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. 2019 Aug 14;5(4):545–555. doi: 10.1002/vms3.190

Seroprevalence and characterization of Brucella species in cattle slaughtered at Gauteng abattoirs, South Africa

Francis B Kolo 1,✉, Abiodun A Adesiyun 2,3, Folorunso O Fasina 1, Charles T Katsande 4, Banenat B Dogonyaro 1, Andrew Potts 5, Itumeleng Matle 5, Awoke K Gelaw 5, Henriette van Heerden 1
PMCID: PMC6868451  PMID: 31414558

Abstract

Background

Brucellosis is an infectious and contagious zoonotic bacterial disease of both humans and animals. In developing countries where brucellosis is endemic, baseline data on the prevalence of brucellosis, using abattoir facilities, is important.

Objectives

The aim of this study was to determine the seroprevalence of antibodies against Brucella in slaughter cattle at Gauteng province, South Africa and to characterize isolates of Brucella spp.

Methods

In this cross‐sectional study, un‐clotted blood samples with corresponding organ tissue samples were collected from slaughtered cattle. Serological [Rose Bengal test (RBT), complement fixation test (CFT) and indirect ELISA (iELISA)], molecular (PCR) and bacteriological methods were used to detect Brucella antibodies and Brucella spp. from 200 slaughtered cattle in 14 abattoirs.

Results

The RBT revealed a seroprevalence of brucellosis as 11.0% (22 of 200) and iELISA confirmed 5.5% (11 of 200). The estimated seroprevalence from RBT and iELISA was 5.5% while RBT and CFT was 2.0% (4 of 200). Brucella melitensis (n = 6) and B. abortus (n = 5) were isolated from 11 cattle tissues (5.5%) as confirmed to species level with AMOS PCR and differentiated from vaccine strains with Bruce‐ladder PCR. Seven of the 11 isolates originated from seropositive cattle of which five were biotyped as B. abortus bv 1 (n = 2) and B. melitensis bv 2 (n = 1) and B. melitensis bv 3 (n = 2).

Conclusions

This is the first documentation of B. melitensis in cattle in South Africa. The zoonotic risk of brucellosis posed by Brucella‐infected slaughter cattle to abattoir workers and consumers of improperly cooked beef cannot be ignored.

Keywords: abattoirs, Brucella abortus, Brucella melitensis, Brucellosis, cattle, seroprevalence

Seroprevalence and characterization of brucellosis in cattle slaughtered at Gauteng abattoirs, South Africa.

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1. INTRODUCTION

Brucellosis is an infectious and contagious zoonotic bacterial disease of humans and a wide range of domestic animals and wildlife, particularly the ruminants (Corbel, 2006; Radostits, Gay, Hinchcliff, & Constable, 2006; Smirnova et al., 2013) and some marine animals (Foster, Osterman, Godfroid, Jacques, & Cloeckaert, 2007; Scholz & Vergnaud, 2013). The Brucella species are Gram‐negative, non‐capsulated, facultative intracellular, non‐spore forming, cocco‐bacilli bacteria (Godfroid, 2012; Seleem, Boyle, & Sriranganathan, 2010; Smirnova et al., 2013). Brucella spp. infecting farm animals include B. abortus, B. melitensis, B. suis and B. ovis (Godfroid, Nielsen, & Saegerman, 2010; Smirnova et al., 2013). The Brucella species are known to have host preferences, although there could be cross‐infection with other hosts. Brucella abortus has a host preference for the cattle but it can cause infection in other hosts including humans (undulant fever). Brucella melitensis, has a host preference for sheep and goats and it is the most pathogenic of the Brucella spp. that causes infection in humans (Malta fever). Brucella canis and B. suis has host preferences for dogs and pigs, respectively, and can cause brucellosis in humans (Alton, 1990; Carmichael, 1990; Godfroid et al., 2005; Pappas, 2013). Brucellosis has been eradicated or well controlled in developed countries (Pappas, Papadimitriou, Akritidis, Christou, & Tsianos, 2006). However, in many of the low and middle income countries (LMICs) such as in Africa, South and Central America, Middle East, Asia, Mediterranean Basin and the Caribbean, brucellosis is still common and high in occurrence both in the animal and human populations (Adesiyun & Cazabon, 1996; Godfroid et al., 2005; Olsen & Palmer, 2014; Pappas et al., 2006). Brucellosis may have existed in South Africa as an ancient disease as suggested by a paleopathological analysis study on the fossil of the late Pliocene hominin species (D'Anastasio, Zipfel, Moggi‐Cecchi, Stanyon, & Capasso, 2009). It had been suspected in 1898 that goats may have been the source of suspected cases of undulant fever which was also called “camp fever” in 40 patients around the Kimberley area of South Africa, an area where diamond mines were operating (Strachan, 1932; Van Drimmelen, 1949). Serological diagnosis, as well as cultural isolation, which identified B. melitensis, were relied upon for diagnosis from 1902 to 1911 (Strachan, 1932; Zammit, 1905) from samples including human blood, goat serum and milk samples in South Africa (Strachan, 1932). Brucella melitensis outbreaks have been documented in sheep in 1965 (Van Drimmelen, 1965), in goats in 1989 that was identified as B. melitensis bv 1 (Ribeiro, Herr, & Chaparro, 1990), 1994 (Reichel, Nel, Emslie, & Bishop, 1996), 2000 (Emslie & Nel, 2002), 2007 and 2015 (DAFF, 2015). Brucella abortus infection in South Africa was reported in 1913, when contagious abortion was observed to spread across the country in cattle (Van Drimmelen, 1949). In South Africa, B. abortus bv 1 predominantly cause infection in cattle (90%) and B. abortus bv 2 to a lesser extent (Godfroid, Bishop, Bosman & Herr, 2004). Mauff (1980) reported fives cases of brucellosis associated with by‐products (condemned meat and unborn calves) at a new abattoir in South Africa. These were confirmed cases based on serological tests and culture and it was reported that the affected individuals did not wear any protective clothing at the by‐product facility (Mauff, 1980). Despite this risk and threat of exposure of humans to brucellosis at abattoirs, the only published report of an abattoir‐based study on bovine brucellosis was in 1984 where a prevalence rate of 1.5% was reported for cattle sampled at Cato Ridge abattoir in KwaZulu‐Natal province (Bishop, 1984). In the Eastern Cape province, a 9.2% prevalence rate of B. abortus (of which 0.8% B. abortus S19 vaccine strain) was isolated from cattle, 2.9% B. melitensis from sheep and 6.3% B. melitensis from goats using different samples (blood, milk and lymph nodes) followed by species specific confirmation using PCR (Caine, Nwodo, Okoh, & Green, 2017).

In South Africa, brucellosis is a reportable disease. Control measures have been instituted to prevent the spread of brucellosis in the country with the focus mainly on bovine brucellosis through the animal diseases Act 35 of 1984 and the bovine brucellosis scheme (R.2483 of 9 Dec 1988) which is regulated by the Director of Animal Health at the Department of Agriculture, Forestry and Fisheries (DAFF). Currently the testing scheme for bovine brucellosis (established under section 10 of the Animal Disease act 35) is compulsory for only high‐risk herds that have been confirmed or suspected of infection using Rose Bengal test (RBT) and the complement fixation test (CFT). Entering the brucellosis testing scheme is voluntary for all other bovine herds and livestock owners. Vaccination is practised in South Africa according to the stipulated standards with mainly B. abortus S19 and to a lesser extent B. abortus RB 51 in cattle while B. melitensis Rev 1 is used in sheep and goats (OIE, 2016). These control measures amongst others are instituted to prevent a spillover of the disease to other domestic animals and wildlife in areas close to the wildlife parks (Simpson et al., 2018).

In South Africa, an estimation of over 3,476,000 of cattle were slaughtered from September 2015 to August 2016 in the abattoirs (DAFF, 2016). These abattoir facilities can also be used to monitor disease control policies, detect newly introduced disease agents and to assess intervention programmes, such as brucellosis vaccination, and most importantly, abattoir survey may also facilitate early intervention to mitigate the epidemic loss of animals (Alton, Pearl, Bateman, McNab, & Berke, 2015; Fasina et al., 2015; Kaneene, Miller, & Meyer, 2006). As such, an abattoir surveillance study on brucellosis can generate baseline data on the occurrence of the disease among the animal population, especially when the animals come from various farms to be processed into wholesome meat products for human consumption (Alton et al., 2015; Fasina et al., 2015; Kaneene et al., 2006). However, abattoir data may not produce reliable prevalence estimates because the population of slaughtered cattle tends not to correctly represent the target population which may affect the validity of results from such facilities. Animals and carcasses of Brucella‐infected slaughtered animals can be a source of infection to susceptible abattoir workers, as these workers may be exposed to infection through direct contact with infected animal's secretions or blood, or indirectly through the consumption of raw meat or undercooked meat (Corbel, 2006). It has been documented in the literature that the risk of carcass contamination by bacteria, including Brucella spp., increases with the throughput, i.e. the number of animals slaughtered (Sadler, 1960). Due to the predominant voluntary nature of the brucellosis testing scheme in South Africa, known and unknown brucellosis infected cattle can be slaughtered at abattoirs. The aim of this study was to determine the seroprevalence of antibodies against Brucella and characterize Brucella spp. from slaughtered cattle from abattoirs.

2. MATERIALS AND METHODS

2.1. Study area, study design and sample size

A cross‐sectional study was conducted to determine the seroprevalence of brucellosis and to detect, isolate and characterize Brucella spp. in cattle slaughtered at the Gauteng province abattoirs from April 2016 to April 2017.

The study area was the Gauteng Province of South Africa. The Province is the smallest province in South Africa with approximately 1.5% (surface area of 1,219,602 km2) of the land area, yet it remains most populated, accounting for approximately 23.7% of the country's population. Although the most recent estimated number of cattle in the Gauteng province in May 2018 was 246,395, it is known that a large number of cattle from other provinces move to Gauteng on regular basis (DAFF, 2018).

Fourteen abattoirs in the Gauteng Province (Figure 1), which were operational and consented to participate in the study, were randomly selected from 28 operational abattoirs identified. These abattoirs were categorized into high throughput (n = 7) slaughtering more than 20 cattle per day and low throughput (n = 7) where 20 or less cattle are slaughtered daily. Among the selected abattoirs for the study, 13 operated as multi‐species and 1 was mono‐species facilities.

Figure 1.

Figure 1

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Locations of abattoirs sampled in the Gauteng Province of South Africa

2.2. Sample size and collection

In this cross‐sectional study, a systematic simple random sampling method was used to determine the sample size and power. Sample size was estimated at 50.0% expected frequency, with 5.0% acceptable margin of error and a design effect of 1 and clusters equals to 1 in Epi‐Info 7 version 10. At a 95.0% confidence level, the sample size of 200 animals was achieved. A systematic sampling method was used at selected abattoirs. For all selected animals, blood samples were collected as follows:

Unclotted blood was collected from 200 cattle at the point of slaughter using sterile 50 ml cups with approximately 5 ml of the blood aliquoted into yellow‐capped vacutainer tubes. Corresponding tissue samples comprising of lymph nodes, spleen and liver were collected from each of the cattle. The lymph nodes of each animal were pooled and comprised the retropharyngeal, parotid, submandibular and mesenteric lymph nodes. The excised tissues were processed according to set laboratory protocols in a bio‐safety laboratory level 2 hood. The homogenized tissues were used for DNA extraction (PCR) and bacterial isolation. Demographic data comprising of the animal species, breed, sex, age, the farm origin and the identity of the abattoir where the animal was slaughtered were collected for each of the animals slaughtered. The age of the animals was determined using the dental formula as described by Eubanks (2012). The vaccination status of the animals could not be ascertained.

2.3. Serological test methods on serum samples

Serological procedures were carried out using three different procedures including the RBT, the CFT and the indirect enzyme‐linked immunosorbent assay (iELISA). For this study, because RBT is unable to differentiate between antibodies produced in response to vaccination or natural exposure, its results were not used as stand‐alone percentage estimates but only as part of a serial test in a combination with either CFT or iELISA results.

Rose Bengal test was conducted as described by OIE (2009). The diagnostic sensitivity and specificity for the RBT have earlier been confirmed to be 100.0% and 75.0%, respectively, based on previous validation studies (Nielsen et al., 2005; Stemshorn et al., 1985).

CFT was performed on the cattle sera at the Agricultural Research Council‐ Onderstepoort Veterinary Research (ARC‐OVR) laboratory, South Africa using the OIE protocol (OIE, 2009). The cut‐off value for this test was ≥30 IU/ml as an indication of infection and the obtained values were compared with the positive and negative controls. The IDEXX brucellosis serum ×2 ELISA Test kit from Pourquier®, IDEXX, Switzerland were used according to the manufacturer's instruction. The cut‐off value for determination of antibody‐positive status in cattle recommended by IDEXX is 80.0%.

2.4. Bacterial isolation from tissue samples

Homogenate (200 μl) from each tissue (lymph nodes, spleen and liver) was inoculated onto Farrell's and modified CITA media, respectively, and incubated at 37°C with 5.0% CO2. Plates were observed for bacterial colony growth for 10 days. Brucella organisms were identified presumptively by morphology using the Stamp's modified Ziehl‐Neelsen staining method (OIE, 2009). Morphologically identified Brucella colonies were purified by sub‐culturing on the sheep blood agar media. Mixed or contaminated cultures were subjected to serial dilution 1:1,000 using buffered peptone water and inoculated onto the selective media until purified colonies were obtained. Pure Brucella cultures were biotyped at ARC‐OVR, South Africa according to the methods described by Ribeiro and Herr (1990).

2.5. Molecular detection

Genomic DNA was extracted from cultures of each homogenized tissue (lymph nodes, spleen and liver) and thereafter, the cells of Brucella cultures were purified using the set protocol according to Isolate II Genomic DNA kit by Bioline (South Africa). Detection of DNA and isolation procedure for Brucella spp. isolation was performed on all animals, regardless of their serological status. Genus‐specific 16S‐23S rRNA interspacer region (ITS) PCR was used to amplify Brucella region and B. abortus strain 544 and B. melitensis Rev 1 served as positive controls as described by Keid et al. (2007) for detection Brucella DNA in tissue samples (lymph nodes, spleen and liver). The process amplified a 214 bp fragment using primers (ITS66: ACATAGATCGCAGGCCAGTCA and ITS279: AGATACCGACGCAAACGCTAC). Primers were used at a final concentration of 0.2 μM with 1× DreamTaq Green PCR Master Mix (ThermoFisher Scientific, South Africa) and 2 μl DNA in a 15 μl PCR reaction. The PCR cycling condition consisted of 95°C for 3 min, followed by 35 cycles at 95°C for 1 min, 60°C for 2 min, 72°C for 2 min and a final extension at 72°C for 5 min. PCR products were analysed by electrophoresis using a 2.0% agarose gel stained with ethidium bromide and viewed under UV light.

The multiplex AMOS PCR assay that identifies and differentiates B. abortus, B. melitensis, B. ovis and B. suis was conducted as described (Bricker & Halling, 1994; Weiner, Iwaniak, & Szulowski, 2011) using DNA extraction from cultures. Four species‐specific forward primers were used at a final concentration of 0.1 μM with 0.2 μM reverse primer IS711 (Table 1) with 1× MyTaq™ Red PCR Mix (Bioline, South Africa) and 2 μl of template DNA in 25 μl PCR reaction. PCR cycling condition was initial denaturation at 95°C for 5 min followed by 35 cycles at 95°C for 1 min, 55.5°C for 2 min, 72°C for 2 min and a final extension step at 72°C for 10 min. PCR products were analysed by electrophoresis using a 2.0% agarose gel stained with ethidium bromide and viewed under UV light.

Table 1.

Sequences and characteristics of the oligonucleotide primers used for different Brucella species in the AMOS PCR assay

PCR name Primer name Sequence (5′‐ 3′) DNA Targets Amplicon (bp) Concentration (μM) Reference
AMOS B. abortus GAC GAA CGG AAT TTT TCC AAT CCC IS711 498 0.1 (Bricker, Ewalt, Olsen, & Jensen, 2003; Bricker & Halling, 1994)
B. melitensis AAA TCG CGT CCT TGC TGG TCT GA 731 0.1
B. ovis CGG GTT CTG GCA CCA TCG TCG GG 976 0.1
B. suis GCG CGG TTT TCT GAA GGT GGT TCA 285 0.1
IS711 TGC CGA TCA CTT AAG GGC CTT CAT   0.2
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A multiplex Bruce‐ladder PCR assay to identify and differentiate between vaccine strains and field isolates of Brucella spp. was conducted as described (García‐Yoldi et al., 2006; Lopez‐Goñi et al., 2008; Weiner et al., 2011) (Table 2). Eight species‐specific forward and reverse primers were used at a final concentration of 6.25 μM with 1× MyTaq™ Red PCR Mix (Bioline, South Africa) and 2 μl of template DNA in a 25 μl PCR reaction. The PCR cycling condition included an initial denaturation cycle at 95°C for 5 min followed by 25 cycles at 95°C for 30 s, at 64°C for 45 s, and at 72°C for 3 min and a final extension step at 72°C for 10 min. PCR products were analysed by electrophoresis using a 2.0% agarose gel stained with ethidium bromide and viewed under UV light.

Table 2.

Sequences and characteristics of the Bruce‐ladder PCR assay primers used in the study

PCR name Primer name Sequence (5′‐ 3′) DNA targets Amplicon (bp) Concentration (μM) Reference
Bruce‐ladder BMEI0998f ATC CTA TTG CCC CGA TAA GG wboA 1682 6.25 (Lopez‐Goñi et al., 2008)
BMEI0997r GCT TCG CAT TTT CAC TGT AGC
BMEI0535f GCG CAT TCT TCG GTT ATG AA bp26 450 6.25
BMEI0536r CGC AGG CGA AAA CAG CTA TAA
BMEII0843f TTT ACA CAG GCA ATC CAG CA omp31 1071 6.25
BMEII0844r GCG TCC AGT TGT TGT TGA TG
BMEI1436r ACG CAG ACG ACC TTC GGT AT Deacetylase 794 6.25
BMEI1435r TTT ATC CAT CGC CCT GTC AC
BMEII0428f GCC GCT ATT ATG TGG ACT GG eryC 587 6.25
BMEII0428r AAT GAC TTC ACG GTC GTTCG
BR0953f GGA ACA CTA CGC CAC CTT GT ABC 272 6.25
BR0953r GAT GGA GCA AAC GCT GAA G Transporter
BMEI0752f CAG GCA AAC CCT CAG AAG C rpsL 218 6.25
BMEI0752r GAT GTG GTA ACG CAC ACC AA
BMEII0987f CGC AGA CAG TGA CCA TCA AA CRP 152 6.25
BMEII0987r GTA TTC AGC CCC CGT TAC CT Regulator
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2.6. Statistical analysis

Data collected were managed using the Microsoft Excel version 2007. The R software (RCoreTeam, 2013) was used to analyse the data and to conduct descriptive analysis, and Epi‐Info 7 version 10 was used to conduct analyses of frequency with 95% confidence interval, and calculated odds ratio and Chi‐Square tests to examine differences between groups.

Animal prevalence was determined by the number of positive animals divided by the total number of animals sampled.

2.7. Ethical approval

Ethics approval for the study was obtained from ARC‐OVI Animals Ethics Committee (AEC12‐16), University of Pretoria Animal ethics Committee (V089‐16). Section 20 approval was granted according to Act 35 of 1984 by the Directorate of Animal Health, South Africa.

3. RESULTS

Out of the cattle samples tested, 57.5% (115/200) were from high throughput (HT) abattoirs, and 42.5% (85/200) originated from low throughput (LT) abattoirs. Of the 200 heads of cattle, 41.0% (82/200) were female while 59.0% (118/200) were male. The distribution of the cattle stratified by age was 92.0% (184/200) for adult and 8.0% (16/200) for young cattle; the distribution stratified by breed was Bonsmara, 69.5% (139/200), Nguni, 16.5% (33/200), Brahman, 5.5% (11/200), Jersey, 5.0% (10/200) and Holstein, 3.5% (7/200).

Among the 200 cattle tested, 11.0% (22/200) were positive on RBT, 5.5% (11/200) were positive on iELISA while only four of the 22 (18.2%) RBT positive sera were confirmed positive on CFT (2.0%, 4/200). The estimated seroprevalence from RBT and confirmed by iELISA was 5.5%. While the seroprevalence from RBT confirmed with CFT was 2.0%. Brucella DNA detection rate from the screened cattle tissues by ITS‐PCR was 12.5% (25/200) (Table 3). Brucella spp. isolation rate was 5.5% (11/200).

Table 3.

Prevalence and risk of Brucella spp. from abattoirs samples by seropositivity, 16S‐23S rRNA interspacer region (ITS) PCR and the isolation rate

Animal demography Total Serology positives †(%) ‡OR (95% § CI) Chi square p‐value
Seropositivity to Brucella spp. stratified by sex, age of animals and type of abattoirs sampled
Sex   (RBT)      
Female 82 16 (19.5) 4.5 (1.67–12.13) 8.9 0.00
Male 118 6 (5.1)      
Sex   (ELISA)      
Female 74 8 (9.8) 0.2 (0.06–0.94) 3.5 0.05
Male 115 3 (2.5)      
Age   (RBT)      
Adult 184 18 (9.8) 0.3 (0.09–1.11) 2.1 0.15
Young 16 4 (25.0)      
Age   (ELISA)      
Adult 176 8 (4.4) 5.1 (1.20–21.5) 3.4 0.06
Young 13 3 (18.5)      
Animals positive by 16S‐23S rRNA interspacer region (ITS) PCR stratified by sex, age and type of abattoirs sampled
Sex
Female 82 17 (20.7) 3.60 (1.47–8.80) 7.4 0.01
Male 118 8 (6.8)      
Age
Adult 184 22 (12.0) 0.59 (0.15–2.23) 0.2 0.69
Young 16 3 (18.8)      
Abattoir type
High throughput 115 6 (5.2) 0.19 (0.07–0.51) 11.4 0.00
Low throughput 85 19 (22.4)      
Isolation rate by bacteriological method stratified by sex, age of animals and type of abattoirs sampled
Sex
Female 82 8 (9.8) 4.14 (1.06–16.12) 3.6 0.06
Male 118 3 (2.5)      
Age
Adult 184 9 (4.9) 0.36 (0.7–1.83) 0.5 0.48
Young 16 2 (12.2)      
Abattoir type
High throughput 115 1 (0.9) 0.06 (0.01–0.52) 9.2 0.00
Low throughput 85 10 (11.7)      
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† = percentage, ‡ = odds ratio, § = confidence interval, ¶ = polymerase chain reaction.

Of the 25 ITS‐PCR positives, 56.0% (14/25) were from seropositive cattle (RBT or ELISA), while 44.0% (11/25) were from seronegative cattle. The serology positivity, PCR detection and isolation rates among the cattle population according to sex and age with the chi‐square measure of association is shown in Table 3. Type of abattoir is an indication for carcass contamination that can be detected by molecular and bacteriological methods.

The distribution of cattle that tested seropositive by RBT and confirmed with iELISA according to breed was as follows: Bonsmara, 3.6% (5/139); Nguni, 9.1% (3/33); Holstein, 28.6% (2/7); Brahman, 9.1% (1/11) and Jersey, 0.0% (0/10). The differences were not statistically significant (p > 0.05). With the use of the molecular method, the distribution of positivity according to the breed was as follows: Bonsmara 10.8% (15/139), Nguni 12.1% (4/33), Holstein 42.8% (3/7), Brahman 27.3% (3/11) and Jersey 0.0% (0/10). The differences were not statistically significant (p > 0.05). Of the cattle tested with bacteriological methods, the distribution of isolation positivity according to the breed of the animals was as follows: Bonsmara, 4.3% (6/139), Nguni 3.0% (1/33), Holstein 42.8% (3/7), Brahman 9.1% (1/11) and Jersey 0.0% (0/10). The differences were not statistically significant (p > 0.05).

Of the tissue samples from the 200 slaughtered cattle tested using the ITS‐PCR for the detection of Brucella DNA, the frequencies of detection were 11.5% (23/200, 95%CI = 7.43–16.6) from the lymph nodes, 7.5% (15/200, 95%CI = 4.26–12.1) from the spleen and 7.0% (14/200, 95%CI = 3.88–11.5) from the liver. The detection rates from the three tissue samples among the 25 ITS‐PCR‐positive cattle are lymph nodes 92.0% (23/25), 64.0% (16/25) and 56.0% (14/25) for the lymph nodes, spleen and liver, respectively.

Of the 25 ITS‐PCR positive cattle, the frequency of detection was 68.0% (17/25, 95%CI = 46.5–85.1) in the females and 32.0% (8/25, 95%CI = 14.9–53.5) in the males. Of the 25 ITS‐PCR positive cattle, the distribution by age classification was 88.0% (22/25, 95%CI = 68.8–97.4) for adults and 12.0% (3/25, 95%CI = 2.5–31.2) for young cattle; and by abattoir types was 24.0% (6/25, 95%CI = 9.36–45.1) for HT abattoirs and 76.0% (19/25, 95%CI = 54.9–90.6) for LT abattoir.

Among the 23, 15 and 14 ITS‐PCR‐positive lymph nodes, spleen and liver tissues, respectively, the frequency of isolation was 26.1% (6/23) from the lymph nodes, 40.0% (6/15) from the spleen and 42.8% (6/14) from the liver samples. Of the total 25 PCR‐positive cattle, the frequency of Brucella spp. isolation was 44.0% (11/25, 95%CI = 24.4–65.1). The isolation rate from all the three tissue samples among the 11 culture‐positive cattle was 26.1% (6/23) for each tissue.

Of the 11 confirmed Brucella isolates from cattle by the “gold standard” method of culture and isolation, 63.6% (7/11) were from the confirmed iELISA‐positive cattle. The other Brucella isolates were from RBT‐positive cattle (n = 3) except one from a seronegative cow. AMOS‐PCR assay characterized the seven isolates from the iELISA‐positives as B. melitensis (four isolates) and B. abortus (three isolates). The remaining four isolates, three of which were RBT‐positives, were classified as B. melitensis (two) and B. abortus (one); while the only single isolate that was negative on serology was characterized as B. abortus. In total, the AMOS‐PCR characterized the 11 isolates as six B. melitensis and five B. abortus. Two of these isolates had amplification for both B. abortus and B. melitensis (Figure 2). All isolates were differentiated from the vaccine strains using the Bruce‐ladder PCR assay (Figure 3).

Figure 2.

Figure 2

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Gel electrophoresis of products from AMOS‐PCR of the IS711 gene using species‐specific primers. Lanes 1 and 2 show amplification products of 498 bp in length for Brucella abortus and Lanes 3 to 6 show amplification products of 710 bp for B. melitensis, while Lanes 4 and 6 show double amplification as a result of mixed infection of both B. abortus and B. melitensis, in the smaples. Lanes 7 and 8 show the positive and negative control, lane L shows 100 bp DNA ladder (Invitrogen, ThermoFisher® scientific, South Africa)

Figure 3.

Figure 3

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Gel electrophoresis of Bruce‐Ladder PCR amplification product using species specific primers. Lanes 1 and 2 show amplification product of B. abortus, lane 3 is B. abortus strain 544 (control), lanes 4 and 5 show amplification for B. melitensis (cattle) and lanes 6 and 7 show amplification for B. melitensis (sheep), lanes 8 and 9 show Rev 1 positive and negative controls, respectively, lane L shows 100 bp DNA ladder (Invitrogen, ThermoFisher® scientific, South Africa)

From the 11 Brucella culture‐positive cattle, the frequency of isolation of Brucella spp. by sex was 72.7% (8/11, 95%CI = 39.0–94.0) in the females and 27.3% (3/11, 95%CI = 6.02–61.0) in the males and by age was 81.8% (9/11, 95%CI = 48.2–97.7) in adults and 18.2% (2/11, 95%CI = 2.28–51.8) in the young cattle. The frequency of isolation was 9.1% (1/11, 95%CI = 0.23–41.3) and 90.9% (10/11, 95%CI = 58.7–99.9) in the HT and LT abattoirs respectively.

Five of the 11 Brucella isolates were purified and biotyped. Two were determined to be B. melitensis bv 3, one as B. melitensis bv 2 and the remaining two as B. abortus bv 1. The five Brucella isolates were from iELISA‐positive cattle.

4. DISCUSSION

In this study, antibodies as well as the pathogen, were detected by a polyphasic approach using serology (RBT, CFT and iELISA), PCR and culturing. The brucellosis estimated seroprevalence and culture prevalence (gold standard) among the slaughter cattle at the Gauteng abattoirs is at 5.5%. Brucella abortus bv 1 was isolated in this study, which was expected since B. abortus bv 1 is the most dominant biovar causing infection in cattle (Coetzer, Thomson, & Tustin, 1994). For the first time B. melitensis bv 2 and 3 were also isolated from cattle tissue. In the past B. melitensis bv 1 has been isolated from goats (Reichel et al., 1996) and humans (Wojno et al., 2016), while B. melitensis bv 3 was isolated from sable in 2007 and 2015 as documented by DAFF (2015) and van Heerden (unpublished results).

As mentioned above, the testing scheme for bovine brucellosis (established under section 10 of the Animal Disease act 35) is compulsory for only high‐risk herds, while entering the brucellosis testing scheme is voluntary for all other bovine herds and livestock owners in South Africa. Due to the predominant voluntary nature of the brucellosis testing scheme in South Africa, known and unknown brucellosis infected cattle can be slaughtered at abattoirs. Therefore, it is concerning that B. melitensis that has been detected in sheep and goats in South Africa have now been identified in cattle. This information should be taken into consideration in the revision of the brucellosis testing scheme that is currently ongoing (DAFF, 2017). This abattoir survey provided useful baseline data on the prevalence of brucellosis in the tested animal population within a region or the country.

The vaccination status of the slaughtered animals in this study was unknown. Vaccination of 3–8 months heifers with B. abortus S19 is compulsory. However, the percentage of cattle vaccinated with S19 is unknown as the vaccine producer of S19 (Onderstepoort Biological Products) only indicate overall total vaccine sold. There was also a period when B. abortus S19 vaccine was unavailable (DAFF, 2017). There is no compulsory testing to monitor B. melitensis in sheep and goats in South Africa (DAFF, 2017). Brucella melitensis outbreaks reported in South Africa in sheep and goats have mostly been noticed only when human brucellosis was detected in individuals associated with sheep and goats (Emslie & Nel, 2002).

The detection of B. abortus bv 1 has been documented to be the common species and biotype infecting the cattle population in South Africa (Coetzer et al., 1994; Gradwell, 1977) and in other countries especially in the Southern African region and the Caribbean (Fosgate et al., 2002; Matope, Bhebhe, Muma, Skjerve, & Djønne, 2009; Muendo et al., 2012). The isolation of B. melitensis from cattle in this study is however significant because it is considered the first documentation of B. melitensis in the cattle population in South Africa (Kolo et al., 2018). The occurrence of B. melitensis in cattle in this study may be as a result of rearing cattle together with sheep, goats and wildlife on the same farm or sharing of grazing land with sheep or goats (Radostits et al., 2006; Verger, Garin‐Bastuji, Grayon, & Mahé, 1989). This has huge implication to infection of other cattle in the herd and most importantly spill over to other farms and other species around that geographical areas (Godfroid et al., 2014). Brucella melitensis bv 2 and 3 have never been isolated in livestock in the country. This has significant implications for brucellosis control in South Africa as the brucellosis scheme only focus on testing bovine high risk herds. As this study report B. melitensis bv 2 and 3 to be present in cattle, there is a high probability of its presence in the sheep and goat populations. B. melitensis has been isolated from sable antelope (DAFF, 2015) and biotyped as B. melitensis bv 3 (unpublished results). Brucella melitensis was isolated and reported in humans in the Western Cape province in 2015 (Wojno et al., 2016) and biotyped as B. melitensis bv 1 (van Heerden unpublished data). Furthermore, Caine et al. (2017) detected B. melitensis from the tissues and blood samples of sheep at abattoirs in the Eastern Cape province, but was not biotyped.

In our study, the detection of co‐infection by B. melitensis and B. abortus is being reported for the first time in cattle in South Africa. AMOS‐PCR indicated a mixed infection of B. abortus and B. melitensis (Figure 2) in an impure culture, but Bruce‐ladder is unable to differentiate a mixed infection of both organisms since it will only show the profile of B. melitensis. Mixed infection is plausible if morphological identical Brucella colonies of different species grow on the same plate. In future studies, the impure culture could be purified by picking multiple colonies resulting from the same animal and speciating the pure cultures using PCR followed by biotyping.

The possibility exists that a higher frequency of false positives and tests occurs when RBT is used relative to other tests which led to the recommendation that the RBT should not be used as a stand‐alone test to determine the seroprevalence of brucellosis (OIE, 2009). Since the current study was unable to confirm the vaccination status of the slaughter animals tested, the possibility of vaccine‐induced antibody production, unrelated to natural exposure to Brucella spp. by both tests cannot be ignored. However, molecular (Bruce‐ladder PCR) and isolation methods were used to differentiate vaccine strains from our isolates (Figure 3) and no vaccine strain was identified.

It is of diagnostic significance that Brucella DNA was detected using ITS‐PCR in 11 seronegative cattle along with 14 seropositive cattle, an overall prevalence of 12.5% (25/200). The ITS‐PCR (genus‐specific) is a good option as a screening tool for Brucella DNA as the PCR is capable of detecting very small amount of DNA in the tissue samples even if as little as 3.8 fg of Brucella DNA mixed with 450 ng of host DNA (Keid et al., 2007). This is significant because the application of PCR to detect Brucella DNA in animal tissues can be used to diagnose brucellosis in immune‐compromised animals that are unable to seroconvert following exposure and infection, and in animals where the Brucella organism, which is an intra‐cellular organism, is localized in the tissues, such that serology may not be able to diagnose the infection from the serum samples (Radostits et al., 2006). However, the ITS‐PCR sensitivity and specificity should be validated in South Africa especially to ensure the specificity of this PCR, as it could react with Brucella‐like organisms in this region that could not be tested in the initial validation by Keid et al. (2007). The PCR method used in our study has proven to be fast, safe and does not require specialized laboratory as required in the bacteriological methods. Based on the detection rate of 92.0% in the lymph node tissues in our study, we recommend the pooling of the lymph nodes initially to assay for Brucella DNA. The strategic application of bacteriology, serology and PCR was shown to result in increased detection frequency of brucellosis in cattle.

5. RECOMMENDATIONS

It is recommended that abattoir‐based study be conducted at provincial and national levels, to ascertain the frequency of brucellosis detection in all the nine provinces of the country. A serological testing strategy based on RBT and iELISA may be used in series considering the fact that CFT is very laborious to conduct, not as robust as other serological tests, such as the iELISA and difficult to standardize (OIE, 2009; Padilla, Nielsen, Ernesto, & Ling, 2010). This will provide baseline data for policy makers to proffer solutions and interventions to mitigate the risk of economic losses to livestock in the country and to mitigate the public health impact of the disease in the human population. It should be mandatory for livestock farmers to permit trace back to their farms if slaughtered animals from their farms are seropositive for brucellosis. This can be achieved by enforcing existing regulations.

6. LIMITATIONS

The use of serology alone can only be presumptive because other pathogens can cross‐react to yield false‐positive results. In our study, many of the RBT‐positive sera were negative on iELISA. Major inferences could not be made on serological data generated because of the unavailability of important variables such as the vaccination history. This is because some serological tests are unable to distinguish between antibodies produced in response to exposure to vaccination or natural exposure to Brucella spp. Although the data generated from abattoir studies may be invaluable to surveillance (passive and active), the animals sampled may not be representative of the animal population in the province, based on the sample size used and the movement of livestock from other provinces in the country to Gauteng province where the study was conducted from other provinces in the country. A trace back study to the farm sources from where abattoir‐tested seropositive cattle originated was not conducted due to the lack of cooperation by the farmers who were apprehensive of potential quarantine, with associated economic losses, due to anticipated enforcement of control measures.

7. CONCLUSIONS

In conclusion, this study has provided data that B. melitensis may be circulating in the cattle population in South Africa. Brucella melitensis bv 2 and 3 have never been isolated in the cattle population in the country. This suggests a potential risk to abattoir workers and to a lesser extent to the consumers of raw or under cooked meat and meat products. The study also provided a current data on the prevalence (isolation, serology and PCR) of brucellosis in slaughter cattle in Gauteng province, South Africa. The seroprevalence of brucellosis detected in slaughter cattle in this study emphasizes the importance of using abattoirs for passive and active surveillance of diseases of public health and economic importance. Our study also confirmed that as a diagnostic strategy, it is imperative to institute more than one diagnostic method or test for the diagnosis of brucellosis in animals.

CONFLICT OF INTEREST

There is not conflict of interest with regards to this research.

AUTHORS’ CONTRIBUTIONS

FK, carried out all of the experimental work and manuscript preparation. AA, FF and HvH played equal roles as the study leaders and in carrying out data analysis and manuscript preparation. BD, CK, IM, AG and AP were involved in the generation and analysis of the biochemical tests and amplicon data. All authors read and approved the final manuscript.

ETHICAL STATEMENT

Permission to perform the research was granted regarding Section 20 Animal Diseases Act, 1984 (Act number 34 of 1984), by the Department of Agriculture Fisheries and Forestry, Reference number 12/11/1/1/6. Ethics approval was also granted by the University of Pretoria's Animal Ethics Committee, project number AEC12‐16.

ACKNOWLEDGEMENTS

The authors acknowledge Gauteng Department of Agriculture and Rural Development for funding this project. We also acknowledge our collaborators; Agricultural Research Council‐ Onderstepoort Veterinary Research (ARC‐OVR), Faculty of Veterinary Sciences, University of Pretoria and all abattoirs that participated in the study.

Kolo FB, Adesiyun AA, Fasina FO, et al. Seroprevalence and characterization of Brucella species in cattle slaughtered at Gauteng abattoirs, South Africa. Vet Med Sci. 2019;5:545–555. 10.1002/vms3.190

Funding Information

This research is supported by the Gauteng Department of Agriculture and Rural Development (GDARD). The funder had no role in the study design, data collection and interpretation, or the decision to submit the work for publication. Any opinion, finding and conclusion or recommendation expressed in this material is that of the author(s) and GDARD do not accept any liability in this regard.

REFERENCES

  1. Adesiyun, A. A. , & Cazabon, E. (1996). Seroprevalence of brucellosis Q‐fever and toxoplasmosis in slaughter livestock in Trinidad. Review of Livestock and Veterinary Medicine in Tropical Countries, 49(1), 28–30. [PubMed] [Google Scholar]
  2. Alton . (1990). Brucella suis: In Animal Brucellosis (pp. 411–422). Nielsen K. & Duncan J. R. (Eds.). Boca Raton, FL: CRC Press. [Google Scholar]
  3. Alton, G. D. , Pearl, D. L. , Bateman, K. G. , McNab, W. B. , & Berke, O. (2015). Suitability of sentinel abattoirs for syndromic surveillance using provincially inspected bovine abattoir condemnation data. BMC Veterinary Research, 11, 37 10.1186/s12917-015-0349-1 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bishop, G. (1984). A brucellosis serological survey on beef cattle slaughtered at Cato Ridge Abattoir. Journal of the South African Veterinary Association, 55(4), 185–186. [PubMed] [Google Scholar]
  5. Bricker, B. J. , Ewalt, D. R. , Olsen, S. C. , & Jensen, A. E. (2003). Evaluation of the Brucella abortus species–specific polymerase chain reaction assay, an improved version of the Brucella AMOS polymerase chain reaction assay for cattle. Journal of Veterinary Diagnostic Investigation, 15, 374–378. [DOI] [PubMed] [Google Scholar]
  6. Bricker, B. J. , & Halling, S. M. (1994). Differentiation of Brucella abortus bv. 1, 2, and 4, Brucella melitensis, Brucella ovis, and Brucella suis bv. 1 by PCR. Journal of Clinical Microbiology, 32(11), 2660–2666. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Caine, L. A. , Nwodo, U. U. , Okoh, A. I. , & Green, E. (2017). Molecular characterization of Brucella species in cattle, sheep and goats obtained from selected municipalities in the Eastern Cape, South Africa. Asian Pacific Journal of Tropical Disease, 7(5), 293–298. 10.12980/apjtd.7.2017D6-431 [DOI] [Google Scholar]
  8. Carmichael, L. E. (1990). Brucella canis . Animal Brucellosis, 1, 336–350. [Google Scholar]
  9. Coetzer, J. , Thomson, G. , & Tustin, R. (1994). Infectious diseases of livestock with special reference to Southern Africa. [Google Scholar]
  10. Corbel, M. J. (2006). Brucellosis in humans and animals in Corbel M.J., Elberg S. S., and Cosivi O. WHO/CDS/EPR/2006.7. Geneva, Switzerland: World Health Organization. [Google Scholar]
  11. DAFF . (2015). Brucella melitensis : A forgotten disease?
  12. DAFF . (2016). Abstract of agricultural statistics. (9783709117828). Department of Agriculture Forestry and Fisheries. Republic of South Africa, https://www.daff.gov.za/Daffweb3/Portals/0/Statistics.
  13. DAFF . (2017). Discussion Paper on the Review of Bovine Brucellosis Control in South Africa. Department of Agriculture, Forestry and Fisheries, https://www.daff.gov.za/&/Discussion%20paper%20on%20the%20review%20of%20b.
  14. DAFF . (2018). Department of Agriculture Forestry and Fisheries, Livestock numbers Fact Sheet: Agroecology. New letter livestock, https://www.daff.gov.za/daffweb3/Home/Crop-Estimates/.
  15. D'Anastasio, R. , Zipfel, B. , Moggi‐Cecchi, J. , Stanyon, R. , & Capasso, L. (2009). Possible Brucellosis in an early hominin skeleton from sterkfontein, South Africa. PLoS ONE, 4(7), e6439 10.1371/journal.pone.0006439 [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Emslie, F. R. , & Nel, J. R. (2002). An overview of the eradication of Brucella melitensis from KwaZulu‐Natal. Onderstepoort Journal of Veterinary Research, 69(2), 123–127. [PubMed] [Google Scholar]
  17. Eubanks, D. L. (2012). Dental considerations in cows and goats. Journal of Veterinary Dentistry, 29(3), 200–203. [DOI] [PubMed] [Google Scholar]
  18. Fasina, F. , Njage, P. , Ali, A. , Yilma, J. , Bwala, D. , Rivas, A. , & Stegeman, A. (2015). Development of disease‐specific, context‐specific surveillance models: Avian influenza (H5N1)‐related risks and behaviours in African countries. Zoonoses and Public Health, 63(1), 20–33. [DOI] [PubMed] [Google Scholar]
  19. Fosgate, G. T. , Adesiyun, A. A. , Hird, D. W. , Johnson, W. O. , Hietala, S. K. , Schurig, G. G. , & Ryan, J. (2002). Comparison of serologic tests for detection of Brucella infections in cattle and water buffalo (Bubalus bubalis). American Journal of Veterinary Research, 63(11), 1598–1605. [DOI] [PubMed] [Google Scholar]
  20. Foster, G. , Osterman, B. S. , Godfroid, J. , Jacques, I. , & Cloeckaert, A. (2007). Brucella ceti sp. nov. and Brucella pinnipedialis sp. nov. for Brucella strains with cetaceans and seals as their preferred hosts. International Journal of Systematic and Evolutionary Microbiology, 57(11), 2688–2693. [DOI] [PubMed] [Google Scholar]
  21. García‐Yoldi, D. , Marín, C. M. , de Miguel, M. J. , Muñoz, P. M. , Vizmanos, J. L. , & López‐Goñi, I. (2006). Multiplex PCR assay for the identification and differentiation of all Brucella species and the vaccine strains Brucella abortus S19 and RB51 and Brucella melitensis Rev1. Clinical Chemistry, 52(4), 779–781. [DOI] [PubMed] [Google Scholar]
  22. Godfroid, J. (2012). What is brucellosis? Infectious Diseases of Wild Mammals and Birds in Europe, 1, 318–328. [Google Scholar]
  23. Godfroid J., Bishop G. C., Bosman P. P. & Herr S. (2004). Bovine brucellosis. In Coetzer J.A.W. & Tustin R.C. (Eds.) Infectious diseases of livestock (vol. 3, 1510–1527). Cape Town, South Africa: Oxford University Press. [Google Scholar]
  24. Godfroid, J. , Cloeckaert, A. , Liautard, J.‐P. , Kohler, S. , Fretin, D. , Walravens, K. , … Letesson, J.‐J. (2005). From the discovery of the Malta fever's agent to the discovery of a marine mammal reservoir, brucellosis has continuously been a re‐emerging zoonosis. Veterinary Research, 36(3), 313–326. [DOI] [PubMed] [Google Scholar]
  25. Godfroid, J. , De Bolle, X. , Roop, R. , O'Callaghan, D. , Tsolis, R. M. , Baldwin, C. L. , … Nymo, I. H . (2014). The quest for a true One Health perspective of brucellosis. [DOI] [PubMed]
  26. Godfroid, J. , Nielsen, K. , & Saegerman, C. (2010). Diagnosis of brucellosis in livestock and wildlife. Croatian Medical Journal, 51(4), 296–305. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Gradwell, D. (1977). The isolation of Brucella abortus biotype I from African buffalo in the Kruger National Park. Journal of the South African Veterinary Association, 48(1), 41–44. [PubMed] [Google Scholar]
  28. Kaneene, J. B. , Miller, R. , & Meyer, R. M. (2006). Abattoir surveillance: The US experience. Veterinary Microbiology, 112(2), 273–282. [DOI] [PubMed] [Google Scholar]
  29. Keid, L. , Soares, R. , Vieira, N. , Megid, J. , Salgado, V. , Vasconcellos, S. , … Richtzenhain, L. (2007). Diagnosis of canine brucellosis: Comparison between serological and microbiological tests and a PCR based on primers to 16S‐23S rDNA interspacer. Veterinary Research Communications, 31(8), 951–965. [DOI] [PubMed] [Google Scholar]
  30. Kolo, F. B. , Fasina, F. O. , Ledwaba, B. , Glover, B. , Dogonyaro, B. B. , Adesiyun, A. A. , … van Heerden, H. (2018). Isolation of Brucella melitensis from cattle in South Africa. The Veterinary Record, 182(23), 668. [DOI] [PubMed] [Google Scholar]
  31. Lopez‐Goñi, I. , Garcia‐Yoldi, D. , Marin, C. , De Miguel, M. , Munoz, P. , Blasco, J. , … Ferreira, A. (2008). Evaluation of a multiplex PCR assay (Bruce‐ladder) for molecular typing of all Brucella species, including the vaccine strains. Journal of Clinical Microbiology, 46(10), 3484–3487. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Matope, G. , Bhebhe, E. , Muma, J. B. , Skjerve, E. , & Djønne, B. (2009). Characterization of some Brucella species from Zimbabwe by biochemical profiling and AMOS‐PCR. BMC Research Notes, 2(1), 261. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Mauff, A. (1980). Acute brucellosis in Johannesburg. South African Medical Journal, 58(12), 477–479. [PubMed] [Google Scholar]
  34. Muendo, E. N. , Mbatha, P. M. , Macharia, J. , Abdoel, T. H. , Janszen, P. V. , Pastoor, R. , & Smits, H. L. (2012). Infection of cattle in Kenya with Brucella abortus biovar 3 and Brucella melitensis biovar 1 genotypes. Tropical Animal Health and Production, 44(1), 17–20. [DOI] [PubMed] [Google Scholar]
  35. Nielsen, K. , Smith, P. , Yu, W. , Nicoletti, P. , Elzer, P. , Robles, C. , … Ruiz, A. (2005). Towards single screening tests for brucellosis. Revue Scientifique et Technique‐Office International des Épizooties, 24(3), 1027. [PubMed] [Google Scholar]
  36. OIE (2009). Bovine brucellosis. Terrestrial Manual. Paris: Office International des Epizooties. [Google Scholar]
  37. OIE (2016). Brucellosis manual of diagnostic tests and vaccines for terrestrial animals. Paris: Office International des Epizooties. [Google Scholar]
  38. Olsen, S. , & Palmer, M. (2014). Advancement of knowledge of Brucella over the past 50 years. Veterinary Pathology, 51(6), 1076–1089. [DOI] [PubMed] [Google Scholar]
  39. Padilla, P. F. , Nielsen, K. , Ernesto, S. , & Ling, Y. (2010). Diagnosis of brucellosis. The Open Veterinary Science Journal, 4(1), 46–60. [Google Scholar]
  40. Pappas, G. (2013). 70 ‐ Brucellosis In Magill A. J., Hill D. R., Solomon T., & Ryan E. T. (Eds.), Hunter's tropical medicine and emerging infectious disease, 9th ed. (pp. 577–579). London: W.B. Saunders. [Google Scholar]
  41. Pappas, G. , Papadimitriou, P. , Akritidis, N. , Christou, L. , & Tsianos, E. V. (2006). The new global map of human brucellosis. The Lancet Infectious Diseases, 6(2), 91–99. [DOI] [PubMed] [Google Scholar]
  42. Radostits, O. M. , Gay, C. C. , Hinchcliff, K. W. , & Constable, P. D. (2006). Veterinary Medicine: A textbook of the diseases of cattle, horses, sheep, pigs and goats. (pp. 969–984). Edinburgh: Sanders. [Google Scholar]
  43. RCoreTeam . (2013). R Foundation for Statistical Computing. Vienna, Austria, 3(0).
  44. Reichel, R. , Nel, J. R. , Emslie, R. , & Bishop, G. C. (1996). Brucella melitensis biotype 1 outbreak in goats in northern KwaZulu‐Natal. Onderstepoort Journal of Veterinary Research, 63(2), 183–185. [PubMed] [Google Scholar]
  45. Ribeiro, L. , & Herr, S . (1990). The use of filter paper discs impregnated with thionin acetate, basic fuchsin and thionin blue in the identification of Brucella species. [PubMed]
  46. Ribeiro, L. , Herr, S. , & Chaparro, F. (1990). The isolation and serology of Brucella melitensis in a flock of goats in central RSA. The Onderstepoort Journal of Veterinary Research, 57(2), 143–144. [PubMed] [Google Scholar]
  47. Sadler, W. W. (1960). Present evidence on the role of meat in the epidemiology of human brucellosis. American Journal of Public Health and the Nations Health, 50(4), 504–514. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Scholz, H. , & Vergnaud, G. (2013). Molecular characterisation of Brucella species. Revue Scientifique et Technique, 32(1), 149–162. [DOI] [PubMed] [Google Scholar]
  49. Seleem, M. N. , Boyle, S. M. , & Sriranganathan, N. (2010). Brucellosis: A re‐emerging zoonosis. Veterinary Microbiology, 140(3), 392–398. [DOI] [PubMed] [Google Scholar]
  50. Simpson, G. J. , Marcotty, T. , Rouille, E. , Chilundo, A. , Letteson, J.‐J. , & Godfroid, J. (2018). Immunological response to Brucella abortus strain 19 vaccination of cattle in a communal area in South Africa. Journal of the South African Veterinary Association, 89(1), 1–7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Smirnova, E. A. , Vasin, A. V. , Sandybaev, N. T. , Klotchenko, S. A. , Plotnikova, M. A. , Chervyakova, O. V. , … Kiselev, O. I. (2013). Current methods of human and animal brucellosis diagnostics. Advances in Infectious Diseases, 3(03), 177. [Google Scholar]
  52. Stemshorn, B. , Forbes, L. , Eaglesome, M. , Nielsen, K. , Robertson, F. , & Samagh, B. (1985). A comparison of standard serological tests for the diagnosis of bovine brucellosis in Canada. Canadian Journal of Comparative Medicine, 49(4), 391. [PMC free article] [PubMed] [Google Scholar]
  53. Strachan, P. (1932). Undulant fever. South African Medical Journal, 6(9), 291–294. [Google Scholar]
  54. Van Drimmelen, G. (1949). The brucellosis survey in South Africa. Journal of the South African Veterinary Association, 20(3), 178–188. [Google Scholar]
  55. Van Drimmelen, G. (1965). The presence of brucella melitensis infection in sheep in the transvaal. Bulletin de l'Office International Des Épizooties, 64, 745–756. [PubMed] [Google Scholar]
  56. Verger, J. , Garin‐Bastuji, B. , Grayon, M. , & Mahé, A . (1989). La brucellose bovine à Brucella melitensis en France. [PubMed]
  57. Weiner, M. , Iwaniak, W. , & Szulowski, K. (2011). Comparison of PCR‐based AMOS, Bruce‐Ladder and MLVA assays for typing of Brucella species. Bulletin of the Veterinary Institute in Pulawy, 55, 625–630. [Google Scholar]
  58. Wojno, J. M. , Moodley, C. , Pienaar, J. , Beylis, N. , Jacobsz, L. , Nicol, M. P. , … Bamford, C. (2016). Human brucellosis in South Africa: Public health and diagnostic pitfalls. SAMJ ‐ South African Medical Journal, 106(9), 883–885. 10.7196/SAMJ.2016.v106i9.11020 [DOI] [PubMed] [Google Scholar]
  59. Zammit, T. (1905). A preliminary note on the examination of the blood of goats suffering from Mediterranean fever. Reports of the Royal Society of London, Mediterranean fever commission, London, Harrison and Sons, part, 3, 83. [Google Scholar]

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