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Applied and Environmental Microbiology, September 1999, p. 3888-3895, Vol. 65, No. 9
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Analysis of the Gene Family Encoding Lipases in Candida
rugosa by Competitive Reverse Transcription-PCR
Guan-Chiun
Lee,1
Shye-Jye
Tang,2,*
Kuang-Hui
Sun,3 and
Jei-Fu
Shaw1,4,*
Institute of Marine Biotechnology, National Taiwan Ocean
University, Keelung, Taiwan 20224,2
Institute of Biochemistry,1 and
Department of Medical Technology3, National Yang-Ming University,
Taipei, Taiwan 11211, and Institute of Botany,
Academia Sinica, Taipei, Taiwan 115294
Received 8 March 1999/Accepted 6 June 1999
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ABSTRACT |
Synthesis of multiple extracellular lipases in Candida
rugosa has been demonstrated. However, it is difficult to
characterize the expression spectrum of lip genes, since
the sequences of the lip multigene family are very closely
related. A competitive reverse transcription-PCR assay was developed to
quantify the expression of lip genes. In agreement with the
protein profile, the abundance of lip mRNAs was found to be
(in decreasing order) lip1, lip3, lip2, lip5, and lip4. To analyze
the effects of different culture conditions, the transcript
concentrations for these mRNA species were normalized relative to the
values for gpd, encoding glyceraldehyde-3-phosphate dehydrogenase. In relative terms, lip1 and lip3
were highly and constitutively expressed (about 105
molecules per µg of total RNA) whereas the other inducible
lip genes, especially lip4, showed significant
changes in mRNA expression under different culture conditions. These
results indicate that differential transcriptional control of
lip genes results in multiple forms of lipase proteins.
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INTRODUCTION |
Lipases (EC 3.1.1.3) from the
nonsporogenic yeast Candida rugosa (formerly C. cylindracea) are very important industrial enzymes which have been
widely used in biotechnological applications such as the production of
fatty acid, synthesis of various esters, and kinetic resolution of
racemic mixtures (10, 14, 22, 29, 40, 43, 44). Crude enzyme
preparations are used in most applications, and enzymes from various
suppliers have been reported to show variations in their catalytic
efficiency and stereospecificity in various applications such as
resolution of racemic 2-(4-hydroxyphenoxy) propionic acid
(2). After our discovery of multiple enzyme forms with
different substrate specificities and thermostabilities in a commercial
C. rugosa lipase preparation (39), two to six
enzyme forms were detected in subsequent studies (4, 9, 36,
37). More recently, we discovered that three commercial C. rugosa lipase preparations differed in protein composition, which
accounted for the difference in their catalytic efficiency and
specificity (5). This was related to the different culture conditions used. It was supported by the observation that different inducers in the culture media of C. rugosa changed the
multiple form patterns and therefore the specificity and
thermostability of crude lipase preparations (5).
The multiplicity of extracellular lipases in fungi (3a, 17-19,
32) has been attributed to a change in gene expression, variable
glycosylation, partial proteolysis, or other posttranslational modification. After the cloning of five lipase genes (lip1
to lip5) from the C. rugosa genome (24,
26), a change in gene expression has been suggested to be the
most probable mechanism for the enzyme multiplicity. However, the high
similarity of same-sized deduced amino acid sequences in these mature
proteins (66% identity and 84% similarity) makes it difficult to
purify and identify the lipase gene products (27).
For highly related genes, the conventional methods in mRNA analysis are
not specific and sensitive enough to distinguish and quantitate
individual mRNAs. It is difficult to distinguish the transcription
pattern of genes with a high degree of identity by Northern blot
analysis. Although the nuclease protection assay has the ability to
discriminate among closely related genes, this method, like Northern
blotting, is not sensitive enough to detect small amounts of mRNA and
permits only crude quantitation. The competitive reverse
transcription-PCR (RT-PCR) technique (3, 12) may be a
feasible alternative to obtain quantitative information on the highly
related lip genes at the transcriptional level owing to its
high sensitivity and specificity (35, 41).
Although the lip1 cDNA has been isolated from C. rugosa (21), there is no evidence that the other four
genomic lipase-encoding sequences isolated are functional. Moreover,
the molecular mechanisms of the individual lip gene
regulation remain unclear. In this report, we describe a modification
of the competitive RT-PCR method to detect and quantitate the five
lip mRNA transcripts and thus confirm the functional
expression of the five lip genes. By using this technique,
it is possible to demonstrate the differential expression of the five
lip genes in the presence of different inducers that are
known to be able to increase C. rugosa lipase production
(5, 13, 42).
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MATERIALS AND METHODS |
Microorganism and medium.
C. rugosa ATCC 14830 was
cultured in YM growth medium (0.3% yeast extract, 0.3% malt extract,
0.5% peptone, 1% dextrose) at 30°C for 24 h. The
concentrations of additives are specified for the different experiments.
Bacterial transformation.
Plasmid DNA was transformed into
Escherichia coli TOP10 (Invitrogen) by the CaCl2
method and extracted from ampicillin-resistant colonies by the alkali
lysis method (38).
RNA preparation.
Total RNA from C. rugosa was
isolated by a modification of the method of Köhrer and Domdey
(23). Cultured cells (5 ml) were collected by centrifugation
(3,000 × g at 4°C for 5 min), resuspended in 300 µl of sodium acetate buffer (50 mM sodium acetate, 10 mM EDTA [pH
5.0]), and then transferred to a 1.5-ml microtube containing 0.3 g of glass beads (pretreated with 1 M HCl and autoclaved). After two
cycles of hot-phenol extraction, total RNA was collected by ethanol precipitation.
To eliminate contaminating genomic DNAs, two purification methods were
used. (i) RNase-free DNase I (20 U; Promega) was mixed with 50 µg of
the total RNA in RT buffer (50 mM Tris-HCl, 75 mM KCl, 3 mM
MgCl2, 10 mM dithiothreitol [pH 8.3]) and incubated at
37°C for 30 min in a total volume of 50 µl. The RNA was purified by
phenol-chloroform extraction followed by ethanol precipitation. (ii)
The RNA pellets obtained from the hot-phenol extraction were dissolved
in 500 µl of prechilled denaturing solution (4 M guanidine thiocyanate, 42 mM sodium citrate, 0.83%
N-lauroylsarcosine, 0.2 mM 2-mercaptoethanol), and 50 µl
of 2 M sodium acetate (pH 4.0) was then added to provide acidity. After
500 µl of a mixed organic solvent (phenol-chloroform-isoamyl alcohol,
25:24:1) was added, the mixture was vigorously vortexed for 1 min and
chilled on ice for 3 min. After centrifugation at 10,000 × g and 4°C for 15 min, the aqueous phase was transferred to a new
microtube and RNA was precipitated twice with an equal volume of
isopropanol at 
20°C for 30 min.
RT-PCR.
Total RNA (5 µg) was reverse transcribed into
first-strand cDNA in a 20-µl reaction mixture by using oligo(dT)
primers, deoxyribonucleoside triphosphates, and SuperScript II enzyme
as specified by the manufacturer (Life Technologies, Gaithersburg,
Md.). To increase the efficiency of PCR initiation, RNase H (2 U; Life
Technologies) was added and the mixture was incubated at 37°C for 20 min. PCR amplification was performed in a 25-µl reaction volume
containing 0.5 µl of RT reaction solution, each deoxyribonucleoside
triphosphate at 100 µM, 5 pmol each of 5' and 3' primer, 10 mM
Tris-HCl (pH 8.8), 1.5 mM MgCl2, 50 mM KCl, 0.01% (wt/vol)
gelatin, and 0.25 U of DynaZyme (Finnzymes Oy, Espoo, Finland). The
reagents for RT and PCR were always prepared as a single reaction
mixture and then divided among different tubes. PCR was carried out in
an Omnigene thermal cycler (Hybaid, Teddington, United Kingdom) on the
following cycle program: one cycle of 94°C for 3 min and 72°C for 1 min; 40 cycles of 95°C for 30 s, 57°C for 30 s, and
72°C for 90 s; and a final 10-min extension step at 72°C.
Cloning of template DNAs.
The full-length lipase-encoding
cDNA fragments were obtained by RT-PCR with specific primers based on
the published sequences (24, 26). The PCR products with
created flanking restriction sites were digested with restriction
enzymes and ligated into pET-23a (Novagen) or pGEMT vector (Promega). A
schematic presentation of the plasmids is given in Fig.
1.
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FIG. 1.
Plasmids for deletion-containing competitive templates
and relative positions of PCR primers. Open arrows indicate primers for
the generation of competitor DNAs. Solid arrows indicate those for
competitive PCR. The scale shows the nucleotide size of the DNA
fragment.
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To obtain the
C. rugosa gpd gene, consensus sequences were
determined by aligning multiple related DNA sequences from a database.
The partial
C. rugosa gpd DNA fragment was obtained by
RT-PCR
with degenerative primers designed from the consensus sequences
indicated in Table
1. This DNA fragment
was then cloned into
the pGEMT vector (Promega), and its sequence (327 bp) was determined
on both strands.
Competitive RT-PCR.
Competitive RT-PCR was conducted as
previously described for RT-PCR, except that known concentrations of
competitor DNA, an exogenous template as an internal PCR standard, were
spiked into a series of PCR tubes containing constant amounts of cDNA
generated from total RNA. The competitor DNA has the same primer
recognition sites as the target and thus competes with the target for
the same primers during the amplification. It is important to select the appropriate lip competitor DNAs and the specific primer
sets used for competitive RT-PCR, since one primer set must amplify only one lipase gene among the highly related gene family. Competitor DNA vectors (Fig. 1) were constructed by restriction digestion within
each lipase-encoding region followed by self-ligation; therefore, after
competitive PCR, the products of target and competitor DNA could be
distinguished by their size. The competitor DNAs were obtained by PCR
with primer pairs present in the vectors. To remove PCR template and
primers, the competitor DNA fragment was eluted after agarose
electrophoresis. The specific primer set, as shown in Table 1, had a
unique and specific 5' primer and a common 3' primer for each lipase
gene. These primer sets were tested for specificity and efficiency by
PCR (data not shown).
Determination of the quantity of the competitor DNA used to assess the
amount of target cDNA is important for precise quantitation
by
competitive RT-PCR. An accurate quantity of the competitor
DNA was
determined by capillary gel electrophoresis (CGE). All
CGE analyses
were conducted on the P/ACE system 5510 (Beckman
Instruments,
Fullerton, Calif.). Separations were carried out
in the reversed mode
(anode at detector side) with the Beckman
eCAP dsDNA 20,000 kit. UV
on-line detection was set at 254 nm
with a running temperature of
20°C. Samples were injected for
10 s by pressure, and separation
was performed at 9.0 kV for 25
min in a 47-cm capillary tube. Data was
collected and analyzed
automatically using Gold Software, Version 8.1 (Beckman). The
peak of the competitor DNA was identified by the
retention time.
The concentration of the competitor DNA was determined
from the
calibration curve derived from the simultaneous injection of a
molecular weight standard. The precise amount of the competitor
DNA was
calculated by integrating the area of its peak on a CGE
chromatogram.
The same quantity of competitor DNA was rechecked
by electrophoresis
with ethidium bromide
staining.
Detection and quantification of competitive PCR products.
Each PCR mixture (10 µl) was immediately loaded onto 1% agarose gels
in TAE buffer (40 mM Tris-acetate, 1 mM EDTA). Electrophoresis was then
performed at 100 V and room temperature for 60 min. After migration,
the gels were stained with a fluorescent double-stranded DNA-specific
stain (SYBR Green I; Molecular Probes, Eugene, Oreg.) for 20 min and
then directly scanned and visualized with the STORM system (Molecular
Dynamics, Sunnyvale, Calif.). The SYBR Green I-bound DNA bands in the
gel were excited at 497 nm and emitted at 520 nm.
The chemifluorescence data were obtained by computer-based video image
analysis (ImageQuaNT software; Molecular Dynamics).
The fluorescence
intensities were proportional to the amount of
DNA in the samples. To
correct for differences in molecular weight
and to enable direct
comparison of the corrected intensities of
the competitor bands with
the target bands, the fluorescence intensity
of the competitor band was
multiplied by a factor of target size
(bp)/competitor size (bp). In the
determination of the competition
equivalence point (EP), the
log
10 of the intensity ratio of competitor
to target was
plotted as a function of the log
10 of the initial
amount of
competitor (
33). By linear regression analysis, the
fluorescence intensity of the competitor band should be equal
to that
of the target band at the EP, and their ratio should be
equal to 1. Interpolation on the plot for a
y axis value of 0
(log
10 1 = 0) gives the copy number of target present
in the test
sample.
Nucleotide sequence accession number.
The DNA sequence of
C. rugosa gpd has been submitted to the GenBank database
under accession number AF025307.
| |
RESULTS |
Validation of competitive PCR analysis.
To ensure a successful
analysis of gene expression within the highly conserved lip
gene family, the quality of the RNA, the design of competitor DNAs and
specific primer set, the choice of control gene, the quantitation of
competitor DNAs, and the evaluation of competitive PCR results were considered.
Because all the five cloned
C. rugosa lip sequences are
intronless genes (
24,
26) frequently discovered in
yeast-like
fungi (
15), any genomic DNA contamination during
the RNA preparation
will lead to false-positive and ambiguous results
in RT-PCR. Therefore,
an accurate quantitation with the competitive
RT-PCR substantially
relies on the quality of the RNA preparation. The
RNA was purified
by a hot-phenol extraction method as well as the
additional procedures,
including treatment with either DNase I or
guanidine thiocyanate
(Gdn/HSCN) and
N-lauroylsarcosine
(
6,
8) in RNA preparation
prior to RT. RNA samples prepared
under various culture conditions
by different procedures were
demonstrated to have good integrity
(Fig.
2A); however, no genomic DNA
contamination was observed
only in the RNA samples isolated by the
Gdn/HSCN procedure with
RNA-PCR (Fig.
2B). Consequently, the lipase
gene could be PCR
amplified only from the first-strand cDNA generated
from the RNA
isolated by the Gdn/HSCN procedure, instead of the genomic
DNA,
as shown in lane 3 to 6 of Fig.
2C. In conclusion, we have
established
an improved protocol for isolating high-quality RNA, free
of contaminating
DNA from fungi, which can be readily used in
competitive RT-PCR
to detect mRNA expression.
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FIG. 2.
Analysis of RNA integrity, genomic DNA contamination,
and lip gene expression by RT-PCR. C. rugosa was
cultured in YM alone (lanes 1 to 3) or containing 1% olive oil (lane
4), 1% oleic acid (lane 5), or 1% Tween 20 (lane 6). RNA was isolated
or treated by different methods as indicated. (A) RNA samples (5 µg
per lane) were electrophoresed on a 1% native agarose gel and stained
with ethidium bromide. Arrows indicate the positions of the 25S and 17S
rRNA bands. (B) Genomic DNA contamination was detected by RNA-PCR with
prepared RNA as the template and the lipase-specific primers (LIP1sp-5'
and LIP-3' in Table 1). The position of lip1 is indicated by
the arrow. (C) Gene expression of lip1 was analyzed by
RT-PCR with cDNA generated from related RNA as the template. M,
 /HindIII DNA size markers.
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The choice of the copy number range of competitor DNA is critical for
quantitative sensitivity. The appropriate range for
the competitor DNA
added to the PCR mixture was inferred from
a comparison between a broad
range of fivefold serial dilutions
(4 to 0.00128 amol) and a narrow
range of twofold serial dilutions
(0.8 to 0.025 amol) (Fig.
3A). The results showed that a fine-tuned
narrow range of competitor DNA would be coamplified with the target
cDNA and that the patterns of PCR products (the ratios of target
to
competitor) would properly measure the quantity of
lip
transcripts.
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FIG. 3.
Validation of competitive PCR analysis. (A) Titer
determination of lip1 competitor with a constant amount of
cDNA generated from RNA isolated from YM-cultured C. rugosa.
Fivefold (lanes 1 to 6) (4 to 0.00128 amol) and twofold (lanes 7 to 12)
(0.8 to 0.025 amol) serial dilutions of lip1 competitor were
coamplified with a constant amount of cDNA. After 40 cycles of
amplification, the products were resolved on a 1% agarose gel and
stained with ethidium bromide. The positions of the 1,188-bp
lip1 target (T) and 813-bp competitor (C) PCR products are
indicated. Lane M contains a HindIII digest of DNA
as a size marker. (B) Analysis of relative changes in lip1
target levels by competitive PCR. cDNA samples generated from 125 and
62.5 ng of total RNA were amplified in the presence of twofold
dilutions of lip1 competitor (the same as in panel A). A
total of 40 cycles of PCR and electrophoresis on a 1% agarose gel were
performed. The gels were stained with SYBR Green I. The DNA products
visualized by chemifluorescence were scanned and quantitatively
analyzed as described in Materials and Methods. The open and solid
circles denote data derived from 62.5 and 125 ng of RNA,
respectively.
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We reconfirmed the sensitivity of the competitive PCR method by
determining the effect of the increasing amount of cDNA. cDNA
samples
generated from 125 and 62.5 ng of total RNA were coamplified
with
twofold serial dilutions of the
lip1 competitor. The copy
numbers of
lip1 transcript were calculated by determining
the
competition equivalence points in linear regression plots as
described
in Materials and Methods (Fig.
3B). The correlation
coefficients
(
r2) of the lines determined by
least-square regression analysis
were very close to 1. From
mathematical considerations of competitive
PCR (
34), the
slopes of the lines, ranging from 0.9 to 1.0,
suggested that the
amplification efficiencies of the target and
the competitor were
similar in the same reaction tube. The copy
numbers of
lip1
cDNA determined from the 125- and 62.5-ng RNA
plots (Fig.
3B) were
2.60 ± 0.02 × 10
4 and 1.42 ± 0.01 × 10
4 molecules, respectively, giving a 1.83-fold difference
(very
close to the predicted value of 2.0-fold). In consequence,
competitive
PCR can be used to accurately analyze the
lip
gene expression
in a quantitative
manner.
Differential expression of C. rugosa lip genes under
different culture conditions.
The expression of the different
lip genes in various culture conditions was examined by
competitive RT-PCR with specific primers and running for 40 PCR cycles.
As shown in Fig. 4, abundant amounts of
lip1, lip3, and gpd transcripts were
observed. The copy numbers of lip1, lip3, and
gpd transcripts were calculated by determining the
competition EPs in regression plots (Fig. 4 and Table
2). Correlation coefficients
(r2 values) and slopes of the regression lines
were within the confidence limits. gpd, a housekeeping gene,
was used as an experimental control because it was highly expressed
(approximately 100-fold higher than lip1) and its expression
pattern was seldom affected by various treatments (giving a mean of
2.12 × 107 ± 0.19 × 107
molecules per µg of total RNA). Experimental errors in the sample preparation or assay condition can be ruled out by normalizing the
measured copy number of each lip transcript to that of the gpd transcript.
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FIG. 4.
Quantitation of lip1, lip3, and
gpd mRNAs by competitive RT-PCR. Twofold serial dilutions of
the competitors (lanes 1 to 6) (0.8 to 0.025 amol for lip1
and lip3; 250 to 7.8 amol for gpd) were added to
PCR mixtures containing a constant amount of cDNA generated from RNA
isolated from C. rugosa cultured in YM, YM containing 1%
olive oil, YM containing 1% oleic acid, or YM containing 1% Tween 20. The gpd gene was used as an endogenous standard for
calibration among different culture conditions. After 40 cycles of
amplification, the products were resolved on a 1% agarose gel stained
with SYBR Green I. The gels were then scanned and quantitatively
analyzed as described in Materials and Methods. Quantitative plots are
shown (insets). The positions of the corresponding target (T) and
competitor (C) PCR products are indicated.
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Expression of
lip1 at 2.08 × 10
5 ± 0.02 × 10
5 molecules per µg of total RNA was
observed when
C. rugosa was cultured in YM,
and the amount
did not change after adding 1% olive oil or Tween
20. However,
addition of oleic acid (1%) to the medium did reduce
the number of
lip1 transcripts by 44%.
lip3 expressing
1.42 ×
10
5 ± 0.03 × 10
5
molecules per µg of total RNA (68% of
lip1 expression)
was detected
when
C. rugosa was cultured in YM. When olive
oil or oleic acid
was added to the cultures, the
lip3
transcripts increased by a
significant 2.1-fold and 1.53-fold,
respectively. In contrast,
the
lip3 transcripts decreased
12% when Tween 20 was added to
the
YM.
The levels of
lip2,
lip4, and
lip5
were much lower (Fig.
5). Since the
quantities of these three
lip genes were too low to
reach
the plateau phase within 40 PCR cycles, it may not be possible
to
accurately quantitate them in regression plots. The EPs of
various
culture conditions in individual gene could be estimated
from lanes
with similar target/competitor ratios (Table
2). When
C. rugosa was cultured in YM, the amounts of
lip2,
lip4, and
lip5 expressed relative to
lip1 were estimated to be 0.5, 0.1 and 0.4%,
respectively.
Olive oil also enhanced
lip2,
lip4, and
lip5 expressions
by about twofold, and oleic acid promoted
the mRNA expression
of
lip4 and
lip5 by four- and
twofold, respectively. Only
lip4 was highly induced by Tween
20, by a dramatic 7.9-fold.
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FIG. 5.
Quantitation of lip2, lip4, and
lip5 mRNAs by competitive RT-PCR. Twofold serial dilutions
of the competitors (lanes 2 to 7) (0.032 to 0.001 amol for
lip2 and lip5; 0.0128 to 0.0004 amol for
lip4) (lane 1 contains a HindIII-digest of
DNA as a size marker) were added to PCR mixtures containing a
constant amount of cDNA generated from RNA isolated from C. rugosa cultured in YM, YM containing 1% olive oil, YM containing
1% oleic acid, or YM containing 1% Tween 20. After 40 cycles of
amplification, the products were resolved on a 1% agarose gel stained
with SYBR Green I. The gels were then scanned in the same way as in the
experiment in Fig. 4 and quantitatively analyzed as described in the
text. The positions of the corresponding target (T) and competitor (C)
PCR products are indicated.
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DISCUSSION |
Studies on the differential expression of C. rugosa lip
gene have been hampered by the difficulties of quantitation of
individual mRNAs due to the highly related DNA sequences among this
gene family. In the present study, we developed a sensitive and
specific competitive RT-PCR method to quantitate the individual mRNA
transcript of the five C. rugosa lip genes. The results
demonstrated that the amount of lip transcripts follows the
descending order of lip1, lip3, lip2,
lip5, and lip4. lip1 and lip3 achieve
higher expression, whereas expression of lip2,
lip4, and lip5 is only 0.1 to 0.5% of the
expression of lip1 transcript under YM culture conditions.
These expression profiles are consistent with the findings that LIP1
and LIP3 are the major lipase proteins obtained by purification methods
(9, 21, 36).
Lotti and Alberghina (25) suggested that C. rugosa constitutively produces LIP1 as the major isoform whereas
expression of the other isoenzymes could be modulated according to the
growth condition. Recently, Lotti et al. (28) hypothesized
that lip genes might be subjected to different regulations.
Some of them are constitutively expressed, and others are switched on
by fatty acids. However, the DNA probe and antibody they used were
nonspecific and were not sensitive enough to determine which members of
the lip multigene family are in fact expressed under each
condition at either the RNA or protein level. In this study, we have
developed a sensitive and specific competitive RT-PCR method for
precisely quantitating the steady-state levels of mRNAs for the
different lipase genes. We showed that all the five lip
genes are transcriptionally active and that different inducers may
change the expression profile of individual genes. The constitutively
expressed lip1 and lip3 showed few changes at the
transcriptional level in various culture media, while suppression of
lip1 by oleic acid and induction of lip3 by olive
oil was demonstrated. Olive oil and oleic acid also promoted the
expression of inducible lip2, lip4, and
lip5 even in the presence of glucose, which previously was
reported to be a repressing carbon source (28). Obviously,
Tween 20 had a significant inducing effect on lip4 only.
These results elucidate that the different lipase enzyme profiles under
various culture conditions that we observed previously (5)
were due to the differential expression of lip genes.
Previously, lipase isozymes purified from a commercial preparation
(Lipase type VII [Sigma, St. Louis, Mo.]) were deduced to be products
of different genes based on partial peptide sequencing (9,
36). However, LIP3, purified by Rúa et al. (36)
as a major component of this lipase preparation, was not detected in
any of the purified lipase preparations obtained by Diczfalusy et al.
(9). This may be explained by our conclusion that the expression profile of C. rugosa lip genes can be altered by
different culture conditions and even by batch-to-batch culture
differences. We previously reported that different lipase isoforms from
C. rugosa displayed quite different substrate specificities
and thermostabilities (5, 39). Therefore, the production of
different lipase isoforms in response to different growth conditions is
physiologically important for C. rugosa, enabling it to grow
on various substrates and in different environments. Traditionally, the
culture conditions in fermentation are optimized for the maximal
production of enzyme activity units. Our results indicate that quality
is as important as quantity in enzyme preparations, since different
culture conditions might result in production of heterogeneous
compositions of the isozymes, which display different catalytic
activities and specificities. By engineering the culture conditions, we
can obtain enzyme preparations enriched in selected isozymes for
particular biotechnological applications.
The multiplicity of genes encoding isozymes has been reported for many
other yeast species (1, 3a, 16, 30, 31). The competitive
RT-PCR method developed in this study can be used to examine the
possible differential regulation of other yeast gene families (11,
20). The expression level of the mRNA transcript may be affected
by many factors, such as promoter activity, upstream regulatory
elements, and stability of the mRNA. Although elements such as the CAAT
and TATAA boxes characteristic of eukaryotic promoters for
transcriptional initiation have been found in the conserved regions
upstream from the lip genes (26), we anticipate that transcriptional regulatory elements of C. rugosa lip
promoters should be localized upstream from the conserved regions.
Recently, nutrient-related transcriptional controlling elements have
been identified in Saccharomyces cerevisiae (7).
By assaying the 
-galactosidase activities of
promoter-lacZ fusions in S. cerevisiae, our
unpublished data showed that the promoter activities of lip3 under various culture conditions were much higher than those of lip4 promoter. These findings suggest that the expression
profile of lip genes could be accompanied by different
regulation of the lip promoter activities. Studies of the
transcriptional controlling elements of C. rugosa lip genes
to further elaborate the mechanism of differential regulation of
lip genes by various inducers are under way.
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ACKNOWLEDGMENTS |
This work was supported by grants NSC-83-0409-B-019-008,
NSC-85-0409-B-019-008, and NSC-87-2311-B-001-047 from the National Science Council, Republic of China.
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FOOTNOTES |
*
Corresponding authors. Mailing address for Dr. Shaw:
Institute of Botany, Academia Sinica, Nankang, Taipei, Taiwan 11529. Phone: 886-2-27899590 ext. 226. Fax: 886-2-27821605. E-mail:
boplshaw{at}gate.sinica.edu.tw. Mailing address for Dr.
Tang: Institute of Marine Biotechnology, National Taiwan Ocean
University, Keelung, Taiwan 20224. Phone: 886-2-24622192 ext. 5510. Fax: 886-2-24622320. E-mail:
tsj{at}ntou-66.ntou.edu.tw.
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Applied and Environmental Microbiology, September 1999, p. 3888-3895, Vol. 65, No. 9
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Abstract
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