals allowed between two consecutive pulses (Figure 5.4).
13. Althouh it is observed that a low medium conductivity is required for the
electro-secretion or release of intracellular proteins and enzymes (chapter 4),
this study revealed that the viability of M. glutamicus affected greatly while
cells suspended in H2O or low conductivity buffers were pulsed since a lowconductivity medium allows more pulses to be applied, and results in high cell
mortality (Figure 5.5).
14. This investigation demonstrated that the cell wall of M. glutamicus is not as
flexible as that of E. coli to electric pulses although the electric field strength
and the number of pulses were kept similar in both conditions. The relative cell
viability of M. glutamicus after 5 repetitive pulses at 12.5kV cm-1, 200 and
25µF was about 13-fold higher than that of E. coli (Figure 5.6). Hence, careful
optimization of electroporation factors is required in order to achieve the
maximum efficiency of permeabilization with reduced cells mortality.
15. This examination clearly demonstrated that addition of NaCl in Seed Medium
decreased the specific growth rate of M. glutamicus and reduced the biomass
production, irrespective of the growth phases at which salt was added.
Furthermore, the degree of cell growth inhibition is dependent on the osmotic
strength of medium in which M. glutamicus was grown.
202
16. This research showed that addition of NaCl (at the start of fermentation) in
CGXII medium markedly reduced the L-glutamate production by M.
glutamicus confirming that the osmotic pressure of industrial media is one of
the limiting factors during amino acid fermentation.
17. In summary, this study confirmed that the pulse electric field treatment can
successfully be applied for the extraction of intracellular protein or enzymes
that are not usually excreted to the extracellular medium. It is expected that
high intensity electric pulses could provoke a considerable release of
intracellular L-glutamate (that is not secreted by biotin limitation) if M.
glutamicus cells suspended in a low conductivity medium are pulsed by an
appropriate electric field strength. However, the release of intracellular product
by electropermeabilization is found to be greatly dependent on the electrical
parameters, the composition of pulsing buffers and the physiological state of
cells. Although further works are needed to investigate intracellular product
secretion by electropermeabilization, it is anticipated that this preliminary data
may contribute significant indication for the enhancement of industrial
bioprocesses through transient or irreversible electroporation.
18. It is almost half a century since the discovery that M. glutamicus is capable of
excreting L-amino acids under certain growth conditions. Since then, numerous
attempts by classical strain breeding involving repeated random mutation and
selection have been conducted in order to meet the demand of these amino
acids for human and livestock consumption. Rapid progress in biochemistry
and genetics has increased our knowledge and understanding of carbon
metabolism and metabolic regulation of this bacterium. Furthermore, the
advancement in molecular biology tools in recent years enabled researchers to
improve microbial strains by rational approaches i.e., investigation of their
metabolic pathways, transport functions and regulatory mechanisms based on
the knowledge of fundamental aspects of physiology, biochemistry, molecular
biology and bioprocess engineering. In addition to genome sequencing, modern
techniques, such as transcriptomics, proteomics, metabolomics and metabolic
203
flux analysis have recently been introduced in order to identify new and
important target genes and to quantify metabolic activities. Combination of
these techniques is expected not only to increase the yield and productivity but
also to provide a mechanistic understanding of amino acids production by
Corynebacteria.
The following future works can be conducted-
1. In this study, L-glutamate was produced by shake flask cultivation of M.
glutamicus that resulted in only 57mM of L-glutamate. Since the demand of
this amino acid for human and livestock consumption is enormous and any
production process needs to be developed for industry, it will be worthwhile to
investigate the effect of those agents to glutamic acid production through the
use of batch, continuous and fed-batch fermentation.
2. Due to the discovery of the genome sequence of C. glutamicum and the recent
of improvements of metabolic engineering and functional genomics, industrial
production of amino acids have already been started. Hence, the further work
of this research will be the recognition of pathways responsible for amino acids
production, rigid branch points that result in low excretion of amino acids, the
identification of their respective genes from genome database and constructing
the recombinants enhancing the yield of amino acids production.
3. It is well established the yield of any product (for instance, amino acid) is
limited due to the effective recovery from the fermentation broth. Hence, it is
necessary to develop chromatographic based separation procedures by which a
particular amino acid will be separated from the fermentation broth.
4. In this work, the viability of pulsed cells was measured by the traditional agar
plate method. However, a long incubation time (48-72hr) was required to form
countable colonies, and it was too tough to get the reproducible results by this
approach. Moreover, colonies were often observed to clump together and
204
formed a chain like structure, and this technique showed very sensitivity to
cross contamination with airborne microorganisms. The determination of cell
viability by flow cytometry in combination with several fluorescent probes will
be an alternative approach for further research.
5. Although an attempt was taken for evaluating the cell membrane
permeabilization by means of a nonpermeant cytotoxic agent (Bleomycin), the
amount of information gather from this work was not sufficient to establish this
technique for the determination of electropermeabilized cells of both E. coli
and M. glutamicus. There was no strong reason why the bacterial cells
exhibited resistance to this particular antibiotic or why the addition of BLM
killed the non-permeabilized cells. Proteomic analysis may discover the
responsible proteins conferring resistance to this cytotoxic agent and explain
the reasons why the expression of these proteins inhibits the entry of BLM
molecules into the electropermeabilized bacterial cells.
6. In this study, electric pulses were generated through the Bio-Rad Gene Pulser
that generates an exponential decay pulse by discharging a capacitor loaded to
a specific voltage. Since the control over pulse parameters is not absolute in a
capacitor discharge instrument, it may be useful to investigate applying squarewave pulses (for example, BK Precision 3003 10MHz Sine/Square Wave
Generator) where both the voltage and pulse duration can be precisely
controlled.
7. Although the continuous production of any intracellular protein or enzymes
and
its
subsequent
secretion
(during
fermentation)
through
the
electropermeabiliztion is limited by the conductivity of fermentation medium,
the release of total protein (intracellular) increased considerably while M.
glutamicus cells suspended in a least conductivity medium (H2O) were pulsed
by electric pulses, confirming the applicability of this approach in intensified
bioprocessing. Hence, the future work might be the construction of
recombinant strain of Corynebacteria, which produce a particular heterologous
205
protein that does not secrete into the extracellualr medium during fermentation.
After completing the fermentation, cells are required to suspend in low
conductivity buffer and thereafter are pulsed with the high electric field
strengths resulting the secretion of protein of interest into the extracellualr
medium.
8. Since the heating generated during pulses may affect the extracted proteins or
product of interest, the development of a cooling system in conjunction with
the pulsing chamber will be an important breakthrough in using this approach
in bioprocessing.
206
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