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Erysipelothrix rhusiopathiae contamination in the poultry house environment during erysipelas outbreaks in organic laying hen flocks a
b
b
c
Helena Eriksson , Elisabeth Bagge , Viveca Båverud , Claes Fellström & Désirée S. a
Jansson a
Department of Animal Health and Antimicrobial Strategies, National Veterinary Institute, Uppsala, Sweden b
Department of Bacteriology, National Veterinary Institute, Uppsala, Sweden
c
Department of Clinical Sciences, Swedish University of Agricultural Sciences, Uppsala, Sweden Accepted author version posted online: 24 Mar 2014.Published online: 22 Apr 2014.
To cite this article: Helena Eriksson, Elisabeth Bagge, Viveca Båverud, Claes Fellström & Désirée S. Jansson (2014): Erysipelothrix rhusiopathiae contamination in the poultry house environment during erysipelas outbreaks in organic laying hen flocks, Avian Pathology, DOI: 10.1080/03079457.2014.907485 To link to this article: http://dx.doi.org/10.1080/03079457.2014.907485
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Avian Pathology, 2014 http://dx.doi.org/10.1080/03079457.2014.907485
ORIGINAL ARTICLE
Erysipelothrix rhusiopathiae contamination in the poultry house environment during erysipelas outbreaks in organic laying hen flocks Helena Eriksson1*, Elisabeth Bagge2, Viveca Båverud2, Claes Fellström3 and Désirée S. Jansson1 Department of Animal Health and Antimicrobial Strategies, National Veterinary Institute, Uppsala, Sweden, 2Department of Bacteriology, National Veterinary Institute, Uppsala, Sweden, and 3Department of Clinical Sciences, Swedish University of Agricultural Sciences, Uppsala, Sweden
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This study investigated organic laying hen farms for the presence of Erysipelothrix rhusiopathiae in the house environment and from potential carriers (i.e. insects and mice) during ongoing erysipelas outbreaks, and compared the obtained isolates with those from laying hens. The samples were investigated by selective culture followed by species-specific polymerase chain reaction on cultures. E. rhusiopathiae was isolated from the spleen, jejunal contents, manure, dust and swabs from water nipples. Three more samples from the house environment tested positive by polymerase chain reaction compared with selective culture alone. Selected isolates were investigated by pulsed-field gel electrophoresis (PFGE) and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS). One farm was represented by isolates from laying hens only, and one of these isolates differed in one PFGE band from the others. Different banding patterns were observed for isolates from laying hens and manure on one farm. On the remaining two farms, the isolates from the house environment and laying hens were identical but differed between farms. Outbreaks reoccurred in the next flock on two of the farms, and different PFGE types were isolated from consecutive flocks. Our results suggest an external source of infection, which would explain the previously reported increased risk of outbreaks in free-range flocks. Contaminated manure and dust may represent sources of transmission. For the isolates, MALDI-TOF MS and biochemical typing results were in agreement but, since the type strain of Erysipelothrix tonsillarum was typed as E. rhusiopathiae using MALDI-TOF MS, further studies into this method are needed.
Introduction Erysipelas, caused by the Gram-positive bacterium Erysipelothrix rhusiopathiae, is an emerging cause of mortality in non-caged laying hens in Europe (Mazaheri et al., 2005; Eriksson et al., 2010; Stokholm et al., 2010). The bacterial colonies are small (0.5 mm) with a narrow alpha-haemolytic zone on blood agar plates and may be difficult to detect when more profusely growing bacteria are present in the sample. Isolation is therefore often performed using selective media (Packer, 1943; Wood, 1965; Harrington & Hulse, 1971). Several polymerase chain reaction (PCR) methods have been established to accelerate diagnostics and to differentiate E. rhusiopathiae from the closely related species Erysipelothrix tonsillarum, which is considered apathogenic in poultry (Takahashi et al., 1994; Shimoji et al., 1998; Takeshi et al., 1999; Pal et al., 2010). Pulsed-field gel electrophoresis (PFGE) has been used previously to study the molecular epidemiology of poultry isolates of E. rhusiopathiae (Købke et al., 2005; Eriksson et al., 2009, 2010). Although the species is diverse by PFGE analysis, homogeneous banding patterns were observed during clinical outbreaks, which suggest that
they may be of a clonal nature (Købke et al., 2005; Eriksson et al., 2010). In recent years, matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS) has been introduced for bacterial identification to species level (Wieser et al., 2012). However, reports on the use of this method for E. rhusiopathiae species determination are missing. To the authors’ knowledge, this is the first study on the presence of E. rhusiopathiae in the poultry house environment of affected laying hen flocks during ongoing outbreaks. Therefore, and due to the reported tendency for free-range flocks to be at higher risk of an outbreak (Eriksson et al., 2013), the aims of this study were to determine whether E. rhusiopathiae is present in the environment on affected organic laying hen farms, to identify where it occurs and to compare the environmental isolates with those obtained from laying hens on the same farm. Materials and Methods Selection of flocks. A total of six Swedish commercial organic laying hen farms were included in the study. The farms had been diagnosed with
*To whom correspondence should be addressed. Tel: +46 18 67 40 00. Fax: +46 18 30 91 62. E-mail: [email protected] (Received 17 December 2013; accepted 28 February 2014) © 2014 Houghton Trust Ltd
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H. Eriksson et al. sample were subcultured on agar plates, and were stored in beef broth with 10% equine serum and 15% glycerol at −70°C until further analyses.
ongoing erysipelas outbreaks in 2011 (Farms A to C) and 2012 (Farm D), based on necropsy findings and bacterial culture at the National Veterinary Institute (SVA), Uppsala, Sweden. For comparison, two clinically healthy flocks on other farms with no previous history of erysipelas were used as controls (Farms E and F). These matched the other farms in terms of flock size and housing system. The studied flocks consisted of 2500 birds (Farm B), 2600 birds (Farm F) and 3000 birds (Farms A and C to E), whereas the total hen numbers varied between 2600 laying hens (Farm F) and 27,000 laying hens (Farm D). No other livestock was present on Farms A and B and D to F. On Farm C, cattle were grazing 40 m from the affected flock. Birds found dead were collected at least once a day on all six farms and stored in a freezer before transferring to separate containers that were collected by a cadaver transport company. Flocks affected by erysipelas were euthanized by gassing with carbon dioxide in the house (Farms B to D) or were vaccinated and subsequently sent for slaughter (Farm A). Consecutive flocks on two of the farms (Farms A and C) were affected by erysipelas outbreaks in 2012 and isolates from laying hens in these flocks were also included. The chickens were affected at around 55 weeks of age, despite vaccination with an inactivated vaccine containing E. rhusiopathiae serotype 2 (Nobilis®Erysipelas vet; Intervet International B. V., Boxmeer, the Netherlands). Instructions were given to administer the vaccine subcutaneously to each bird. For practical reasons the vaccine was administered once, on arrival at the farm at around 16 weeks of age.
Mice Mice were caught using snap traps placed during the visits, in areas where there were signs of rodent activity (i.e. presence of faeces) or at locations identified by the farmer as sites of previous mouse activity. The snap traps were attached to wooden boards (five traps per board) and baited with peanut butter, and the farmers were asked to inspect the traps every day. The procedures used were approved by the Ethical Committee for Scientific Experiments (protocol C361/10). The farmers were instructed that all mice caught during a month after the visit should be sent on the same day as they were found, by overnight mail, to SVA together with an ice pack. At SVA, necropsies were performed and samples from the nasopharynx and caecocolonic junction were cultured as described above. In addition, caecocolonic junction samples from mice on Farm D were also streaked onto horse blood agar plates supplemented with kanamycin (400 µg/ml) and neomycin (50 µg/ml) (KN-agar) (Wood, 1965; Bender et al., 2010). The selective properties of the KN-agar plates were tested and confirmed in a pilot study before the samples were analysed (data not shown). The nasopharyngeal samples from mice were spread on horse blood agar plates without prior incubation in SACVB.
Samples from the poultry house environment Samples from the house environment were obtained for bacteriological analysis on all six farms (Table 1). These samples were obtained by sock swabs (gauze moistened with sterile water) from the ground outside the entrance door of the house, the floor between the entrance door and the hygiene barrier, the floor of the anteroom, the litter bed, the winter garden (if present) and the outdoor pen, according to procedures described previously (Widgren et al., 2013). Dust samples from the ventilation exhaust fans were collected by scraping dust from the interior surface. On Farm B, samples were also obtained from the air inlets. E-Swabs (Copan, Brescia, Italy) were used to sample 25 evenly distributed nipple drinkers (five nipples per swab). Manure (approximately 0.2 dm3 manure per sample) was collected from the manure heap outside the house on all farms except Farm E, where samples were obtained from the manure storage below the slats in the house. Soil samples from pasture were obtained using a stainless steel cylinder (0.2 dm3) from five locations in the outdoor pen, two near the house, two further away in the pen and one in the middle of these sampling points. Attempts to net flying insects were performed over the manure heap on all farms except Farm E. Sticky fly traps (Flyson® FlyTube; Pharmaxim, Markaryd, Sweden) were mounted in the poultry house to collect flying insects from the house. Other insects observed during the visit were collected by hand. The samples from the house environment were placed on ice immediately and transported to the laboratory on the same day. They were stored at 20°C
Isolation of Erysipelothrix rhusiopathiae. The study was conducted between August 2011 and September 2012 when all farms were visited by the first author to collect samples from chickens, environment and potential vectors, except mice that were caught after the visits.
Laying hens The affected farms were visited for sampling and data collection during ongoing outbreaks: Farms A to C in August and September 2011, and Farm D in September 2012. The two clinically healthy flocks (Farms E and F) were visited in September and October 2011 respectively. During these visits, five dead hens were collected, transported to SVA and refrigerated until the next day, when samples from spleen and jejunal contents were cultured. The samples were inoculated in 5 ml broth containing 0.2 mg/ml sodium azide and 5 µg/ml crystal-violet (SACVB; SVA) and were incubated at 37°C for 48 h. One loopful (approximately 10 µl) of the broth was spread on horse blood agar plates (Blood Agar Base no. 2 [Oxoid, Basingstoke, UK] supplemented with 5% citrated horse blood [Håtunalab, Bro, Sweden]) and incubated at 37°C for 48 h. Growth of E. rhusiopathiae was confirmed based on colony morphology, Gram staining and biochemical characteristics as described previously (Eriksson et al., 2009). One colony from each positive spleen and a maximum of five colonies from each positive jejunal
Table 1.
Culture of organ and samples from the poultry house environment.
Material Laying hen—spleen Laying hen—jejunum Mouse (Mus musculus)—nasopharynx Mouse (M. musculus)—caecocolonic junction Sock sample Nipple drinkers (swab) Dust from air inlet Dust from exhaust fans Soil from outside pen Manure from heap/storage Insects captured above manure heap Sticky fly trap Other arthropods Total
Farm A
Farm B
Farm C
Farm D
Farm E
Farm F
Total
5/5 0/5 NA NA 0/5 0/5 NA 0/2 0/5 0/3 0/1 0/2 0/1 5/34
5/5 1/3 NA NA 0/6 0/5 0/6 0/3 0/5 1/3 0/1 0/1 0/2 7/40
5/5 3/5 NA NA 0/6 0/5 NA 0/2 0/5 2/3 0/1 0/2 0/1 10/35
5/5 4/5 0/5 0/5 0/5 4/5 NA 2/3 0/6 2/4 0/1 NA 0/2 17/46
0/5 0/2 0/5 0/5 0/6 0/5 NA 0/3 0/5 0/3 NA 0/1 NA 0/40
0/5 0/2 NA NA 0/6 0/5 NA 0/2 0/5 0/3 NA NA NA 0/28
20/30 8/22 0/10 0/10 0/34 4/30 0/6 2/15 0/31 5/19 0/4 0/6 0/10 39/223
Data presented as number of samples testing positive for E. rhusiopathiae/number of investigated samples. Farms A to D represent farms with ongoing outbreaks of erysipelas, and Farms E and F represent farms with no signs of diseased birds. NA, not available.
E. rhusiopathiae in the house environment
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for 6 to 8 months before analysis (Farms A to C and E to F). The samples from Farm D were cultured on the day after the visit, without prior freezing. Insects and swab samples from nipple drinkers were incubated in 5 ml SACVB. One of the sock swabs in each pair was placed in a plastic jar and immersed in SACVB, and 10 g of the other samples were incubated in 90 ml SACVB. After incubation at 37°C for 48 h, 10 µl of the broth were streaked onto horse blood agar and KN-agar. The plates were incubated at 37°C for 48 h and suspected Erysipelothrix colonies were confirmed as described previously (Eriksson et al., 2009). For each positive sample, a maximum of five colonies were stored at −70°C.
Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Fifteen isolates from laying hens and six isolates from the house environment were subjected to typing by MALDI-TOF MS using a Bruker-Microflex MALDI-TOF MS spectrometer and the supplied software Maldi Biotyper 4.0 real-time classification (Bruker Daltonics, Bremen, Germany). The isolates were selected to represent different PFGE banding patterns (see below) and various sources. The samples were prepared by the direct transfer method according to the instructions from the manufacturer. Scores over 2.0 were considered as positive identification to species level. Isolates with scores below 2.0 were investigated using the formic acid method according to the instructions from the manufacturer, and if still with scores below 2.0 the isolates were investigated using ethanol–formic acid extraction as described by the manufacturer. Type strains E. rhusiopathiae obtained from the Culture Collection at the University of Göteborg (CCUG) as CCUG 221T and E. tonsillarum (CCUG 31352T) were included as reference strains and were investigated using ethanol–formic acid extraction.
Polymerase chain reaction analysis. DNA preparation Templates for PCR were prepared from all agar plate cultures except for those from the spleen. Each plate with bacterial growth was harvested by rinsing with 1 ml sterile phosphate-buffered saline (PBS) without Ca2 + and Mg2 + , pH 7.4 (SVA). PBS consisted of Super-Q water (Merck Millipore, Darmstadt, Germany) with NaCl (8 g/l), KCl (0.2 g/l), KH2PO4 (0.2 g/l) and Na2HPO4 × 2 H2O (1.44 g/l) all from Merck (Darmstadt, Germany). After centrifugation at 16,200 × g (Fresco 21; Thermo Electron Co., Osterode, Germany) for 10 min, the supernatant was discarded and one well-filled white loop (1 µl) of the pellet was transferred to 200 µl PBS without Ca2 + and Mg2 + . The pellet was washed twice in PBS and resuspended in 50 µl ultrapure water (Sigma-Aldrich Co., St Louis, Missouri, USA). Bacterial cells were lysed on a heating block (98 to 99°C) for 10 min before rapid chilling on ice. After centrifugation at 6200 × g for 4 min, the supernatant was transferred to a new tube and stored at −20°C. Polymerase chain reaction A 937-base-pair fragment of a gene coding for a capsular biosynthesisassociated polypeptide was amplified with the E. rhusiopathiae specific primers ER1 and ER2 (Shimoji et al., 1998). The PCR mixture (25 µl total volume) consisted of 10 mM Tris–HCl (pH 8.3), 50 mM KCl, 1.75 mM MgCl2, 0.2 mM (each) dNTP, 0.2 µM each primer, 1.25 u AmpliTaq® DNA Polymerase and 2 µl template DNA. The templates were used undiluted and diluted 1:10, and all reactions were carried out in duplicate. The PCR reaction was performed in a Thermal Biocycler 2720 (Applied Biosystems, Foster City, CA, USA). The amplification programme consisted of an initial denaturation at 94°C for 3 min, followed by 30 cycles of denaturation at 94° C for 30 sec, primer annealing at 63°C for 30 sec and extension at 72°C for 1 min. After the cycles, final annealing at 63°C for 1 min and a last extension at 72°C for 10 min were performed. The PCR products were separated on a 1.5% agarose gel (GE Healthcare Bio-Sciences, Uppsala, Sweden), stained with SYBR® Safe DNA gel stain (Invitrogen, Eugene, OR, USA) and visualized by ultraviolet transillumination. Prior to investigation of the samples, the specificity of the PCR was investigated with reference strains and clinical isolates. The strains used were E. rhusiopathiae (CCUG 221T), E. tonsillarum (CCUG 31352T), Staphylococcus aureus subsp. aureus (ATCC 29213), Enterococcus faecalis (ATCC 29212), Lactobacillus acidophilus (CCUG 5917T), Escherichia coli (ATCC 25922) and Pasteurella multocida subsp. multocida (CCUG 224). From the strain collection at SVA, 18 Swedish field isolates of E. rhusiopathiae from clinical outbreaks of erysipelas in chickens, one isolate of E.
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rhusiopathiae from a mink (CCUG 56461), two isolates of sucrose-negative presumed E. tonsillarum, one isolate from a dog (CCUG 36858) and another from a rat (CCUG 56462) (Eriksson et al., 2009), and one clinical isolate of P. multocida subsp. multocida from a pheasant were also included. Sequencing of six randomly chosen PCR products from manure and jejunal contents was performed for confirmation of specificity. The amplicons were purified by the Illustra GFXTM PCR DNA and Gel Band Purification Kit (GE Healthcare, Uppsala, Sweden) and sequence reactions were performed with the BigDye® Terminator v.3.1 kit (Applied Biosystems) in an ABI PRISM® 2700 Genetic Analyzer (Uppsala Genome Center, Uppsala University, Uppsala; http://genpat.uu.se/node462). The data were analysed by the CLC Sequence Viewer version 6.4 (CLC Bio, Arhus N, Denmark; http://www.clcbio.com/index.php?id = 28) and were compared by the BLAST application provided by the GenBank database against previously deposited sequences.
Pulsed-field gel electrophoresis analysis. Three horse blood agar plates with overnight cultures of bacteria were harvested for each isolate, and preparation of genomic DNA for PFGE analysis was performed as described by Eriksson et al. (2010). The purified DNA was digested with 40 u restriction enzyme Smal (New England BioLabs, Ipswich, MA, USA) according to the manufacturer’s instructions. DNA fragments were separated in 1% agarose gel (Agarose NA; GE Healthcare) in 0.5 × TBE buffer (45 mM Tris-borate, 1 mM ethylenediamine tetraacetic acid) for 24 h at pulse switch time ramped from 0.5 to 40 sec in a CHEF DRII apparatus (BioRad, Hercules, CA, USA). PFGE patterns were analysed with the BioNumerics version 6.6 software (Applied Maths, Sint-Martens-Latem, Belgium). Cluster analysis was performed with the unweighted pair group method with the arithmetic mean, Dice coefficient and 0.5% optimization with 1.4% band position tolerance.
Sequence data. The sequences of the six PCR amplicons were deposited in GenBank under the accession numbers KC986835 to KC986840.
Results Isolation of Erysipelothrix rhusiopathiae. Laying hens Culture results from laying hens are presented in Table 1. E. rhusiopathiae was isolated from the spleen of all sampled hens from farms with ongoing disease outbreaks (Farms A to D). Jejunal contents from hens from three of the four affected flocks were also culture-positive (8/18 samples analysed). All samples from laying hens from the two clinically healthy flocks (Farms E and F) were culturenegative. Mice Mice were present on all farms according to the farmers, but they were only caught 3 weeks after the visit on Farm D and 1 week after the visit on Farm E. No gross lesions were found in any of the mice at necropsy and all organ samples from mice were culture-negative (Table 1). Samples from the poultry house environment E. rhusiopathiae was isolated from manure, dust and swab samples from nipple drinkers on farms with ongoing outbreaks, but not from the two clinically healthy flocks (Table 1). Selective culture conditions (KN-agar) produced more isolates from manure and dust samples than nonselective culture (Table 2). Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Using the direct transfer method, the 21 isolates previously confirmed as E. rhusiopathiae by biochemical tests were identified as E. rhusiopathiae by MALDI-TOF MS, but the scores ranged from 1.718 to 2.38
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H. Eriksson et al. Table 2.
Comparison of bacteriological culture and species-specific PCR for E. rhusiopathiae.
Horse blood agar (non-selective) Sample
Culture
Nipple drinkers (n = 30) Manure (n = 19) Dust from exhaust fans (n = 15) Laying hen—jejunum (n = 22)
4/30 1/19 0/15 8/22
KN-agar (selective)
PCR a
4/19 2/19 0/15 8/19a
Culture
PCR
4/30 5/19 2/15 ND
4/11a 7/19 3/14a ND
(16 isolates with scores over 2.0 and five isolates with scores below 2.0). After investigations using the formic acid method, two of the five isolates with low scores were identified as E. rhusiopathiae with scores of 2.06 and 2.179. Renewed preparation by ethanol–formic acid extraction was performed on the remaining three isolates, which were again identified as E. rhusiopathiae with scores of 2.138, 2.232 and 1.99. The reference strains were both identified as E. rhusiopathiae with scores of 2.219 (E. rhusiopathiae CCUG 221T) and 2.275 (E. tonsillarum CCUG 31352T) respectively.
products was 890 nucleotides (GenBank accession numbers KC986835 to 986840). The five nucleotide sequences (Farms A and C) were identical to each other but differed in three positions from the sixth sequence (Farm B) and in four positions from the sequence of the target gene of the type strain E. rhusiopathiae Fujisawa (GenBank accession number AP012027, gene ERH_0856). One of the sequences (isolate from manure on Farm B) differed in one position from the type strain. The five amino acid sequences differed in two positions from the type strain, whereas that from Farm B was identical to the type strain.
Polymerase chain reaction analysis. Specificity control produced PCR fragments of the expected size for all E. rhusiopathiae isolates but not for the other bacterial species investigated, including the presumed E. tonsillarum isolates (Eriksson et al., 2009). All samples (n = 193, excluding those from the spleen in Table 1) were cultured as described above and the plates (horse blood agar and KNagar) with visible bacterial growth (n = 313) were used for DNA preparation. Among these plates, 28 were positive by PCR, corresponding to 22 samples. All agar plates with confirmed growth of E. rhusiopathiae (by biochemical tests) were PCR-positive but E. rhusiopathiae isolates were only obtained from 19 out of the 22 PCR-positive samples. Table 2 presents a comparison of results of culture and PCR analysis. The PCR on manure and dust samples was positive to a higher extent with templates from KN-agar plates than from horse blood agar plates. Sequence analysis of the six PCR amplicons confirmed their identity. The length of the six deposited PCR
Pulsed-field gel electrophoresis analysis. In total, 93 E. rhusiopathiae isolates were analysed by PFGE; 24 isolates from the spleen (one isolate/hen), 30 isolates from the jejunal contents from laying hens (one to five isolates/ hen) and 39 isolates from environmental samples. Seven different PFGE types were observed among the isolates from the four farms (Figure 1). Different PFGE types were obtained from all four farms. On Farm A, all isolates from laying hens showed an identical banding pattern except one from a spleen, which differed by a single band. For the three farms where samples from laying hens and the environment could be compared (Farms B, C and D), identical banding patterns were observed in samples from laying hens and the house environment on Farms C and D, whereas on Farm B different banding patterns were obtained (Figure 1). Furthermore, on Farms A and C, which were represented by isolates obtained in 2011 and in 2012, different banding patterns were identified between years.
100
80
Degree of similarity (%) 60
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Cultures were performed using non-selective horse blood agar plates and horse blood agar plates supplemented with kanamycin (400 µg/ml) and neomycin (50 µg/ml) (KN-agar). PCR templates were prepared from all plates with visible growth (n = 313) (spleen samples not included), and 28 templates (from 14 horse blood agar and 14 KN-agar plates) were PCR-positive. Only sample categories with confirmed growth of E. rhusiopathiae by biochemical tests are included. ND, not determined. a Fewer samples were analysed by PCR than by culture because PCR was only performed from agar plates with visible growth.
Farm A / 2011 Laying hen Farm A / 2011 Laying hen Farm C / 2012 Laying hen Farm B / 2011 Laying hen Farm C / 2011 Laying hen & Manure Farm A / 2012 Laying hen Farm D / 2012 Laying hen, Manure, Dust & Nipple drinker Farm B / 2011 Manure Figure 1. The seven PFGE types obtained (farm designation/year of outbreak and isolation source) based on original comparison of 93 E. rhusiopathiae isolates from laying hens and the poultry house environment of four organic laying hen flocks on different farms. Banding patterns produced by SmaI restriction digestion provided the data for the dendrogram.
E. rhusiopathiae in the house environment
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Discussion Knowledge of sources of E. rhusiopathiae infection and transmission routes in laying hen flocks is of the utmost importance to prevent and manage outbreaks but few epidemiologic data are currently available. To our knowledge, this is the first study to investigate intestinal samples and samples from the house environment from laying hen farms for the presence of E. rhusiopathiae during an ongoing outbreak. In previous studies, the bacterium has been detected in manure and soil from pig and cattle farms (Wood & Packer, 1972; Nørrung et al., 1987). In addition, Bender et al. (2010) isolated the bacterium from nipple drinkers, walls, fans, feed and central line water in affected pig herds. The detection in this study of E. rhusiopathiae in manure from three of the four affected chicken flocks and in dust and on water nipples from one of the farms suggests that the bacterium is widespread in the environment during ongoing outbreaks. The dust samples that were positive for E. rhusiopathiae were collected inside the exhaust fans, so the possibility that contaminated dust could leave the house via this route must be taken into consideration. However, whether the bacteria can survive in dust in the longer term and whether dust can act as a source of transmission remain to be determined. Based on our results, the risk for transmission with manure must be considered greater than that for dust. E. rhusiopathiae was detected in eight of 18 jejunal samples from infected flocks when cultured on blood agar, which indicates a high number of bacteria in the contents of the jejunum. Furthermore, the bacteria were detected in the manure heaps from three out of four affected flocks. In 1965, Ostašev showed that E. rhusiopathiae could survive for more than 290 days when inoculated into pig slurry and stored at 18 to 20°C (reviewed by Mitscherlich & Marth, 1984). However, the composition of manure differs between species and the survival ability of E. rhusiopathiae in poultry manure requires further study. Until such knowledge becomes available, special precautions are needed when handling manure from an infected flock. The manure from affected flocks should preferably not be stored near poultry houses, as vectors may potentially transmit the infection from the heap. Measures should also be taken to minimize the risk of exposure to other animals, including wildlife, especially when manure from an infected flock is spread on arable land. Manure should therefore be ploughed down immediately and should not be used on pasture. The fact that the isolates from manure on Farm B in this study were of a different PFGE type than the isolates from the organs of the hens on the same farm is interesting. This could be explained by the limited number of samples or contamination of the manure heap from an external source. Another possibility is that the flock was infected by two distinctly different PFGE types, but one of these could have been introduced earlier or may be better adapted for in vitro growth (Eriksson et al., 2010). The possibility of a carrier state in chickens must also be considered. Pigs have been shown to be carriers of E. rhusiopathiae (Opriessnig & Wood, 2012), but studies in chickens are scarce. Nakazawa et al. (2008) isolated E. rhusiopathiae from broiler chickens in an abattoir and suggested that chickens may be a potential reservoir. However, in our study the samples from the two clinically healthy flocks all tested negative, which suggests that the bacterium is not normally present, is only present at low numbers or is only present in few individuals.
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The routes of transmission of E. rhusiopathiae within a poultry flock have not been determined fully, but in previous studies chickens have been infected both by oral and intramuscular experimental challenge (Takahashi et al., 1994; Mazaheri et al., 2005). It has been suggested that the bacteria gain entry through wounds and damaged mucous membranes and that behaviours such as cannibalism and feather pecking in flocks increase transmission (Bricker & Saif, 2013). Proper culling practices, including prompt removal of dead and diseased birds from the flock several times a day, are therefore important to limit the spread and contamination rate of E. rhusiopathiae in the poultry house environment. Our finding that the bacterium is present in jejunal contents and manure from affected flocks suggests that a faecal–oral transmission route could be important in field outbreaks, but further research is needed. This finding also indirectly supports a previous report of higher risks of an outbreak in laying hen flocks in litter-based systems than those in cages (Eriksson et al., 2013), because caged chickens are less exposed to faeces than chickens kept on litter or as free range. Despite vaccination of pullets on arrival at the farms, outbreaks occurred in the consecutive flocks on Farms A and C in 2012. Previous experiences from Sweden have shown that a single vaccination at placement of pullets protects the flock from erysipelas for the duration of the production period (approximately 50 to 60 weeks). However, this particular vaccine was developed for use in turkeys and the manufacturer recommends that birds should be vaccinated twice. For practical reasons this is difficult and laborious in pullets, but should be considered to avoid future outbreaks. Along with aspects of vaccine handling such as transport, storage and administration, there is a potential risk that the outbreaks in the consecutive flocks were caused by strains to which the vaccine offered no or only limited protection. Studies in non-avian species have shown a variable degree of cross-protection between serotypes and, more recently, also between the surface protective antigen types (Sawada & Takahashi, 1987; Ingebritson et al., 2010). The isolates obtained from the flocks in this study were not further investigated. Crossprotection studies in poultry should preferably be performed. Interestingly, the outbreaks on Farms A and C in 2012 were caused by different PFGE types to those in 2011. In a previous study we found that isolates from an organic laying hen farm affected in 1998, 1999, 2001 and 2004 were of identical (in 2001 and 2004) or dissimilar (in 1998 and 1999) PFGE types (Eriksson et al., 2009). The fact that different PFGE types were found in some of these cases suggests that the birds were infected by an external source rather than by residual bacteria in the house environment. In the case of identical PFGE types in consecutive flocks, re-introduction of the same bacterial genotype from a common source in the external environment should also be considered. The incidence of E. rhusiopathiae in wild fauna and its ability to survive in the environment, including outside pens, are important aspects to address in the future. Until further knowledge has been obtained, a high level of biosecurity, including effective hygiene barriers, is recommended. In addition, farmers should ensure that fences effectively keep wild animals outside the pen. Interestingly, during a survey in this study (data not shown) several farmers reported problems with wild animals (e.g. wild boars) around, and sometimes even inside, the outside pen.
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With the exception of spleen samples from laying hens in affected flocks, a relatively small proportion of the samples in this study was positive by culture combined with biochemical identification and/or PCR from agar plate cultures. This may have been due to low sensitivity of the diagnostic methods used, suboptimal sampling procedures and/or a negative effect of frozen storage. In fact, our results suggest that the bacterium does not survive long-term storage in frozen samples from the poultry house environment. However, in a previous study on environmental samples from pig farms affected by erysipelas, E. rhusiopathiae was successfully isolated from samples stored at −20°C when cultured within 3 months of collection (Bender et al., 2010). Further, survivability of the bacterium during frozen storage needs to be further investigated. The PCR from agar plate cultures seemed slightly more sensitive than culture combined with biochemical identification, most probably because E. rhusiopathiae grows in very small colonies, which can be difficult to detect on agar plates with other more profusely growing bacteria. The negative sampling results from mice and arthropods indicate that these populations were not heavily infected. However, early experimental studies have indicated that E. rhusiopathiae may be transmitted by flies and mosquitoes (Wellmann 1950, 1955), although no study of insects on a farm during an outbreak could be found in the literature. Neither could any report on natural E. rhusiopathiae infection in mice be found; however, mice are sometimes used as a challenge model for E. rhusiopathiae infection. In our study, mice were only caught on one of the infected farms 3 weeks after the visit. Further studies with extensive sampling of arthropods and mice caught during ongoing outbreaks are needed. Finally, our investigation into the use of MALDI-TOF MS as a method for species confirmation of E. rhusiopathiae was inconclusive. However, based on the fact that the reference strain of E. tonsillarum (CCUG 31352T) was typed as E. rhusiopathiae with a high score, further studies are clearly needed. The problem of misidentification of E. tonsillarum in this study may be due to the fact that MALDI-TOF MS species identification is based on investigations of the ribosomal protein spectra and that E. rhusiopathiae and E. tonsillarum are very similar in their 16S rRNA sequences (Eriksson et al., 2009; Wieser et al., 2012). The number of available spectra in the database was also very limited. If MALDI-TOF MS can replace conventional biochemical typing, this would shorten the time to diagnosis and would be of importance since the farmer needs to take appropriate measures as soon as possible to reduce bird suffering. In conclusion, we have shown that E. rhusiopathiae is widespread in the poultry house environment during ongoing erysipelas outbreaks in organic laying hen flocks.
Acknowledgements The authors wish to thank the six farmers for providing data, time and help during sampling, Ricardo Feinstein for performing necropsies of the mice, Sigbrit Mattsson for helping with sampling on the farms, Helena Ljung, Anna Eriksson and Lena Lundgren for assistance with laboratory work, and Anna Aspán for Figure 1.
Funding The Ekhaga foundation is acknowledged for funding (under Grant 2010-64) sampling and bacteriological analyses, and SLU EkoForsk is acknowledged for funding PCR analyses.
References Bender, J.S., Shen, H.S., Irwin, C.K., Schwartz, K.J. & Opriessnig, T. (2010). Characterization of Erysipelothrix species isolates from clinically affected pigs, environmental samples, and vaccine strains from six recent swine erysipelas outbreaks in the United States. Clinical and Vaccine Immunology, 17, 1605–1611. Bricker, J.M. & Saif, Y.M. (2013). Erysipelas. In D.E. Swayne, J.R Glisson, L.R McDougald, L.K. Nolan, D.L. Suarez, & V. Nair (Eds.). Diseases of Poultry 13th edn (pp. 986–994). Ames, IA: John Wiley & Sons. Eriksson, H., Brännström, S., Skarin, H. & Chirico, J. (2010). Characterization of Erysipelothrix rhusiopathiae isolates from laying hens and poultry red mites (Dermanyssus gallinae) from an outbreak of erysipelas. Avian Pathology, 39, 505–509. Eriksson, H., Jansson, D.S., Johansson, K.E., Båverud, V., Chirico, J. & Aspán, A. (2009). Characterization of Erysipelothrix rhusiopathiae isolates from poultry, pigs, emus, the poultry red mite and other animals. Veterinary Microbiology, 137, 98–104. Eriksson, H., Nyman, A.-K., Fellström, C. & Wallgren, P. (2013). Erysipelas in laying hens is associated with housing system. The Veterinary Record, 173, 18. Harrington, R. & Hulse, D.C. (1971). Comparison of two plating media for the isolation of Erysipelothrix rhusiopathiae from enrichment broth culture. Applied Microbiology, 22, 141–142. Ingebritson, A.L., Roth, J.A. & Hauer, P.J. (2010). Erysipelothrix rhusiopathiae: association of Spa-type with serotype and role in protective immunity. Vaccine, 28, 2490–2496. Købke, B., Eigaard, N.M., Christensen, J.P. & Bisgaard, M. (2005). Investigations on clonality and stability of clones of Erysipelothrix rhusiopathiae in turkeys and free range layers flocks affected by erysipelas. In 14th World Veterinary Poultry Congress Final Program & Abstract Book (p. 313). Istanbul, Turkey. Mazaheri, A., Lierz, M. & Hafez, H.M. (2005). Investigations on the pathogenicity of Erysipelothrix rhusiopathiae in laying hens. Avian Diseases, 49, 574–576. Mitscherlich, E. & Marth, E.H. (1984). Microbial Survival in the Environment. Berlin: Springer-Verlag. Nakazawa, H., Hayashidani, H., Higashi, J., Kaneko, K.-I., Takahashi, T. & Ogawa, M. (1998). Occurrence of Erysipelothrix spp. in broiler chickens at an abattoir. Journal of Food Protection, 61, 907–909. Nørrung, V., Munch, B. & Errebo Larsen, H. (1987). Occurrence, isolation and serotyping of Erysipelothrix rhusiopathiae in cattle and pig slurry. Acta Veterinaria Scandinavica, 28, 9–14. Opriessnig, T. & Wood, R.L. (2012). Erysipelas. In J.J. Zimmerman, L.A. Karriker, A. Ramirez, K.J. Schwartz, & G.W. Stevenson (Eds.). Diseases of Swine 10th edn (pp. 750–759). Ames, IA: John Wiley & Sons. Packer, R.A. (1943). The use of sodium azide (NaN3) and crystal violet in a selective medium for streptococci and Erysipelothrix rhusiopathiae. Journal of Bacteriology, 46, 343–349. Pal, N., Bender, J.S. & Opriessnig T. (2010). Rapid detection and differentiation of Erysipelothrix spp. by a novel multiplex real time PCR assay. Journal of Applied Microbiology, 108, 1083–1093. Sawada, T. & Takahashi, T. (1987). Cross protection of mice and swine inoculated with culture filtrate of attenuated Erysipelothrix rhusiopathiae and challenge exposed to strains of various serovars. The American Journal of Veterinary Research, 48, 239–242. Shimoji, Y., Mori, Y., Hyakutake, K., Sekizaki, T. & Yokomizo, Y. (1998). Use of an enrichment broth cultivation-PCR combination assay for rapid diagnosis of swine erysipelas. Journal of Clinical Microbiology, 36, 86–89. Stokholm, N.M., Permin, A., Bisgaard, M. & Christensen, J.P. (2010). Causes of mortality in commercial organic layers in Denmark. Avian Diseases, 54, 1241–1250.
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Takahashi, T., Takagi, M., Yamaoka, R., Ohishi, K., Norimatsu, M., Tamura, Y. & Nakamura, M. (1994). Comparison of the pathogenicity for chickens of Erysipelothrix rhusiopathiae and Erysipelothrix tonsillarum. Avian Pathology, 23, 237–245. Takeshi, K., Makino, S., Ikeda, T., Takada, N., Nakashiro, A., Nakanishi, K., Oguma, K., Katoh, Y., Sunagawa, H. & Ohyama, T. (1999). Direct and rapid detection by PCR of Erysipelothrix sp. DNAs prepared from bacterial strains and animal tissues. Journal of Clinical Microbiology, 37, 4093–4098. Wellmann, G. (1950). Transmission of erysipelas to pigeons by various species of blood-sucking insects. Zentralblatt für Bakteriologie, Parasitenkunde, Infektionskrankheit und Hygiene, 155, 109–115. Wellmann, G. (1955). Transmission of pig erysipelas by the housefly (Musca domestica). Zentralblatt für Bakteriologie, Parasitenkunde, Infektionskrankheit und Hygiene, 162, 261–264.
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Widgren, S., Eriksson, E., Aspán, A., Emanuelson, U., Alenius, S. & Lindberg, A. (2013). Environmental sampling for evaluating verotoxigenic Escherichia coli O157: H7 status in dairy cattle herds. Journal of Veterinary Diagnostic Investigation, 25, 189–198. Wieser, A., Schneider, L., Jung, J. & Schubert S. (2012). MALDI-TOF MS in microbiological diagnostics–identification of microorganisms and beyond (mini review). Applied Microbiology & Biotechnology, 93, 965–974. Wood, R.L. (1965). A selective liquid medium utilizing antibiotics for isolation of Erysipelothrix insidiosa. The American Journal of Veterinary Research, 26, 1303–1308. Wood, R.L. & Packer, R.A. (1972). Isolation of Erysipelothrix rhusiopathiae from soil and manure of swine-raising premises. The American Journal of Veterinary Research, 33, 1611–1620.
Avian Pathology Publication details, including instructions for authors and subscription information: http://www.tandfonline.com/loi/cavp20
Erysipelothrix rhusiopathiae contamination in the poultry house environment during erysipelas outbreaks in organic laying hen flocks a
b
b
c
Helena Eriksson , Elisabeth Bagge , Viveca Båverud , Claes Fellström & Désirée S. a
Jansson a
Department of Animal Health and Antimicrobial Strategies, National Veterinary Institute, Uppsala, Sweden b
Department of Bacteriology, National Veterinary Institute, Uppsala, Sweden
c
Department of Clinical Sciences, Swedish University of Agricultural Sciences, Uppsala, Sweden Accepted author version posted online: 24 Mar 2014.Published online: 22 Apr 2014.
To cite this article: Helena Eriksson, Elisabeth Bagge, Viveca Båverud, Claes Fellström & Désirée S. Jansson (2014): Erysipelothrix rhusiopathiae contamination in the poultry house environment during erysipelas outbreaks in organic laying hen flocks, Avian Pathology, DOI: 10.1080/03079457.2014.907485 To link to this article: http://dx.doi.org/10.1080/03079457.2014.907485
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Avian Pathology, 2014 http://dx.doi.org/10.1080/03079457.2014.907485
ORIGINAL ARTICLE
Erysipelothrix rhusiopathiae contamination in the poultry house environment during erysipelas outbreaks in organic laying hen flocks Helena Eriksson1*, Elisabeth Bagge2, Viveca Båverud2, Claes Fellström3 and Désirée S. Jansson1 Department of Animal Health and Antimicrobial Strategies, National Veterinary Institute, Uppsala, Sweden, 2Department of Bacteriology, National Veterinary Institute, Uppsala, Sweden, and 3Department of Clinical Sciences, Swedish University of Agricultural Sciences, Uppsala, Sweden
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This study investigated organic laying hen farms for the presence of Erysipelothrix rhusiopathiae in the house environment and from potential carriers (i.e. insects and mice) during ongoing erysipelas outbreaks, and compared the obtained isolates with those from laying hens. The samples were investigated by selective culture followed by species-specific polymerase chain reaction on cultures. E. rhusiopathiae was isolated from the spleen, jejunal contents, manure, dust and swabs from water nipples. Three more samples from the house environment tested positive by polymerase chain reaction compared with selective culture alone. Selected isolates were investigated by pulsed-field gel electrophoresis (PFGE) and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS). One farm was represented by isolates from laying hens only, and one of these isolates differed in one PFGE band from the others. Different banding patterns were observed for isolates from laying hens and manure on one farm. On the remaining two farms, the isolates from the house environment and laying hens were identical but differed between farms. Outbreaks reoccurred in the next flock on two of the farms, and different PFGE types were isolated from consecutive flocks. Our results suggest an external source of infection, which would explain the previously reported increased risk of outbreaks in free-range flocks. Contaminated manure and dust may represent sources of transmission. For the isolates, MALDI-TOF MS and biochemical typing results were in agreement but, since the type strain of Erysipelothrix tonsillarum was typed as E. rhusiopathiae using MALDI-TOF MS, further studies into this method are needed.
Introduction Erysipelas, caused by the Gram-positive bacterium Erysipelothrix rhusiopathiae, is an emerging cause of mortality in non-caged laying hens in Europe (Mazaheri et al., 2005; Eriksson et al., 2010; Stokholm et al., 2010). The bacterial colonies are small (0.5 mm) with a narrow alpha-haemolytic zone on blood agar plates and may be difficult to detect when more profusely growing bacteria are present in the sample. Isolation is therefore often performed using selective media (Packer, 1943; Wood, 1965; Harrington & Hulse, 1971). Several polymerase chain reaction (PCR) methods have been established to accelerate diagnostics and to differentiate E. rhusiopathiae from the closely related species Erysipelothrix tonsillarum, which is considered apathogenic in poultry (Takahashi et al., 1994; Shimoji et al., 1998; Takeshi et al., 1999; Pal et al., 2010). Pulsed-field gel electrophoresis (PFGE) has been used previously to study the molecular epidemiology of poultry isolates of E. rhusiopathiae (Købke et al., 2005; Eriksson et al., 2009, 2010). Although the species is diverse by PFGE analysis, homogeneous banding patterns were observed during clinical outbreaks, which suggest that
they may be of a clonal nature (Købke et al., 2005; Eriksson et al., 2010). In recent years, matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS) has been introduced for bacterial identification to species level (Wieser et al., 2012). However, reports on the use of this method for E. rhusiopathiae species determination are missing. To the authors’ knowledge, this is the first study on the presence of E. rhusiopathiae in the poultry house environment of affected laying hen flocks during ongoing outbreaks. Therefore, and due to the reported tendency for free-range flocks to be at higher risk of an outbreak (Eriksson et al., 2013), the aims of this study were to determine whether E. rhusiopathiae is present in the environment on affected organic laying hen farms, to identify where it occurs and to compare the environmental isolates with those obtained from laying hens on the same farm. Materials and Methods Selection of flocks. A total of six Swedish commercial organic laying hen farms were included in the study. The farms had been diagnosed with
*To whom correspondence should be addressed. Tel: +46 18 67 40 00. Fax: +46 18 30 91 62. E-mail: [email protected] (Received 17 December 2013; accepted 28 February 2014) © 2014 Houghton Trust Ltd
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H. Eriksson et al. sample were subcultured on agar plates, and were stored in beef broth with 10% equine serum and 15% glycerol at −70°C until further analyses.
ongoing erysipelas outbreaks in 2011 (Farms A to C) and 2012 (Farm D), based on necropsy findings and bacterial culture at the National Veterinary Institute (SVA), Uppsala, Sweden. For comparison, two clinically healthy flocks on other farms with no previous history of erysipelas were used as controls (Farms E and F). These matched the other farms in terms of flock size and housing system. The studied flocks consisted of 2500 birds (Farm B), 2600 birds (Farm F) and 3000 birds (Farms A and C to E), whereas the total hen numbers varied between 2600 laying hens (Farm F) and 27,000 laying hens (Farm D). No other livestock was present on Farms A and B and D to F. On Farm C, cattle were grazing 40 m from the affected flock. Birds found dead were collected at least once a day on all six farms and stored in a freezer before transferring to separate containers that were collected by a cadaver transport company. Flocks affected by erysipelas were euthanized by gassing with carbon dioxide in the house (Farms B to D) or were vaccinated and subsequently sent for slaughter (Farm A). Consecutive flocks on two of the farms (Farms A and C) were affected by erysipelas outbreaks in 2012 and isolates from laying hens in these flocks were also included. The chickens were affected at around 55 weeks of age, despite vaccination with an inactivated vaccine containing E. rhusiopathiae serotype 2 (Nobilis®Erysipelas vet; Intervet International B. V., Boxmeer, the Netherlands). Instructions were given to administer the vaccine subcutaneously to each bird. For practical reasons the vaccine was administered once, on arrival at the farm at around 16 weeks of age.
Mice Mice were caught using snap traps placed during the visits, in areas where there were signs of rodent activity (i.e. presence of faeces) or at locations identified by the farmer as sites of previous mouse activity. The snap traps were attached to wooden boards (five traps per board) and baited with peanut butter, and the farmers were asked to inspect the traps every day. The procedures used were approved by the Ethical Committee for Scientific Experiments (protocol C361/10). The farmers were instructed that all mice caught during a month after the visit should be sent on the same day as they were found, by overnight mail, to SVA together with an ice pack. At SVA, necropsies were performed and samples from the nasopharynx and caecocolonic junction were cultured as described above. In addition, caecocolonic junction samples from mice on Farm D were also streaked onto horse blood agar plates supplemented with kanamycin (400 µg/ml) and neomycin (50 µg/ml) (KN-agar) (Wood, 1965; Bender et al., 2010). The selective properties of the KN-agar plates were tested and confirmed in a pilot study before the samples were analysed (data not shown). The nasopharyngeal samples from mice were spread on horse blood agar plates without prior incubation in SACVB.
Samples from the poultry house environment Samples from the house environment were obtained for bacteriological analysis on all six farms (Table 1). These samples were obtained by sock swabs (gauze moistened with sterile water) from the ground outside the entrance door of the house, the floor between the entrance door and the hygiene barrier, the floor of the anteroom, the litter bed, the winter garden (if present) and the outdoor pen, according to procedures described previously (Widgren et al., 2013). Dust samples from the ventilation exhaust fans were collected by scraping dust from the interior surface. On Farm B, samples were also obtained from the air inlets. E-Swabs (Copan, Brescia, Italy) were used to sample 25 evenly distributed nipple drinkers (five nipples per swab). Manure (approximately 0.2 dm3 manure per sample) was collected from the manure heap outside the house on all farms except Farm E, where samples were obtained from the manure storage below the slats in the house. Soil samples from pasture were obtained using a stainless steel cylinder (0.2 dm3) from five locations in the outdoor pen, two near the house, two further away in the pen and one in the middle of these sampling points. Attempts to net flying insects were performed over the manure heap on all farms except Farm E. Sticky fly traps (Flyson® FlyTube; Pharmaxim, Markaryd, Sweden) were mounted in the poultry house to collect flying insects from the house. Other insects observed during the visit were collected by hand. The samples from the house environment were placed on ice immediately and transported to the laboratory on the same day. They were stored at 20°C
Isolation of Erysipelothrix rhusiopathiae. The study was conducted between August 2011 and September 2012 when all farms were visited by the first author to collect samples from chickens, environment and potential vectors, except mice that were caught after the visits.
Laying hens The affected farms were visited for sampling and data collection during ongoing outbreaks: Farms A to C in August and September 2011, and Farm D in September 2012. The two clinically healthy flocks (Farms E and F) were visited in September and October 2011 respectively. During these visits, five dead hens were collected, transported to SVA and refrigerated until the next day, when samples from spleen and jejunal contents were cultured. The samples were inoculated in 5 ml broth containing 0.2 mg/ml sodium azide and 5 µg/ml crystal-violet (SACVB; SVA) and were incubated at 37°C for 48 h. One loopful (approximately 10 µl) of the broth was spread on horse blood agar plates (Blood Agar Base no. 2 [Oxoid, Basingstoke, UK] supplemented with 5% citrated horse blood [Håtunalab, Bro, Sweden]) and incubated at 37°C for 48 h. Growth of E. rhusiopathiae was confirmed based on colony morphology, Gram staining and biochemical characteristics as described previously (Eriksson et al., 2009). One colony from each positive spleen and a maximum of five colonies from each positive jejunal
Table 1.
Culture of organ and samples from the poultry house environment.
Material Laying hen—spleen Laying hen—jejunum Mouse (Mus musculus)—nasopharynx Mouse (M. musculus)—caecocolonic junction Sock sample Nipple drinkers (swab) Dust from air inlet Dust from exhaust fans Soil from outside pen Manure from heap/storage Insects captured above manure heap Sticky fly trap Other arthropods Total
Farm A
Farm B
Farm C
Farm D
Farm E
Farm F
Total
5/5 0/5 NA NA 0/5 0/5 NA 0/2 0/5 0/3 0/1 0/2 0/1 5/34
5/5 1/3 NA NA 0/6 0/5 0/6 0/3 0/5 1/3 0/1 0/1 0/2 7/40
5/5 3/5 NA NA 0/6 0/5 NA 0/2 0/5 2/3 0/1 0/2 0/1 10/35
5/5 4/5 0/5 0/5 0/5 4/5 NA 2/3 0/6 2/4 0/1 NA 0/2 17/46
0/5 0/2 0/5 0/5 0/6 0/5 NA 0/3 0/5 0/3 NA 0/1 NA 0/40
0/5 0/2 NA NA 0/6 0/5 NA 0/2 0/5 0/3 NA NA NA 0/28
20/30 8/22 0/10 0/10 0/34 4/30 0/6 2/15 0/31 5/19 0/4 0/6 0/10 39/223
Data presented as number of samples testing positive for E. rhusiopathiae/number of investigated samples. Farms A to D represent farms with ongoing outbreaks of erysipelas, and Farms E and F represent farms with no signs of diseased birds. NA, not available.
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for 6 to 8 months before analysis (Farms A to C and E to F). The samples from Farm D were cultured on the day after the visit, without prior freezing. Insects and swab samples from nipple drinkers were incubated in 5 ml SACVB. One of the sock swabs in each pair was placed in a plastic jar and immersed in SACVB, and 10 g of the other samples were incubated in 90 ml SACVB. After incubation at 37°C for 48 h, 10 µl of the broth were streaked onto horse blood agar and KN-agar. The plates were incubated at 37°C for 48 h and suspected Erysipelothrix colonies were confirmed as described previously (Eriksson et al., 2009). For each positive sample, a maximum of five colonies were stored at −70°C.
Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Fifteen isolates from laying hens and six isolates from the house environment were subjected to typing by MALDI-TOF MS using a Bruker-Microflex MALDI-TOF MS spectrometer and the supplied software Maldi Biotyper 4.0 real-time classification (Bruker Daltonics, Bremen, Germany). The isolates were selected to represent different PFGE banding patterns (see below) and various sources. The samples were prepared by the direct transfer method according to the instructions from the manufacturer. Scores over 2.0 were considered as positive identification to species level. Isolates with scores below 2.0 were investigated using the formic acid method according to the instructions from the manufacturer, and if still with scores below 2.0 the isolates were investigated using ethanol–formic acid extraction as described by the manufacturer. Type strains E. rhusiopathiae obtained from the Culture Collection at the University of Göteborg (CCUG) as CCUG 221T and E. tonsillarum (CCUG 31352T) were included as reference strains and were investigated using ethanol–formic acid extraction.
Polymerase chain reaction analysis. DNA preparation Templates for PCR were prepared from all agar plate cultures except for those from the spleen. Each plate with bacterial growth was harvested by rinsing with 1 ml sterile phosphate-buffered saline (PBS) without Ca2 + and Mg2 + , pH 7.4 (SVA). PBS consisted of Super-Q water (Merck Millipore, Darmstadt, Germany) with NaCl (8 g/l), KCl (0.2 g/l), KH2PO4 (0.2 g/l) and Na2HPO4 × 2 H2O (1.44 g/l) all from Merck (Darmstadt, Germany). After centrifugation at 16,200 × g (Fresco 21; Thermo Electron Co., Osterode, Germany) for 10 min, the supernatant was discarded and one well-filled white loop (1 µl) of the pellet was transferred to 200 µl PBS without Ca2 + and Mg2 + . The pellet was washed twice in PBS and resuspended in 50 µl ultrapure water (Sigma-Aldrich Co., St Louis, Missouri, USA). Bacterial cells were lysed on a heating block (98 to 99°C) for 10 min before rapid chilling on ice. After centrifugation at 6200 × g for 4 min, the supernatant was transferred to a new tube and stored at −20°C. Polymerase chain reaction A 937-base-pair fragment of a gene coding for a capsular biosynthesisassociated polypeptide was amplified with the E. rhusiopathiae specific primers ER1 and ER2 (Shimoji et al., 1998). The PCR mixture (25 µl total volume) consisted of 10 mM Tris–HCl (pH 8.3), 50 mM KCl, 1.75 mM MgCl2, 0.2 mM (each) dNTP, 0.2 µM each primer, 1.25 u AmpliTaq® DNA Polymerase and 2 µl template DNA. The templates were used undiluted and diluted 1:10, and all reactions were carried out in duplicate. The PCR reaction was performed in a Thermal Biocycler 2720 (Applied Biosystems, Foster City, CA, USA). The amplification programme consisted of an initial denaturation at 94°C for 3 min, followed by 30 cycles of denaturation at 94° C for 30 sec, primer annealing at 63°C for 30 sec and extension at 72°C for 1 min. After the cycles, final annealing at 63°C for 1 min and a last extension at 72°C for 10 min were performed. The PCR products were separated on a 1.5% agarose gel (GE Healthcare Bio-Sciences, Uppsala, Sweden), stained with SYBR® Safe DNA gel stain (Invitrogen, Eugene, OR, USA) and visualized by ultraviolet transillumination. Prior to investigation of the samples, the specificity of the PCR was investigated with reference strains and clinical isolates. The strains used were E. rhusiopathiae (CCUG 221T), E. tonsillarum (CCUG 31352T), Staphylococcus aureus subsp. aureus (ATCC 29213), Enterococcus faecalis (ATCC 29212), Lactobacillus acidophilus (CCUG 5917T), Escherichia coli (ATCC 25922) and Pasteurella multocida subsp. multocida (CCUG 224). From the strain collection at SVA, 18 Swedish field isolates of E. rhusiopathiae from clinical outbreaks of erysipelas in chickens, one isolate of E.
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rhusiopathiae from a mink (CCUG 56461), two isolates of sucrose-negative presumed E. tonsillarum, one isolate from a dog (CCUG 36858) and another from a rat (CCUG 56462) (Eriksson et al., 2009), and one clinical isolate of P. multocida subsp. multocida from a pheasant were also included. Sequencing of six randomly chosen PCR products from manure and jejunal contents was performed for confirmation of specificity. The amplicons were purified by the Illustra GFXTM PCR DNA and Gel Band Purification Kit (GE Healthcare, Uppsala, Sweden) and sequence reactions were performed with the BigDye® Terminator v.3.1 kit (Applied Biosystems) in an ABI PRISM® 2700 Genetic Analyzer (Uppsala Genome Center, Uppsala University, Uppsala; http://genpat.uu.se/node462). The data were analysed by the CLC Sequence Viewer version 6.4 (CLC Bio, Arhus N, Denmark; http://www.clcbio.com/index.php?id = 28) and were compared by the BLAST application provided by the GenBank database against previously deposited sequences.
Pulsed-field gel electrophoresis analysis. Three horse blood agar plates with overnight cultures of bacteria were harvested for each isolate, and preparation of genomic DNA for PFGE analysis was performed as described by Eriksson et al. (2010). The purified DNA was digested with 40 u restriction enzyme Smal (New England BioLabs, Ipswich, MA, USA) according to the manufacturer’s instructions. DNA fragments were separated in 1% agarose gel (Agarose NA; GE Healthcare) in 0.5 × TBE buffer (45 mM Tris-borate, 1 mM ethylenediamine tetraacetic acid) for 24 h at pulse switch time ramped from 0.5 to 40 sec in a CHEF DRII apparatus (BioRad, Hercules, CA, USA). PFGE patterns were analysed with the BioNumerics version 6.6 software (Applied Maths, Sint-Martens-Latem, Belgium). Cluster analysis was performed with the unweighted pair group method with the arithmetic mean, Dice coefficient and 0.5% optimization with 1.4% band position tolerance.
Sequence data. The sequences of the six PCR amplicons were deposited in GenBank under the accession numbers KC986835 to KC986840.
Results Isolation of Erysipelothrix rhusiopathiae. Laying hens Culture results from laying hens are presented in Table 1. E. rhusiopathiae was isolated from the spleen of all sampled hens from farms with ongoing disease outbreaks (Farms A to D). Jejunal contents from hens from three of the four affected flocks were also culture-positive (8/18 samples analysed). All samples from laying hens from the two clinically healthy flocks (Farms E and F) were culturenegative. Mice Mice were present on all farms according to the farmers, but they were only caught 3 weeks after the visit on Farm D and 1 week after the visit on Farm E. No gross lesions were found in any of the mice at necropsy and all organ samples from mice were culture-negative (Table 1). Samples from the poultry house environment E. rhusiopathiae was isolated from manure, dust and swab samples from nipple drinkers on farms with ongoing outbreaks, but not from the two clinically healthy flocks (Table 1). Selective culture conditions (KN-agar) produced more isolates from manure and dust samples than nonselective culture (Table 2). Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Using the direct transfer method, the 21 isolates previously confirmed as E. rhusiopathiae by biochemical tests were identified as E. rhusiopathiae by MALDI-TOF MS, but the scores ranged from 1.718 to 2.38
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H. Eriksson et al. Table 2.
Comparison of bacteriological culture and species-specific PCR for E. rhusiopathiae.
Horse blood agar (non-selective) Sample
Culture
Nipple drinkers (n = 30) Manure (n = 19) Dust from exhaust fans (n = 15) Laying hen—jejunum (n = 22)
4/30 1/19 0/15 8/22
KN-agar (selective)
PCR a
4/19 2/19 0/15 8/19a
Culture
PCR
4/30 5/19 2/15 ND
4/11a 7/19 3/14a ND
(16 isolates with scores over 2.0 and five isolates with scores below 2.0). After investigations using the formic acid method, two of the five isolates with low scores were identified as E. rhusiopathiae with scores of 2.06 and 2.179. Renewed preparation by ethanol–formic acid extraction was performed on the remaining three isolates, which were again identified as E. rhusiopathiae with scores of 2.138, 2.232 and 1.99. The reference strains were both identified as E. rhusiopathiae with scores of 2.219 (E. rhusiopathiae CCUG 221T) and 2.275 (E. tonsillarum CCUG 31352T) respectively.
products was 890 nucleotides (GenBank accession numbers KC986835 to 986840). The five nucleotide sequences (Farms A and C) were identical to each other but differed in three positions from the sixth sequence (Farm B) and in four positions from the sequence of the target gene of the type strain E. rhusiopathiae Fujisawa (GenBank accession number AP012027, gene ERH_0856). One of the sequences (isolate from manure on Farm B) differed in one position from the type strain. The five amino acid sequences differed in two positions from the type strain, whereas that from Farm B was identical to the type strain.
Polymerase chain reaction analysis. Specificity control produced PCR fragments of the expected size for all E. rhusiopathiae isolates but not for the other bacterial species investigated, including the presumed E. tonsillarum isolates (Eriksson et al., 2009). All samples (n = 193, excluding those from the spleen in Table 1) were cultured as described above and the plates (horse blood agar and KNagar) with visible bacterial growth (n = 313) were used for DNA preparation. Among these plates, 28 were positive by PCR, corresponding to 22 samples. All agar plates with confirmed growth of E. rhusiopathiae (by biochemical tests) were PCR-positive but E. rhusiopathiae isolates were only obtained from 19 out of the 22 PCR-positive samples. Table 2 presents a comparison of results of culture and PCR analysis. The PCR on manure and dust samples was positive to a higher extent with templates from KN-agar plates than from horse blood agar plates. Sequence analysis of the six PCR amplicons confirmed their identity. The length of the six deposited PCR
Pulsed-field gel electrophoresis analysis. In total, 93 E. rhusiopathiae isolates were analysed by PFGE; 24 isolates from the spleen (one isolate/hen), 30 isolates from the jejunal contents from laying hens (one to five isolates/ hen) and 39 isolates from environmental samples. Seven different PFGE types were observed among the isolates from the four farms (Figure 1). Different PFGE types were obtained from all four farms. On Farm A, all isolates from laying hens showed an identical banding pattern except one from a spleen, which differed by a single band. For the three farms where samples from laying hens and the environment could be compared (Farms B, C and D), identical banding patterns were observed in samples from laying hens and the house environment on Farms C and D, whereas on Farm B different banding patterns were obtained (Figure 1). Furthermore, on Farms A and C, which were represented by isolates obtained in 2011 and in 2012, different banding patterns were identified between years.
100
80
Degree of similarity (%) 60
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Cultures were performed using non-selective horse blood agar plates and horse blood agar plates supplemented with kanamycin (400 µg/ml) and neomycin (50 µg/ml) (KN-agar). PCR templates were prepared from all plates with visible growth (n = 313) (spleen samples not included), and 28 templates (from 14 horse blood agar and 14 KN-agar plates) were PCR-positive. Only sample categories with confirmed growth of E. rhusiopathiae by biochemical tests are included. ND, not determined. a Fewer samples were analysed by PCR than by culture because PCR was only performed from agar plates with visible growth.
Farm A / 2011 Laying hen Farm A / 2011 Laying hen Farm C / 2012 Laying hen Farm B / 2011 Laying hen Farm C / 2011 Laying hen & Manure Farm A / 2012 Laying hen Farm D / 2012 Laying hen, Manure, Dust & Nipple drinker Farm B / 2011 Manure Figure 1. The seven PFGE types obtained (farm designation/year of outbreak and isolation source) based on original comparison of 93 E. rhusiopathiae isolates from laying hens and the poultry house environment of four organic laying hen flocks on different farms. Banding patterns produced by SmaI restriction digestion provided the data for the dendrogram.
E. rhusiopathiae in the house environment
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Discussion Knowledge of sources of E. rhusiopathiae infection and transmission routes in laying hen flocks is of the utmost importance to prevent and manage outbreaks but few epidemiologic data are currently available. To our knowledge, this is the first study to investigate intestinal samples and samples from the house environment from laying hen farms for the presence of E. rhusiopathiae during an ongoing outbreak. In previous studies, the bacterium has been detected in manure and soil from pig and cattle farms (Wood & Packer, 1972; Nørrung et al., 1987). In addition, Bender et al. (2010) isolated the bacterium from nipple drinkers, walls, fans, feed and central line water in affected pig herds. The detection in this study of E. rhusiopathiae in manure from three of the four affected chicken flocks and in dust and on water nipples from one of the farms suggests that the bacterium is widespread in the environment during ongoing outbreaks. The dust samples that were positive for E. rhusiopathiae were collected inside the exhaust fans, so the possibility that contaminated dust could leave the house via this route must be taken into consideration. However, whether the bacteria can survive in dust in the longer term and whether dust can act as a source of transmission remain to be determined. Based on our results, the risk for transmission with manure must be considered greater than that for dust. E. rhusiopathiae was detected in eight of 18 jejunal samples from infected flocks when cultured on blood agar, which indicates a high number of bacteria in the contents of the jejunum. Furthermore, the bacteria were detected in the manure heaps from three out of four affected flocks. In 1965, Ostašev showed that E. rhusiopathiae could survive for more than 290 days when inoculated into pig slurry and stored at 18 to 20°C (reviewed by Mitscherlich & Marth, 1984). However, the composition of manure differs between species and the survival ability of E. rhusiopathiae in poultry manure requires further study. Until such knowledge becomes available, special precautions are needed when handling manure from an infected flock. The manure from affected flocks should preferably not be stored near poultry houses, as vectors may potentially transmit the infection from the heap. Measures should also be taken to minimize the risk of exposure to other animals, including wildlife, especially when manure from an infected flock is spread on arable land. Manure should therefore be ploughed down immediately and should not be used on pasture. The fact that the isolates from manure on Farm B in this study were of a different PFGE type than the isolates from the organs of the hens on the same farm is interesting. This could be explained by the limited number of samples or contamination of the manure heap from an external source. Another possibility is that the flock was infected by two distinctly different PFGE types, but one of these could have been introduced earlier or may be better adapted for in vitro growth (Eriksson et al., 2010). The possibility of a carrier state in chickens must also be considered. Pigs have been shown to be carriers of E. rhusiopathiae (Opriessnig & Wood, 2012), but studies in chickens are scarce. Nakazawa et al. (2008) isolated E. rhusiopathiae from broiler chickens in an abattoir and suggested that chickens may be a potential reservoir. However, in our study the samples from the two clinically healthy flocks all tested negative, which suggests that the bacterium is not normally present, is only present at low numbers or is only present in few individuals.
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The routes of transmission of E. rhusiopathiae within a poultry flock have not been determined fully, but in previous studies chickens have been infected both by oral and intramuscular experimental challenge (Takahashi et al., 1994; Mazaheri et al., 2005). It has been suggested that the bacteria gain entry through wounds and damaged mucous membranes and that behaviours such as cannibalism and feather pecking in flocks increase transmission (Bricker & Saif, 2013). Proper culling practices, including prompt removal of dead and diseased birds from the flock several times a day, are therefore important to limit the spread and contamination rate of E. rhusiopathiae in the poultry house environment. Our finding that the bacterium is present in jejunal contents and manure from affected flocks suggests that a faecal–oral transmission route could be important in field outbreaks, but further research is needed. This finding also indirectly supports a previous report of higher risks of an outbreak in laying hen flocks in litter-based systems than those in cages (Eriksson et al., 2013), because caged chickens are less exposed to faeces than chickens kept on litter or as free range. Despite vaccination of pullets on arrival at the farms, outbreaks occurred in the consecutive flocks on Farms A and C in 2012. Previous experiences from Sweden have shown that a single vaccination at placement of pullets protects the flock from erysipelas for the duration of the production period (approximately 50 to 60 weeks). However, this particular vaccine was developed for use in turkeys and the manufacturer recommends that birds should be vaccinated twice. For practical reasons this is difficult and laborious in pullets, but should be considered to avoid future outbreaks. Along with aspects of vaccine handling such as transport, storage and administration, there is a potential risk that the outbreaks in the consecutive flocks were caused by strains to which the vaccine offered no or only limited protection. Studies in non-avian species have shown a variable degree of cross-protection between serotypes and, more recently, also between the surface protective antigen types (Sawada & Takahashi, 1987; Ingebritson et al., 2010). The isolates obtained from the flocks in this study were not further investigated. Crossprotection studies in poultry should preferably be performed. Interestingly, the outbreaks on Farms A and C in 2012 were caused by different PFGE types to those in 2011. In a previous study we found that isolates from an organic laying hen farm affected in 1998, 1999, 2001 and 2004 were of identical (in 2001 and 2004) or dissimilar (in 1998 and 1999) PFGE types (Eriksson et al., 2009). The fact that different PFGE types were found in some of these cases suggests that the birds were infected by an external source rather than by residual bacteria in the house environment. In the case of identical PFGE types in consecutive flocks, re-introduction of the same bacterial genotype from a common source in the external environment should also be considered. The incidence of E. rhusiopathiae in wild fauna and its ability to survive in the environment, including outside pens, are important aspects to address in the future. Until further knowledge has been obtained, a high level of biosecurity, including effective hygiene barriers, is recommended. In addition, farmers should ensure that fences effectively keep wild animals outside the pen. Interestingly, during a survey in this study (data not shown) several farmers reported problems with wild animals (e.g. wild boars) around, and sometimes even inside, the outside pen.
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H. Eriksson et al.
With the exception of spleen samples from laying hens in affected flocks, a relatively small proportion of the samples in this study was positive by culture combined with biochemical identification and/or PCR from agar plate cultures. This may have been due to low sensitivity of the diagnostic methods used, suboptimal sampling procedures and/or a negative effect of frozen storage. In fact, our results suggest that the bacterium does not survive long-term storage in frozen samples from the poultry house environment. However, in a previous study on environmental samples from pig farms affected by erysipelas, E. rhusiopathiae was successfully isolated from samples stored at −20°C when cultured within 3 months of collection (Bender et al., 2010). Further, survivability of the bacterium during frozen storage needs to be further investigated. The PCR from agar plate cultures seemed slightly more sensitive than culture combined with biochemical identification, most probably because E. rhusiopathiae grows in very small colonies, which can be difficult to detect on agar plates with other more profusely growing bacteria. The negative sampling results from mice and arthropods indicate that these populations were not heavily infected. However, early experimental studies have indicated that E. rhusiopathiae may be transmitted by flies and mosquitoes (Wellmann 1950, 1955), although no study of insects on a farm during an outbreak could be found in the literature. Neither could any report on natural E. rhusiopathiae infection in mice be found; however, mice are sometimes used as a challenge model for E. rhusiopathiae infection. In our study, mice were only caught on one of the infected farms 3 weeks after the visit. Further studies with extensive sampling of arthropods and mice caught during ongoing outbreaks are needed. Finally, our investigation into the use of MALDI-TOF MS as a method for species confirmation of E. rhusiopathiae was inconclusive. However, based on the fact that the reference strain of E. tonsillarum (CCUG 31352T) was typed as E. rhusiopathiae with a high score, further studies are clearly needed. The problem of misidentification of E. tonsillarum in this study may be due to the fact that MALDI-TOF MS species identification is based on investigations of the ribosomal protein spectra and that E. rhusiopathiae and E. tonsillarum are very similar in their 16S rRNA sequences (Eriksson et al., 2009; Wieser et al., 2012). The number of available spectra in the database was also very limited. If MALDI-TOF MS can replace conventional biochemical typing, this would shorten the time to diagnosis and would be of importance since the farmer needs to take appropriate measures as soon as possible to reduce bird suffering. In conclusion, we have shown that E. rhusiopathiae is widespread in the poultry house environment during ongoing erysipelas outbreaks in organic laying hen flocks.
Acknowledgements The authors wish to thank the six farmers for providing data, time and help during sampling, Ricardo Feinstein for performing necropsies of the mice, Sigbrit Mattsson for helping with sampling on the farms, Helena Ljung, Anna Eriksson and Lena Lundgren for assistance with laboratory work, and Anna Aspán for Figure 1.
Funding The Ekhaga foundation is acknowledged for funding (under Grant 2010-64) sampling and bacteriological analyses, and SLU EkoForsk is acknowledged for funding PCR analyses.
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Widgren, S., Eriksson, E., Aspán, A., Emanuelson, U., Alenius, S. & Lindberg, A. (2013). Environmental sampling for evaluating verotoxigenic Escherichia coli O157: H7 status in dairy cattle herds. Journal of Veterinary Diagnostic Investigation, 25, 189–198. Wieser, A., Schneider, L., Jung, J. & Schubert S. (2012). MALDI-TOF MS in microbiological diagnostics–identification of microorganisms and beyond (mini review). Applied Microbiology & Biotechnology, 93, 965–974. Wood, R.L. (1965). A selective liquid medium utilizing antibiotics for isolation of Erysipelothrix insidiosa. The American Journal of Veterinary Research, 26, 1303–1308. Wood, R.L. & Packer, R.A. (1972). Isolation of Erysipelothrix rhusiopathiae from soil and manure of swine-raising premises. The American Journal of Veterinary Research, 33, 1611–1620.
