Environmental risk factors in the incidence of Johne's disease.

http://informahealthcare.com/mby ISSN: 1040-841X (print), 1549-7828 (electronic) Crit Rev Microbiol, Early Online: 1–20 ! 2014 Informa Healthcare USA, Inc. DOI: 10.3109/1040841X.2013.867830

REVIEW ARTICLE

Environmental risk factors in the incidence of Johne’s disease Geoffrey N. Elliott1, Rupert L. Hough1, Lisa M. Avery1, Charlotte A. Maltin2, and Colin D. Campbell1 The James Hutton Institute, Craigiebuckler, Aberdeen, UK and 2Quality Meat Scotland, The Rural Centre, Newbridge, United Kingdom

Critical Reviews in Microbiology Downloaded from informahealthcare.com by Australian National University on 03/14/15 For personal use only.

1

Abstract

Keywords

This review addresses the survival and persistence of Mycobacterium avium subsp. paratuberculosis (MAP), the causative pathogen of Johne’s disease (JD), once it has left its ruminant host. JD has significant economic impact on dairy, beef and sheep industries and is difficult to control due to the long-term sub-clinical nature of the infection, intermittent or persistent MAP shedding during and after this period, inadequate test effectiveness, and the potential for MAP to exist for extended periods outside the host. The role that environmental factors play in the persistence and spread of MAP and consequent disease is assessed. Published risk factor analysis, organism survival across various environmental media and conditions, presence and spread in ruminant and non-ruminant wildlife, and the general potential for survival and multiplication of MAP ex-host both on and off-farm are discussed and knowledge gaps highlighted. An inclusive approach to disease management that takes into account the persistence and transport of the causative organism in on-farm soils and waters, land use and management, dispersal by domestic and non-domestic host species, as well as general animal husbandry is required on those farms where more traditional approaches to disease management have failed to reduce disease prevalence.

Fecal shedding, Mycobacterium avium subsp. paratuberculosis, pathogen survival and transport, paratuberculosis, soil

Introduction Mycobacterium avium subsp. paratuberculosis (MAP) is the causative microbial agent of paratuberculosis or Johne’s disease (JD), a chronic wasting disease first described in cattle by Johne & Frothingham (1895). In affected ruminants, normal absorption of food is impeded due to a thickening of the intestinal wall as a result of intracellular incorporation of MAP cells, leading to persistent diarrhoea and progressive weight loss for the animal and eventual animal death, and the consequent economic impacts on dairy, beef and sheep industries through loss of milk and reduced meat and wool production are significant (Bush et al., 2006; Ott et al., 1999). A central recognition of most JD control programs in place across the globe is the concept that MAP is an obligate intracellular pathogen, and so the removal of any and all infected animals from the herd should, by definition, largely lead to the eradication of the resulting disease in the remaining herd stock (Silva, 2012). JD control programs have been shown in some cases to show significant levels of JD prevalence reduction but have met with much less success in terms of disease eradication (e.g. Benedictus et al., 2008; Collins et al., 2010; Nielsen & Toft, 2011) with further measures or longer monitoring periods suggested for assessment of potential further reductions in JD prevalence.

Address for correspondence: Geoffrey N. Elliott, The James Hutton Institute, Craigiebuckler, Aberdeen, AB15 8QH United Kingdom. E-mail: [email protected]

History Received 6 September 2013 Revised 16 November 2013 Accepted 18 November 2013 Published online 25 March 2014

The potential for infection to exist sub-clinically in the animal host for years before symptoms appear (Nielsen & Ersbøll, 2006), intermittent or persistent shedding of the infective organism during this prolonged latent phase of infection (Sweeney, 2006) and the low sensitivity of available tests for JD (Timms et al., 2011; Weber, 2006) are factors generally included in the design of such control programs. However, MAP remains widespread in ruminant livestock worldwide, including in those under JD control programs, and continues to be discovered in new species. MAP is an acid-fast bacterium with a unique cell wall structure, giving the organism an increased persistence in harsh environments and an ability to survive for extended periods outside the host (Grewal et al., 2006; Whittington et al., 2004, 2005). Thus JD eradication may be confounded by long-term reservoirs of shed, viable MAP and therefore infection, in both the immediate farm and wider natural environment. Although the improvement of on-farm management measures has been modeled to be the only approach to reduce true herd prevalence of JD (Bennett et al., 2010, 2012; Kudahl et al., 2007, 2008), the potential for infection via the wider environment needs to be considered for effective disease eradication. This review discusses the spread, survival and persistence of MAP once it has left its ruminant host, and addresses the role that environmental and wider farm factors play in the persistence and spread of MAP leading to further JD infection in livestock and beyond. The potential for these factors to be included in the design for future studies and control programs is discussed.

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G. N. Elliott et al.


Review methodology This was a purposive review of peer-reviewed literature related to environmental risk factors for JD. In a few cases, this was supplemented with literature from web-accessible documents and other grey literature. Computerized database searching of a range of international databases was carried out: MEDLINE (MEDLINE Database, National Library of Medicine, Bethesda, MD, USA www.biomednet.com), BIDS (Join Information Systems Committee, University of Bath, Bath, UK, www.bids.ac.uk), PubMed (National Library of Medicine, www.ncbi.nlm.nih.gov/PubMed) and ISI Web of Knowledge (Mimas, University of Manchester, UK, wok.mimas.ac.uk). A systematic, staged search strategy was employed using the following search terms: ‘‘Johne’s’’ OR ‘‘paratuberculosis’’ OR ‘‘Mycobacterium avium subsp. paratuberculosis’’ OR ‘‘MAP’’ AND ‘‘risk factors’’ OR ‘‘environment’’ OR ‘‘survival’’ OR ‘‘soil’’ OR ‘‘water’’ OR ‘‘climate’’ OR ‘‘management’’ OR ‘‘husbandry’’. Over 600 peer-reviewed references were found. The search was further refined to papers that investigated explicit potential links between environmental factors and JD. Original full texts were obtained for all studies. Wider, more descriptive studies and accounts were also included. Studies whose focus was to demonstrate or develop a specific molecular analysis were excluded, except where a method was used that showed principles relevant to the context of this review. The remaining 179 studies formed the basis of this review, although other papers within the database were used as background material. Immediate host infection and dispersal during pre-clinical and clinical disease phases By far the most common route of entry for MAP into the host animal is via the intestinal tract, with the infective dose investigated by numerous studies. However, despite illustrations of infection using a dose of only 103 or 104 MAP cells in sheep (Brotherston et al., 1961, Reddacliff & Whittington, 2003), the exhibition of clinical disease had only been demonstrated in sheep with an infective dose of 108 MAP cells or more, with MAP strain variation and host compatibility as well as a lack of standardization implicated in the varied responses found in such studies (for a review, see Begg & Whittington, 2008). Studies assessing infective dose in cattle have not been able to demonstrate clinical disease onset, likely due to the time required to keep the animal under controlled observation before such onset. Infection has been shown to occur through the colostrum of milk carrying the organism (Streeter et al., 1995) or in utero (Whittington & Windsor, 2009), but it is the MAP-containing manure-contaminated environment to which animals are exposed that is key. Cattle of less than 6 months of age are considered most, but not exclusively, susceptible (Mortier et al., 2013; Windsor & Whittington, 2010), becoming infected as calves but not manifesting clinical symptoms, including diarrhoea and associated weight loss, until sometimes several years later (Manning & Collins, 2001). The increased likelihood of infection is thought to be due to the reduced immunity of the very young calf, with

Crit Rev Microbiol, Early Online: 1–20

older cattle manifesting macrophage-based reduction of multiplication of MAP in the intestinal mucosa and thus increased resistance (Bendixen et al., 1981; Merkal et al., 1975), although sheep and other domestic ruminants may never develop this (Koets et al., 2000). In cattle, after a latent period of generally 2 to 5 years following infection (for a review of JD pathogenesis, see Sweeney, 2011), during which MAP survives and multiplies in the mucosa of the intestinal tract, infected animals begin to shed the organism in large numbers in their feces (Benedictus et al., 2008; Figure 1). Shedding has been shown to also occur in low numbers in calves only 8 months of age or younger (Antognoli et al., 2007; Collins & Zhao, 1994; McDonald et al., 1999). Such pre-clinical shedding can be intermittent and at a low level (Chiodini, 1996) and so attempted isolation of MAP by fecal culture from such individuals is unreliable as a definitive diagnostic tool for the demonstration of infection status. Further, the humoral response may not occur until after the onset of significant shedding (Sweeney et al., 2006), therefore the utility of serological testing (via enzyme-linked immunosorbent assay, i.e. ELISA) may be compromised in the early stages of the disease in the demonstration of infection status (Milner et al., 1987) and may have low diagnostic sensitivity (Elzo et al., 2006). This said, other studies have shown antibodies to be detectable prior to significant MAP shedding if the test is calibrated to be sensitive and specific (Nielsen, 2008), and ELISA has been shown to be successful in the reduction of JD prevalence in a herd if repeated confirmatory testing is performed prior to selective culling (Nielsen & Toft, 2011). However, whilst infected animals remain in the herd, shedding can significantly contaminate the surrounding environment causing exposure of susceptible animals to the disease-causing organism (Figure 1). Fecal shedding in high numbers has been reported to occur most frequently in adult cattle at, or shortly before, the onset of clinical symptoms (Sweeney, 2011). This clinical phase is commonly manifested at a younger age in deer and wild ruminants (Mackintosh and Griffin, 2010). However, the length of the pre-clinical phase in sheep and goats is more difficult to assess without laboratory testing, due to a lack of definitive symptoms other than weight loss (which can be due to multiple other causes) (Robbe-Austerman, 2011) and has been reported to be as short as 1 month or as long as 5 years, possibly dependent on exposure dose (McGregor et al., 2012). During the subsequent clinical disease stage, shedding of the organism can exceed 108 organisms per gram of feces, with a single animal having the potential to excrete over 1012 organisms per day (Chiodini et al., 1984). Thickening of the intestinal mucosa causes nutrient absorption to be decreased leading to chronic diarrhoea in cattle, resulting in rapid weight loss, diffuse oedema, decreased milk production and infertility. Although MAP has been detected in both the milk and the semen as well as from other tissues of infected cattle (Ayele et al., 2004; Slana et al., 2008), it is the environmental fecal contamination resulting from such shedding that is considered a primary source of infection (Benedictus et al., 2008; Marce´ et al., 2011; Windsor & Whittington, 2010). Shedding of MAP and consequent spread of the organism

Johne’s disease

DOI: 10.3109/1040841X.2013.867830

3

Wider environment

Waters Nematodes Invertebrates

Pasture

Farmyard d Removal/culling

Domesc cale Latent period

Silent

Uninfected

Sub-clinical

Clinical

Advanced

M I L K


FAECAL SHEDDING

DORMANCY?

Wild herbivores

Temp/UV exposure

STORAGE

Soil pH/ Chemistry

Arable land

Temp/UV exposure DORMANCY?

Wild carnivores ores DORMANCY?

SPREADING

Slurry/Waste /Silage

Nematodes Invertebrates

Amoebae?

pH/ Chemistry

Figure 1. Distribution of MAP within the farming and wider environment.

within the immediate farm environment and therefore between animals has been shown to be exacerbated (perhaps unsurprisingly) by the regular close proximity and relatively high density of stock animals on farms, with e.g. the risk of incidence of JD found to increase by more than 2-fold for every 100 animals in the herd (Pillars et al., 2009; Table 1; Figure 2). Epidemiologic studies have repeatedly highlighted husbandry practices as important risk factors for the disease, with hygiene being a key factor (e.g. Johnson-Ifearulundu & Kaneene, 1999; Ansari-Lari et al., 2009). Many hygiene measures, such as the use of gutter cleaner or the cleaning of calf pens after each use (Johnson-Ifearulundu & Kaneene, 1998; Table 1; Figure 2), have been shown to be protective against JD, indicating that basic control measures in the immediate farm environment can have a significant effect, although this is not always the case e.g. the washing of udders prior to parturition were reported as increasing the chances of JD, suggested as indicating a general lack of good hygiene, as well as possibly enhancing teat contamination (JohnsonIfearulundu & Kaneene, 1998; Table 1; Figure 2). However, if the ‘‘wider’’ environment (i.e. external to the farmyard) is also acting as a source of infection then control measures such as good hygiene practice may be undermined. Detection in the immediate farm environment As shedding can fluctuate in terms of numbers of MAP expelled (Nielsen, 2008; Stewart et al., 2004, 2006), MAP culture using communal or pooled samples from the farmyard has been shown by many as a viable and less expensive alternative to individual animal fecal sampling for the determination of within-herd infection prevalence (Kalis

et al., 2004; Whittington et al., 2000). Raizman et al. (2004) showed that farmyard areas such as alleyways and pens when tested gave comparable results to laboratory-pooled fecal samples from individual animals and could be used as an effective proxy for herd infection, and sampling of water from a farm wastewater lagoon has been found to be more likely to give a positive result than composite manure samples (Berghaus et al., 2006; Figure 1). A nationwide US study found that using an immediate farm-environment testing approach, 70% of herds were identified as being infected (Lombard et al., 2006). By 2008, standardized environmental sampling (pooling of multiple fecal samples taken from communal areas where adult cow manure regularly accumulates) and culture-based MAP testing of the composite samples had indicated that 68% of American dairy herds were positive for MAP. A quarter of these farms were estimated as having high numbers of infected cows with disease prevalence increasing with herd size (USDA:APHIS:VS, 2008, 2010). A similar UK survey estimated 35% herd prevalence in UK dairy farms (Cook et al., 2009). Similar significant levels of infection have been found in the Netherlands, Canada, Denmark, the Republic of Ireland and Australia amongst others, although testing approaches (and associated uncertainties) vary, making direct comparison difficult (Nielsen & Toft, 2009). Sampling of wastewater lagoons, fecal slurry and communal areas has now become a common strategy for the classification of herds, based on MAP infection (Aly et al., 2010; Cook et al., 2009; Raizman et al., 2011). Pooled environmental testing such as this may, however, become less sensitive in low prevalence herds as small numbers of positive samples become diluted to the point of

Soil % organic carbon (paddock)(S) % sand (paddock)(S) Depth of water table low (53 m) Drainage poor or worse Entisol versus alfisol Inceptisol versus alfisol Loam Sand content550 % Sands Sandy loam Silt 5or ¼ 50 % Silt loam Slope 5or ¼ 1% Soil iron 1 ppm increase Soil iron 1 ppm increase Soil pH57 Soil pH 0.1 increase Surface water high in Fe (S) Climate Altitude4or ¼ 750 m (S) Concrete corrosion low/moderate Frost low/moderate Non-arid climate Steel corrosion low/moderate Land use/management Boreal forest (versus montane) Boreal forest (versus parkland) Fertilizer applied to pastures Grassland (vs. montane) Grassland (vs. parkland) Lime applied to pasture Lime applied to pasture (1) Lime applied to pasture (2) Manure spread on pastures Manure stored outside Montane (versus parkland) Parkland (versus grassland) Parkland (versus montane) Pasture (non-lactating cows) Rotate pasture (lactating cows) Slurry spread on fields Animal husbandry 45% of cows purchased in herd Addition of new animals (G) Addition of new animals (S) Barn scraped (heifers) Barn scraped (lactating cows) Barn scraped (non-lactating cows)

Risk factor 1.5 0.9 * * 1.6 0.4 1 * 0.210 1.4 2.1 0.1 * 1.002 1.003 1.14 0.922 1.1 1.31 * * 1.05 * 0.38 0.37 0.325 0.02 0.14 0.147 0.008 0.02 0.066 1.676 0.32 1.62 0.31 0.368 0.331 1.472 0.74 1.8 1.1 1.188 1.473 0.114

3.01 1.18 1.74 1.94 0.76 3.64 0.87 0.556 0.37 0.33 0.282 0.063 0.1 0.129 2.536 0.55 5.85 2.85 0.569 0.524 5.79 1.34 2.9 1.9 2.162 3.756 0.22

95% CI

2.2 0.97 0.5 0.3 25.9 3.5 3.6 0.32 1.05 6.2 7.2 0.2 1.09 1.004 1.014 1.92 0.951 2.64

OR

2.4 4.6 3.6 3.936 9.574 0.424

NS 0.04 0.04 0.0117 0.0560 0.0001

0.044 0.05 0.0327 0.044 0.05 50.001 0.007 0.008 0.129 0.0001 0.05 0.05 0.044 0.0117 0.0055 0.012

NP 0.791 0.323 0.04 0.612

50.001 0.03 0.219 0.051 NP NP PS 0.059 0.950 PS 0.002 PS 0.866 0.0015 0.013 0.03 0.0019 NP

p

Group Group Group Individual Individual Individual

Individual Individual Individual Individual Individual Individual Group Group Individual Individual Individual Group Individual Individual Individual Group

Individual Group Group Individual Group

Group Group Group Group Group Group Group Group Group Group Group Group Group Individual Group Individual Individual Individual

Level

Cattle Goats Sheep Cattle Cattle Cattle

Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle

Sheep Cattle Cattle Cattle Cattle

Sheep Sheep Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Sheep

Animal

Denmark Jordan Jordan USA USA USA

Canada Canada USA Canada Canada USA USA USA USA USA Canada Canada Canada USA USA USA

Australia USA USA Canada USA

Australia Australia USA USA Spain Spain USA USA USA USA USA USA USA USA USA Canada USA Australia

Country

ELISA ELISA ELISA ELISA ELISA ELISA

ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA

Reporting scouring ELISA ELISA ELISA ELISA

Pooled fecal culture Pooled fecal culture ELISA ELISA Serum AGID Serum AGID ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA Reporting scouring

Test

Nielsen & Toft, 2007 Al-Majali et al., 2008 Al-Majali et al., 2008 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1999

Scott et al., 2007 Scott et al., 2006 Johnson-Ifearulundu & Kaneene, 1999 Scott et al., 2007 Scott et al., 2006 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1998 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1999 Scott et al., 2006 Scott et al., 2006 Scott et al., 2007 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1999

Lugton, 2004a Ward & Perez, 2004 Ward & Perez, 2004 Scott et al., 2006 Ward & Perez, 2004

Dhand et al., 2009a Dhand et al., 2009a Ward & Perez, 2004 Ward & Perez, 2004 Reviriego et al., 2000 Reviriego et al., 2000 Ward & Perez, 2004 Ward & Perez, 2004 Johnson-Ifearulundu & Kaneene, 1999 Ward & Perez, 2004 Ward & Perez, 2004 Ward & Perez, 2004 Ward & Perez, 2004 Johnson-Ifearulundu & Kaneene, 1999 Johnson-Ifearulundu & Kaneene, 1999 Scott et al., 2006 Johnson-Ifearulundu & Kaneene, 1999 Lugton, 2004a

Reference


35.27 2.06 0.953 6.97 0.76 0.539 0.472 0.56 0.252 3.838 0.95 21.1 26.66 0.882 0.827 22.8

6.9 * * 3.57 *

3.2 0.99 * * 411 45 13.2 * 5.28 27.4 24.5 0.7 * 1.007 1.026 3.22 0.982 6.29

þ95% CI

Table 1. Risk factors (apart from those considering age or sex of host animal) addressed in the literature – data described in final appropriate analyses only presented.


4 Crit Rev Microbiol, Early Online: 1–20

Bulk and high SCC milk (versus replacer) Bulk tank milk (versus replacer) Pooled high SCC milk and replacer (versus replacer) Pooled high SCC milk (versus replacer) Calf feed grown where manure spread Calves on pasture in summer time Calves separated from cows at 52 months Calves separated from cows at 2 - 12 months Calving in individual pens Cleaning of calf pens after each use Colostrum from dam and multiple cows (versus dam) Colostrum from multiple cows (versus dam only) Common feed site for adults and calves Cows (after 1st calving) have exercise facility Cows (after 1st calving) on pasture in summer Dam MAP positive 5or ¼ 2 years post-calving Dam MAP positive 42 years post-calving Dam was MAP positive Dry cows in exercise facility Dry cows on pasture in summer time Exercise lot for lactating cows (1) Exercise lot for lactating cows (2) Hay offered at 510 days (versus none) Hay offered at 410 days age (versus none) Herd size 590 Herd size 117 adults Herd size (per 100 cows) History of clinical signs in herd History of JD in the herd History of test positive animals Mixed farming (goats and sheep) PTB test-positive in past 3yr Purchased cows in herd Purchased cows in past 5yr Replacements purchased privately Same equipment for feed and manure Suckling with foster cows (versus milk replacer) Tested for MAP in past 5yr Use of gutter cleaner Used purchased course mix feed Udder contamination Wash udder Wash udder prior to parturition Winter shearing (S) (versus at other times) Young stock 412 months separated from cows Young stock 412 months on pasture in summer Wildlife Deer present Genetic factors Brahman Angus versus Holstein (BolFNG) Brahman Angus versus Holstein (SLC11A1-275) 0.89 1.31 1.54

2.45 2.72

1.000 0.895 0.910 0.961 1.16 0.53 0.33 0.84 0 0.08 0.882 1.089 0.093 0.11 0.38 1.4 1.55 2.05 0.13 0.42 1.03 1.053 0.11 0.1 0.27 0.62 1.09 1.24 1.02 1.217 1 1.46 0.65 1.01 1.77 0.036 1.37 1.59 0.022 0 1.29 2.392 1.87 1.62 0.85 0.29

15.15

1.137 1.070 1.056 1.097 1.704 0.98 0.84 1.67 0.21 0.28 1.003 1.243 0.257 0.52 0.77 9 5.98 6.8 0.38 0.93 3.01 3.9 0.29 0.3 0.51 1.42 2.24 1.91 6.7 7.35 1.1 6.71 1.27 3.09 4.19 0.16 2.012 4.62 0.1 0.3 6.38 3.84 8.66 6.2 1.54 0.81

4.75 4.78

258.7

1.292 1.279 1.225 1.153 2.503 2 2.3 3.8 0.93 0.89 1.140 1.418 0.709 2.3 1.51 57.9 23.07 22.51 1.06 2.1 8.8 14.48 0.71 0.88 0.96 3.28 4.6 2.941 44.2 44.35 2.9 30.9 2.9 9.47 9.93 0.74 2.956 13.47 0.48 0.88 31.5 6.164 40.08 23.6 2.8 1.92

0.005 0.0005

0.06

0.0498 0.4575 0.4740 0.1701 0.0066 NS NS NS 0.04 0.030 0.9669 0.0012 0.0087 NS NS 50.01 50.01 50.01 NS NS 0.040 0.042 0.007 0.03 0.05 0.406 0.281 0.0033 0.04 0.03 0.05 0.010 NS 0.0480 0.001 0.019 0.0004 0.0050 0.003 0.03 0.02 0.0001 0.006 NP NS NS

Individual Individual

Group

Individual Individual Individual Individual Individual Group Group Group Group Group Individual Individual Individual Group Group Individual Individual Individual Group Group Group Group Group Group Individual Individual Group Individual Group Group Group Group Group Group Group Group Individual Group Group Group Group Individual Group Individual Group Group

Cattle Cattle

Cattle

Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Zoo Zoo Zoo Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Goats Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle cattle Cattle Cattle Sheep Cattle Cattle

USA USA

UK

Denmark Denmark Denmark Denmark USA Denmark Denmark Denmark UK USA Denmark Denmark USA Denmark Denmark USA USA USA Denmark Denmark USA USA UK UK Canada Canada USA USA Iran USA Jordan USA Denmark USA UK USA Denmark USA USA UK Iran USA USA Australia Denmark Denmark

Any one of multiple. Any one of multiple.

Reporting

ELISA ELISA ELISA ELISA ELISA ELISA ELISA ELISA Reporting ELISA ELISA ELISA ELISA ELISA ELISA Culture/PCR Culture/PCR Culture/PCR ELISA ELISA ELISA ELISA Reporting Reporting ELISA ELISA Pooled fecal culture ELISA Bulk tank milk PCR ELISA ELISA ELISA ELISA Pooled fecal culture Reporting ELISA ELISA Pooled fecal culture ELISA Reporting Bulk tank milk PCR ELISA ELISA Reporting scouring ELISA ELISA


Pinedo et al., 2009 Pinedo et al., 2009

Cetinkaya et al., 1997

Nielsen et al., 2008 Nielsen et al., 2008 Nielsen et al., 2008 Nielsen et al., 2008 Johnson-Ifearulundu & Nielsen & Toft, 2007 Nielsen & Toft, 2007 Nielsen & Toft, 2007 Cetinkaya et al., 1997 Johnson-Ifearulundu & Nielsen et al., 2008 Nielsen et al., 2008 Johnson-Ifearulundu & Nielsen & Toft, 2007 Nielsen & Toft, 2007 Witte et al., 2009 Witte et al., 2009 Witte et al., 2009 Nielsen & Toft, 2007 Nielsen & Toft, 2007 Johnson-Ifearulundu & Johnson-Ifearulundu & Cetinkaya et al., 1997 Cetinkaya et al., 1997 Scott et al., 2006 Scott et al., 2007 Pillars et al., 2009 Johnson-Ifearulundu & Ansari-Lari et al., 2009 Johnson-Ifearulundu & Al-Majali et al., 2008 Johnson-Ifearulundu & Nielsen & Toft, 2007 Pillars et al., 2009 Cetinkaya et al., 1997 Johnson-Ifearulundu & Nielsen et al., 2008 Pillars et al., 2009 Johnson-Ifearulundu & Cetinkaya et al., 1997 Ansari-Lari et al., 2009 Johnson-Ifearulundu & Johnson-Ifearulundu & Lugton, 2004a Nielsen & Toft, 2007 Nielsen & Toft, 2007

(continued )

Kaneene, 1999 Kaneene, 1998

Kaneene, 1999

Kaneene, 1999

Kaneene, 1998

Kaneene, 1999

Kaneene, 1999

Kaneene, 1998 Kaneene, 1999

Kaneene, 1999

Kaneene, 1998

Kaneene, 1999

DOI: 10.3109/1040841X.2013.867830

Johne’s disease 5

G. N. Elliott et al. OR – Odds ratio; CI – Confidence interval; P – Probability; Level – Point of assessment of infection; NS – Stated in publication as ‘non-significant’, but P not specified; PS – Stated in publication that data ‘significant’, but P not specified; NP -Data not presented; Factors in bold are represented graphically (Figure 2).

multiple. multiple. multiple. multiple.

Pinedo et al., 2009 Pinedo et al., 2009 Cetinkaya et al., 1997 Scott et al., 2007 Scott et al., 2007 Scott et al., 2007 Scott et al., 2007 Scott et al., 2007 Scott et al., 2007 Cetinkaya et al., 1997 Pinedo et al., 2009 Pinedo et al., 2009 Pinedo et al., 2009 Pinedo et al., 2009 Jakobsen et al., 2000 Jakobsen et al., 2000 multiple. multiple.

Any one of Any one of Reporting ELISA ELISA ELISA ELISA ELISA ELISA Reporting Any one of Any one of Any one of Any one of ELISA ELISA USA USA UK Canada Canada Canada Canada Canada Canada UK USA USA USA USA Denmark Denmark Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Individual Individual Group Individual Individual Individual Individual Individual Individual Group Individual Individual Individual Individual Individual Individual 50.001 0.005 50.001 0.659 0.659 0.659 0.659 0.659 0.659 50.001 0.02 0.03 0.01 0.01 50.001 50.001 Brahman Angus versus Holstein (SLC11A1-279) Brahman Angus versus Holstein (SLC11A1-281) Breed ¼ Channel Islands Breed ¼ Angus Breed ¼ Red Angus Breed ¼ Charolais Breed ¼ Hereford Breed ¼ Limousin Breed ¼ Simmental Channel Island breeds predominate Jersey versus Holstein (BolFNG) Jersey versus Holstein (SLC11A1-275 allele) Jersey versus Holstein (SLC11A1-279 allele) Jersey versus Holstein (SLC11A1-281 allele) Jersey versus large breed, tested mth1 post-calving Large breed þ tested mth1 post-calving

2.84 2.39 12.9 0.16 0.37 0.38 0.19 0.23 0.27 10.89 0.32 0.34 0.31 0.31 7.4 12.8

1.57 1.29 4.7 0.03 0.05 0.07 0.03 0.02 0.05 4.25 0.12 0.13 0.12 0.12 4.1 6.1

5.10 4.39 35.37 1.04 2.76 1.92 1.43 2.78 1.35 27.91 0.83 0.91 0.82 0.82 13.4 26.9

Level þ95% CI 95% CI OR Risk factor

Table 1. Continued


p

Animal

Country

Test


Reference

6

non-detection via culture (Smith et al., 2011). In many cases, culture-based approaches have been supplemented or replaced with the use of molecular methods, particularly PCR-based approaches. When both PCR and culture-based methodologies have been used on the same samples, in most cases the molecular method has resulted in more ‘‘MAP-positive’’ samples than has culture. Although this may indicate the detection of low numbers of viable cells missed by culture, the PCR-based approach would also detect non-viable cells from past contamination events or the presence of ‘‘naked’’ DNA (DNA present outside a cell) and neither of these constitutes an epidemiological risk in terms of JD. However, in sampling areas where there is a rapid turnover of fecal material, it has been shown that a molecular testing approach compares favorably with culture, as this approach can give greater test sensitivity and the presence of DNA indicates the presence of recent viable MAP (Alinovi et al., 2009; Aly et al., 2010). Several studies have also discussed the possibility that MAP may be present in a viable but noncultivable or dormant state, both of which would also arguably lead to detection only by the ‘‘molecular’’ method (Lamont et al., 2012).

MAP persistence outside the host Persistence in the environment MAP survival in the extremes of conditions has been demonstrated in various studies, with survival in highly alkaline sheep dip (pH 12.4) for over 14 days (Eamens et al., 2001), in feces at 70 C for over 15 weeks (Richards, 1981) and in infected tissue mixed with compost kept at 80 C for 90 days (Tkachuk et al., 2013). Such resistance is in large part conferred by the thick cell wall which MAP possesses, comprising 60% lipids and giving increased resistance to physical and chemical hardships (Grant et al., 2002; Whan et al., 2001). However, such a resistant cell wall also leads to a restriction in nutrient uptake and consequent slow growth, with 12–16 weeks generally required for primary in vitro culture (Stevenson, 2010). The survival and persistence of MAP in more moderate conditions once it has left its animal host has been intermittently studied for many years, with the first questions regarding the continued infectivity of manure-contaminated pastures published over a century ago (Penberthy, 1912). In 1944, Lovell et al. published the results of the first series of survival experiments on ‘‘Johne’s bacilli’’, due to their belief that JD was spread mainly by contamination of water, foodstuffs and pastureland with infected feces (Table 2). Assessment of the persistence of MAP within naturally infected bovine feces exposed to rain and sunlight showed viable MAP remaining in the samples after 21 weeks, with samples kept wet remaining MAP viable after 29 weeks, during which time the material had undergone periods of freezing (Lovell et al., 1944). Even longer periods of survival were shown in MAP-inoculated feces mixed with soil (Vishnevskii et al., 1940; Table 2) although the conditions under which these experiments were performed are unclear. Conversely, the duration of MAP survival in bovine feces in the presence of bovine urine or in mixed urine and feces was found to be less than 30 days at 38 C (Larsen et al., 1956) and less than 1 week in an earlier study (Vishnevskii et al.,

Johne’s disease

7


DOI: 10.3109/1040841X.2013.867830

Figure 2. Published risk factors in terms of odds ratios, with 95% confidence intervals shown. Factors with p40.05, with 95% confidence interval inclusive of 1.000, or with a p value not specified are not included (see Table 1). Factor relating to herd genetics, sex, breed or non-farm animal infection are not included. All data is with respect to cattle, unless labelled as follows: (S), Sheep; (G), Goats. Data corresponds to bold data in Table 1.

1940; Table 2). Data for MAP survival in slurry (liquid manure) indicates increasing temperature to have a profound negative effect on MAP viability in this medium (Jorgensen 1977; Olsen et al., 1985; Table 2), but similar studies on non-liquified feces are lacking, although some evidence for the utility of thermophilic composting for the removal of viable MAP exists (Grewal et al., 2006; Table 2).

Considering the importance of these basic survival data, the lack of similar examinations of MAP survival in this most basic and epidemiologically important of substrates is surprising and new studies are warranted. Experiments using caprine feces, again by Lovell et al. (1944), showed a lesser survival time for MAP in relation to similar experiments in the same study using bovine

Excreta Feces Feces Feces Feces Feces/soil Feces Urine/feces Urine Urine Slurry Slurry Slurry Slurry Slurry Slurry Compost Wider environment Desiccated culture Desiccated culture Desiccated culture Distilled water Tap water Tap water Pond water/mud Lake water River water River water River flood-water Dam water in trough Dam water in trough Sediment (dam water) Sediment (dam water) Soil core (top 10 cm) Natural pasture plots Natural pasture plots Natural pasture plots Natural pasture plots Natural pasture plots

Substrate NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS NS Type NS NS NS Type Type Type Type Type Type Type Type Type Type

Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Sheep Sheep Sheep Sheep Cattle Sheep Sheep Sheep Sheep Sheep

Culture Culture Culture Culture Culture Culture Culture Culture Intestine Intestine Intestine Feces Feces Feces Feces Culture/slurry Slurry Feces Feces Feces Feces

Feces Feces Feces Feces Culture Feces Culture Culture Culture Feces Culture Culture Culture Culture Feces/culture Feces/culture

Medium

200 mg dw 200 mg dw 100 mg dw NS 25 mg dw NS NS NS NS NS NS 103 ml1 103 ml1 103 ml1 103 ml1 106 cm2 106 cm2 105 cm2 104–105 cm2 104 cm2 104 cm2

NS NS NS 102–106 g1 NS NS 25 mg dw 25 mg dw NS NS 106 g1 106 g1 104 g1 104 g1 106 g1 106 g1

CFU

to 23 C

to 21 C

to 23 C to 30 C to 18 C

38 C Room temp. Up to 44 C Amb., 9–26 C 38 C Amb., 9–26 C Amb., 9–26 C NS Amb., 7 to 18 C Amb., 7 to 18 C Amb. (LW UK) Amb. (Aust.) Amb. (Aust.) Amb. (Aust.) Amb. (Aust.) Amb. (Chile) Amb. (Aust.) Amb. (Aust.) Amb. (MS Aust.) Amb. (LS Aust.) Amb. (Aust.)

Amb., 3 Amb., 3 Amb., 7 70 C NS Amb., 3 38 C 38 C NS Amb., 3 5 C 15 C 35 C 53–55 C 525 C 25–55 C

Temperature*

0–70% shade,plant cover 0–70% shade,plant cover No shade/plant cover No shade/plant cover 70% shade, no plant cover

Sun shaded,rain exposed Sun/rain exposed Sun shaded,sealed No shade 70% shade No shade 70% shade

Dark Not in dark Directly sun exposed Sun shaded,sealed Dark, pHs 5.0,7.0,8.5 Sun shaded,sealed Sun shaded,sealed

Sun/rain exposed Anaerobic Anaerobic Anaerobic Anaerobic Aerobic Aerobic

Dark Dark

Sun/rain exposed Sun/rain exposed,wet Allowed to dry

Further conditions (where specified)

517 1430 5–8# 273 425–578 273 273 632 135 163 113 112 252 252 336 425 182–224 182–224 14–35 70 70–84

152 208 163 105 334 67 530 530 57 246 252 98 21 51 56 53

Last measured survival (days)

Larsen et al., 1956 Larsen et al., 1956 Larsen et al., 1956 Lovell et al., 1944 Larsen et al., 1956 Lovell et al., 1944 Lovell et al., 1944 Pickup et al., 2005 Lovell et al., 1944 Lovell et al., 1944 Lovell et al., 1944 Whittington et al., 2005 Whittington et al., 2005 Whittington et al., 2005 Whittington et al., 2005 Salgado et al., 2011a Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004

Lovell et al., 1944 Lovell et al., 1944 Lovell et al., 1944 Richards, 1981 Vishnevskii et al., 1940 Lovell et al., 1944 Larsen et al., 1956 Larsen et al., 1956 Vishnevskii et al., 1940 Lovell et al., 1944 Jorgensen, 1977 Jorgensen, 1977 Olsen et al., 1985 Olsen et al., 1985 Grewal et al., 2006 Grewal et al., 2006

Reference


I/III I/III I/III I/III II I/III I/III I/III I/III I/III

II

Strain

Cattle Cattle Cattle Cattle Cattle Goat Cattle Cattle Cattle Cattle Cattle Cattle Cattle Cattle Sheep/human Sheep/human

Source

Inoculum

Table 2. Survival of MAP ex-host detected using culture-based approaches.



Johne’s disease

II

CFU ¼ Colony forming units (approximated); NS ¼ not specified; dw ¼ total dry weight of culture added. *LW ¼ Late winter; MS ¼ Midsummer; LS ¼ Late summer; Amb. ¼ Ambient temperature; UK ¼ United Kingdom. Aust. ¼ Australia. # Assumes a mean of 12 h sunlight day-1.

Anaerobic

70% shade 70% shade No shade,some plant cover No shade,some plant cover Complete shade Complete shade 70% shade Complete shade 70% shade ‘High’ pH (after 14 days) ‘Low’ pH (after 14 days)

Amb. (MS Aust.) Amb. (LS Aust.) Amb. (MS Aust.) Amb. (LS Aust.) Amb. (MS Aust.) Amb. (LS Aust.) Amb. (Aust.) Amb. (LS Aust.) Amb. (LS Aust.) 5 C 30–37 C 10–20 C 41–42 C 22 C 104 cm2 104 cm2 104 cm2 104 cm2 104 cm2 104 cm2 104 cm2 104 cm2 104 cm2 NS NS 4106 g1 NS 103 ml1 Feces Feces Feces Feces Feces Feces Feces Feces Feces Culture Culture Culture Feces Feces I/III I/III I/III I/III I/III I/III I/III I/III I/III

Type Type Type Type Type Type Type Type Type NS NS Type NS NS Sheep Sheep Sheep Sheep Sheep Sheep Sheep Sheep Sheep Cattle Cattle Cattle Cattle Cattle Pasture soil (boxed) Pasture soil (boxed) Pasture soil (boxed) Pasture soil (boxed) Pasture soil (boxed) Pasture soil (boxed) Pasture soil (boxed) Grass (pasture) Grass (pasture) Silage Silage Silage Biogas fermenter Dip fluid (pH 12.4)


84 112–168 14–84 84–112 196 385 84 168 72 14 514 54 60 14

Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Whittington et al., 2004 Katayama et al., 2000 Katayama et al., 2000 Khol et al., 2010 Slana et al., 2011 Eamens et al., 2001

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9

feces (Table 2). Although it had been suggested around this time that JD may be caused by either of two closely related organisms, it was only decades later it was established that MAP isolates could be genetically and phenotypically designated as one of two major strain types, S type (sheep) and C type (cattle) (Collins et al., 1990). These groups were subsequently reassigned as Type I/III and Type II strains, respectively, as it became apparent that the species of origin was not always an accurate indicator of strain type (Stevenson et al., 2002). The lack of strain type identification in the majority of the studies described in Table 2 highlight again the lack of coverage of this basic area of survival, and this makes extrapolation of particular studies difficult, especially as speed of growth is a key differentiator between these strain types, Type II being the faster of the two (Stevenson et al., 2002). Pure cultures of untyped MAP placed into previously sterilized pond water have been shown to survive for over 9 months at room temperature and MAP was able to be cultured from river water containing intestinal scrapings kept outside and exposed to rain and cold weather for up to 163 days (Lovell et al., 1944). Further studies showed similar lengthy survival periods, with cultured bacilli surviving for between 14 and 19 months in tap water at 38 C under a variety of pHregimes and in desiccated culture for over 47 months (Larsen et al., 1956; Table 2). More recent studies have highlighted the survival of Type II MAP inoculated into sterile freshwater lake water columns, finding cultivable MAP even after 632 days (Pickup et al., 2005; Table 2). Tests by Whittington et al. (2004, 2005) on Type I/III MAP strain survival performed in Australia in dam water and sediment showed survival of the strain for 48 weeks under shade versus 36 weeks when partially exposed, and that survival in sediment was between 12 and 26 weeks longer than that in the corresponding water column (Table 2). Similar survival experiments, but assessing Type I/III MAP strain persistence on soils and grasses, showed survival for a maximum of 55 weeks when covered, but less than 2 weeks when fully exposed to sunlight in the absence of even plant cover. The fact that this period was extended by a full 30 weeks in the presence of herbage revealed the importance of cover in these harsh exposed conditions (Whittington et al., 2004). Although it is well known that exposure to UV radiation can harm cells, the use of sheep pellets with only the surface layer exposed to such radiation suggested that it was thermal flux rather than UV that likely was most significant in reducing the viability of the MAP cells present (Whittington et al., 2004). Survival on plant material itself has been little studied, but Whittington et al. (2004) found that a TypeI/III MAP strain could be cultured for up to 24 weeks from aerial parts of grasses following inoculation of the surrounding soil. However, as these inocula could have been deposited on the leaf at any time from during germination through the soil surface to during recent rainfall (i.e. through splash deposition), the actual period of persistence on the plant surface itself was undetermined. The storage of MAP-containing manure and its subsequent use in materials for bedding, feed and fertiliser has led to research on MAP survival during and at the end of these processes. Such studies have shown the potential for MAP


10


survival in silage should the ensilage process not be properly executed (Katayama et al., 2000; Table 2). A study using DNA-intercalating dye (propidium monoazide – PMA) to selectively bind and effectively remove any exposed DNA (i.e. that not contained within an intact cell) from analysis (Nocker et al., 2007) to attempt to assess the presence of viable MAP using molecular techniques following ensilage did indicate that MAP survived the process when properly performed, but also found that the complexity of the substrate and its microbial community compromised the PMA approach and so these data should be viewed with caution (Cook et al., 2013). However, such methodologies are increasingly used and detection of MAP in simpler matrices is possible (Kralik et al., 2010) and may be preferable in future molecular studies to avoid the lengthy MAP culture periods required. The survival of MAP within a farm-scale biogas plant, producing digestate for possible application on land or for use as feed or bedding showed the survival of MAP after 2 months but not after 6 months (Slana et al., 2011), and the general storage of manure outdoors has been shown to be a significant risk factor in the presence of JD within herds (Johnson-Ifearulundu & Kaneene, 1999; Table 1; Figure 2) although the reasons for this were not discussed. However, the detection of viable MAP after 175 days of thermophilic composting may indicate both time and temperature to be a factor here (Grewal et al., 2006; Table 2). Again, more studies are required to elucidate this matter, especially with respect to ‘on-the-ground’ real-life treatment of manures in both smaller and larger scale farms across a range of climates. pH and soil composition The effects of the chemical composition of soils on the survival of MAP have been debated for many years, with both soil pH and iron content regularly proposed as factors affecting MAP persistence in the environment. However, studies performed in a way which precludes the introduction of new MAP organisms into the system are few in comparison with those that relate environmental factors to continued JD presence in herds and flocks. Persistence of the disease in cattle herds in the US was found to be elevated in regions with acid soils relative to those with alkaline, calcareous soils (Kopecky, 1977) and similar associations have been found in other studies (Michel & Bastianello, 2000; Reviriego et al., 2000). JohnsonIfearulundu & Kaneene (1999) found associations between both pH and iron of the soils on which dairy herds in Michigan were reared and herd risk for JD infection, showing a 1.4% increase in the risk of a herd being positive for every 1 ppm increase in soil iron content, and a 4% increase in the numbers of MAP-positive cows associated with every 10 ppm increase in soil iron content. Similarly, an increase of 0.1 pH units was associated with a 5% decrease in the number of test positive cows, and the application of lime to pasture (to increase soil pH) was shown to be protective against JD (Table 1) and associated with a 72% reduction in the number of positive cattle. A herd raised on lime-treated pasture was one-third as likely to test positive compared to a herd raised on untreated pasture (Johnson-Ifearulundu & Kaneene, 1998,


1999; Table 1; Figure 2). This said, these studies showed neither pasture soil pH nor iron content to be significantly associated with the application of lime to pasture soils, perhaps indicating a non-pH related action of lime application, and others showed no effect of the prior application of lime to soils in terms of subsequently inoculated MAP survival in those soils (Whittington et al., 2004), although again these studies can be expected to have assessed different strain types. Scott et al. (2006, 2007) found a moderate association between reduced risk of infection and increasing pH and aridity in Canada, and suggested a non-linear relationship between organism viability and soil factors. However, studies in Australia and the Netherlands found that pH was not significant in terms of MAP presence in soil and feces on sheep farms (Whittington et al., 2004) or in the prediction of herd seroprevalence for MAP in cattle (Muskens et al., 2003) or sheep (Dhand et al., 2009a). Subsequent studies evaluated these associations in greater depth (analyzing 87 sheep farms in Australia for 20 differing soil characteristics) found a positive linear relationship between JD prevalence in sheep and organic C content of the soil, while parent soil type was not significantly related to JD prevalence (Dhand et al., 2009a; Table 1; Figure 2). It was suggested that this might be due to increases in pasture growth and therefore ground-cover for shed MAP (increasing its potential for survival), or to the positive relationship between the waterholding capacity of soil and organic carbon. However, the possibility of increased pasture growth leading to increased use by the farmer for grazing and therefore greater JD prevalence in the grazing stock was similarly valid. Higher clay content has also been associated with higher JD levels, and high sand content and nitrogen levels associated with the opposite (Dhand et al., 2009a). In this case, the authors suggested that the larger surface area of clay soils allowed for increased binding of MAP in the upper layers of soils, such that they remained available for ingestion by livestock. Stock animals normally consume significant amounts of topsoil along with surface pasture (McGrath et al., 1982) and this has impacts in terms of direct consumption of MAP along with implications for surface run-off and wider dissemination of the organism (Pickup et al., 2006; Figure 1). The hydrophobic nature of MAP allows the organism to bind to like-charged particles to a greater degree than many other organisms and this reduces its potential to be transported away from the soil surface. Further, under slightly acidic conditions, MAP has been shown to carry a strong negative charge, and the presence of positively charged functional groups associated with iron coatings has been hypothesized to further increase MAP retention in such soils (Bolster et al., 2009). The use of lysimeters to assess MAP transport under rainfall in a loamy and a sandy soil showed that MAP was retained in upper soil layers and that even under high rainfall it took 2 months for MAP to be detected in the soil leachate, collected 90 cm below the surface. Transport through sandy soil under ‘‘slightly’’ acidic conditions was more rapid than in the more acidic loamy soil. In all cases, more MAP was detected on the surface grass than in the leachate. On dismantling of the experiment, at which point

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MAP had not been found in leachate for over 10 months, MAP was found only in the upper layers of soil (Salgado et al., 2011a). Other researchers have theorized that the presence of iron aids MAP survival, helping to compensate for the organisms’ poor iron chelating ability, demonstrated by the general requirement for the iron chelator mycobactin for in vitro culture of MAP (Merkal & Curran, 1974). However, as an obligate pathogen, it is persistence outside the host rather than multiplication that is minimally required and so the necessity of a higher Fe availability for MAP growth could be expected to be minimal (Dhand et al., 2009a; Thorel et al., 1990) and the macro and micro-nutrient make-up of soils and consequently of the animals feeding on them and on the process of the disease once acquired have been investigated elsewhere (reviewed by Lugton, 2004b). However, it is clear in terms of soil chemistry that a lower pH combined with reducing conditions increases Fe availability. Whether or not this leads to more Fe for direct use by the organism or a specific increase in binding of MAP to soil particles, the same effect is apparent in most reported cases: a higher retention of MAP in soils at lower values of pH (Dhand et al., 2009b) and/or under reducing conditions (Lugton, 2004a; Table 1). It should also be considered that under in vitro culture, the growth characteristics, acid resistance and protein expression of MAP have been shown to vary when conditions including pH are altered (Sung & Collins, 2003), and these changes may further affect adsorption to soil particles, although specific effects have yet to be demonstrated. An additional wide variety of biotic and abiotic factors undoubtedly play roles in the varied rates of survival of MAP in soils and waters and these can be expected to include variations in temperature (e.g. freeze-thaw cycles), soil structure, porosity and particle size, soil moisture content, lateral movement, biofilm potential, predation and competition, amongst others. For other microbial pathogens these have been reviewed elsewhere (Bradford et al., 2013; Falkinham, 2009; Ferguson et al., 2003; Jamieson et al., 2002; Mawdsley et al., 1995; Unc & Goss, 2004), but additional specifically MAP-related studies in these areas are lacking. This said, it has recently been established that MAP (or due to the molecular approaches used, at least MAP-derived DNA) is widely distributed in UK soils and waters, and various characteristics of the samples and sites tested were positively associated to MAP DNA presence (Rhodes et al, 2013). Although these factors included cattle (but not sheep) density, there were additional factors related to latitude and longitude that could not be explained by cattle density alone. These included significant associations between increases in MAP presence and higher pH, temperature and soil bulk density, and with lower rainfall and organic matter content, although many of these were inter-related (Rhodes et al, 2013). Further confounding issues such as land selection for grazing, selecting against the use of nutrient poor, acid grasslands were suggested as influencing these data and were discounted in favor of cattle density (Rhodes et al., 2013), thus removing the discrepancy between this and earlier studies (JohnsonIfearulundu & Kaneene, 1999; Scott et al., 2006; Table 1; Figure 2) indicating higher pHs to be detrimental to JD presence. Land use comparisons in the same study showed

11

statistically significant differences between MAP DNA presence in soils from land used for agricultural/horticultural purposes and from both land classified as bog and as shrub heath, although no significant differences were shown in relation to agricultural use/management when all sites were taken into account (Rhodes et al., 2013). Thus, although somewhat limited by the complexity of the sample type and geography as well as the molecular methodologies required to be used, this study indicates that that assessment of the levels of movement of viable MAP into the general environment from JD-infected farms is something that requires further investigation and similar data from other regions would be of interest. The standing or background levels of MAP in pristine environments should also be investigated. Dormancy and sporulation Inconsistencies in MAP detection via culture from both waters and soils tested within studies of strain survival in Australia (Whittington et al., 2004; 2005) were suggested to be possibly due to MAP existing in a dormant state. Dormancy genes have been found in the MAP genome and it is well known that other species of Mycobacteria can respond to stress such as acidic conditions, nutrient deprivation or hypoxia by entering and persisting in a dormant state (Lamont et al., 2012). Studies describing the formation of endospores in other Mycobacteria (Ghosh et al., 2009) have not been able to be repeated by other laboratories, but Lamont et al. (2012) were successful in identifying and describing a previously undocumented spore-like morphotype of MAP, formed under conditions of nutrient starvation. These spores were able to retain both infectivity and ability to germinate, findings which were independently verified. Although it is clear that more studies are required to investigate these phenomena and their implications in terms of persistence and infectivity, there appears to be significant potential for MAP to achieve longevity in the environment via sporulation or dormancy.

Wildlife vectors Prevalence in ruminants Although the existence of MAP infection and paratuberculosis in wildlife has long been established, with a recent metaanalysis of studies reporting MAP prevalence in wildlife in general to be at 2.41% of those animals tested (Carta et al., 2013), its importance in the spread of disease within and between livestock herds has not been definitively determined. JD has always been considered a disease of both wild and domestic ruminants, and natural JD infection is well documented across the globe in a range of ruminants including red, white-tailed, sika, fallow and roe deer, as well as in alpaca, tule elk, bison, ibex, saiga antelope, moufflon and bighorn sheep (Table 3). Although cases have not been reported in all ruminant species to date, all ruminants are believed to be susceptible to MAP infection (Manning, 2011). Whereas ruminants such as wild goat and antelope have been reported to become emaciated but with no evidence of diarrhoeal symptoms (Griffin, 1988), MAP-associated diarrhoea has been repeatedly reported in red deer (Griffin, 1988;

Bovidae Bovidae Bovidae Bovidae Bovidae Bovidae Bovidae Bovidae Bovidae Bovidae Bovidae Camelidae Camelidae Camelidae Camelidae Camelidae Cervidae Cervidae Cervidae Cervidae Cervidae Cervidae Cervidae Cervidae Cervidae Cervidae

Barbary sheep Bison Yak Zebu Water buffalo Ibex Gnu Rocky mountain goat Bighorn sheep Mouflon Saiga antelope Bactrian camel Dromedary camel Llama Guanaco Alpaca Moose Axis deer Roe deer Red deer Tule elk Sika deer Fallow deer White tailed deer Key deer Reindeer

Order

Artiodactyla Artiodactyla Carnivora Carnivora Carnivora Carnivora Carnivora Carnivora Carnivora Carnivora Carnivora Carnivora Cingulata Didelphimorphia

Common name

Non-ruminant mammals Wild boar Domestic pig Coyote Cat (feral) Badger Striped skunk Stoat Weasel Ferret Raccoon Brown bear Red fox Nine-banded armadillo Opossum

Non-ruminant animals MAP infected as confirmed by culture

Family

Common name

Ruminants (all Order Artiodactyla)

YES YES YES YES YES YES YES YES YES YES YES YES YES YES

Tissues

MAP presence

Boever & Peters, 1974 Buergelt et al., 2000 Almejan, 1958 Katic, 1961 Katic, 1961 Ferroglio et al., 2000 Rankin, 1958 Williams et al., 1979 Williams et al., 1979 Machackova et al., 2004 Dukes et al., 1992 Katic, 1961 Amand, 1974 Appleby & Head 1954 Salgado et al., 2009 Ridge et al., 1995 Soltys et al., 1967 Riemann et al., 1979 Machackova et al., 2004 Pavlik et al., 2000 Jessup et al., 1981 Thoen et al., 1977 Riemann et al., 1979 Chiodini & Vankrunigen, 1983 Quist et al., 2002 Katic,1961

Example reference publication

NT NT NT YES NT NO YES NT NT YES NO YES YES YES

Feces

NT YES NT NT NO NT YES YES YES NT NT YES NT NT

lesions

JD consistent

Notario et al., 2010 Miranda et al., 2011 Anderson et al., 2007 Corn et al., 2005 Beard et al., 2001a Corn et al., 2005 Beard et al., 2001a Beard et al., 2001a De Lisle et al., 2003 Corn et al., 2005 Kopecna et al., 2006 Beard et al., 2001a Corn et al., 2005 Corn et al., 2005

example

Reference


Sus scrofa Sus scrofa domesticus Canis latrans Felis familiaris Meles meles Mephitis mephitis Mustela erminea Mustela nivalis Mustela putorius furo Procyon lotor Ursus arctus Vulpes vulpes Dasypus novemcinctus Didelphys virginiana

Species

Ammotragus lervia Bison bison Bos grunniens Bos indicus Bubalus bubalis Capra ibex Connochaetes albojubatus Oreamnos americanus Ovis canadensis Ovis mousimon Saiga tatarica Camelus bactrianus Camelus dromedarius Llama glama Llama guanicoe Lama pacos Alces alces Axis axis Capreolus capreolus Cervus elaphus Cervus elaphus nannodes Cervus nippon Dama dama Odocoileus virginianus Odocoileus virginianus clavium Rangifer tarandus

Species

Table 3. Wildlife or exotic species with evidence of PTB. These are wild free-ranging or captive (includes herds) confirmed MAP infected via culture-based methods.



Macropus eugenii Macropus fuliginosus Trichosurus sp. Erinaceus europaeus Lepus europeus Oryctolagus cuniculus Sylvilagus floridanus Diceros bicornis Equus asinus Macaca arctoides Papio sphinx Apodemus sylvaticus Microtus arvalis Mus musculus Rattus norvegicus Rattus rattus Sigmodon hispidus Thomomys talpoides Blarina brevicauda Crocidura suaveolens Tadorna variegata Gallinago gallinago Corvus corone Corvus frugilegus Corvus monedula Passer domesticus Sturnus vulgaris

Diprodontia Diprodontia Diprodontia Erinaceomorpha Lagomorpha Lagomorpha Lagomorpha Perissodactyla Perissodactyla Primates Primates Rodentia Rodentia Rodentia Rodentia Rodentia Rodentia Rodentia Soricomorpha Soricomorpha

Anseriformes Charadriiformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes

YES YES (GI tract) YES YES YES YES (GI tract) YES (GI tract)

YES YES YES YES YES YES NT NT YES YES (GI tract) YES YES YES YES YES YES YES (GI tract) YES (GI tract) YES YES NO NT YES YES NT NT NT

NO NO YES YES YES YES YES YES NT NT YES YES NT NT NO NT NT NT NO NT

SLIGHT NO NO

NO

NT NT

YES YES NT NT NO YES#

YES YES YES YES YES YES NT NT YES

Nugent et al., 2011 Corn et al., 2005 Beard et al., 2001a Beard et al., 2001a Beard et al., 2001a Corn et al., 2005 Corn et al., 2005

Cleland et al., 2010 Cleland et al., 2010 Nugent et al., 2011 Nugent et al., 2011 Nugent et al., 2011 Beard et al., 2001b Raizman et al., 2005 Bryant et al., 2012 Van Ulsen, 1970 McClure et al., 1987 Zwick et al., 2002 Beard et al., 2001a Pavlik et al., 2010 Florou et al., 2008 Beard et al., 2001a Florou et al., 2008 Corn et al., 2005 Corn et al., 2005 Corn et al., 2005 Pavlik et al., 2010

NT: Not tested. GI: Gastro-intestinal. *Data shown for culture or microscopic visualisation from feces of viable MAP. # Acid fast bacteria-positive macrophages identified within intestinal lamina propria. Species not specified in the reference. Although the species listed above would appear the most likely when sampling region is considered, there are many alternative species both with and outside the genus that may in fact be correct.

Tammar wallaby Grey kangaroo Brushtail possum Hedgehog European brown hare Rabbit Eastern cottontail Black rhinocerous Pygmy ass Stumptail macaque Mandrill Wood mouse Common vole House mouse Norway rat Black Rat Hispid cotton rat Pocket gopher Northern short-tailed shrew White toothed shrew Birds Paradise shelduck Common snipe Crow Rook Jackdaw House sparrow European starling


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Macintosh & Griffin, 2010) and the core features of infection are believed to be generally consistent across all ruminants. Rapid development of the farming of deer and other wild ruminants in some parts of the world has led to JD problems in free-range systems, with red deer being particularly susceptible, individuals sometimes developing profuse diarrhoea in their first year with death following the observation of clinical signs within as little as 2 weeks (Macintosh & Griffin, 2010). As with other domestic livestock, the direct effect of herd population density on exposure of young animals to the organism is a primary risk factor (Macintosh et al., 2004). This is exacerbated by the non-host-specific nature of MAP strains (Stevenson et al., 2009) if wild ruminants come into contact with land or water contaminated by MAP from domestic livestock (Whittington et al., 2005) and the reverse can also be expected to be the case. Prevalence in non-ruminants The prevalence of MAP in non-ruminants has been reported increasingly over the past decade or so, with suspicion that non-ruminant reservoirs of MAP may be present and linked to its transmission to domestic stock. As with ruminant infection, grazers ingest the organism through feeding on pasture shared with already infected animals or by sharing housing and the infection is further passed on through predation and scavenging to non-grazing animals and birds. In 1997, 67% of wild rabbits culled near Perth, Scotland, were found to be infected with MAP and also displayed pathology consistent with paratuberculosis (Greig et al., 1997, 1999). Shortly afterwards, multiple predatory and scavenging animals and birds in the same region were shown to harbour MAP, possibly due to feeding on infected rabbits (Beard et al., 2001a; Table 3). A study based in the US (Anderson et al., 2007) looking at the tissues of 212 scavenging mammals across six genera found an average of 38% of animals to be PCR positive when tested using a variety of sequences specific to MAP, the authors attributing this to the presence of viable MAP in the digestive tracts of these animals at some time. However, only one of the 212 animals tested was found to be culture-positive, indicating that in the majority of cases the MAP cells had been digested without them (a) infecting the new ‘‘host’’ or (b) retaining the potential to be shed in viable form. Other researchers have obtained MAP-positive cultures from non-ruminant animals and birds (Table 3) with one study finding MAP-positive cultures across a range of nonruminants, encompassing nine species of mammal and three species of bird; the organism being isolated from liver and fecal samples as well as from the gastrointestinal tract. From these species, shedding was demonstrated in raccoons, armadillos, opossum and a feral cat (Corn et al., 2005). Due to the low prevalence and volume of wildlife-derived shedding, additional contamination of already infected farm environments was deemed to be of negligible impact and only infection of pristine farms had epidemiological significance (Corn et al., 2005). This said, of the 41 non-ruminant species


from which MAP has been isolated to date, viable MAP has been confirmed in the feces of 16 of these species (of 21 tested) with a further 6 species showing evidence of MAP infection in tissues (Table 3). Repeated discovery of MAP in the lymph nodes of carnivores (Beard et al., 2001a) coupled with associated changes in the histopathology of the host indicates the likelihood of chronic infection of predatory animals and other non-ruminants, with the organism residing in the lymphoid tissue of the gut as observed in the early stages of infection within ruminants, and numerous nonruminant species have been shown to have the potential to spread MAP throughout the environment via passive, mechanical or multiplicative infective transmission. Thus, the literature is somewhat contradictory with respect to the significance of the potential for transmission of MAP via socalled ‘dead-end’ hosts, at least in the absence of culturepositive samples (Carta et al., 2013; Salgado et al., 2011b; Stevenson et al., 2009). Associations between MAP strains derived from different species have shed some light on this issue. Although it was found that there was no apparent difference between cattle and rabbit strains during a farm survey in Scotland, only 3 of 14 farms with a history of JD yielded infected rabbits (Greig et al., 1999). These results were supported a decade later by the finding that MAP isolates exhibited relatively little genetic diversity in comparison to similar pathogens, and found that MAP isolates from multiple different host species on the same property, both wild and domestic, had identical genotypes (Stevenson et al., 2009), supporting the potential for interspecies transmission. Isolate typing showed all isolates to be Type II, despite isolation from multiple species including sheep. Studies assessing the potential for cattle and sheep to become infected by MAPcarrying rabbits through stock grazing preferences in the presence of varying accumulations of rabbit feces showed that both sheep and cattle consumed rabbit feces when grazing and unlike other species, cattle showed no aversion to grazing in areas with high numbers of rabbit feces (Daniels et al. 2001, 2003; Judge et al., 2005). This was regardless of sward height, which has been found previously to affect grazing preference of cattle with respect to badger latrines and other animal deposits (Forbes & Hodgson, 1985; Hutchings & Harris, 1997; Hutchings et al., 1998). As infected rabbits have been shown to shed large numbers of MAP in their feces (estimated to contribute over 106 CFU of MAP per hectare per day (Daniels et al., 2003)) the consumption of MAP via rabbit feces by both sheep and cattle appears likely. However, infection leading to JD via this route has yet to be shown and further research to examine this would be of benefit. Thus although it is likely that transmission between livestock and wildlife occurs at some level, with potential for reservoirs of infection in both the natural and the immediate farm environment, apart from those animals that may be infected and shedding (Table 3; Figure 1), the majority are simple mechanical vectors that are expected to have little to no relative impact in terms of spread within an infected herd. This impact might reasonably be expected to increase when herd to herd spread is considered but more studies in this area are again required.

DOI: 10.3109/1040841X.2013.867830


Amoebal, protozoan, nematode and invertebrate interactions Due to its obligate intracellular pathogen status, persistence outside the animal host has generally been considered as a matter of endurance, and unless indefinite dormancy or conversion to a spore-like persistent form is involved, it has been accepted that it is simply a matter of time (outside the animal host) before the pathogen succumbs and dies (even though this might take years). In other words, it is survival rather than multiplication that takes place once MAP has been shed. However, the interaction of Mycobacterium avium with environmental amoebae such as Acanthamoeba spp. has been shown to lead to MAP replication within the protective walls of amoebae (Cirillo et al., 1997). Normally these protozoa will prey upon such bacteria, but there are certain microbes that have shown resistance to amoebae, and these have been termed amoeba resistant microbes. More specifically it has become clear recently that amoeba-resistant MAP exist and these have been shown to survive in free-living amoebae (FLA) that can exist in both waters and soils (Rowe & Grant, 2006; Salah et al., 2009). The benefits of intracellular life in the environment for MAP are multiple, as free-living amoeba can form persistent cysts, potentially protecting the MAP from both physical and chemical perturbations and stresses such as UV exposure and desiccation, as well as from chemical treatments administered to disinfect water such as chlorine and hydrogen peroxide. Due to the potential for the amoebae to supply nutrients in low nutrient environments such as freshwater, this association has been suggested to be an effective adaptive mechanism to both the dispersal and persistence of pathogens such as MAP. Thus free-living amoebas have been referred to as ‘Trojan Horses’ in microbial ecology (Barker and Brown, 1994) and in this way free-living amoebae could act as protectors and vectors for MAP, facilitating survival and replication within soil or water bodies and subsequent entry into the host (Salah et al., 2009). This said, no work has been published to date regarding this action in vivo and studies are required to investigate whether the presence and multiplication of MAP within FLA occurs in the environment, and further whether it is significant, specifically with respect to infection of higher animals in the natural environment. MAP has also been found in or on a range of invertebrates collected from farms, pastures and slaughterhouses contaminated with MAP. These include adult and larval diptera (Fischer et al., 2001; 2005), and earthworms (Fischer et al., 2003). Other studies have shown uptake of MAP under in vitro conditions by other organisms including nematodes, which have been suggested to have the potential to increase the infectivity of MAP through their ability to invade the mucosa of ingesting animals (Whittington et al., 2001). However, in vivo data are lacking and other similar studies have tried and failed to find invertebrate carriers in MAP-infected environments (Fischer et al., 2004).

Implications and knowledge gaps Research has shown that pooled fecal sampling followed by MAP culture is the most effective method for the monitoring of JD status of dairy herds and as such, when possible and

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practicable, this should be the primary method for detection of JD within non-clinical samples in general. Molecular methods can be extremely useful, especially when used to complement rather than replace other methods. However, in low prevalence herds or areas, molecular methods may be preferable over culture due to their potential for higher sensitivity and rapidity. Recognition of the limitations of molecular techniques in their ability to differentiate DNA from viable MAP from ‘‘naked’’ DNA from non-viable MAP has not always been considered in studies and inferences which do not follow sometimes made, especially as it is known that DNA can persist for extended periods outside the cell (Dale et al., 2002). The use of intercalating dyes such as propidium monoazide shows potential at least in some cases for the amelioration of this ‘‘problem’’ (Kralik et al., 2010), but field studies with respect to the use of such methods with MAP and/or JD are lacking. The use of immunological testing (ELISA) of serum and bulk milk is commonly used and allows for a much more rapid diagnostic of JD infection, but can be unreliable when performed in isolation i.e. without confirmatory testing, and with respect to the focus of this review, cannot be applied to environmental samples. Thus it is likely that a combination of methodologies will continue to be used on and off-farm, and although MAP culture may be the gold standard, more rapid and sometimes less reliable tests are likely to remain the primary method for MAP detection. Although the more direct factors in Johne’s transfer have long been considered in the management of JD infection, such as the particular susceptibility of young animals and the occurrence of intermittent pre-clinical shedding, it is clear that (although extremely important) hygienic practice alone cannot be used to effectively control the spread of JD within a herd. The reduction in survival of MAP in slurry in the presence of urine as opposed to in its absence may well inform waste storage approaches as may the clear reduction in survival of MAP in conditions of higher temperature over extended periods in most media. The storage and use of on farm waste is obviously key to reduction of the spread of disease and the spreading of liquid manure slurry on fields should be considered in terms of MAP distribution. The long-term survival of MAP in both soils and waters indicates that wider farm land use and management have significant implications for the incidence of JD. Indeed the rotation of pasture has been shown to reduce JD incidence by roughly half (Johnson-Ifearulundu & Kaneene, 1999). An increase in pH, possibly with a concomitant reduction in iron availability appears to be a viable approach for the reduction of MAP persistence, although to date this has only been shown in correlative terms for pH and Fe in relation to herd presence. Thus application of lime to soils and pastures appears appropriate in terms of reducing JD prevalence in cattle herds, where we might assume Type II strains predominate (Johnson-Ifearulundu et al., 1998), but with others showing no effect of prior liming on TypeI/III MAP survival (Whittington et al., 2004) its precise action, if any remains unclear. In addition to pH change it may well also be that a top-dressing treatment of lime acts in a general antibacterial manner prior to interspersal throughout the soil, although no MAP-specific evidence exists for this to date for


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either strain type. Should this be the case, timing of the application of lime to fields is relevant. Movement of MAP away from the surface of pasture/soil in low clay, low organic C, high sand and silt soils appears to reduce on-farm infection, at least at the grazing surface, and so selection of land for use as pasture should consider this, and good drainage should be considered in all areas of land to be used for grazing, with a reduction in ‘‘natural’’ standing water available for consumption by cattle appropriate, reducing both extended MAP survival in feces-contaminated waters and the potential impact of amoebae-based MAP replication/persistence, which requires further study. This said, we can expect ‘removal’ of MAP from farms via e.g. runoff to lead to increased levels of the organism in the natural environment and the implications of this spread remain unclear, both for farm stock and for humans (Rhodes et al., 2013). However, whether the detection of MAP DNA in the environment reflects the presence of viable MAP remains to be seen and the effects of sporulation or dormancy are similarly unknown as are strain type effects. The testing of manure prior to spreading as organic fertilizer should be considered when only one or perhaps two years will separate crop growth and land use for pasture. The batch testing of silage prior to feeding may also be prudent. The potential for non-ruminant involvement in MAP spread is to be considered, but it would appear that the direction of movement of the pathogen is from cattle to wildlife in the main and mechanical transfer of MAP between non-ruminants is minimal at most. This said, areas of poor quality grazing should be limited to restrict rabbit feces ingestion (areas with the greatest rabbit numbers had the greatest risk of cattle consuming rabbit droppings and these areas corresponded largely with poor grazing (Daniels et al., 2003)). Non-domestic ruminant herds should be considered as are other domestic herds in terms of potential cross-herd infection. It is clear that a systematic approach to the assessment of environmental survival and persistence is lacking for MAP. Accurate and reliable estimates for clinical disease-causing infective doses, covering both strain types and host species and climate combinations are surprisingly absent in most cases and the assessment of the potential for wildlife-based distribution of MAP cannot be made with any certainty until such basic data are established, despite the existing extensive yet circumstantial evidence. Perhaps the lack of ‘‘standard’’ type strains for Type I/III and Type II MAP and a standardized approach would allow a better appraisal of the effects of factors influencing MAP spread and JD prevalence both on and off farm. Current approaches to control JD have not always taken a systems approach to the problem, instead focusing on aspects of husbandry or herd management. An inclusive approach to disease management that takes into account the persistence and transport of the organism within manure, derived waste products and in manure-contaminated on-farm soils and waters, land management, dispersal by domestic and non-domestic host species as well as general animal husbandry and land use is required on those farms where more traditional approaches to disease management have failed.


Additional implications of the wider spread and persistence of MAP in the environment should also be considered by the wider community.

Declaration of interest The authors report no declaration of interest.

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