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Clostridium difficile in neonatal piglets in three Dutch
farms.
Onderzoeksstage by: drs. Famke E. Reeuwijk
In the period: September 2007 – December 2007
supervisors:
and
prof. dr. Aldert A. Bergwerff,
Institute for Risk Assessment Sciences
Veterinary Public Health division
Utrecht University
Utrecht, The Netherlands
dr. Leo A.M.G. van Leengoed
Faculty of Veterinary Medicine
Department of Farm Animal Health
Internal Medicine division
Utrecht University
Utrecht, The Netherlands
1
Index
page
Summary
3
Introduction
3
Material and Methods
4
Results
7
From Hospital to Farm
8
Discussion
10
Literature
13
2
Summary
Clostridium difficile is a known cause of diarrhea in neonatal piglets. The objective of
this study was twofold. The first objective was to determine the sensitivity of the
diagnostic procedures used. At three sow herds fecal samples of neonatal piglets
suspected from Clostridium difficile- associated diarrhea, and their dams, were tested for
both A and B toxins produced by Clostridium difficile. In addition, samples were cultured
to isolate Clostridium difficile. Our diagnostic procedures and/or sample handling
resulted in probable false negative results. This suspicion arose because in many piglets
which showed the typical clinical signs, the bacteria could not be found. Further research
is needed to increase sensitivity and of tests used, both isolation of Clostridium difficile
and ELISA, for diagnosis of Clostridium difficile in neonatal piglets and to evaluate toxin
stability in piglets’ feces.
The second objective was to translate hospital protocols to the actual farm situation. How
should farmers act when confronted with a Clostridium difficile in their pigs.
Under present farm conditions it is difficult to eradicate or minimize Clostridium difficile
related disease and further research is needed to design farm procedures.
Introduction
Clostridium difficile is a Gram positive, sporeforming, anaerobic bacillus that causes
Clostridium difficile associated diarrhea (CDAD), antibiotic-associated diarrhea in
hospitalized patients or pseudomembranous colitis (PMC) in humans (1, 12, 17, 18, 21).
This bacterial species also causes enteritis in horses and foals, calves, ostriches,
elephants, dogs and neonatal piglets (1, 6, 4, 12, 17, 18, 22, 23, 24, 26, 27, 28, 29).
The first report of PMC dated from 1893, when Finney described pseudomembranes
found in the gut of a deceased woman (9, 11).
In 1935 Bacillus difficile, as it was named then, was found in stools of healthy newborn
babies by Hall and O’Toole (11). As these babies were healthy, the bacillus was believed
to be nonpathogenic (11).
In the 1960s, anaerobic bacteria were identified as important pathogens, and new research
on Clostridium difficile started (11).
Clostridium difficile was found to produce at least two toxins: toxin A and toxin B, which
are responsible for the damage to the epithelium in the gut (9, 11). For a long time it was
believed that toxin B could only be active if toxin A was present (9, 11), but recently
pathogenic subtypes have been found that were toxin A negative and toxin B positive.
This supports the hypothesis that toxin B is a potent toxin on its own (8, 15, 20, 24).
Nearly always is PMC, an antibiotic-associated diarrhea in hospitals (25% of the cases in
the US), caused by Clostridium difficile (2, 15).
Due to antibiotic use, the incidence of CDAD has increased dramatically, and recently
various hospitals in many countries faced an outbreak by very virulent strains (5, 9, 10,
11, 15, 19, 25). Especially, use of fluoroquinolones and cephalosporines increase the risk
for CDAD in humans (14).
3
In 1983 the bacillus was found for the first time in pigs (29). Human pathogen ribotypes
017 and 027 have been found in pigs, and opens the question whether these ribotypes are
present in the Dutch pig population. When present, does Clostridium difficile in pigs
cause/induce a public health hazard?
In pigs, most Clostridia-related enteritis is caused by Clostridium perfringens type A and
C. Enteritis due to Clostridium difficile is seldomly diagnosed in The Netherlands, as
routine procedures do not include Clostridium difficile isolation, or detection of its toxins
(22, 29). As Clostridium difficile is also found in healthy piglets, the diagnostic value of a
single isolation of this bacterium is questionable (22).
Clostridium difficile associated diarrhea (CDAD) may develop in neonatal piglets. The
neonatal diarrhea, that starts on 1 to 7 days of age, is characterized by a low mortality,
high morbidity, and limited reaction on treatment. The diarrhea may vary in aspect from
pasty to watery and is yellow to orange of color (22, 23). Recently, it was found that
Clostridium difficile also produce toxins in healthy piglets (30).
Because there might be an exchange between human and animal strains, it is possible that
animals might act as a reservoir. For example, in calves in Canada human pathogen
ribotypes O27 and O17 were found. Transmission between humans and animals might
occur through direct contact or through the food chain, i.e. in Canada Clostridium difficile
was found in retail ground meat (18, 19, 21).
We are unaware how many farms are infected with Clostridium difficile, and whether
there is a connection with use of antibiotics on these farms and if transmission is possible
between piglets and humans.
With respect to control, basic knowledge is lacking how piglets become infected, by sows
that carry the bacteria, or by direct contact with a contaminated farrowing pen.
In addition, other questions come to mind as well, i.e. how should farmers deal with this
infection? In hospitals a strict protocol is to be used when a patient is suspected to have
or diagnosed with Clostridium difficile.
The purpose of this study was twofold. Firstly, sows and piglets on farms suspected of a
Clostridium difficile infection were tested to determine the sensitivity of the toxin-assay
used. Secondly, a protocol used in hospitals to contain Clostridium difficile infections
was studied to reflect and transform described measures for control of Clostridium
difficile outbreaks in a swine farm situation. .
Material and Methods
Sampling:
Three farms were visited. In Farm 1 (W from Y), 22 piglets from 4 sows (5 – 7 piglets
per litter) were sampled from a herd of 240 sows. The average number of live birth
piglets was 12.2 per sow, and the average number of weaned piglets was 10.7.
In Farm 2, 41 piglets from 7 sows (5 piglets per litter) were sampled from a herd of 520
sows. The average number of live born piglets was 13.0, of the average number of
4
weaned piglets was 11.5. In this farm, 2 litters (12 piglets) were sampled with a fecal
swab that did not show diarrhea.
In Farm 3, 21 piglets from 4 sows (5 per litter) were sampled in a herd of 700 sows. The
average number of live born piglets was 12.8, of the average number of weaned piglets
was 11.1. Samples were taken from neonatal piglets (1 to 4 days old) with diarrhea
(varying from pasty, slimy to watery).
Prior to sampling, piglets were squeezed in the abdomen, and feces was then collected in
10 ml tubes. When piglets did not produce feces easily, samples were obtained directly
from the rectum with a fecal swab.
In Farm 1 and Farm 2, 8 sows in total were sampled. The sample was obtained with a
glove directly from the rectum. Over all 96 piglets and 8 sows were sampled, 84 piglets
showed diarrhea, 12 did not show diarrhea, and none of the sows showed diarrhea
Samples were stored during transportation in a cool box and immediately put in the
refrigerator after arrival at the laboratory. Samples were processed within 48 hours, if the
time between sampling and processing was expected to be longer, samples were frozen
after arrival at the laboratory.
Toxin test:
Samples were tested with ImmunoCard Toxins A&B®, a commercially available single
EIA test to detect toxins A and/or B. The test could only be performed on samples that
contained at least 2 grams feces and not on samples taken with a fecal swab.The
manufacturer’s instruction was followed. Fecal samples were stored also at 2-8 ºC and
tested at room temperature. When all reagents, samples and the immunocard were at
room temperature, a suspension was made with 200 µl Specimen Diluent, 3 drops of
Enzyme Conjugate and 25 µl of sample. If the sample was not fluid, a tiny bit of sample
was put in the mixture with a wooden applicator stick. This mixture was vortexed and
then incubated for 5 min. When toxins were present in the sample, they were bound to
the toxin antibodies in the Enzyme Conjugate. After incubation, the mixture was
vortexed again and 125 µl was put in both sample ports of the immunocard (see Figure
1).
Figure 1: The Immunocard
5
After 5 min incubation, both reaction ports should be completely wet. During this time,
the toxin-conjugate complex is separated from al the particulates still present in the
sample and only the fluid flows through the membrane in the ‘test’ and ‘control’ reaction
ports.
Within the ‘test’ reaction port, antitoxin immobilizes the toxin-conjugate complex, the
‘control’ reaction port is an internal control to check whether the right procedure is
followed and the Immunocard is still active.
After the second incubation, both reaction ports are washed with 3 drops Wash Reagent.
This reagent washes away contaminating proteins which may interfere with the Substrate
Reagent. When the Wash Reagent was absorbed, the Substrate Reagent (3 drops) was put
in the reaction ports, and after 5 min the results on the immunocard was read.
The ‘control’ port should turn blue, showing the immunocard and reagents were active
and the right amount of sample has flown in the reaction ports.
The ‘test’ port can turn blue or stay white: blue indicated toxins were present in the
sample, white indicated toxins were not present in the sample.
When toxin test was found positive, the spare sample diluent was further tested on
presence of Clostridium difficile bacteria by culturing.
Ethanol Shock
All samples were treated with the ethanol-shock method before culturing, because this is
considered to be a good method to isolate Clostridium difficile spores (13). An amount of
5 g of sample was mixed with 5 ml of 96% ethanol. The mixture was incubated for
60 min at room temperature. Thereafter, the mixture was inoculated on Columbia ANC
agar + 5% sheep blood plates (CNA, Biomerieux) and Clostridium difficile agar plates
(CLO, Biomerieux). Plates were incubated at 37°C for 48 h in an anaerobic environment,
which was created within a closed container (Oxoid HP) by adding a Gaspack (BD) that
binds the oxygen present in the container. Clostridium difficile colonies were recognized
by their specific smell, like horse dung, and were non-hemolytic and appeared to be
white, grey to green.
Non-hemolytic colonies grown on CNA plates and all colonies grown on CLO plates
were incubated anaerobically for another 48 hours at 37°C on Schadler agar + 5% sheep
blood (SCS, Biomerieux). Colonies grown on SCS plates were Gram-stained and
examined microscopically. Finally, suspected colonies from ICTAB-positive piglets were
picked, cultured, and sent to LUMC Hospital (Clostridium Refernce Laboratory, Leiden,
The Netherlands) for confirmation through PCR ribotyping. PCR ribotyping is
considered a quick and reliable method to identify and classify Clostridium difficile (3,
7).
6
Results
When piglets were sampled on all 3 farms, there were no toxins found in the toxin test
and no Clostridium difficile could be determined after culturing. Colonies were typed
through smell, shape, color and Gram staining.
The control-strains grew like expected , the Clostridium difficile colonies have a specific
smell, like horse dung, are non-hemolytic and appear to be white, grey to green, and
Gram staining of these colonies had results as expected, they appear as little pink tennis
rackets under the microscope.
It was decided to sample Farm 1 and Farm 2 another time, because the suspicion arose
that the ICTAB and culture results were false negative. In previous internships (by drs. E.
Willems and drs. A. Klooster) both farms were already confirmed to be positive for
Clostridium difficile.
The farms were especially expected to have neonatal infections of Clostridium difficile
as on both farms piglets with neonatal diarrhea had colitis, which is characteristic for
CDAD. All other known causes for neonatal diarrhea cause enteritis in the small
intestine. Furthermore the clinical signs on all farms were also an indication that
Clostridium difficile was the cause of the problems, namely the neonatal diarrhea that
starts on 1 to 2 days of age, low mortality, high morbidity, and limited effect of treatment.
These last samples will be used to evaluate the tests used.
On Farm 1, three sows were sampled and in total 15 of their piglets were sampled (5 per
litter). All sows tested negative on toxins and culture. Of the 15 piglets, 10 (67%) tested
positive on toxins and 5 negative (33%). After culture and Gram-staining, 23 SCS plates
(11 transferred from CNA, 12 transferred from CLO plates), with samples from 11
piglets had colonies that were suspicious to be Clostridium difficile. There were 9 piglets
that tested both positive by ELISA and by culturing, 2 piglets tested negative by ELISA
and positive by culturing, 1 piglet tested positive by EIA and negative by culturing and 3
piglets tested both negative by EIA and by culturing.
After confirmation by ribotyping of the 9 piglets that tested both positive by EIA and by
culturing, 7 samples were confirmed Clostridium difficile ribotype O78-positive, while in
2 last samples no Clostridium difficile was found.
On Farm 2, five sows were sampled and in total 31 of their piglets were sampled (6 per
litter) All sows tested negative on toxins and culture. Because fecal extraction from these
piglets was hampered after squeezing the abdomen, it was difficult to obtain sufficient
material for ICTAB testing, so it was decided to take swabs. Out of the 5 litters sampled,
2 litters didn’t show clinical signs. Out of the 30 piglets sampled, the EIA toxin test
could only be performed on 6 piglets, because of the limited material obtained and the
swabs used on the rest of the piglets.
Of the 6 piglets, 2 (33%) tested positive on toxins and 4 negative (67%).
Because a new culture medium became available during this study, not all the swabs
were used for culture, but of every litter three swabs (15 piglets in total) were stored in
the freezer, so results can be compared in continuing studies.
7
Out of the 16 samples used in this study, after culturing and Gram staining, 4 piglets
(25%) were suspected positive of Clostridium difficile. One piglet tested both positive by
ELISA and by culturing, 4 piglets tested negative by ELISA and positive by culturing, 1
piglet tested positive by ELISA and negative by culturing and 10 piglets tested both
negative by ELISA and culturing.
After confirmation by ribotyping of the suspected colonies, in a single colony ribotype
O78 was found, in 5 colonies (from 2 piglets) Clostridium difficile was found and in 2
colonies no Clostridium difficile was found. The combined results of Farm 1 and Farm 2
are shown in Table 1.
Table 1: Combined C. difficile results of Farms 1 and 2. All eight sampled sows were
negative. n Farm 2, the positive piglets were in two litters, one of the litters showed
diarrhea, the other litter did not show diarrhea.
Culture positive
Culture Negative
Total piglets
ICTAB positive
10
6
16
ICTAB negative
2
13
15
Total piglets
12
19
31
From Hospital to Farm
In hospitals a strict protocol should be followed when a patient is suspected of or
confirmed with Clostridium difficile-associated diarrhea (CDAD). Prevention is directed
on one hand on hygiene, on the other hand it is directed on restricted use of antibiotics
(Workgroup Infection Prevention, 2006; the 36th Outbreak Management team, 2005).
The hygiene measures imply:
 Isolating patient to a one-person room with bathroom. The patient is not allowed
to use the common bathrooms.
 Nursing personnel should wear non-sterile gloves and protective apron that are to
be removed when leaving the chamber. When leaving, hands should be washed
with water and soap and wiped off thoroughly. Because the spores of Clostridium
difficile are resistant to alcohol, the use of hand-alcohol is not useful to prevent
spread of spores (it can help to prevent spread of other bacteria).
 The patient should be instructed about the appropriate toilet hygiene (washing
hands and flush the toilet with the lid on)
 The room where the patient was before isolation and the isolation room should be
cleaned and disinfected, though it is important to realize that the spores of
Clostridium difficile are resistant for the commonly used disinfectants. It is
momentarily not workable to kill the spores with surface disinfectants; therefore
the advice is to keep the surroundings of the patient as clean as possible by daily
cleaning and disinfecting, so at least the vegetative form of the bacteria is taken
care off.
 Instruments should be used patient-bound and cleaned and disinfected thoroughly
after use.
8
The antibiotic restrictions imply:
 Preferred therapy for the patient is 4 times daily 500 mg of vancomycine orally
during 14 days.
 Other antibiotic treatment of the patient should be stopped immediately (if
possible).
 In the institution the use of fluoroquinolones should be restricted to the minimum.
 The use of clindamycine, macrolides and cefalosporines (especially the third
generation) are debatable, so a restriction on these antibiotics in the hospital or
nursinghome could be useful.
On the farm:
To make a useful and clear protocol for farms where CDAD is a problem, the hospital
situation can be extrapolated to the farm situation. Because there is a difference between
a large hospital and a local nursing home, a difference should also be made between an
intensive rearing farm and an extensive rearing farm.
The guidelines used in hospitals, can not be used just like that on any pig farm. On the
basis of the protocols used in hospitals (see above) the measures that can be taken on the
farm are:
 Isolation
When neonatal piglets develop diarrhea, these piglets and their mother have to be
isolated. This can be achieved by putting them in another stable although this is not
feasible in most farms. Alternatively, the whole section can be considered as
contaminated. In the field situation, this will lead to the treatment of the whole farm as
being contaminated.
 Personnel
The infected section has to be entered in section-bound clothes and boots, to prevent the
spread of spores to other sections. A bucket with disinfectants does not kill the spores on
the boots and is therefore not effective. Personnel has to handle the piglets on several
occasions (castration, ear marking, iron injection) which can incorporate a risk of
infection for the person handling the piglets. Because of the nature of contact between
farmer and animal, it is impossible to prevent exposure of the farmer or personnel.
Clostridium difficile can cause disease in humans, but in healthy individuals this is not
very likely. It is important that farmers and personnel are informed about the bacteria, so
that they are aware of the possibility of infection when a farmer has to get antibiotic
treatment himself.
 Surroundings
In the hospital the isolation room is cleaned and disinfected daily to kill the vegetative
bacteria and reduce the risk of infection. In a pig farm, it is not manageable to clean and
disinfect farrowing crates daily. What can be done, is to perform a strict all-in all-out
system within the farrowing crates. If it is possible, the farrowing crates can be cleaned
thoroughly, and it is important to remove the overall dust and fat layers, because spores
can accumulate in them. Then, the crates can be disinfected to reduce the number of
9
vegetative bacteria. It is not possible to get the crates free of spores with cleansers and
disinfectants, because they only work with contact times of 6 hours or longer, and that is
not achievable. The risk of infection can only be reduced by removing as much dust and
fat as possible. This is intensive work and the effect gained by this has to be evaluated.
 Means
It is recommended that material and equipment used within the infected areas remain
within the infected area (e.g. vaccination syringes, shovels, warmth lamps etc.).
On large farms these measures are easier to imply than on small farms. On large farms a
single area can be closed off of the rest of the farm, and then one staff member can be
responsible for this area. By doing this the risk of spreading spores all over the farm can
be reduced. The question remains how the spread of the bacteria within the farm occur. If
the sows carry the bacteria with them and spread the spores where they go, little effect is
to be expected of closing off a single area of the farm, because the sows will have to
move around the farm, thus taking the bacteria with them.
 Antibiotic use
The farmer can treat the piglets with antibiotics according to the formularium of the farm
that was made in consultation with his veterinarian. Preferably piglets are treated after a
culture and antibiogram is made with an antibiotic that has a small working spectrum and
that is working well against Clostridium difficile.
An example of treatment could be orally given amoxicilin, but effectiveness should be
evaluated. The use of antibiotics on the entire farm can be reconsidered, and perhaps the
use can be reduced.
The relationship between the incidence of CDAD and the antibiotic use on farms is not
yet clear, but when extrapolated to the human situation, a reduction of antibiotic use to a
minimum is a condicio sine qua non. It is important that the veterinarian decides which
antibiotics at what dose is used on a pig farm, and that the farmer looses his degrees of
freedom in this.
In conclusion, it is most difficult to get free of Clostridium difficile if not impossible, but
infection risk can be lessened by thorough cleaning, careful hygiene and use of material
and clothing and perhaps by reducing the amounts of administered antibiotics.
Discussion
The first four times samples were taken, Farms 1, 2 and 3 were all found negative on
toxins and on Clostridium difficile culturing. Toxin tests were negative possibly due to
denaturalization or proteinolysis of the toxins. It must be noted here that opinions differ
on whether samples should be frozen after collecting (26, 29).
The EIA used has a specificity of 98,9% and a sensitivity of 93,1% in stool samples
(manufacturers guide). The test is developed for use in human stool samples, and is not
validated for use in pigs.
In human stools, toxins remain detectable at least 24 hours, but for this has never been
confirmed for piglet feces. Testing of the toxin stability in piglet feces will give more
information on how the samples should be treated after collection. Furthermore, there is
10
the question of what the relation is between the detection of toxins and the clinical signs
in piglets. Research on the levels of toxins in feces from piglets with and without clinical
signs could provide an answer to this.
It is remarkable, that the two farms that were visited by the author, gave positive results,
while samples that were taken by others, gave negative results. Perhaps the transport and
storage of the samples by the others affected the quality of the samples.
Here, positive samples were tested within 5 h after sampling. Negative samples were
always re-tested after 24 h. Again testing of the toxin stability might be the solution here.
This can be done by testing positive samples on different times and monitor how long a
sample remains positive.
It is strange though, that in the cultures on the farms visited by others no Clostridium
difficile was cultured, because the spores of Clostridium difficile are extremely resistant.
It would also be interesting to know how many spores or bacteria are present in the
piglets’ diarrhea, to get to know something about the chance of actually putting spores on
the plates. The sensitivity of the method used should be evaluated.
Too little farms have been tested to draw general conclusions from the results. It can be
concluded that Clostridium difficile O78 is present in piglets in the Netherlands, but many
questions remain. The prevalence in the Netherlands is unknown and what the
consequences are for public health, the farmer and other farms. Though veterinarians in
several practices were approached with the request to collect samples when they
encountered neonatal piglets with diarrhea, little response and no samples came to the
laboratory. One veterinary practice responded with the message that there were no piglets
with suspected diarrhea at the moment. Further reasons for the little response remain
unclear. Perhaps Clostridium difficile associated diarrhea has a low prevalence in piglets
in the Netherlands, but no data are available on this subject.Furthermore the disease
causes little economic losses (morbidity and mortality are low), so priorities lie with other
more urgent problems.
On the two farms that were already confirmed to have a problem with CDAD positive
toxin tests were found. The 67% positive result on the first farm was as expected from
earlier testing on the same farm (reports drs. E. Willems and drs. A. Klooster). The 33%
positive results on the second farm was lower as expected, but too little piglets were
tested to draw conclusions about sensitivity and specificity of the used tests.
When a litter of piglets shows diarrhea, it is expected that Clostridium difficile is present
within every piglet. The results of the ICTAB and the culturing show that the agent is
only found in 73% of the sampled piglets on Farm 1, and in 31% of the sampled piglets
on Farm 2.
Especially the 31% is a low score, a new culture medium might be the solution to get a
higher positive percentage. Further research has to explore this option.
A rapid, reliable screening test is needed to know if Clostridium difficile is present on the
farm. To diagnose the disease in the piglets, dissection is necessary to prove causality
between the presence of the bacteria on the farm and the diarrhea within the piglets. Only
then a certain diagnose can be made.
11
For the farmers the biggest problem is of course how to deal with an infection of
Clostridium difficile.
To control the infection on the farm several elements are necessary. It is of vital
importance to know all risk factors that influence the survival, multiplication and spread
of the bacteria. The most important question here is: how does Clostridium difficile
maintain itself within the farm? Are the sows carrier of the bacteria and do they infect the
piglets during or short after birth? Or is the surrounding of the piglets the only route of
infection? These questions still remain unanswered.
Further more there is the question whether Clostridium difficile is easily transmitted from
the pigs to the farmer/caretaker. How can he be protected and what can he do to prevent
infection of himself and his newborn piglets?
In hospitals a strict protocol is followed when a patient is diagnosed with Clostridium
difficile. Following an interpretation of these protocols to implement measures in the
practical situation farmers are in, the conclusion must be made that it is quite impossible
to get free of Clostridium difficile. Even if the crates can be thoroughly cleaned and
disinfected, the chance that spores survive seems substantial, and when the newborn
piglets take in the spores, and develop the disease, huge amounts of spores are produced
and the whole section is contaminated as bad as it was before. The most important step to
take into the controlling of the infection is to detect the source of the infection. How do
the piglets get infected? Only when the source can be revealed and eliminated or
controlled, protocols and measures for controlling the infection can be successful and
have effect.
It seems impossible to get rid of the infection on a farm, but perhaps with good
management the disease will be manageable in the future. To reach this, the farmer has to
be extremely motivated and further research is necessary for new methods to get the
infection risk between animals and between farmer and animal as low as possible on the
farm.
Perhaps if antibiotic use is evaluated and the use can be reduced to a minimum other (non
pathogenic) bacteria can take in the niche Clostridium difficile is filling now.
An idea for getting rid of more spores than can be achieved by cleaning and disinfecting,
is perhaps by making the spores sporulate and then kill them off with disinfectants
(because vegetative bacteria can be killed with disinfectants). When the farrowing crates
are empty, a high temperature and moist climate can be created by spraying the crates
with water and putting the heaters on. It has to be evaluated when most of the spores
sporulate and at the optimal time the disinfectant can be sprayed within the crates so the
vegetative bacteria will be killed.
12
Literature
1. Arroyo LG, Weese JS, Stämpfli HR (2004) Experimental Clostridium difficile
enterocolitis in foals. Journal of Veterinary Internal Medicine 18: 734-738
2. Asha NJ, Tompkins D, Wilcox MH (2006) Comparative analysis of prevalence,
risk factors and molecular epidemiology of antibiotic-associated diarrhea due to
Clostridium difficile, Clostridium perfringens and Staphylococcus aureus. Journal
of Clinical Microbiology 44-8: 2785-2791
3. Belanger SD, Boissinot M, Clairoux N, Picard FJ, Bergeroni MG (2003) Rapid
detection of Clostridium difficile in feces by real-time PCR. Journal of Clinical
Microbiology 41-2: 730-734
4. Bojesen AM, Olsen KEP, Bertelsen MF (2006) Fatal enterocolitis in Asian
elephants (Elephas maximus) caused by Clostridium difficile. Veterinary
Microbiology 116: 329-335
5. Delaney JAC, Dial S, Barkun A, Suissa S (2007) Antimicrobial drugs and
community acquired Clostridium difficile-assosiated disease, UK. Emerging
infectious diseases 13-5: 761-763
6. Frazier KS, Herron AJ, Hines ME, Gaskin JM, Altman NH (1993) Diagnosis of
enteritis and enterotoxemia due to Clostridium difficile in captive ostriches
(Struthio camelus). Journal of Veterinary Diagnostic Investigation 5: 623-625
7. Kato N, Ou CY, Kato H, Bartley SL, Brown VK, Dowell jr VR, Ueno K (1991)
Identification of toxigenic Clostridium difficile by the Polymerase Chain
Reaction. Journal of Clinical Microbiology 29-1: 33-37
8. Kato H, Kato N, Watanabe K, Iwai N, Nakamura H, Yamamoto T, Suzuki K,
Shin-Moo K, Chong Y, Wasito EB (1998) Identification of toxin A-negative,
toxin B-positive Clostridium difficile by PCR. Journal of Clinical Microbiology
36-8: 2178-2182
9. Knoop FC, Owens M, Crocker IC (1993) Clostridium difficile: Clinical disease
and diagnosis. Clinical Microbiology Reviews 6-3: 251-265
10. Kuijper EJ, van den Berg RJ, Debast S, Visser CE, Veenendaal D, Troelstra A,
van der Kooi T, van den Hof S, Notermans DW (2006) Clostridium difficile
ribotype 027, toxinotype III, the Netherlands. Emerging Infectious Diseases 12-5:
827-830
11. Lyerly DM, Krivan HC, Wilkins TD (1988) Clostridium difficile: Its disease and
toxins. Clinical Microbiology Reviews 1-1: 1-18
12. Marks JL, Kather EJ (2003) Antimicrobial susceptibilities of canine Clostridium
difficile and Clostridium perfringens isolates to commonly utilized antimicrobial
drugs. Veterinary Microbiology 94: 39–45
13. Marler LM, Siders JA, Wolters LC, Pettigrew Y (1992) Comparison of five
cultural procedures for isolation of Clostridium difficile from stools. Journal of
Clinical Microbiology 30-2: 514-516
14. McCusker ME, Harris AD, Perencevich E, Roghmann MC (2003)
Fluoroquinolone use and Clostridium difficile-associated diarrhea. Emerging
Infectious Diseases 9-6: 730-735
13
15. Moncrief JS, Zheng L, Neville LM, Lyerly DM (2000) Genetic Characterization
of toxin A-negative, toxin B-positive Clostridium difficile isolates by PCR.
Journal of Clinical Microbiology 38-8 3072-3075
16. Post KW, Jost BH, Songer JG (2002) Evaluation of a test for Clostridium difficile
toxins A and B for the diagnosis of neonatal swine enteritis. Journal of Veterinary
Diagnostic Investigation 14: 258-259
17. Post KW, Songer JG (2004) Antimicrobial susceptibility of Clostridium difficile
isolated from neonatal pigs with enteritis. Anaerobe 10: 47-50
18. Rodriguez-Palacios A, Stämpfli HR, Duffield T, Peregrine AS, Trotz-Williams
LA, Arroyo LG, Brazier JS & Weese JS (2006) Clostridium difficile PCR
ribotypes in Calves, Canada. Emerging infectious diseases 12-11: 1730-1736
19. Rodriguez-Palacios A, Staempfli HR, Duffield T, Weese JS (2007) Clostridium
difficile in retail ground meat, Canada. Emerging Infectious Diseases 13-3: 485487
20. Rupnik M (2001) How to detect Clostridium difficile variant strains in a routine
laboratory. Clinical Microbiology and Infectious Diseases 7: 417-420
21. Rupnik M (2007) Is Clostridium difficile-associated infection a potentially
zoonotic and foodborne disease? Clincial Microbiology and Infectious Diseases
13: 457-459
22. Songer JG, Uzal FA (2005) Clostridial enteric infections in pigs. Journal of
Veterinary Diagnostic Investigation 17: 528-536
23. Songer JG, Anderson MA (2006) Clostridium difficile: An important pathogen of
food animals. Anaerobe 12: 1-4
24. Taha S, Johansson O, Jonsson SR, Heimen D, Krovacek K (2007) Toxin
production by and adhesive properties of Clostridium difficile isolated from
humans and horses with antibiotic-associated diarrhea. Comparative Immunology,
Microbiology & Infectious Diseases 30: 163-174
25. Vonberg RP, Schwab F, Gastmeier P (2007) Clostridium difficile in discharged
inpatients, Germany. Emerging Infectious Diseases 13-1: 179-180
26. Waters EH, Orr JP, Clark EG, Schaufele CM (1998) Typhlocolitis caused by
Clostridium difficile in suckling piglets. Journal of Veterinary Diagnostic
Investigation 10: 104-108
27. Weese JS, Stämpfli HR, Prescott JF, Kruth SA, Greenwood SJ, Weese HE (2001)
The roles of Clostridium difficile and Enterotoxigenic Clostridium perfringens in
diarrhea in dogs. Journal of Veterinary Internal Medicine 15: 374-378
28. Weese JS, Armstrong J (2003) Outbreak of Clostridium difficile-associated
disease in a small animal veterinary teaching hospital. Jounal of Veterinary
Internal Medicine 17: 813-816
29. Yaeger M, Funk N, Hoffman L (2002) A survey of agents associated with
neonatal diarrhea in Iowa swine including Clostridium difficile and porcine
reproductive syndrome virus. Journal of Veterinary Diagnostic Investigation 14:
281-287
30. Yaeger MJ, Kinyon JM, Songer JG (2007) A prospective, case control study
evaluating the association between Clostridium difficile toxins in the colon of
neonatal swine and gross and microscopic lesions. Journal of Veterinary
Diagnostic Investigation 19: 52-59
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