Previous Article | Next Article 
Applied and Environmental Microbiology, October 2001, p. 4781-4788, Vol. 67, No. 10
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4781-4788.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Development of a Combined Selection and Enrichment
PCR Procedure for Clostridium botulinum Types B, E, and
F and Its Use To Determine Prevalence in Fecal Samples from
Slaughtered Pigs
Maria
Dahlenborg,1,2
Elisabeth
Borch,2 and
Peter
Rådström1,*
Applied Microbiology, Center for Chemistry
and Chemical Engineering, Lund Institute of Technology, Lund
University, SE-221 00 Lund,1 and Swedish
Meats R&D, SE-222 40 Kävlinge,2 Sweden
Received 6 April 2001/Accepted 1 August 2001
 |
ABSTRACT |
A specific and sensitive combined selection and enrichment PCR
procedure was developed for the detection of Clostridium
botulinum types B, E, and F in fecal samples from slaughtered
pigs. Two enrichment PCR assays, using the DNA polymerase
rTth, were constructed. One assay was specific for the
type B neurotoxin gene, and the other assay was specific for the type E
and F neurotoxin genes. Based on examination of 29 strains of C.
botulinum, 16 strains of other Clostridium spp.,
and 48 non-Clostridium strains, it was concluded that
the two PCR assays detect C. botulinum types B, E, and F
specifically. Sample preparation prior to the PCR was based on heat
treatment of feces homogenate at 70°C for 10 min, enrichment in
tryptone-peptone-glucose-yeast extract broth at 30°C for 18 h,
and DNA extraction. The detection limits after sample preparation were
established as being 10 spores per g of fecal sample for nonproteolytic
type B, and 3.0 × 103 spores per g of fecal sample
for type E and nonproteolytic type F with a detection probability of
95%. Seventy-eight pig fecal samples collected from slaughter houses
were analyzed according to the combined selection and enrichment PCR
procedure, and 62% were found to be PCR positive with respect to the
type B neurotoxin gene. No samples were positive regarding the type E
and F neurotoxin genes, indicating a prevalence of less than 1.3%.
Thirty-four (71%) of the positive fecal samples had a spore load of
less than 4 spores per g. Statistical analysis showed that both rearing conditions (outdoors and indoors) and seasonal variation (summer and
winter) had significant effects on the prevalence of C.
botulinum type B, whereas the effects of geographical location
(southern and central Sweden) were less significant.
| |
INTRODUCTION |
Clostridium
botulinum is an obligate anaerobic,
endospore-forming bacterium that is ubiquitous in the environment. The
pathogenicity of the organism is associated with the production of
serologically distinct neurotoxins, types A to G (15).
Types A, B, E, and F, which are responsible for food-borne botulism in
humans, contain both proteolytic and nonproteolytic strains. Only a
limited number of surveys have been performed in Europe on the
distribution of C. botulinum in the environment and in raw
materials for foodstuffs (16, 18-20). In Sweden, the most
recent survey was carried out in 1963 by Johannsen (22) in
soil and coast sediment. Both C. botulinum types
B and E were found.
Today, there is an increasing demand from the consumer for convenient
foods of high quality. This has resulted in the development of
refrigerated processed foods of extended durability that require minimal preparation time and contain low concentrations of
preservatives. Nonproteolytic strains of C. botulinum are
considered to be a hazard in these foods due to lower heat treatment
than, for example, canned foods and the anaerobic atmosphere used
during processing of the foods (29). These conditions
favor germination of spores and toxin formation from nonproteolytic
C. botulinum, which can multiply and produce
neurotoxins of types B, E, or F at temperatures as low as 3.3°C
(8, 9, 32). The potential presence of C. botulinum spores in these foods represents the most severe hazard of food-borne poisoning (15, 29). Therefore,
information about the incidence and levels of spores in environmental
samples and in foods is necessary for an assessment of the botulinum hazard.
Many previous investigations into the prevalence of C. botulinum in the environment and in food samples are based
on enrichment for 5 to 10 days and subsequent detection by in vivo
mouse bioassay of the toxin produced (24, 33). A drawback
of the in vivo mouse bioassay (23) is the use of
experimental animals. Conventional isolation and identification
methods, based on phenotypic characteristics, have also been developed,
but their performance is often time consuming and they have been found
to be insufficient in identifying strains of C. botulinum correctly (7, 27). In recent years, a
number of PCR assays have been developed for the detection of botulinal
neurotoxin genes (5, 36). However, the amplification capacity of many PCR assays is limited due to the presence of PCR-inhibitory components, low concentration of target cells or DNA,
and high concentration of microorganisms within the complex biological
sample (25). Thus, the probability of detecting the target
organism with a PCR assay at different cell concentrations needs to be
established both in a pure system and in a system containing the
biological sample to be analyzed.
The first objective of this study was to construct a highly specific
and sensitive PCR detection method for C. botulinum types B, E, and F in pig fecal samples. As a
sample preparation step prior to PCR, we developed a combined selection
and enrichment procedure including activation, germination, and
transformation of bacterial endospores into active vegetative bacteria
by heat treatment and multiplication of the bacterium in enrichment
broth to a PCR-detectable concentration. Two nested PCR assays were constructed using rTth, a DNA polymerase that has been found
to exhibit high levels of resistance to various known PCR-inhibitory components (1-3). One PCR assay is specific for the type
B neurotoxin gene, and the other assay is specific for both the type E
and F neurotoxin genes. The second objective of this study was to determine the prevalence of C. botulinum types B,
E, and F in slaughtered pigs in Sweden.
| |
MATERIALS AND METHODS |
Statistical analyses.
In the development of the combined
selection and enrichment PCR procedure, the data obtained from the
different experiments were analyzed by replicate analyses. The number
of replicates in the different experiments varied between 8 and 24. Each replicate resulted in an individual PCR detection pattern from
which a probability of detecting the bacteria at different
concentrations was calculated (P = number of positive
PCR results/number of replicates tested). The detection probability
curves were adjusted using a model described by Baranyi and Roberts
(4). The statistical analysis of the results from the
prevalence study was performed using the software SYSTAT 227 (version
7.0.1; SPSS INC., Evanston, Ill.). Both analysis of variance (ANOVA)
and the Pearson chi-square test were used for statistical analysis. The
following model, in which Y represents prevalence, was
used when ANOVA was performed:
Y = season + rearing condition + geography + season × rearing
condition + season × geography + rearing condition × geography + season × rearing condition × geography
Bacteria and culture conditions.
Twenty-nine strains
of C. botulinum, 16 strains of other
Clostridium spp., and 48 non-Clostridium strains
were used (Table 1). All the
C. botulinum strains were grown anaerobically in tryptone-peptone-glucose-yeast extract (TPGY) broth at 30°C. The TPGY
broth contained tryptone (50 g/liter; Oxoid Ltd., Basingstoke, United Kingdom), Proteose Peptone (5 g/liter; Oxoid Ltd.), yeast extract (20 g/liter; Oxoid Ltd.), D-glucose (4 g/liter; Merck, Darmstadt, Germany), sodium thioglycolate (1 g/liter;
Merck), and soluble starch (1 g/liter; Merck). Before sterilization
(121°C for 15 min), anaerobic conditions were created by boiling the medium for 10 min and, during cooling, flushing the medium with nitrogen gas. All the other strains of Clostridium spp. were
grown anaerobically in TPGY broth at 37°C. The
non-Clostridium strains were cultivated at 37°C in brain
heart infusion broth (Oxoid Ltd.) except as follows.
Lactobacillus strains were grown in MRS broth (Oxoid Ltd.);
Brochothrix, Pediococcus, and
Streptococcus strains were grown in APT broth (BBL, Becton
Dickinson Microbiology Systems, Cockeysville, Md.); and
Escherichia strains were grown in tryptone soy broth (Oxoid
Ltd.).
Preparation of spores.
Spores from two proteolytic strains
of C. botulinum type B (ATCC 17841 and ATCC 7949)
were produced in a sporulation medium as described by Gaze and Brown
(14). Spores from three nonproteolytic strains of
C. botulinum type B (Eklund 2B, Eklund 17B, and
Johannesson 105-66), C. botulinum type E
(CB-S3-E), and C. botulinum type F (Craig
610B8-6F) were produced in a two-phase medium as described by Peck et
al. (30). During the production of spores, contamination was checked on blood agar plates {blood agar base (37 g/liter; Lab M,
Bury, United Kingdom) and citrate-treated horse blood (4% [vol/vol];
SVA, Uppsala, Sweden)} after aerobic and anaerobic incubation (Gas
Pak Plus Anaerobic system; BBL, Becton Dickinson Microbiology Systems)
at 30°C for 48 h. The spores were finally resuspended in water
and stored at 1 to 2°C. The enumeration of vegetative cells and
spores of C. botulinum was performed using a
Bürker chamber in a phase-contrast microscope. The bacterium or
spore suspension was diluted so that each square contained approximately 10 to 15 cells or spores. Counting was performed three
times per suspension, and the mean value was calculated. The
concentration was expressed as cells or spores per milliliter.
Primer design.
Alignment (Clustal X 1.64b Multiple Sequence
Alignment Program) (35) of sixteen published nucleotide
sequences of the botulinal neurotoxin gene of all seven, A to G,
serotypes was performed. The nucleotide sequences were collected from
the GenBank Sequence Database (http://www.ncbi.nlm.nih.gov) for
C. botulinum type A (accession no. X52066,
X73423, and M30196), C. botulinum type B
(accession no. X71343 and M81186), C. botulinum
types C and D (accession no. D38442, X54254, and D49440), C. botulinum type E (accession no. X62683 and X62089),
C. botulinum type F (accession no. L35496,
X81714, and M92906), C. botulinum type G
(accession no. X74162), Clostridium butyricum type E
(accession no. X62088), and Clostridium baratii type F (accession no. X68262). Two sets of primers were designed, one
specific for the type B neurotoxin gene and one specific for the type E
and F neurotoxin genes (Table 2).
Conditions in the PCR assays.
A nested PCR strategy was used
for the two PCR assays, one specific for the type B neurotoxin gene and
one specific for both the type E and F neurotoxin genes. In the first
step of the assays, amplification was performed with the following set
of primers: primers fB and rB for type B and primers fEF and rEF for
types E and F (Table 2). The total volume of the PCR master mixture was
25 µl. The PCR mixture consisted of 1× chelating buffer (Perkin Elmer Applied Biosystems, Foster City, Calif.); 1.5 mM
MgCl2 (Perkin Elmer Applied Biosystems); 0.5 µM
concentrations of each primer (Scandinavian Gene Synthesis AB,
Köping, Sweden); 0.2 mM (each) dATP, dTTP, dCTP, and dGTP
(Amersham Pharmacia Biotech, Inc., Piscataway, N.J.); 1.25 U of
rTth (Perkin Elmer Applied Biosystems); and 5 µl of
template solution (DNA or cells). The water used in the assay was
autoclaved Millipore water (Millipore S. A., Molsheim, France).
Each amplification commenced with a denaturation step at 94°C for 5 min, followed by 30 cycles for the type B-specific primer and 35 cycles
for the type E-and-F-specific primer, consisting of heat denaturation
at 94°C for 40 s, primer annealing at 58°C for 40 s, and
extension at 72°C for 40 s. Finally, extension was performed at
72°C for 7 min to complete the synthesis of all strands. The second
step consisted of amplification with an internal set of primers,
primers fBn and rBn for the first assay, and primers fEFn and rEFn for
the second assay, using the same conditions as those described above.
The template added to the second step (1 to 25 µl), was diluted 1:500
in autoclaved Millipore water. All amplifications were carried out in a
Gene Amp 9700 thermal cycler (Perkin-Elmer Cetus, Norwalk, Conn.). The
PCR products were visualized by agarose gel electrophoresis. The 1.5%
(wt/vol) agarose gel was obtained by dissolving ultrapure DNA grade
agarose (Bio-Rad Laboratories, Hercules, Calif.) in 1× TBE buffer (108 g of Tris [ICN Biochemicals, Inc., Aurora, Ohio], 55 g of
boric acid [Merck], 40 ml of 0.5 M EDTA [Sigma Chemical Co., St.
Louis, Mo.], 10 liters of Millipore water) and stained with
ethidium bromide (Sigma Chemical Co.) (31). Five
microliters of 6× loading buffer (40% [wt/vol]
D-sucrose [Sigma Chemical Co.] and 0.025% [wt/vol] bromophenol blue [Merck] dissolved together in 1× TBE buffer) was added to each PCR tube. The gel was analyzed using a gel
documentation system (Bio-Rad Laboratories), and Molecular Analyst
software (Bio-Rad Laboratories) was used to analyze the gel images. To
determine the size of the amplicons, a DNA molecular size marker
(100-bp ladder; Amersham Pharmacia Biotech, Inc.) was included in the gel.
In order to optimize the reaction conditions for the two assays, pure
DNA was used as a template. DNA was extracted from
C. botulinum Eklund 2B,
C. botulinum
CB-S3-E, and
C. botulinum Craig
610B8-6F in
accordance with standard protocol (
31) and modified
by the
addition of 30 U of mutanolysine (Sigma Chemical Co.) per
ml of lysis
solution. The DNA was further purified by using
phenol-chloroform-isoamyl
alcohol (25:24:1) (Sigma Chemical Co.)
extraction. The DNA was
resuspended in 500 µl of TE buffer containing
1 µl of 100-mg/ml
RNase A (Sigma Chemical Co.) The concentration and
purity of the
DNA were measured spectrophotometrically at 260 and 280 nm. When
Taq DNA polymerase (Roche Diagnostics GmbH,
Mannheim, Germany)
was added to the PCR mixture, the same reaction
conditions were
used as for
rTth, except for the use of 1×
PCR buffer (Roche Diagnostics
GmbH) as the buffer system and a
concentration of 0.75 U of
Taq DNA polymerase in a
total volume of 25 µl.
Preparation of standardized pig fecal samples.
The PCR assay
was evaluated using inoculated pig feces. Approximately 2.5 kg of pig
feces was sampled at the slaughter line in a slaughterhouse. The feces
were transported to the laboratory within 2 h at a temperature
below 4°C. The feces were homogenized in a sterilized meat grinder,
mixed, distributed into stomacher bags in portions of approximately 0.1 kg, and stored at 
20°C. Before use, the samples were thawed at
4°C for 16 h.
Fecal samples were prepared by mixing 10 g of feces with 90 ml of
TPGY broth in a stomacher for 2 min. Homogenates (9 ml)
were
transferred to six sterile test tubes. Large debris was avoided
when
withdrawing the samples. Each tube was spiked with cells
of
C. botulinum type B (Eklund 2B) to a final
concentration of
1.6 × 10
2 to 1.6 × 10
7 cells per ml of homogenate. Ten 1-ml samples
from each test tube
were subjected to DNA extraction using a PrepMan
kit for gram-positive
bacteria (Perkin Elmer Applied Biosystems).
Finally, after DNA
precipitation, the pellet was resuspended in 100 µl of autoclaved
Millipore
water.
Preparation of microorganisms from pig feces.
Ten grams of
feces was mixed with 90 ml of 0.9% (wt/vol) NaCl solution in a
stomacher for 2 min. Eight samples of the homogenate (0.83 ml) were
transferred to sterile Eppendorf tubes, and the microorganisms in each
homogenate were extracted using a buoyant density centrifugation
technique (Percoll; Amersham Pharmacia Biotech, Inc.) (26)
with the following modifications. The bacterial cells were washed in
0.9% (wt/vol) NaCl at least five times. The remaining supernatant (83 µl) was prepared so that the final volume of 100 µl corresponded to
1 ml of feces homogenate (1:10 dilution).
Development of the combined selection and enrichment
procedure.
All enrichment was performed in TPGY medium at 30°C
for up to 48 h. In order to select for C. botulinum in pig feces, two different methods were
evaluated: (i) heat treatment of sample homogenate at 70°C for 10 min
prior to enrichment and (ii) addition of
D-cycloserine to the enrichment broth. When heat
treatment was used, 90 ml of TPGY was preheated to 70°C and
inoculated anaerobically with 10 g of feces to make a 1:10
dilution and the mixture was vortexed for 1 min and reheated. After the
temperature reached 70°C, the homogenate was inoculated with spores
of nonproteolytic C. botulinum type B (Eklund 2B
and Eklund 17B) at a low concentration (10 spores per sample of
homogenate) and heating was continued for 10 min. A logger in a
reference bottle, connected to a computer, monitored the temperature.
When D-cycloserine (25 mg/ml) was used, it was
mixed in the TPGY medium before the feces and the spores were added.
The control samples were enriched in TPGY medium at 30°C without
additional selection. Two 1-ml samples were withdrawn after 0, 18, 24, 30, 36, and 48 h of enrichment and subjected to analysis with the
PrepMan DNA extraction kit for gram-positive bacteria (Perkin Elmer
Applied Biosystems). Each sample was amplified three times with the
type B PCR assay.
An additional experiment was performed at 70°C for 10 min combined
with enrichment for 18 h at 30°C. The homogenate samples
were
inoculated with spores from
C. botulinum strain
Eklund 2B
and strain Eklund 17B (both nonproteolytic type B),
C. botulinum strain ATCC 17841 (proteolytic type
B),
C. botulinum strain CB-S3-E
(type E), and
C. botulinum strain Craig 610B8-6F
(nonproteolytic
type F) at the following concentrations:
10
1, 10
2,
10
3, and 10
4 spores per
homogenate sample. The growth experiment was performed
twice with each
strain. Three 1-ml samples were withdrawn after
18 h of incubation
and treated in the same way as described above.
Each sample was
amplified twice with the PCR
assay.
Collection and analysis of naturally contaminated pig fecal
samples.
Seventy-eight pig fecal samples were collected at the
slaughter line of two slaughterhouses, one in southern Sweden (37 samples) and one in central Sweden (41 samples). The two
slaughterhouses are situated 600 km away from each other. Approximately
10 g of feces was collected from the rectum, transferred to
sterile tubes, and subsequently stored at 20°C until analysis.
Forty-five of the samples were collected during the summer (June to
September), and 33 of the samples were collected during the winter
(December to March). Thirty-six of the 78 fecal samples collected were
obtained from pigs reared outdoors, and the other 42 samples were
collected from pigs reared indoors.
Before analysis, the fecal samples were thawed at 4°C for 7 h
and mixed thoroughly in each tube. Three grams of feces was
subsequently inoculated into 27 ml of preheated TPGY broth (1:10
dilution) and analyzed according to the combined selection and
enrichment PCR procedure: (i) selection at 70°C for 10 min, (ii)
enrichment at 30°C for 18 h, (iii) DNA extraction with a PrepMan
DNA extraction kit for gram-positive bacteria (Perkin Elmer Applied
Biosystems), and (iv) amplification with the two nested PCR assays
with
the DNA polymerase
rTth (Perkin Elmer Applied Biosystems).
A
spore suspension of 10
3 spores (Eklund 2B) was
used to inoculate 30 ml of feces homogenate
(1:10 dilution in TPGY
broth) to be included as a positive control,
and 30 ml of TPGY broth
was included as a negative control in
each experiment. Two 1-ml samples
were withdrawn after enrichment,
and each sample was amplified twice
with the type B and type E
and F PCR
assays.
| |
RESULTS |
Specificity and detection limits of PCR assays.
Two sets of
primers were designed, one specific for part of the type B neurotoxin
gene and the other specific for part of both the type E and F
neurotoxin genes (Table 2). The specificity of the two nested PCR
assays was evaluated using 29 strains of C. botulinum neurotoxin serotypes A, B, E, and F; 16 strains of other Clostridium spp.; and 48 non-Clostridium
strains (Table 1). The assay for the type B neurotoxin gene resulted in
a PCR product at the predicted size, 0.22 kb, for all six C. botulinum type B strains. The Atlanta 2204-4 strain, a type
A strain, also resulted in a PCR product of the same size with the type
B assay. The assay for type E and F neurotoxin genes resulted in a
0.20-kb PCR product, as predicted, when tested on all 12 type E strains and all 6 type F strains. None of the other Clostridium spp.
strains or the non-Clostridium strains gave amplified PCR
products with the two tested PCR assays.
The detection probability of
C. botulinum types
B, E, and F in Millipore water was established at different cell
concentrations
(Fig.
1). The probability
curves display a difference of 2 log
units in the detection limit
between the two PCR assays at a detection
probability of 95%. The
detection limit for
C. botulinum type
B in water,
with a detection probability of 95%, was 1.6 × 10
3 cells per ml of water, which is approximately
8 cells per reaction
tube. The corresponding values for
C. botulinum types E and F
were 1.6 × 10
5 cells per ml of water and approximately
8.0 × 10
2 cells per reaction tube. By
adding equal amounts of purified
cells of the microorganisms from feces
to an increasing concentration
of
C. botulinum
Eklund 2B cells in water, a 3 log unit decrease
in sensitivity, at the
detection probability of 95%, to 1.6 ×
10
6
cells per ml, was found, that is, 8.0 × 10
3
cells per reaction tube (Fig.
1). Cells of
C. botulinum Eklund
2B were inoculated in fecal homogenates and
subsequently treated
with the PrepMan DNA extraction kit prior to PCR
to determine
the concentration of
C. botulinum
required to obtain detection
with a probability of 95%. The
concentration was established to
be 1.6 × 10
6 cells per ml of homogenate.
View larger version (13K):
[in this window]
[in a new window]
|
FIG. 1.
Detection probability curves of C.
botulinum types B, E, and F at different cell
concentrations obtained using the two PCR assays and testing the
effects of fecal microflora on the detection limit of the type B PCR
assay. P, no. of positive PCR results/no. of replicates
tested (n = 8 to 15);  , serial dilution of
C. botulinum type B cells (Eklund 2B and
Eklund 17B) in water (n = 15);  , serial dilution
of C. botulinum type B cells (Eklund 2B)
in water with the addition of purified fecal microbial flora
(n = 8);  , fecal homogenate (1:10 in TPGY broth)
spiked with a serial dilution of C.
botulinum type B cells (Eklund 2 B) and subsequently
treated with PrepMan DNA extraction kit (n = 10);
 , serial dilution of C. botulinum type
E and F cells (CB-S3-E and Craig 610B8-6F) in water
(n = 15).
|
|
Enrichment PCR procedure and detection limits in pig fecal
samples.
From the combinations of different selection procedures
and different enrichment times, the probability of detecting 10 spores of C. botulinum type B per 10 g of pig feces
was determined (Table 3). This low
concentration of spores was used based on the predicted low numbers of
spores in naturally contaminated fecal samples. Overall, heat treatment
at 70°C for 10 min resulted in a higher probability of positive PCR
results during 48 h of enrichment than did not performing
selection or the addition of 25-mg/ml D-cycloserine to the enrichment medium. Regarding
heat treatment, the highest probabilities (P) were
calculated after the following incubations: 18 h
(P = 0.42), 36 h (P = 0.50), and
48 h (P = 0.42). A similar probability
(P = 0.42) of positive PCR results was observed with
the combination of 24 h of enrichment and no selection.
View this table:
[in this window]
[in a new window]
|
TABLE 3.
Probability of detecting 10 spores of C. botulinuma type B per sampleb
using different selection methods and enrichment times prior to PCR
|
|
The fecal samples were inoculated with spores of proteolytic and
nonproteolytic
C. botulinum types B, E, and F to
determine
the detection limit in pig fecal samples after analysis with
the
combined selection and enrichment PCR procedure. The inhibitory
effect of the microorganisms in pig feces on the amplification
capacity
of two different DNA polymerases,
rTth and
Taq,
in the
PCR assay was studied (
n = 9). The results
showed a decrease in
sensitivity by 1 log unit when using
Taq DNA polymerase instead
of
rTth. All nine
replicates gave the same result (data not shown).
The detection limit
when using
rTth in the PCR assay for nonproteolytic
C. botulinum type B with a detection probability
of 95% was 10
spores per g of fecal sample (Fig.
2). The corresponding value
for
C. botulinum type E and nonproteolytic type F was
3.0 × 10
3 spores per g of fecal sample. For
the proteolytic
C. botulinum type B strain, no
positive PCR detection was observed at any of
the inoculated spore
concentrations after 18 h of incubation.
When increasing the
enrichment temperature from 30 to 37°C and
incubating the mixture for
18 h, a detection pattern similar to
that of the nonproteolytic
type B strains was observed (data not
shown).
View larger version (8K):
[in this window]
[in a new window]
|
FIG. 2.
Detection probability curves of C.
botulinum types B, E, and F at different spore
concentrations in pig fecal samples after analysis with the combined
selection and enrichment PCR procedure. P, no. of
positive PCR results/no. of replicates tested; , fecal homogenate
(1:10 in TPGY broth) spiked with a serial dilution of C.
botulinum type B spores (Eklund 2B and Eklund 17B)
(n = 24); , fecal homogenate (1:10 in TPGY
broth) spiked with a serial dilution of C.
botulinum type E and F spores (CB-S3-E and Craig
610B8-6F) (n = 24).
|
|
Prevalence of C. botulinum in pig
fecal samples.
Of the 78 pig fecal samples collected, 48 (62%)
gave a positive PCR result with respect to the type B neurotoxin gene.
No samples were positive regarding the type E and F neurotoxin genes, indicating that, if present, the prevalence of types E and F is less
than 1.3%. ANOVA showed that both variation in season (summer and
winter) and rearing conditions (outdoors and indoors) had significant
effects on the prevalence of C. botulinum type B
in pigs (Table 4). The distribution of
positive samples in the groups from the southern and central parts of
Sweden were similar. In the interplay between rearing conditions and
seasonal variation a larger difference in prevalence was observed
between the summer and winter periods in the group of outdoor pigs than
between the corresponding periods in the group of indoor pigs (Table
5).
View this table:
[in this window]
[in a new window]
|
TABLE 4.
Prevalence of C. botulinum type B in pig feces
regarding variations in geography, season, and rearing condition
|
|
Four replicates of each of the 78 fecal samples were analyzed using PCR
to semiquantify the number of spores for each sample
category (Table
6). The probability of detection was
determined
for each positive fecal sample as described above. This
ratio,
combined with the detection probability curve of
C. botulinum type B (Fig.
2), provided an indication of the
spore load in each
fecal sample. Thirty-four of the 48 positive fecal
samples (71%)
had a spore load of less than 40 spores per 10 g.
Within each
category (rearing condition, season) the distribution of
positive
samples, regarding the calculated probability, displayed a
shift
in the group of outdoor pigs from lower levels of spores during
the summer (
15 spores/10 g) to higher levels during the winter
(
40
spores/10 g). For the group of indoor pigs, the distribution
of spore
load remained approximately constant during the two sampling
periods.
View this table:
[in this window]
[in a new window]
|
TABLE 6.
Distribution of positive pig fecal samples (total 48 samples) within each category (rearing condition, season) with regard
to detection probabilitya
|
|
| |
DISCUSSION |
Various sets of primers for the detection of C. botulinum, either degenerated primers (5, 10)
or specific primers (34, 36), have been described
previously. In recent years, a number of nucleotide sequences of the
different types of the neurotoxin genes (A to G) have been published,
increasing the possibility of designing more specific primers. The
results of our study indicate that the two sets of primers, constructed
for type B and types E and F, are specific. The Atlanta 2204-4 strain,
a type A strain, resulted in a PCR product of the same size as the type
B-specific primers (0.22 kb). In the literature, the presence of a
silent or unexpressed type B gene is known to exist in several type A strains (6, 12), in which the presence of a stop codon and deletions has been identified (13, 21).
The detection limits for C. botulinum cells in
water varied by 2 log units between the two PCR assays, at a detection
probability of 95%. The slopes of the probability curves differed from
each other (Fig. 1). At a detection probability of 50%, the range for detecting type B cells with high probability is broader than the corresponding range for types E and F. The variation in the probability curves and detection limits between the two assays is probably due to
the different prerequisites for primer-target annealing efficiency. The
primers for the type B assay were designed from highly conserved
regions of the light chain of the type B neurotoxin gene, whereas the
primers constructed for the type E and F assay were designed from less
conserved regions of the heavy chain of the type E and F neurotoxin genes.
A number of components have been reported to be PCR inhibitors in
feces, namely bile salts, complex polysaccharides, proteinases, and a
high concentration of non-target DNA (25). In pig feces the number of microorganisms is estimated to be around
1010 bacteria per g (28). To
overcome these inhibitory effects when applying the PCR assays to
inoculated fecal samples, an enrichment step, followed by a DNA
extraction step, was included prior to PCR. By adding purified
microorganisms from feces to the PCR, it was assessed that the decrease
in sensitivity was mainly a consequence of the inhibitory effect
constituted by the high concentration of microorganisms on the PCR
amplification (Fig. 1).
Different combinations of selection procedures and different enrichment
times were evaluated with inoculation of the feces homogenate using low
concentrations of spores. This was because of the prediction of
detecting spores, not bacteria, in the pig feces. From the results of
the growth experiments, heat treatment at 70°C for 10 min and
enrichment at 30°C for 18 h was identified as the best
combination (Table 3). This was based on the highest number of positive
PCR results out of the total number of 12 replicates tested
(P = 0.42) together with the aspect of practicable
working hours in the laboratory. Heat activation of spores is well
described in the literature and is dependent on several factors, e.g.,
the heating temperature (15). In our study, the use of
heat treatment at 80°C for 10 min resulted in no PCR product at all
(data not shown), which is why the temperature was lowered to 70°C. A
combined selection and enrichment procedure has several advantages. The heat treatment activates the spores present in the fecal samples and
induces them to germinate and transform into vegetative bacteria. To
some extent, the heat treatment also provides for a reduction in the
numbers of competitive microorganisms. The enrichment in liquid broth
allows the bacteria to grow and multiply to PCR-detectable levels. The
only hitherto reported C. botulinum enrichment
PCR protocol evaluated on spores involves incubating the biological samples for 5 days with no selection, allowing the spores to germinate and transform into vegetative cells in the enrichment broth
(17). Two C. botulinum enrichment
PCR procedures, based on detection after an 18-h enrichment step, have
previously been described (10, 11), but these protocols
were evaluated on food samples inoculated with bacteria, not spores.
Although both Taq DNA polymerase and rTth worked
well in the combined selection and enrichment PCR procedure, the
detection limits were slightly different. A decrease in sensitivity by
1 log unit was observed when using Taq DNA polymerase
instead of rTth, which could be due to the inhibitory effect
of the microorganisms in pig feces on the amplification capacity. This
finding is in accordance with previously performed investigations, in
which rTth was found to exhibit a high level of resistance
to PCR-inhibitory components in biological samples (1, 2,
3). The detection limit for the combined selection and
enrichment PCR procedure, when using rTth in the PCR assay, was approximately 10 spores per g of fecal sample at a detection probability of 95% for nonproteolytic C. botulinum type B. The corresponding value for C. botulinum type E and nonproteolytic type F was 3.0 × 103 spores per g of fecal sample.
No surveys of C. botulinum types A, B, E, or F in
pigs have, to our knowledge, been published. In Scandinavia, the
incidence of the organism in terrestrial samples has been studied, and
both types B and E were found (19, 22), indicating a
potential risk of contamination of animals. In this study, a high
prevalence of C. botulinum type B was established
(62%). The reverse was true for types E and F, where no fecal sample
was found to be positive using the combined selection and enrichment
PCR procedure. This may indicate a prevalence of less than 1.3% of
types E and F in the fecal samples. Another explanation may be the 2 log unit difference in the detection limits between the two PCR assays for type B and types E and F. Furthermore, C. botulinum type F is rarely found in the environment, which
may also explain why no fecal samples were positive regarding this
type. As for type E, this organism is most often found in aquatic
environments and in fish samples (19).
The geographical area studied covers part of the lowlands of Sweden
where almost all pigs are reared. Both rearing conditions and seasonal
variation had a significant effect on the prevalence of C. botulinum type B in pigs. The prevalence of C. botulinum in the group of indoor pigs was markedly higher
than in the group of outdoor pigs (Table 4). As for seasonal variation,
a higher prevalence was observed in the fecal samples collected during the winter than in the samples collected during the summer. In this
article, the term outdoor pigs refers to the ecological production of
pigs; the major difference between this and conventional rearing is
that the animals are always outdoors during the summer and may stay
outdoors or in barns with conditions similar to the outdoor climate
during the winter. In addition, we observed a correlation between
rearing conditions and season. The prevalence of C. botulinum type B was higher in samples collected in the
winter than in samples collected in the summer in both groups. However,
the observed increase in prevalence was greater in the group of outdoor
pigs, increasing from 10% during the summer to 67% during winter,
than the corresponding increase, from 79 to 94%, in the group of
indoor pigs. This may be due to several factors, such as the microbial environment of the barn and/or feed. One theory may be that spores ingested by the outdoor pigs were spread out again in the environment during the summer. However, as the pigs are more often found indoors in
barns during the cold period of the year, the spores may be confined to
the environment of the barn, increasing the risk of recontamination of
the animal. This theory can also explain the higher prevalence in the
group of indoor pigs. The shift in the distribution of positive fecal
samples in the group of outdoor pigs, indicating a lower spore load
during the summer and a higher spore load during the winter, is
evidence of the recontamination of the animal (Table 6). This
difference could not be seen in the group of indoor pigs, where the
level of spores was approximately constant between the summer and
winter. Furthermore, a geographical difference was noticed among the
samples (Table 6). In the southern part of Sweden, the prevalence of
C. botulinum in the group of outdoor pigs
increased from 0% during the summer to 80% during the winter, whereas
the corresponding values in central Sweden were 22 and 40%,
respectively. For the group of indoor pigs, the prevalence of
C. botulinum remained constant at 100% in the
southern part of Sweden during the summer and winter, whereas the
values for central Sweden increased from 72% in the summer to 89% in the winter.
| |
ACKNOWLEDGMENT |
This work was financially supported by the Swedish Foundation for
Strategic Research through a national, industry-oriented program for
research and Ph.D. education, LiFT 
Future Technologies for Food Production.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Applied
Microbiology, Center for Chemistry and Chemical Engineering, Lund
Institute of Technology, Lund University, P.O. Box 124, SE-221 00 Lund, Sweden. Phone: 46 46-222 3412. Fax: 46 46-222 42 03. E-mail:
Peter.Radstrom{at}tmb.lth.se.
| |
REFERENCES |
| 1.
|
Abu Al-Soud, W.,
L. J. Jönsson, and P. Rådström.
2000.
Identification and characterization of immunoglobulin G in blood as a major inhibitor of diagnostic PCR.
J. Clin. Microbiol.
38:345-350[Abstract/Free Full Text].
|
| 2.
|
Abu Al-Soud, W., and P. Rådström.
1998.
Capacity of nine thermostable DNA polymerases to mediate DNA amplification in the presence of PCR-inhibiting samples.
Appl. Environ. Microbiol.
64:3748-3753[Abstract/Free Full Text].
|
| 3.
|
Abu Al-Soud, W., and P. Rådström.
2001.
Purification and characterization of PCR-inhibitory components in blood cells.
J. Clin. Microbiol.
39:485-493[Abstract/Free Full Text].
|
| 4.
|
Baranyi, J., and T. A. Roberts.
1994.
A dynamic approach to predicting bacterial growth in food.
Int. J. Food Microbiol.
23:277-294[CrossRef][Medline].
|
| 5.
|
Campbell, K. D.,
M. D. Collins, and A. K. East.
1993.
Gene probes for identification of the botulinal neurotoxin gene and specific identification of neurotoxin types B, E, and F.
J. Clin. Microbiol.
31:2255-2262[Abstract/Free Full Text].
|
| 6.
|
Cordoba, J. J.,
M. D. Collins, and A. K. East.
1995.
Studies on the genes encoding botulinum neurotoxin type A of Clostridium botulinum from variety of sources.
Syst. Appl. Microbiol.
18:13-22.
|
| 7.
|
Dezfulian, M.,
L. M. McCroskey,
C. L. Hatheway, and V. R. Dowell.
1981.
Selective medium for isolation of Clostridium botulinum from human feces.
J. Clin. Microbiol.
13:526-531[Abstract/Free Full Text].
|
| 8.
|
Eklund, M. W.,
F. T. Poysky, and D. I. Wieler.
1967.
Characteristics of Clostridium botulinum type F isolated from the Pacific coast of the United States.
Appl. Microbiol.
15:1316-1323[Medline].
|
| 9.
|
Eklund, M. W.,
D. I. Wieler, and F. T. Poysky.
1967.
Outgrowth and toxin production of nonproteolytic type B Clostridium botulinum at 3.3 to 5.6°C.
J. Bacteriol.
93:1461-1462[Free Full Text].
|
| 10.
|
Fach, P.,
M. Gibert,
R. Griffais,
J. P. Guillou, and M. R. Popoff.
1995.
PCR and gene probe identification of botulinum neurotoxin A-, B-, E-, F-, and G-producing Clostridium spp. and evaluation in food samples.
Appl. Environ. Microbiol.
61:389-392[Abstract].
|
| 11.
|
Fach, P.,
D. Hauser,
J. P. Guillou, and M. R. Popoff.
1993.
Polymerase chain reaction for the rapid identification of Clostridium botulinum type A strains and detection in food samples.
J. Appl. Bacteriol.
75:234-239[Medline].
|
| 12.
|
Franciosa, G.,
J. L. Ferreira, and C. L. Hatheway.
1994.
Detection of type A, B and E botulism neurotoxin genes in Clostridium botulinum and other Clostridium species by PCR: evidence of unexpressed type B toxin genes in type A toxigenic organisms.
J. Clin. Microbiol.
32:1911-1917[Abstract/Free Full Text].
|
| 13.
|
Franciosa, G.,
C. L. Hatheway, and P. Aureli.
1998.
The detection of a deletion in the type B neurotoxin gene of Clostridium botulinum A(B) strains by two-step PCR.
Lett. Appl. Microbiol.
26:442-446[CrossRef][Medline].
|
| 14.
|
Gaze, J. E., and K. L. Brown.
1988.
The heat resistance of spores of Clostridium botulinum 213B over the temperature range 120 to 140°C.
Int. J. Food Sci. Tech.
23:373-378.
|
| 15.
|
Hauschild, A. H. W., and K. L. Dodds.
1993.
Clostridium botulinum: ecology and control in foods, vol. 26.
Marcel Dekker, Inc, New York, N.Y.
|
| 16.
|
Hielm, S.,
J. Björkroth,
E. Hyytiä, and H. Korkeala.
1998.
Prevalence of Clostridium botulinum in Finnish trout farms: pulsed-field gel electrophoresis typing reveals extensive genetic diversity among type E isolates.
Appl. Environ. Microbiol.
64:4161-4167[Abstract/Free Full Text].
|
| 17.
|
Hielm, S.,
E. Hyytia,
J. Ridell, and H. Korkeala.
1996.
Detection of Clostridium botulinum in fish and environmental samples using polymerase chain reaction.
Int. J. Food Microbiol.
31:357-365[CrossRef][Medline].
|
| 18.
|
Hielm, S.,
E. Hyytiä,
A. B. Andersin, and H. Korkeala.
1998.
A high prevalence of Clostridium botulinum type E in Finnish freshwater and Baltic Sea sediment samples.
J. Appl. Microbiol.
84:133-137.
|
| 19.
|
Huss, H. H.
1980.
Distribution of Clostridium botulinum.
Appl. Environ. Microbiol.
39:764-769[Abstract/Free Full Text].
|
| 20.
|
Huss, H. H.,
A. Pedersen, and D. C. Cann.
1974.
The incidence of Clostridium botulinum in Danish trout farms. I. Distribution in fish and their environment.
J. Food Technol.
9:445-450.
|
| 21.
|
Hutson, R. A.,
Y. Zhou,
M. D. Collins,
E. A. Johnson,
C. L. Hatheway, and H. Sugiyama.
1996.
Genetic characterization of Clostridium botulinum type A containing silent type B neurotoxin gene sequences.
J. Biol. Chem.
271:10786-10792[Abstract/Free Full Text].
|
| 22.
|
Johannsen, A.
1963.
Clostridium botulinum in Sweden and the adjacent waters.
J. Appl. Bacteriol.
26:43-47.
|
| 23.
|
Kautter, D. A.,
H. M. Solomon, and E. J. Rhodehamel.
1992.
Clostridium botulinum, p. 215-225.
In
FDA bacteriological analytical manual, 7th ed. AOAC International, Arlington, Va.
|
| 24.
|
Lalitha, K. V., and K. Gopakumar.
2000.
Distribution and ecology of Clostridium botulinum in fish and aquatic environments of a tropical region.
Food Microbiol.
17:535-541[CrossRef].
|
| 25.
|
Lantz, P.,
W. Abu Al-Soud,
R. Knutsson,
B. Hahn-Hägerdal, and P. Rådström.
2000.
Biotechnical use of polymerase chain reaction for microbiological analysis of biological samples.
Biotechnol. Annu. Rev.
5:87-130[CrossRef][Medline].
|
| 26.
|
Lantz, P.,
R. Knutsson,
Y. Blixt,
W. A. Al-Soud,
E. Borch, and P. Rådström.
1998.
Detection of pathogenic Yersinia enterocolitica in enrichment media and pork by a multiplex PCR: a study of sample preparation and PCR-inhibitory components.
Int. J. Food Microbiol.
45:93-105[CrossRef][Medline].
|
| 27.
|
Lindstrom, M.,
H. M. Jonkola,
S. Hielm,
E. Hyytia, and H. Korkeala.
1999.
Identification of Clostridium botulinum with API 20A, Rapid ID 32A and RapID ANA II.
FEMS Immunol. Med. Microbiol.
24:267-274[Medline].
|
| 28.
|
Melin, L.,
M. Jensen-Waern,
A. Johannisson,
M. Ederoth,
M. Katouli, and P. Wallgren.
1997.
Development of selected faecal microfloras and of phagocytic and killing capacity of neutrophils in young pigs.
Vet. Microbiol.
54:287-300[CrossRef][Medline].
|
| 29.
|
Peck, M. W.
1997.
Clostridium botulinum and the safety of refrigerated processed foods of extended durability.
Trends Food Sci. Technol.
8:186-192[CrossRef].
|
| 30.
|
Peck, M. W.,
D. A. Fairbairn, and B. M. Lund.
1992.
The effect of recovery medium on the estimated heat-inactivation of spores of non-proteolytic Clostridium botulinum.
Lett. Appl. Microbiol.
15:146-151.
|
| 31.
|
Sambrook, J.,
E. F. Fritsch, and T. Maniatis.
1989.
Molecular cloning: a laboratory manual, 2nd ed.
Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
|
| 32.
|
Schmidt, C. F.,
R. V. Lechowich, and J. F. Folinazzo.
1961.
Growth and toxin production by type E Clostridium botulinum below 40°F.
J. Food Sci.
26:626-630[CrossRef].
|
| 33.
|
Schocken-Iturrino, R. P.,
M. C. Carneiro,
E. Kato,
J. O. B. Sorbara,
O. D. Rossi, and L. E. R. Gerbasi.
1999.
Study of the presence of the spores of Clostridium botulinum in honey in Brazil.
FEMS Immunol. Med. Microbiol.
24:379-382[CrossRef][Medline].
|
| 34.
|
Szabo, E. A.,
J. M. Pemberton, and P. M. Desmarchelier.
1993.
Detection of the genes encoding botulinum neurotoxin types A to E by the polymerase chain reaction.
Appl. Environ. Microbiol.
59:3011-3020[Abstract/Free Full Text].
|
| 35.
|
Thompson, J. D.,
T. J. Gibson,
F. Plewniak,
F. Jeanmougin, and D. G. Higgins.
1997.
The CLUSTAL X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools.
Nucleic Acids Res.
25:4876-4882[Abstract/Free Full Text].
|
| 36.
|
Williamson, J. L.,
T. E. Rocke, and J. M. Aiken.
1999.
In situ detection of the Clostridium botulinum type C1 toxin gene in wetland sediments with a nested PCR assay.
Appl. Environ. Microbiol.
65:3240-3243[Abstract/Free Full Text].
|
Applied and Environmental Microbiology, October 2001, p. 4781-4788, Vol. 67, No. 10
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4781-4788.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
Radstrom, P., Lofstrom, C., Lovenklev, M., Knutsson, R., Wolffs, P.
(2008). Strategies for Overcoming PCR Inhibition. CSH Protocols
2008: pdb.top20-pdb.top20
[Abstract]
[Full Text]
-
Artin, I., Bjorkman, P., Cronqvist, J., Radstrom, P., Holst, E.
(2007). First Case of Type E Wound Botulism Diagnosed Using Real-Time PCR. J. Clin. Microbiol.
45: 3589-3594
[Abstract]
[Full Text]
-
Lindstrom, M., Korkeala, H.
(2006). Laboratory Diagnostics of Botulism. Clin. Microbiol. Rev.
19: 298-314
[Abstract]
[Full Text]
-
Gessler, F., Hampe, K., Bohnel, H.
(2005). Sensitive Detection of Botulinum Neurotoxin Types C and D with an Immunoaffinity Chromatographic Column Test. Appl. Environ. Microbiol.
71: 7897-7903
[Abstract]
[Full Text]
-
Lovenklev, M., Holst, E., Borch, E., Radstrom, P.
(2004). Relative Neurotoxin Gene Expression in Clostridium botulinum Type B, Determined Using Quantitative Reverse Transcription-PCR. Appl. Environ. Microbiol.
70: 2919-2927
[Abstract]
[Full Text]
-
Lovenklev, M., Artin, I., Hagberg, O., Borch, E., Holst, E., Radstrom, P.
(2004). Quantitative Interaction Effects of Carbon Dioxide, Sodium Chloride, and Sodium Nitrite on Neurotoxin Gene Expression in Nonproteolytic Clostridium botulinum Type B. Appl. Environ. Microbiol.
70: 2928-2934
[Abstract]
[Full Text]
-
Lofstrom, C., Knutsson, R., Axelsson, C. E., Radstrom, P.
(2004). Rapid and Specific Detection of Salmonella spp. in Animal Feed Samples by PCR after Culture Enrichment. Appl. Environ. Microbiol.
70: 69-75
[Abstract]
[Full Text]
Top
Abstract
Introduction
