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Appl Environ Microbiol, July 1998, p. 2644-2651, Vol. 64, No. 7
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Development of a Lipase Fermentation Process That
Uses a Recombinant Pseudomonas alcaligenes Strain
Gijs
Gerritse,1
Ronald W. J.
Hommes,1 and
Wim J.
Quax1,2,*
Genencor International B.V., 2600 AP
Delft,1 and
Laboratory of Pharmaceutical
Biology, University of Groningen, 9713 AV
Groningen,2 The Netherlands
Received 8 January 1998/Accepted 15 April 1998
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ABSTRACT |
Pseudomonas alcaligenes M-1 secretes an alkaline
lipase, which has excellent characteristics for the removal of fatty
stains under modern washing conditions. A fed-batch fermentation
process based on the secretion of the alkaline lipase from P. alcaligenes was developed. Due to the inability of P. alcaligenes to grow on glucose, citric acid and soybean oil were
applied as substrates in the batch phase and feed phase, respectively.
The gene encoding the high-alkaline lipase from P. alcaligenes was isolated and characterized. Amplification of
lipase gene copies in P. alcaligenes with the aid of
low- and high-copy-number plasmids resulted in an increase of lipase
expression that was apparently colinear with the gene copy number. It
was found that overexpression of the lipase helper gene,
lipB, produced a stimulating effect in strains with high
copy numbers (>20) of the lipase structural gene, lipA. In
strains with lipA on a low-copy-number vector, the
lipB gene did not show any effect, suggesting that LipB is required in a low ratio to LipA only. During scaling up of the fermentation process to 100 m3, severe losses in lipase
productivity were observed. Simulations have identified an increased
level of dissolved carbon dioxide as the most probable cause for the
scale-up losses. A large-scale fermentation protocol with a reduced
dissolved carbon dioxide concentration resulted in a substantial
elimination of the scale-up loss.
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INTRODUCTION |
One of the most persistent problems
with laundry cleaning is the removal of fatty stains. A combination of
high-temperature and high-alkalinity washing conditions can be used to
emulsify and subsequently remove the fat-containing dirt. However,
these conditions lead to damage to the fabric, and they require the input of large amounts of energy. At lower washing temperatures, e.g.,
40°C, fatty-stain removal is very poor unless lipases are added.
The lipase from Pseudomonas alcaligenes, which is
characterized by high alkalinity, has particularly advantageous
characteristics under modern washing conditions (23). The
gene for this P. alcaligenes lipase has been tested for
heterologous expression in a variety of standard industrial production
hosts, such as Bacillus licheniformis, Escherichia
coli, Streptomyces lividans, Aspergillus
niger, and Kluyveromyces lactis (unpublished
observations). However, attempts to express this enzyme to
commercially acceptable levels have been unsuccessful.
Similar observations were made for lipases originating from other
Pseudomonas species. The nature of this problem might reside
in the complex process of folding and secretion of these lipase enzymes
(LipA). These lipases require a specialized helper protein, LipB
(9, 14, 17), and the involvement of proteins of the outer
membrane secretion machinery (Xcp proteins) mediating the secretion
(35). Although the precise role of LipB (also known as LipH,
LimL, or Lif) is unclear at present, it has been suggested that it
functions as a chaperone guiding the formation of correctly folded
lipase. It has been claimed that a molar ratio between LipB and LipA of
1:1 (1) or even 4:1 (15) is needed for optimal
functioning of the helper protein.
In order to develop a fermentation process for this lipase, we decided
to optimize the expression of the gene in its original host,
P. alcaligenes. Starting with very limited knowledge of the genetics and physiology of this organism, we have now made a
large-scale production process. In this contribution the development of
the fermentation process is described. Since P. alcaligenes cannot grow on glucose, a fermentation based on
alternative substrates has been developed. Furthermore, the effects of
amplification of lipase gene (lipA) and the lipase helper
gene (lipB) on lipase production during fermentation have
been tested. Finally, the prevention of scale-up losses by adaptations
in the process conditions is described.
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MATERIALS AND METHODS |
Strains and plasmids.
For this study bacteria from the
species P. alcaligenes and E. coli were
used. The genetic modifications that have been introduced are described
in Table 1. Plasmids used in this study
are described in detail in Table 2.
Media.
For small- and large-scale DNA isolations
P. alcaligenes and E. coli strains were
propagated in 2× TY medium (16 g of Bacto tryptone
liter
1, 10 g of Bacto yeast extract
liter1, 5 g of NaCl liter1, pH 7.0)
supplemented with the appropriate antibiotic (Table 1) (tetracycline,
10 mg liter1; neomycin, 20 mg liter1).
Colonies were plated on 2× TY agar plates or on tributyrin agar
plates. Tributyrin agar plates were composed of buffered (0.2 M HEPES
[Sigma]) minimal medium containing, per liter, 0.6 g of citrate,
0.6 g of K2HPO4, 1.0 g of
(NH4)2SO4, 0.3 g of
MgSO4 · 7H2O, 0.08 g of
CaCl2 · 2H2O, and 10 ml of
tributyrin oil (Merck), supplemented with the appropriate
antibiotics for selection (Table 1) and solidified with 1.5% agar
(Difco).
Recombinant DNA techniques.
Chromosomal DNA from
P. alcaligenes was isolated according to the method of
Andreoli (2). Plasmid DNAs of E. coli and
P. alcaligenes were obtained by the method of Birnboim
and Doly (3) unless otherwise stated.
Isolation of mRNA from P. alcaligenes was done as
follows. An aliquot of 1 ml of fermentation broth was diluted in 9 ml
of milliQ water. The cells were spun down in a centrifuge (4,500 × g). The pellet was homogenized in 10 ml of a solution
containing 3 M LiCl and 6 M urea with an Ultra-turrax blender (20,000 rpm) for 1 min on ice. The suspension was centrifuged (4,500 × g), and the homogenization procedure was repeated for
45 s on ice. Subsequently, the suspension was centrifuged in an
ultracentrifuge at 140,000 × g for 30 min at 4°C.
The pellet was dissolved in 3 ml of 1× FEB, containing 0.1 M NaCl, 10 mM Tris-HCl (pH 9.0), 0.5% sodium dodecyl sulfate, and 5 mM EDTA. To
the suspension was added 3 ml of hot phenol solution
(phenol-chloroform-isoamyl alcohol-3× FEB; 50:48:1:2.5, vol/vol).
The solution was mixed thoroughly and centrifuged (4,500 × g) at room temperature. The upper layer was collected, and 1 volume of chloroform solution (chloroform-isoamyl alcohol; 24:1,
vol/vol) was added, mixed thoroughly, and centrifuged (4,500 × g) at room temperature. The last two steps were repeated
twice. Furthermore, an ethanol precipitation was performed on the water
layer, and an appropriate volume of RNase-free water was added to the
sample.
Restriction endonucleases and other enzymes were purchased from
GIBCO-BRL and used according to the manufacturer's instructions.
For Southern hybridizations, transfers of chromosomal DNA
fragments
and colonies to nitrocellulose filters were done by the
method
of Southern (
31). For Northern hybridizations,
agarose gel electrophoresis
of RNA was performed with addition of
formaldehyde to the gel
matrix, and transfer to nitrocellulose filters
was done by the
method described by Maniatis et al. (
20).
Detection of both
DNA- and RNA-containing filters was executed with
DNA probes labelled
with [
-
32P]dATP (Amersham), using
the random-primer-labelling kit from
Pharmacia according to the
instructions of the supplier.
Sequencing was carried out on an Applied Biosystems 373A DNA sequencer
with the ABI PRISM Dye Termination Cycle Sequencing
Ready Reaction Kit
and AmpliTaq DNA polymerase.
Electroporation and isolation of restriction-negative,
modification-positive strain Ps93.
Electroporation of
P. alcaligenes M-1 with DNA isolated from E. coli gave only 1 to 10 transformants per µg, indicating a very
low transformation frequency, whereas electroporation with plasmid DNA
isolated from P. alcaligenes resulted in a higher transformation frequency. Because of this observation, we reasoned that
there could be a selection for strains that are restriction negative
after electroporation. Therefore, 24 independent transformants obtained
after electroporation of P. alcaligenes M-1 with
plasmids isolated from E. coli were cured from their
plasmid. In these strains an equal amount of plasmid pJRD215
(5) isolated from E. coli was introduced by
electroporation. Two strains showed a transformation frequency of
105 colonies per µg of DNA, and one of the strains was
named strain Ps93. DNA isolated from strain Ps93 and reintroduced in
P. alcaligenes M-1 demonstrated the same high
transformation frequency, indicating that strain Ps93 is a
restriction-negative, modification-positive mutant of P. alcaligenes M-1.
Electroporation of plasmid DNA to
E. coli was carried
out as described by Calvin and Hanawalt (
4). For
electroporation
of
P. alcaligenes strains, the method
of Wirth et al. (
37) was
used, with the modification that
all treatments were performed
at room temperature. Transformants of
E. coli or
P. alcaligenes with plasmid
pLAFR3 were selected on 2× TY agar plates with 10
mg of tetracycline
liter
1. Cells of
E. coli or
P. alcaligenes transformed with pJRD215
were selected on 2× TY agar
plates with 20 mg of neomycin liter
1.
Cloning of the lipase gene.
Purification of the lipase of
P. alcaligenes M-1 from the growth medium was performed
as described by Stuer et al. (33). After gel electrophoresis
and blotting on Immobilon transfer membrane, the N-terminal amino acid
sequence was determined (21). The N-terminal sequence was
found to be GLFGSTGYTKTKYPIVLTHGMLGF. ...
The following oligonucleotide probe, matching residues 6 to 16 (according to the preferential codon usage in
Pseudomonas
aeruginosa [
36]) was prepared: 5'-ACC GGC TAC ACC
AAG ACC AAG TAC CCG/C
ATC GT-3'. This oligonucleotide was used to
hybridize chromosomal
DNA of
P. alcaligenes M-1
digested with several restriction endonucleases.
Chromosomal
PvuII and
BclI fragments (between 1 and 4 kb)
were
cloned in the multiple cloning site of vector pTZ18R. Positive
clones were detected from the library with
PvuII fragments
by
colony hybridization with the oligonucleotide matching the
N-terminal
sequence of M-1 lipase.
Lipase assay.
Lipase activity in growth media was assayed
titrimetrically as described by Gilbert et al. (12) with
some modifications. A pH-Stat unit (Radiometer type ETS 822) was used
at pH 9.0 and at 30°C with a standard olive oil emulsion (Sigma)
diluted to 10% (vol/vol) and supplemented with 20 mM NaCl and 10 mM
CaCl2. One lipase unit (LU) was defined as the amount of
lipase capable of releasing 1 µmol of titratable fatty acid per min
under the assay conditions used.
Lipase detection on plates was done with the use of minimal medium (see
"Media" above) agar with 1% tributyrin oil (Merck).
Plates were
incubated at 37°C during 48 to 72 h, and clearing
zones
indicated lipolytic activity.
Controlled fed-batch fermentation process.
P.
alcaligenes was cultured either in laboratory fermentors with a 10 liter capacity or in 100-liter, 4-m3, or 100-m3
production fermentation vessels. The fermentors were inoculated with
full-grown seed cultures that were grown at 35°C for 16 h on a
yeast extract medium (10 g of yeast extract liter1, pH
7.0). The inoculation percentage used on all scales was 5%. For the
main fermentation a defined minimal salt medium was used. The medium
contained (per liter) K2HPO4, 6.4 g;
(NH4)2SO4, 3.33 g;
MgSO4 · 7H2O, 3.2 g;
CaCl2 · 2H2O, 0.8 g;
CoCl2 · 6H2O, 40 mg;
MnSO4 · 1H2O, 32 mg;
FeSO4 · 7H2O, 20 mg;
ZnSO4 · 7H2O, 12 mg;
CuSO4 · 5H2O, 3 mg;
H3BO3, 3 mg;
Na2MoO4 · 2H2O, 3 mg; and KI, 1 mg. The fermentation was run at 35°C. The pH of the culture was
maintained automatically at 7.0 ± 0.1 by using sulfuric acid and
ammonia as titrants. Foam was controlled automatically by use of a
polyalkylene-silicone mixture (SAG 5693 silicone antifoam; Union
Carbide). As a carbon source, citric acid (6 g liter1)
was added. When the citric acid was fully consumed, 10 to 16 h
after inoculation, a feed of soybean oil (Cargill) was applied at the
rate of 1 g of soybean oil per liter of broth per h. This feed
rate was kept constant for the rest of the fermentation. The
fermentation was run for 48 to 100 h. Fully aerobic conditions (dissolved oxygen tension, >20% air saturation) were maintained throughout by injecting air, at a rate of 1 standard liter of air per
liter of broth per min (= 1 vvm), into the region of the impeller that
was rotating at about 400 rpm.
For scaling up the small-scale process to 100-m
3 production
fermentors, 100-liter and 4-m
3 fermenters were used as
intermediates. The medium, the temperature,
the pH, and the soybean oil
feed rate did not vary with scale.
Only the aeration changed with
varying scale, being 1 vvm at the
10-liter scale, 2 vvm at 100 liters,
0.5 vvm at 4 m
3, and 0.4 vvm at 100 m
3 to
prevent excessive foaming and a high holdup. Except for the
10-liter
scale, a back pressure of approximately 50 kPa was applied.
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RESULTS |
Lipase fermentation process.
In order to develop an
economical production process for the microbial lipase produced by
P. alcaligenes, a fermentation process under fed-batch
control was developed. The inability of P. alcaligenes to metabolize sugars like glucose restricted the choice of suitable (industrial) carbon sources. Citrate was selected as the preferred initial carbon source during the first phase of the fermentation when
the cells were growing batchwise. This was based partly on the fact that citrate enabled the cells to grow at high specific growth rates (µ 
0.9 h1), which resulted in a
relatively short batch phase (10 h). Furthermore, citrate kept the
metal ions in solution during sterilization and the initial stages of
the fermentation. When the citrate was fully consumed, the oxygen
uptake rate sharply decreased (Fig. 1).
At that moment a carbon-limited feed of soybean oil was applied to the
culture to construct a fed-batch process, where the temperature, pH,
and dissolved oxygen concentration were controlled.
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FIG. 1.
Production of lipase by P. alcaligenes
M-1 in a fed-batch fermentation process. After an initial exponential
growth period, the process was controlled by a constant feed rate of
soybean oil, which resulted in the oxygen uptake rate (OUR) (no
symbol), biomass profile ( ), lipase activity ( ), and yield
profile of lipase on soybean oil (YP/S) ( )
shown. The feed was started at 10 h after the initial carbon
source (citrate) was fully consumed and the OUR was down to almost
zero.
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The characteristic parameters of this standard fermentation of the
wild-type
P. alcaligenes M-1 are exemplified in Fig.
1.
Hardly any synthesis of lipase took place during batch phase growth
on citrate. Only after the start of the soybean oil feed was lipase
expression induced. It should also be stressed that a significant
development of biomass occurred during the soybean oil feed phase.
The
highest yield of lipase on soybean oil (
YP/S)
was detected
halfway through the fermentation. Until that time, the
soybean
oil was primarily converted into biomass, lipase, and carbon
dioxide.
Thereafter, the lipase synthesis leveled off rapidly and
the major
part of the soybean oil was used for maintenance purposes
besides
some growth. As a result, the
YP/S
decreased dramatically.
Characterization of the lipase gene.
As a result of the
screening with the oligonucleotide corresponding to the
N-terminal sequence of M-1 lipase, a positively reacting clone
containing a 2.0-kb PvuII fragment (Fig.
2A) was detected. The 2.0-kb
PvuII fragment from the P. alcaligenes M-1 chromosome was cloned in SmaI-opened E. coli
vector pTZ18R (pTZlipA) (Fig. 2A). This fragment was found to contain
an open reading frame that encodes the same NH2-terminal
peptide as determined for the lipase from P. alcaligenes M-1. Therefore, we have named the gene
lipA. The encoded protein shows homology to other
Pseudomonas lipase gene products (Fig.
3), with the highest homology (77% amino
acid identity) to the lipase from P. aeruginosa
(14, 38).
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FIG. 2.
Genetic organization of the lipase operon and plasmid
construction. (A) Construction of plasmid pTZlipAB and map of the
chromosomal fragments of P. alcaligenes that are
present on plasmids pTZlipA, pTZlipB, and pTZlipAB.
Restriction endonuclease sites used for cloning: B, BclI; E,
EcoRI; H, HindIII; P, PvuII;
S, SstI; V, EcoRV. lipA and
'lipA, lipase gene and part of the lipase gene;
lipB and lipB', lipase helper gene and part of
the lipase helper gene; bla,  -lactamase gene. (B) Genetic
organization of the lipase operon. lipA, lipase gene;
lipB, lipase helper gene; PR, lipase promoter sequence; ATT,
attenuator sequence; TER, terminator sequence. The arrows
represent the two lipase mRNAs, one starting from the
transcription start and ending at the attenuator signal (1.2 kb in
length) and the other starting from the transcription start and ending
at the terminator (2.1 kb in length).
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FIG. 3.
Sequence alignment of lipases from
Pseudomonas rRNA group I and group II. All residues
identical to those in the P. alcaligenes protein
sequence are shown as white letters. The alignment was carried out by
using the Clustal method of the WINSTAR software package. LIP-PALC.PRO,
P. alcaligenes (this study); LIP-PAER.PRO,
P. aeruginosa (38); LIP-BGLU.PRO,
B. glumae (8); LIP-BCEP.PRO,
B. cepacia (17).
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The second incomplete open reading frame shows homology to a gene,
lipB, found to be involved in the expression of lipase
in
other pseudomonads. In order to determine the complete
sequence
for
lipB, the 2.0-kb
PvuII fragment
was used as a probe to hybridize
with a library of
BclI
fragments. A colony containing a 1.7-kb
BclI fragment could
be identified as positive in this experiment.
This 1.7-kb
BclI fragment (pTZlipB) (Fig.
2A) harboring the
complete
second open reading frame was isolated and fused to the
overlapping
2.0-kb
PvuII fragment by using a common
EcoRV site (Fig.
2A).
Subsequently, a 2.4-kb fragment with
both
lipA and
lipB could
be subcloned in pTZ18R
(pTZlipAB) (Fig.
2A), and its complete
sequence was determined. The
amino acid sequence of the putative
LipB homolog was derived and was
compared with those of other
lipase helper gene products of
Pseudomonas origin (Fig.
4).
The
N-terminal 25 residues show high hydrophobicity, suggesting
that
LipB is anchored to the membrane.
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FIG. 4.
Sequence alignment of lipase helper proteins from
Pseudomonas rRNA group I and group II. All residues
identical to those in the P. alcaligenes protein
sequence are shown as white letters. The alignment was carried out by
using the Clustal method of the WINSTAR software package. LIPBPALC.PRO,
P. alcaligenes (this study); LIPBPAER.PRO,
P. aeruginosa (16); LIPBBGLU.PRO,
B. glumae (9); LIPBBCEP.PRO,
B. cepacia (17).
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Effect of gene copy number on lipase production.
The 2.0-kb
lipA fragment from pTZlipA was used to construct the
high-copy-number plasmid pJRDlipA (Table 2) and the low-copy-number plasmid pLAFlipA (Table 2) in order to study the effect of gene copy
number on lipase expression.
The transformation of the wild-type
P. alcaligenes
strain by the electroporation method with DNA isolated from the
restriction-negative
P. alcaligenes strain Ps93 (see
Materials and Methods) gave a
transformation frequency of
10
5 colonies per µg of DNA for both plasmids pLAFlipA and
pJRDlipA.
In this way,
P. alcaligenes Ps224 and Ps496,
containing pLAFlipA
and pJRDlipA, respectively, were made. On
tributyrin plates these
strains exhibited significantly larger halos
than the wild-type
strain. The
P. alcaligenes strains
were grown in a fermentor at
a 10-liter scale by using the standard
protocol. The lipase expression
levels are shown in Fig.
5. It can be seen that there is
a clear
gene dose effect, with the high-copy-number plasmid
(pJRDlipA)-containing
strain exhibiting the highest lipase expression.
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FIG. 5.
Lipase production by various plasmid-containing strains
at the 10-liter fermentation scale, showing the effects of low- and
high-copy-number plasmids on lipase production levels. The lipase
production of the wild-type P. alcaligenes strain M-1
(1.5 MLU/liter) is used as the standard and fixed at 100%. Bars: 1, no
plasmid (strain M-1); 2, pLAFlipA (strain Ps224); 3, pLAFlipAB (strain
Ps225); 4, pJRDlipA (strain Ps496); 5, pJRDlipAB (strain Ps495).
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Effect of the helper gene.
In order to investigate the effect
of the helper gene (lipB) on the lipase production, two new
constructs were made. As the 2.0-kb PvuII fragment does
not harbor the complete open reading frame for the putative helper
protein, the 2.4-kb fragment harboring both open reading frames was
made and used to construct pLAFlipAB (Table 2) and pJRDlipAB (Table 2).
This results in P. alcaligenes Ps225 and Ps495,
respectively. Fermentation results for these strains at the 10-liter
scale are shown in Fig. 5.
In order to verify the expression of the second open reading frame, we
have analyzed mRNAs on Northern blots with the 2.4-kb
fragment as a
probe. In Fig.
6 strains Ps496 and Ps495
are compared.
It can be seen that the strain without additional
lipB gene copies
(Ps496) shows only a band of 1.2 kb,
whereas strain Ps495 shows,
besides the strong 1.2-kb band, an
additional longer mRNA of 2.1
kb, which corresponds to the full length
of the
lipAB operon.
The 1.2-kb band corresponds in size
exactly to a transcript running
from the promoter to the hairpin
structure present between the
lipA and
lipB open
reading frames (Fig.
2B). The 2.1-kb fragment
would correspond to a
transcript from the promoter to the terminator
structure downstream of
the
lipB gene (Fig.
2B).
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FIG. 6.
mRNA isolations (see Materials and Methods) from
fermentation broth analyzed by using the Northern blot procedure with
the 2.4-kb insert from plasmid pTZlipAB as a probe. The autoradiograph
was overexposed in order to visualize the 2.1-kb transcript. Lane A,
mRNA of P. alcaligenes Ps496 isolated from a sample
taken at 48 h of fermentation; lane B, mRNA of P. alcaligenes Ps495 isolated from a sample taken at 48 h of
fermentation; lane C, mRNA of P. alcaligenes Ps495
isolated from a sample taken at 72 h of fermentation.
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Scale-up losses.
The small-scale reproducible fed-batch
process, which resulted in a lipase activity of 55.3 ± 4.1 MLU/liter (n = 5), was scaled up from the 10-liter
scale to a 100-m3 production fermentor. During the
scaling up, important parameters did not vary with scale. However, a
significant loss of lipase production was observed, without any obvious
reason. In Fig. 7A it can be seen that
the lipase concentration and thus the production decreased with
increasing scale, accumulating to a loss of production of 65% at the
100-m3 scale. Besides the lipase production, no major
differences between the fermentations at the different scales were
detected.
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FIG. 7.
(A) Lipase production loss during scaling up, showing
fitted lipase production profiles based on five fermentations at a
10-liter scale ( ), four fermentations at a 100-liter scale
(·····), and two
fermentations at a 4-m3 scale (---) and a
lipase production profile at a 100-m3 scale ( ). (B)
Effect of CO2 addition to the inlet air at a 10-liter
scale, showing lipase production without addition of CO2
() (control), with the addition of 5% CO2 ( ), and
with the addition of 10% CO2 (). (C) Lipase production
at 10-liter () and 100-m3 () scales after successful
scaling up with the following modifications: increased ventilation rate
(from 0.4 to 0.8 vvm), decreased back pressure (from 80 to 20 kPa), and
lowered pH (from 7.0 to 6.7).
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A systematic search at the 10-liter scale for possible causes of the
negative scale-up effect revealed the dissolved carbon
dioxide as a
likely candidate. During fermentation, the pH was
regulated at 7.0. At
this pH, CO
2 dissolved in the broth at fairly
high levels,
predominantly as HCO
3. Due to the relatively
high pressure (both hydrostatic and back
pressure) and the low aeration
rate of 0.4 to 0.5 vvm, the dissolved
carbon dioxide concentration
increased with increasing scale.
The total dissolved CO
2
concentration could be estimated from
the fermentation parameters and
the CO
2 equilibrium constants
and was substantially higher
(three- to fivefold) at the larger
scales of 4 and 100 m
3.
In order to obtain dissolved CO
2 concentrations at the
10-liter
scale representative of those for large scales,
CO
2 gas was supplied
to the inlet air to a final
concentration of 5 to 10% (vol/vol).
By doing these experiments, it
could be demonstrated that increasing
dissolved CO
2
concentrations had a negative effect on the lipase
production (Fig.
7B).
The improved large-scale process.
Additional proof that the
dissolved CO2 was responsible for a significant part of
the decrease in lipase production at the large scales was obtained
by using the 100-m3 production fermentor. Decreasing the
broth volume from 100 to 50 m3, increasing the aeration
rate to 1.2 vvm, and decreasing the back pressure to 20 kPa resulted in
a reduction of the CO2 concentration, which had a positive
effect on the production of lipase. The maximal lipase activity
detected was increased by 70%. However, the total output
(lipase concentration × total amount of broth) per fermentor was lowered as a result of a lower occupation of the volume.
These experiments indicated that the prevention of scale-up losses
ought to be done by keeping the dissolved carbon dioxide
concentration as low as possible. At a large scale, the ventilation
rate is usually limited. With a total volume of 100 m
3, the
maximal aeration that could be achieved was 0.8 vvm. Together
with the
reduced back pressure of 20 kPa, a reduction in the CO
2 concentration by almost a factor of 3 could be obtained. This
indeed
gave a beneficial effect on production. Still, it was not
sufficient to
eliminate all of the scale-up losses.
At the lab scale, additional methods of lowering the dissolved
CO
2 concentration during the fermentation were
investigated.
By decreasing the soybean oil feed rate from 1 to
0.5 g of soybean
oil per liter of broth per h, a reduction of the
carbon dioxide
production rate was obtained. However, this had a
negative effect
on lipase production (38.9 compared to 55.3 MLU/liter),
suggesting
that the specific lipase production rate was linked to the
specific
growth rate, which was also affected by changing feed rate.
Further
reduction of the dissolved CO
2 concentration was
achieved by decreasing
the pH. From studies at the lab scale, however,
it was known that
decreasing the pH to 6.5 caused a 10 to 15%
reduction in lipase
production. An optimal balance for production at
large scales
between the decrease of production caused by lowering the
pH and
the increase of production caused by lowering the dissolved
CO
2 concentration was found at pH 6.7. Subsequently, a
combination
of increased ventilation rate (from 0.4 to 0.8 vvm) and
decreased
back pressure (from 80 to 20 kPa) together with the lowered
pH
(from 7.0 to 6.7) was used to minimize the scale-up losses at
100 m
3 considerably (Fig.
7C).
| |
DISCUSSION |
The finding that the P. alcaligenes lipase shows
the highest homology with the P. aeruginosa lipase is
in line with the taxonomic data that both species are classified
as being in rRNA group I of the pseudomonads (29). The
homology with the lipases from rRNA group II
Pseudomonas species (now called Burkholderia
species) is lower but still very significant. The residues in the
vicinity of the catalytic-triad residues Ser111, Asp257, and His279
(Fig. 3) are especially strongly conserved. Interestingly, the two
cysteine residues forming a disulfide bond in Pseudomonas
lipases are also fully conserved (positions 213 and 263 in Fig. 3),
stressing the importance of this single disulfide bond for lipase
folding and stability. Both the P. alcaligenes and
P. aeruginosa lipases lack a stretch of 20 amino acids
(around position 232) as compared to lipases from representatives of
rRNA group II, Burkholderia cepacia and Burkholderia
glumae. From a comparison with the experimentally derived
three-dimensional structure of B. glumae, it can be
seen that this deletion overlaps precisely with two antiparallel
-strands and a -turn in the protein, which seem to be replaceable
in the lipase molecule (25). Another major difference is in
the signal sequence, which is unusually long in the rRNA group II
lipases but not in the rRNA group I lipases.
By using Southern blotting, copy numbers have been determined for
plasmids in P. alcaligenes, resulting in values of 2 to 4 and 20 to 40 copies for pLAFR3 and pJRD215, respectively (data not
shown). These experimental data show that the gene copy number for
pLAFR3 is around 3 and that for pJRD215 is around 30. For pLAFR3 this
is in line with literature data (11). For plasmid RSF1010, which has the same replicon as pJRD215, a copy number of
12 in E. coli has been reported (13).
However, it is known that alterations in the expression of
rep functions can lead to higher copy numbers in plasmid
variants or in other host cells (10). Overexpression of the
lipase from plasmids pLAFlipA and pJRDlipA in P. alcaligenes leads to an increase in the lipase production at
a ratio that seems to correlate with the copy numbers of these
plasmids. Conceivably, LipB is not limiting. However, when
lipA and lipB of P. alcaligenes
were combined on both expression vectors (pLAFR3 and pJRD215), an
additional increase in lipase levels was observed only with the
high-copy-number plasmid, pJRDlipAB. Evidently a single
lipB gene copy, which is still present in the host
chromosome, is sufficient to fully support the efficient expression of
lipA from a low-copy-number plasmid but is insufficient to
support the full expression of lipase from a high-copy-number plasmid,
as judged from the fermentation results for strains Ps496 and Ps495.
The accuracy of plasmid copy number determinations is insufficient to
derive quantitative conclusions, but from the comparison of strains
Ps496 and Ps495 it seems that the helper gene becomes limiting at
lipA gene copy numbers of above 10.
Further support for a model in which the LipB gene product is required
only in catalytic amounts comes from the fact that the amount of
lipB transcript is very low. A hairpin structure situated
between the lipA and lipB genes has all the
characteristics to act as a transcriptional attenuator (for a
review, see reference 18). Alternatively, it could
represent a pausing position for mRNA degradation. The observed
low lipAB/lipA transcript ratio seems to be in contrast to
the report that for B. cepacia (1) the
optimal ratio of LipB to LipA has been measured at 1:1. Also, for
P. aeruginosa a maximal activation in
denaturation-renaturation experiments was found at a LipB/LipA ratio of
1:1 (28). It should be kept in mind, however, that those
measurements were done in a refolding experiment under in vitro
conditions quite distinct from physiological concentrations.
Interestingly, both P. aeruginosa and the two
representatives of rRNA group II, B. cepacia and
B. glumae, have similar hairpin structures, in front of
and within the lipB gene, respectively. Indeed, Frenken et
al. (9) showed that in B. glumae the
dominant transcript is a 1,400-nucleotide-long mRNA that encodes only
lipase. Although the precise mode of action of LipB is yet unknown, the
observations for P. alcaligenes and P. aeruginosa (16) best fit with a chaperoning type of
action, which requires only catalytic amounts of LipB.
Inactivation of the helper gene in the chromosome of P. alcaligenes (unpublished results) leads to a lipase-negative
phenotype, which is in line with the proposed function as a
lipase-specific chaperone.
Although for economic reasons complex substances often are used for
industrial fermentations, for reasons of consistency and quality
control there is a strong tendency nowadays to develop a process with a
medium as defined as possible. This is due mainly to the disadvantages
of the complex raw materials, such as the unknown composition,
the batch-to-batch variability, and the forced combination in a
fixed ratio. Since P. alcaligenes does not
require any special nutrients for the production of lipase, it was
possible to develop a balanced mineral medium for the
fermentation process. Subsequently, the use of this defined medium
significantly facilitated the identification of CO2 as the
major cause for the scale-up problem. It is generally known that
CO2 can be responsible for problems during scaling up.
However, despite numerous studies on the effects of carbon dioxide on
microbial growth and metabolism of microorganisms, knowledge of the
mechanism of CO2 inhibition still remains inconclusive (for
reviews, see references 6 and 26). In this case also, no explanation for the
molecular mechanism by which CO2 affects the lipase
production by P. alcaligenes can be offered. Other than
the lipase production, no major differences in metabolism that
were caused by the increased partial CO2 pressure could be
detected. There were no indications that the increased CO2
concentrations had any effect on the energetics of cell synthesis (34) or on the lipase enzyme itself by affecting its
physicochemical properties (24). The results here could
indicate that there is a CO2-controlled repression of
lipase synthesis. Induction and repression of enzyme synthesis by
CO2 is well known in the process of autotrophic
CO2 fixation, e.g., with photolithotrophically grown
cells of Rhodospirillum rubrum (30).
By a combination of adaptations to the process, the scale-up losses at
100 m3 could be minimized considerably. There is still a
remaining loss of 10 to 15%. Some of this can be attributed to
gradients of soybean oil, ammonia, oxygen, and pH that are generally
known to exist in large-scale fermentors. These gradients are caused by
the lack of ideal mixing that is a characteristic of these large-scale fermentors (7, 27). Sodium dodecyl sulfate-polyacrylamide gel electrophoresis analysis and Western blotting (data not shown) confirmed that the remaining loss in lipase production was not caused
by inactivation of the lipase by shear, as has been reported for a
lipase produced by Candida cylindracea (19).
In a separate paper the purification process leading to the complete
removal of the endotoxins of P. alcaligenes will be
reported.
| |
ACKNOWLEDGMENTS |
We thank Marion Kooman, Lydia Dankmeyer, and Mar van Dam
for technical assistance and Bert Geraats and Manon Cox for stimulating discussions.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Pharmaceutical
Biology, University of Groningen, Antonius Deusinglaan 1, 9713 AV
Groningen, The Netherlands. Phone: 31503632558. Fax:
31503636908. E-mail: w.j.quax{at}farm.rug.nl.
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Abstract
Introduction
