Environmental reservoirs of pathogenic mycobacteria across the Ethiopian biogeographical landscape

Hayley C. King; Tanya Khera-Butler; Phillip James; Brian B. Oakley; Girume Erenso; Abraham Aseffa; Rob Knight; Elizabeth M. Wellington; Orin Courtenay

doi:10.1371/journal.pone.0173811

Abstract

The Mycobacterium genus comprises over one-hundred-and-fifty recognised species, the majority of which reside in the environment and many of which can be pathogenic to mammals. Some species of environmental mycobacteria may interfere with BCG vaccination efficacy and in tuberculin test interpretation. Examining biogeographic trends in the distribution of members of the mycobacteria across a number of physicochemical and spatial gradients in soil and water environments across Ethiopia using oligotyping identified differential distributions of pathogenic and significant species. The tuberculosis complex was identified in more than 90% of water samples and taxonomic groups implicated in lower BCG vaccine efficiency were core in both soil and water Mycobacterium communities. A reservoir of Mycobacterium bovis was identified in water, with up to 7.3×10² genome equivalents per ml. Elevation, temperature, habitat and vegetation type were important predictors of both soil and water Mycobacterium communities. These results represent the first step in understanding the potential risk of exposure to environmental mycobacteria that may undermine efforts to reduce disease incidence.

Citation: King HC, Khera-Butler T, James P, Oakley BB, Erenso G, Aseffa A, et al. (2017) Environmental reservoirs of pathogenic mycobacteria across the Ethiopian biogeographical landscape. PLoS ONE 12(3): e0173811. https://doi.org/10.1371/journal.pone.0173811

Editor: Srinand Sreevatsan, University of Minnesota, UNITED STATES

Received: September 7, 2016; Accepted: February 26, 2017; Published: March 23, 2017

Copyright: © 2017 King et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: Data are available at NCBI BioProject (http://www.ncbi.nlm.nih.gov/bioproject/371412).

Funding: TKB was in receipt of a Natural Environment research Council studentship. Hayley King was in receipt of a Biotechnology and Biological Sciences Research Council studentship.

Competing interests: The authors have declared that no competing interests exist.

Introduction

The Mycobacterium genus contains over one hundred and fifty recognised species which can be broadly grouped into fast-growing and slow-growing species or species complexes, based upon physiological, phenotypic and phylogenetic differences [1]. The Mycobacteria belonging to the slow-growing group are more commonly associated with host pathogenicity than those in the fast growing group [2] however, both groups contain pathogens causing pulmonary, disseminated and cutaneous diseases; particularly in immuno-compromised hosts [3–6]. The majority of the mycobacteria are found in the environment across a range of soil types and water distribution systems [7–13] which act as a reservoir for potential human and animal infection [14–17]. In addition to the potential for disease transmission, prior exposure to certain mycobacteria, including members of the Mycobacterium avium complex, have been implicated in the reduction of BCG vaccine efficacy against adult pulmonary TB in humans [18–22].

The geographical diversity, composition and abundance of environmental mycobacteria reservoirs are not well described, despite the need to address the risk of exposure in the context of disease, BCG vaccine efficacy, and tuberculin skin testing. The few studies that have identified environmental or geographical hotspots are based on relatively small numbers of samples and over limited geographic ranges; often by applying decontamination cultivation techniques considered to only reveal a fraction of the true diversity [10,13,23–25]. To overcome these limitations (particularly for complex environmental microbial communities), targeted amplicon-based pyrosequencing and post-sequencing analyses can provide accurate sequence information. However, the Mycobacteria are a closely related group of organisms showing little variation in the 16 rRNA gene [26] limiting the use of this target as a tool for uncovering diversity of this group. Oligotyping is more specific in targeting regions of conserved entropy in a multiple sequence alignment in order to separate sequences of closely related organisms during down-stream analysis [27].

Based on this oligotyping approach, Mycobacterium-specific pyrosequencing data were analysed to identify biogeographical patterns at the sub genera and species complex level in soil and water samples collected across Ethiopia. Ethiopia encompasses highly diverse and extreme climatic and environmental conditions, with one of the highest prevalences of all forms of tuberculosis worldwide [28] with an incidence rate of 247 per 100,000 [29] and a high rate of extra pulmonary tuberculosis [30]. Ethiopia also has widespread sensitivity to mycobacteria in humans and animals which may be the result of exposure to environmental mycobacteria [28,31–33]. The aims of this study were (i) to quantify the biogeographical variation in slow and fast growing environmental mycobacterial communities, (ii) to evaluate the associations between geographical, climatic and edaphic measures and the observed mycobacterial diversity and composition and (iii) to identify potential biogeographical hotspots of known pathogenic or opportunistic mycobacteria pathogen groups.

Results

Processed sequences results

Mycobacterium genus and slow-growing mycobacteria pyrosequencing primers detected a minimum of 380 sequence counts per soil and water sample after quality control. Using the Mycobacterium specific primers, 57 oligotypes were identified in soil and 41 groups in water samples. The slow-growing mycobacteria specific assay identified an additional 14 species groups in soils and 16 species groups in water.

Core mycobacterial communities and pathogenic groups in soil and water

The core mycobacterial community members found in ≥90% of all soil and water samples were members of the Gordonae clade, Avium complex, and Mycobacterium angelicum. In addition, Mycobacterium riyadhense and Tuberculosis complex were also dominant in water samples (Fig 1).

Download:

Fig 1. Mean proportional abundance of core mycobacteria community members.

Proportional abundance of mycobacteria detected in > 90% of soil and water samples using the genus specific and slow growing specific primers respectively. Groups are ordered from left to right by increasing mean proportional abundance.

https://doi.org/10.1371/journal.pone.0173811.g001

44% and 38% of the Mycobacterium species groups detected in water and soils respectively are known to be pathogenic to humans. The most abundant Mycobacteria in both environments were Mycobacterium moriokaense, members of the Avium complex and the Parafortuitum clade. Mycobacterium barrassiae was as additional pathogen identified in soil. Tuberculosis complex members were present in all 42 water sample locations, and detected in two locations in soil where they contributed <0.2% of the total slow growing community in the latter substrate. QPCR demonstrated that the mean absolute numbers of mycobacteria genome equivalents were greater in the soil (1.60 ×10⁸ cells/gram; range: 9.47×10⁵ to 8.01×10⁸/gram) compared to in water samples (1.17 ×10⁴ cells/ml, range: 3.35×10¹ to 2.68×10⁵/ml) (F[1,71] = 25.33, P<0.001). M. bovis specific qPCR returned counts in the range of 1.5–7.3×10² gene copies per ml of water, and 2.9×10³ gene copies per gram in a single soil sample, indicative of a substantial reservoir source. The pathogens M. tuberculosis and M. canettii were not identified in any sample.

Biogeographical variation in mycobacterium community composition and diversity

The diversity of soil mycobacterial communities at individual geographical locations (Simpsons alpha diversity index: mean 3.24, SD 0.24) was significantly greater than the diversity in the water communities, at both the genus level (mean 2.16, SD 0.64, Welsch’s two sample T-test: t(62) = 1.99, P<0.001) and for the slow growing mycobacteria (mean 1.94, SD 0.64, Welsch’s two sample t-test: t(72) = 1.99, P = 0.07).

The difference in mycobacteria species diversity between locations were significantly greater in soil (mean 101.13, SD 15.05) than across water sites (mean 58.41, SD 24.06, t(62) = 1.66, P<0.001), despite significant geographical heterogeneity in each substrate (soils: ANOVA: F(4,17) = 3.01, P<0.05; water: (F(8,33) = 2.0, P = 0.07). In contrast, the number of slow-growing oligotypes between substrates (F(1,72) = 0.63, P = 0.42) and between geographic regions, were not significantly different (P>0.1 for all comparisons).

Environmental predictors of mycobacteria communities in water

Elevation (R² = 0.28, p<0.001), longitude (R² = 0.27, p = 0.021) and water temperature (R² = 0.37, p<0.001) were significant predictors of Mycobacterium community structure. Isoclines predicting elevation based on community dissimilarity were fitted to the ordination in the form of a GAM model (Adj R² = 0.45, P<0.001) (Fig 2). Of the categorical variables measured, only habitat type was a significant predictor of mycobacteria community structure (R² = 0.30, P<0.001).

Download:

Fig 2. The effect of environmental variables on the presence of slow growing mycobacteria.

NMDS ordination of slow growing mycobacterium water community’s highlighting the effect of each significant environmental variables upon the separation of the data points. GAM fitted isoclines represent predicted values for latitude and the size of each point represents the true value for this variable. A = mycobacteria communities in soil, B = slow growing mycobacteria in soil, C = mycobacteria in water, D = slow growing mycobacteria in water.

https://doi.org/10.1371/journal.pone.0173811.g002

Several known pathogen species and clades containing disease-associated members were found to significantly increase with increasing elevation (Fig 3). Beta diversity increased unimodally with altitude reaching a maximum at intermediate altitudes (2000-2500m) thereafter declining. Alpha diversity was not associated with any numerical variable.

Download:

Fig 3. The relationship between elevation and mycobacteria abundance.

Linear regression of the proportional abundance of mycobacteria in water against elevation.

https://doi.org/10.1371/journal.pone.0173811.g003

Environmental predictors of slow-growing mycobacteria communities in water

Slow growing mycobacteria community structure was significantly predicted by annual season temperature (R² = 0.25, P = 0.005), latitude (R² = 0.48, P< 0.001), pH (R² = 0.21, P = 0.01) and elevation (R² = 0.23, P = 0.026). Isoclines representing predicted values for elevation fitted to the NMDS ordination using a GAM model (R² = 0.22, P<0.001) showed a linear effect across the second NMDS axis (Fig 2). The significant categorical predictors of slow growing mycobacteria diversity included vegetation classification (R² = 0.54, P<0.001) and surrounding habitat type (R² = 0.63, P<0.001) (Fig 2).

Environmental predictors of mycobacteria communities in soil

Elevation (R² = 0.81, P<0.001, GAM R² = 0.80, P<0.001), annual season temperature (linear R² = 0.80, P<0.001), latitude (linear ² = 0.28, P = 0.05) and soil water content (R² = 0.57, P< 0.001) were the numerical predictors of soil mycobacteria community composition when fitted to a NMDS ordination model. The categorical predictors included vegetation classification (linear R² = 0.74, P<0.001), habitat type (R² = 0.50, P = 0.002) and annual season precipitation (R² = 0.28, P = 0.003) (Fig 2).

Alpha diversity was negatively correlated with soil water content (R² = 0.227, P = 0.02). Unimodal relationships were observed between the number of mycobacterium genome equivalents per gram of soil (pseudo R² = 0.317), community richness (pseudo R² = 0.43), and abundance, which was highest at altitudes between 2000m and 3000m, and soil Mycobacterium diversity showed a linear increase with elevation (R² = 0.50, P<0.001).

Several species known to cause disease and clades comprising disease-causing members were observed to significantly change in proportional abundance with changes in elevation: M. confluentis and M. goodie increased in abundance and M. sphagni decreased with increasing elevation (Fig 4).

Download:

Fig 4. The effect of elevation on proportional abundance of soil mycobacteria.

Linear regression of the proportional abundance mycobacteria in soil against elevation.

https://doi.org/10.1371/journal.pone.0173811.g004

Environmental predictors of slow growing mycobacteria communities in soil

Similar to the genus level analysis, elevation (linear R² = 0.30, P = 0.003), soil temperature (linear R² = 0.43, P = 0.002), longitude (R² = 0.48, P<0.001), soil water content (R² = 0.41, P<0.001), vegetation classification (R² = 0.63, P<0.001) and habitat type (R² = 0.45, P<0.001) were significant predictors of the slow-growing mycobacterium community in soils.

Alpha diversity increased with elevation (R² = 0.34, P<0.001) and with longitude (R² = 0.30, P = 0.01), and was correlated with the proportional abundance of Gordonae clade and the Avium complex (Avium complex: Pearson’s r = -0.53, P<0.001; Gordonae Complex: r = 0.52, P = 0.002). Areas with high proportional abundance of the Gordonae complex in the western locations in Ethiopia exhibited low proportional abundances of Avium complex members, and vice versa (Fig 5).

Download:

Fig 5. The proportional abundance of gordonae clade and avium complex in soil.

Linear regression of the proportional abundance of the Avium complex against Gordonae clade in soil.

https://doi.org/10.1371/journal.pone.0173811.g005

Discussion

In this study a combination of oligotyping and qPCR were applied to soil and water ecosystem samples to understand the biogeographical structure and diversity of Mycobacterium communities measured across Ethiopian biomes. Fifty seven unique oligotypes were identified in water and seventy one identified in soils from combining results of the Mycobacterium genus and slow-growing specific assays. Particular significance was attributed to the slow growing groups, as many are known opportunistic pathogens of humans and animals. A high diversity and widespread distribution of mycobacteria was observed, particularly of the Gordonae clade, Avium complex and Tuberculosis complex in these environmental substrates. Particularly surprising, and of potential concern to the public and veterinary health communities, was the presence and relatively high abundance of Tuberculosis complex members identified in 100% of the water sample locations, comprising human and cattle drinking water sources. M. bovis was also detected in 2 village soil samples. M. bovis is the causative agent of tuberculosis in both cattle and humans, and in Ethiopia high prevalences have been recorded [34,35]. The presence of M. bovis in these sources suggests that water may be a reservoir for potential onward transmission. M. bovis is most likely to be present through contamination by visiting livestock which has previously been observed in Spain [36]. The same infected water sources were used by local communities for drinking, washing and bathing, highlighting a potential for cross-contamination to humans in addition to other animals. The relative contribution of water-borne verses air-borne transmission is not currently known, however water and sediment in the water is suspected as infection sources to animals elsewhere [37]. Furthermore, M. bovis residing in soil for over 12 months can colonise mice when eaten [38].

Members of the Mycobacterium Avium Complex (MAC) are also linked to clinical infections in humans and their livestock [39,40]. MAC was widely detected across the Ethiopian locations; though the distribution and proportional abundance varied substantially. Altitude was a significant predictor suggesting that human settlements situated at high latitudes (>5.733m) and at higher altitudes (>2951m) may potentially experiencing increased risk of exposure. MAC is commonly isolated from drinking water before and after water purification treatments [41,42] highlighting that water, in addition to soil, may be an epidemiologically significant source of potentially infectious material. Prior exposure to MAC is thought to “mask” or “block” human BCG vaccine efficacy against M. tuberculosis by interfering with the protective interferon-gamma responses following BCG vaccination [9,11,19]. Globally, the efficacy of BCG tends to decrease with latitude which has been suggested to be a consequence of a corresponding increase in abundance of environmental Mycobacteria. [19]. The current study is the first step in testing for the associations between BCG vaccine efficacy and EM compositions across Ethiopian biomes. In particular, the current data suggest that there is greater risk of exposure to MAC through water at higher altitudes. To further these developments it would be beneficial to characterise the communities of mycobacteria in the environment along a gradient of BCG vaccine efficiencies to determine which species may be producing the vaccine masking effects that have been hypothesised.

Variations in the genus and slow-growing taxa diversity were shown to be significantly associated with altitude, latitude and broad descriptions of the surrounding habitat type and vegetation. Particular species groups dominated water sources at higher elevations, namely M. smegmatis, the Parafortuitum clade, the Simae clade and Vaccae clade, whereas M. rhodesiae and M. goodii dominated high altitude soils. Soil mycobacteria exhibited a monotonic pattern in alpha diversity, oligotype richness and Mycobacterium copy numbers, showing a sharp decline above 4000m altitude indicative of peripheral viable growth conditions. Similar patterns were observed with elevation in the beta diversity within water sources: community variability was highest at intermediate altitudes and where water temperatures ranged between 17–23°C. Opposing trends between comparative water and soil substrates were observed, with the Parafortuitum clade in the water samples showing a significant increase in proportional abundance with elevation and the reverse pattern being observed in soil samples.

Analysis of the soil slow-growing Mycobacterium communities illustrated an inverse relationship between the the Gordonae and Avium complex proportional abundances, which was strongly associated with longitude. Higher Gordonae and lower Avium complex abundance was observed in western regions of Ethiopia, whilst the opposite pattern was observed in eastern Ethiopian sites.

This preliminary study demonstrates the importance of using high resolution techniques such as oligotyping to differentiate closely related taxa towards understanding the distribution of, and exposure risk to potentially pathogenic organisms in the environment. The genus Mycobacterium presents a particular challenge as it is characterised by very limited interspecies genetic variability [43] and robust phylogenies created using 16S rRNA genes can only be achieved for a small number of species groups [43]. The significance of environmental mycobacteria as reservoirs for onward transmission of pathogenic and opportunistic species to humans and livestock is poorly understood. Here heterogeneities in the potential environmental exposure risk were highlighted, particularly in regards to M. bovis and Avium complex members in water shared by human and animals across Ethiopia.

Materials and methods

Environmental sampling sites

Soil and water samples were collected from 42 villages in 21 districts (kebeles) in 9 administrative regions of Ethiopia. These regions were selected to represent the country’s climatic and environmental diversity. Mean annual temperatures ranged from <7.5°C—>27.5°C, varying particularly with elevation (range: 359-2983m). Mean annual rainfall ranged from < 200 mm to 2200 mm, being highest in western areas (1800–2000 mm) and lower in southern and eastern areas (<200 mm) (Ministry of Agriculture & Rural Development, Ethiopia). The sample sites were located between the latitudes 4°42’-12°46’N and longitudes 34°15–37°52’E.

Elevation transects

Eight villages were sampled along each of two elevational transects, one in Woldiya and the other in the Bale region (Fig 6) in order to determine the environmental and climatic effects associated with changes in elevation within a relatively short geographical distance. The eight Woldiya villages were located within 61km over an elevation range of 1414m to 3396m, and the Bale villages were located within 54km over an elevation of 1224m to 3997m.

Download:

Fig 6. Map of sample collection locations.

Map of Ethiopia to show the sampling locations, shaded areas represent climatic descriptions of the nine regions sampled.

https://doi.org/10.1371/journal.pone.0173811.g006

Sample collection and environmental readings

In each village water was sampled from two sampling points including the predominant communal areas for drinking and bathing including rivers, small streams and sealed wells. As sample collection did not involve humans the AHRI-ALERT Ethical review committee deemed ethical approval unnecessary for this study. The University of Warwick approved the study. Informed verbal consent was obtained from all land owners prior to sample collection. Soil was imported to the University of Warwick using DEFRA licence number PHL 208/6138. This study did not involve endangered or protected species.

In each of the 42 villages, soil was sampled from five sites including two households, two household yards and one communal site such as a school or market where the top 3cm of soil and vegetation was removed prior to sample collection to reduce the effect of UV irradiation. At each of the five sites (<2 kilometres apart), soil was collected using a sterile trowel into a plastic bag from three areas (metres apart) and immediately pooled to make a composite sample. The plastic bag was shaken and three replicates of approximately 5g taken immediately and stored in bijou tubes. The composite samples were stored at -20°C. For water sample collection, two sites for each of the 42 villages were chosen comprising of one drinking source and one bathing source (often these were interchangeable). Homes did not have piped running water. Samples were collected from a variety of sources including groundwater, drinking pumps, springs, rivers and lakes. At each sampling point, 100ml of water was collected from the surface and was filtered using a 50ml sterile plastic syringe and the MicrofilV filtration device with 0.22μm mixed cellulose esters white gridded filters (Millipore, MA, USA). After collection the filters were removed from the plastic holder using sterile forceps and air-dried. The filters were then coiled and stored in 5ml bijou tubes in a cool box with ice packs (4°C) for transporting to the laboratory where the filters were stored at -20°C. All samples were collected during the dry season (Jan–March 2010).

Biogeographic measures were recorded for each site using an eTrex Legend H Outdoor GPS (Garmin Ltd., UK). Readings for soil moisture were taken at each soil sampling site using a soil moisture sensor (SM200 Delta-T Devices Ltd., UK) and moisture meter (HH2 Moisture Meter, Delta-T Devices Ltd., UK) where the output was volumetric water content expressed as a percentage. Substrate temperature and pH readings were taken using a Field Scout SoilStik (Spectrum technologies, Inc., IL, USA). The three readings were averaged per sample type (soil and water) and this average used for statistical analysis. The variables tested by multivariate analysis included temperature, elevation, moisture, pH, latitude and longitude.

DNA extraction

Metagenomic DNA was extracted from 0.5–0.6 g of soil using the FastDNA Spin Kit for Soil (MP Biomedicals, OH, USA) according to the manufacturer’s instructions. The soil community DNA was extracted from the five sampling points and pooled to represent the soil community DNA for the village for pyrosequencing. Metagenomic DNA was extracted from the water polycarbonate filters using the PowerWater DNA Isolation Kit (MoBio Laboratories, Inc., CA, USA) according to manufacturer’s instructions. Samples were normalised using readings from the NanoDrop 1000 spectrophotometer (NanoDrop products, Wilmington, DE, USA) so that equal amounts of nucleic acid were represented from each of the sites for one village.

Quantitative PCR assays

A real-time quantitative PCR (qPCR) assay was performed to determine Mycobacterium genus abundance by targeting the internal transcribed spacer and partial 23S gene (23, 31). The total volume was per reaction 25μl, comprising of 12.5μl of TaqMan Environmental master mix 2.0, 0.4 μM of each primer and probe, 0.4 μM bovine serum albumin (BSA), and 1 μl of the total community DNA. Thermocycle conditions were 2 min at 50°C, followed by 10 min denaturation at 95°C, then by 40 cycles of 15 s at 62°C and 95°C for 1 min. Mycobacterium tuberculosis DNA dilutions were prepared to generate a standard curve which ranged from 5.8 x 10⁵ gene copies μl^-1 to 5.8 x10¹ gene copies μl^-1.

A quantitative PCR assay was employed to determine the abundance of M. bovis in environmental samples. The previously designed qPCR assay targets the RD4 scar region specific for M. bovis (33). All procedures and reagents were as previously published (Sweeney et al, 2007).

A multiplex qPCR assay was employed to quantify the abundance of the M. tuberculosis complex and abundance of M. tuberculosis/M. canettii (32). For each reaction, the total volume was 30μl, comprising of 15μl of TaqMan Environmental master mix 2.0 (Applied Biosystems Inc., CA, USA), 0.6μl of each primer and probe, 1μM BSA, 2μl of Mycobacterium smegmatis DNA (1.22 x10³ cell copies per ml) and 1μl of the total community DNA. Primer and probes are referred to as MTC_IAC Fw, MTC_IAC_Rv, MTC probe, IAC probe, wbbl1_Fw, wbbl1_Rv and wbbl1 probe in the previously published paper (32). Thermocycle conditions were 2 min at 50°C, followed by 10 min at 95°C, then by 40 cycles of 15 s at 95°C and 58C for 1 min.

All qPCR reactions were carried out on the ABI 7500 Fast Real-Time PCR System (Applied Biosystems Inc., CA, USA).

Primers targeting mycobacterium 16S rRNA gene for pyrosequencing

The Mycobacterium genus was targeted for amplicon pyrosequencing using published primers (JSY16S) amplifying the variable regions 2–4 of the 16S rRNA gene (34). In a separate reaction a second set of primers specific to a group of slow-growing mycobacteria (APTK16S) was employed as it was shown that the Mycobacterium genus primers (JSY16S) were biased toward the detection of fast-growing mycobacteria. This targets the variable regions 1–3, where it specifically targets the long helix 18 insertion present in the majority of slow-growing mycobacteria (Leclerc et al, 2003).

Bacterial tag-encoded FLX amplicon pyrosequencing

Sequence libraries were constructed using the HotStarTaq Plus Master Mix Kit (Qiagen Ltd., UK) and a one-step PCR with 30 cycles. Bacterial tag encoded amplicon pyrosequencing was carried at the Research and Testing Laboratory (Lubbock, TX) using a Roche 454 FLX instrument with titanium reagents.

Pyrosequencing quality control

The data was quality controlled using QIIME procedures. Sequences were removed if the length was <400bp for the Mycobacterium genus (JSY16S) and <420bp slow-growing mycobacteria (APTK16S) datasets respectively. Sequences were retained if they matched the proximal primer with one mismatch or less and if the sequence had an average quality score of ≥25. Sequences were denoised using the Denoiser in QIIME and chimeric sequences were identified and removed using ChimeriaSlayer. In order to determine the minimum number of sequences needed per sample for sample inclusion, initial rarefaction curves were generated using OTUs at clustered at 97% similarity using UCLUST (Edgar, 2010). Across all datasets, samples were excluded if they had less than 380 sequences after quality control procedures. Samples with > 380 sequences reached an asymptote and were considered comparable.

Oligotype generation

Sequences were aligned using PyNAST (Caporaso et al, 2010) and entropy components were calculated using the Shannon’s entropy script in the oligotyping suite. Entropy components were chosen in a supervised fashion where H exceeded 0.6 and the position in the alignment was overlapped by all sequences.

BLAST assigned taxonomy

Representative sequences were chosen from each oligotype based upon the most frequent sequence and queried using a local Blast algorithm against a customised database containing all mycobacterium sequences in the SILVA database (1019 sequences). Genus level names were altered to reflect the species complex, group or clade where applicable. An acceptable sequence match was retained if the E value was <0.001 and had >90% identity to the candidate sequence.

Statistical analysis

For all pyroseqencing datasets non-metric multidimensional scaling (NMDS) plots were used to determine significant predictors of mycobacteria community structure using oligotype frequency matrices in R. Environmental variables were fitted to the plot in the form of a least squares regression model to determine the best predictors of mycobacteria community structure using the envfit routines in R. Specific environmental variables showing significant linear trends across the NMDS plots were highlighted by adding isoclines of predicted values using a general additive model approach using the ordisurf routines in R. Plotted points were scaled in size to represent the true values of the GAM fitted environmental gradient. Shannon’s diversity index H’ was calculated on rarefied oligotype matrices using vegan in R. Pearson’s correlation coefficient was calculated to statistically test relationships between individual oligotype prevalence and bioclimatic and spatial variables. Resulting probability values were Bonferroni corrected. Beta diversity was calculated after bining the samples on the basis of the predictor variable and calculating the sum of the proportional abundance of each oligotype minus the mean proportional abundance of that oligotype within each group. Beta diversity was calculated by subtracting the proportional abundance of each oligotype within an environmental bin from the mean proportional abundance of that oligotype across all samples within each environmental bin

Acknowledgments

This work was funded by the Natural Environmental Research Council grant awarded to TK. A material Transfer Agreement was made with the Institute of Biodiversity and Conservation, Ethiopia for export of soils. A DEFRA Licence to import, move and keep prohibited soil (Licence No. PHL 208/6138) was issued.

Author Contributions

Conceptualization: OC EMHW AA RK.
Formal analysis: TKB PJ.
Funding acquisition: EMHW OC.
Investigation: TKB.
Methodology: TKB OC EMHW RB BBO.
Resources: OC EMHW RK AA GE.
Supervision: OC EMHW RK.
Visualization: PJ.
Writing – original draft: HK OC PJ.
Writing – review & editing: HK OC.

References

1. Rastogi N, Legrand E, Sola C. The Mycobacteria: an introduction to nomenclature and pathogenesis. Revue scientifique et technique (International Office of Epizootics). 2001;20: 21–54.
- View Article
- Google Scholar
2. Leclerc MC, Thomas F, Guégan JF. Evidence for phylogenetic inheritance in pathogenicity of Mycobacterium. Antonie van Leeuwenhoek 2003;83:265–74. pmid:12776922
- View Article
- PubMed/NCBI
- Google Scholar
3. Cook JL. Nontuberculous Mycobacteria: Opportunistic environmental pathogens for predisposed hosts. Brit Med Bull. 2010; 96:45–59. pmid:20977990
- View Article
- PubMed/NCBI
- Google Scholar
4. Falkinham JO. Epidemiology of infection by non-tuberculous mycobacteria. Clin Microbiol Rev 1996;9:177–215. pmid:8964035
- View Article
- PubMed/NCBI
- Google Scholar
5. Howard ST, Byrd TF. The rapidly growing mycobacteria: saprophytes and parasites. Microbes Infect. 2000;2:1845–53. pmid:11165929
- View Article
- PubMed/NCBI
- Google Scholar
6. Primm TP, Lucero CA, Falkinham JO. Health impacts of environmental mycobacteria. Clin Microbiol Rev 2004;17:98–106. pmid:14726457
- View Article
- PubMed/NCBI
- Google Scholar
7. Adrados B, Julian E, Codony F, Torrents E, Luquin M, Morato J. Prevalence and concentration of non-tuberculous mycobacteria in cooling towers by means of quantitative PCR: a prospective study. Curr Microbiol 2011;62:313–9. pmid:20640853
- View Article
- PubMed/NCBI
- Google Scholar
8. Brooks RW, Parker BC, Gruft H, Falkinham JO. Epidemiology of infection by nontuberculous mycobacteria. Am rev respir dis. 1984;130:630–633 pmid:6486562
- View Article
- PubMed/NCBI
- Google Scholar
9. Dantec C, Duguet JP, Montiel A, Dumoutier N, Dubrou S, Vincent V. Occurrence of mycobacteria in water treatment lines and in water distribution systems. Appl environ microb. 2002;68:5318–5325.
- View Article
- Google Scholar
10. Kirschner R. Epidemiology of infection by nontuberculous mycobacteria—Mycobacterium avium, Mycobacterium intracellulare and Mycobacterium scrofulaceum in acid brown water swamps of the South-Eastern United States and their association with environmental variables. Am rev respir dis. 1992;145:271–275. pmid:1736730
- View Article
- PubMed/NCBI
- Google Scholar
11. Leys NM,Ryngaert A, Bastianens L, Wattiau P, Top EM, Verstraete W, Springael D. Occurrence and community composition of fast-growing Mycobacterium in soils contaminated with polycyclic aromatic hydrocarbons. FEMS microbiol ecol. 2005;51:375–88. pmid:16329885
- View Article
- PubMed/NCBI
- Google Scholar
12. Niva M, Hernesmaa A, Haahtela K, Salkinoja-salonen M, Haukka K. Actinobacterial communities of boreal forest soil and lake water are rich in mycobacteria. Boreal environ res. 2006;11:45–53.
- View Article
- Google Scholar
13. Thorel MF, Falkinham JO, Moreau RG. Environmental mycobacteria from alpine and subalpine habitats. FEMS microbiol ecol. 2004;49:343–7. pmid:19712284
- View Article
- PubMed/NCBI
- Google Scholar
14. Dailloux M, Laurain C, Webber M, Harterman P. Water and nontuberculous Mycobacteria. Water Res. 1999;33:2219–2228
- View Article
- Google Scholar
15. Falkinham J. O. The Biology of environmental mycobacteria. Environ microbial reports. 2009;1:477–487
- View Article
- Google Scholar
16. Ingen J, Bendien SA, de Lange WCM, Hoefsloot W, Dekhuijzen , Boeree MJ, Soolingen D. Clinical relevance of non-tuberculous mycobacteria isolated in the Nijmegen-Arnhem region, The Netherlands. Thorax. 2009;64:502–504. pmid:19213773
- View Article
- PubMed/NCBI
- Google Scholar
17. Vaereqijck MJM, Huys G, Palomino JC, Swings J, Porteals F. Mycobacteria in drinking water distribution systems: ecology and significance for human health. FEMS microbial rev. 2005; 29:911–934
- View Article
- Google Scholar
18. Black GF, Weir RE, Floyd S, Bliss L, Warndorff DK, Crampin AC, Ngwira B, Dockrell HM. BCG-induced increase in interferon gamma response to Mycobacterial antigens and efficacy of BCG vaccination in Malawi and the UK: two randomised control studies. Lancet. 2002;359:1396–1401
- View Article
- Google Scholar
19. Brandt L, Cunha JF, Olsen AW, Chilima BZ, Hirsch P, Appelberg R, Anderson P. Failure of the Mycobacterium bovis BCG vaccine: some species of environmental mycobacteria block multiplication of BCG and induction of protective immunity to tuberculosis. Infect and immun. 2002;70:672–678.
- View Article
- Google Scholar
20. Fine PEM. Variation in protection by BCG: implications of and for heterologous immunity. Lancet. 1995;346:1339–1345 pmid:7475776
- View Article
- PubMed/NCBI
- Google Scholar
21. Flaherty DK, Vesosky B, Beamer GL, Stromberg P, Turner J. Exposure to Mycobacterium avium can modulate established immunity against Mycobacterium tuberculosis infection generated by Mycobacterium bovis BCG vaccination. Journ leukocyte biol. 2006; 80: 1262–1271
- View Article
- Google Scholar
22. Weir RE, Fine PE, Floyd S, Stenson S, Stanley C, Branson K, Dockerell HM. The influence of previous exposure to environmental mycobacteria on the interferon gamma response to Bacilli Calmette-Guerin vaccination in southern England and northern Malawi. Clin exp immunol. 2006;146:390–399 pmid:17100757
- View Article
- PubMed/NCBI
- Google Scholar
23. Kopecky J, Kyselkova M, Omelka M, Cermak L, Novotna J, Grundmann G, Moenne-Loccoz Y, Sagova-Mareckova M. Environmental mycobacteria closely related to the pathogenic species evidenced in an acidic forest wetland. Soil Biol Biochem. 2011;43:697–700
- View Article
- Google Scholar
24. Norby B, Fosgate GT, Manning EJB, Collins MT, Roussel AJ. Environmental mycobacteria in soil and water on beef ranches: association between presence of cultivable mycobacteria and soil and water physicochemical characteristics. Vet microbiol. 2007;124:153–9. pmid:17512144
- View Article
- PubMed/NCBI
- Google Scholar
25. Donoghue HD, Overend E, Stanford JL. A longitudinal study of environmental mycobacteria on a farm in south-west England. Journ appl microbial. 1997;82:57–67
- View Article
- Google Scholar
26. Roth A, Fischer M, Hamid ME, Michalke S, Ludwig W, Mauch H. Differentiation of phylogenetically related slow rowing mycobacteria based on 16S rRNA gene internal transcribed spacer sequences. Journ clin micro. 1998;36:139–147
- View Article
- Google Scholar
27. Eren AM, Maignien L, Sul WJ, Murphy LG, Grim SL, Morrison HG, Sogin ML. Oligotyping: differentiating between closely related microbial taxa using 16S rRNA gene data. Meth ecol evo. 2013.
- View Article
- Google Scholar
28. Jacobs J, Rhodes M, Sturgis B, Wood B. Influence of environmental gradients on the abundance and distribution of Mycobacterium spp. in a coastal lagoon estuary. Appl envirol microbial. 2009;75:7378–84.
- View Article
- Google Scholar
29. Global tuberculosis Report 2013, World Health Organisation (WHO), http://www.who.int/tb/country/en/
30. Berg S, Firdessa R, Habatmu M, Gadisa E, Mangistu A, Yamuah L, Ameni G. The burden of mycobacterial disease in Ethiopian cattle: implications for public health. PloS One. 2009;4:1–8
- View Article
- Google Scholar
31. Chilima BZ, Clark IM, Floyd S, Fine PEM, Hirsch PR. Distribution of environmental mycobacteria in Karonga District, northern Malawi. Appl enviro microbiol 2006;72:2343–2350
- View Article
- Google Scholar
32. Kamala T, Paramasivan CN, Herbert D, Venkatesan P, Prabhakar R. Isolation and Identification of Environmental Mycobacteria in the Mycobacterium bovis BCG Trial Area of South India. Appl enviro microbiol 1994;60:2180–3.
- View Article
- Google Scholar
33. Torvinen E, miklin T, Torkko P, Suomalainen S, Reiman M, Katila ML, Paulin L, Nevalainen A. Mycobacteria and fungi in moisture-damaged building materials. Appl envirl microbial. 2006;72:6822–4.
- View Article
- Google Scholar
34. Shitaye JE, Tsegaye W, Pavlik I. Bovine tuberculosis infection in animal and human populations in Ethipia: A Review. Veterinarni Medicina. 2009;8:317–332.
- View Article
- Google Scholar
35. Shargie EB, Lindtjørn B. Determinants of treatment adherence among smear-positive pulmonary tuberculosis patients in Southern Ethiopia. PLoS Med 2007;4: e37. pmid:17298164
- View Article
- PubMed/NCBI
- Google Scholar
36. Cowie CE, Hutchings MR, Barasona JA, Gortazar C, Vicente J, White PCL. Interactions between four species in a complex wildlife: livestock disease community: implications for Mycobacterium bovis maintenance and transmission. 2016; 62: 51–64
- View Article
- Google Scholar
37. Barasona JA, Vicente J, Diez-Delgado I, Aznar J, Gortazar C, Torres MJ. Environmental presence of Mycobacterium tuberculosis complex in aggregation points at the Wildlife/livestock interface. Transbound emerg disease. 2016;
- View Article
- Google Scholar
38. Ghodbane R, Mba Medie F, Lepidi H, Nappez C, Drancourt M. Long-term survival of tuberculosis complex mycobacteria in soil. Microbuil 2014;160:496–501.
- View Article
- Google Scholar
39. Reed C, Fordham von Reyn C, Chamblee S, Ellerbrock TV, Johnson JW, Marsh BJ, Johnson LS, Trenschel RJ, Hornsburugh CR. Environmental Risk Factors for Infection with Mycobacterium avium Complex. Am. J. Epidemiol. 2006; 164:32–40 pmid:16675537
- View Article
- PubMed/NCBI
- Google Scholar
40. Horsburg CR. Epidemiology of Mycobacterium avium complex. In Mycobacterium avium complex infection: Progress in Research and Treatment. 1^st edition p1–22. Edited by Korvick J. and Benson C.. New York: Mark Dekker 1996
41. Falkinham JO. Nontuberculous mycobacteria from household plumbing of patients with nontuberculous mycobacteria disease. Emerg infect diseases. 2011;17:419–424
- View Article
- Google Scholar
42. Pickup RW, Rhodes G, Arnott S, Sidi-Boumedine K, Bull TJ, Weightman A, Hurley M, Hermon-Taylor J. Mycobacterium avium subsp. Paratuberculosis in the Catchment Area and Water of the River Taff in South Wales, United Kingdom, And its potential relationship to clustering of chrone’s disease cases in the city of Cardiff. Appl environmen microbial. 2005;71:2130–2139
- View Article
- Google Scholar
43. Tortoli E. Phylogeny of the genus Mycobacterium: Many doubts, few certainties. Infect Gen Evol. 2012; 12:827–831
- View Article
- Google Scholar

[ref1] 1. Rastogi N, Legrand E, Sola C. The Mycobacteria: an introduction to nomenclature and pathogenesis. Revue scientifique et technique (International Office of Epizootics). 2001;20: 21–54.
View Article
Google Scholar

[2] View Article

[3] Google Scholar

[ref2] 2. Leclerc MC, Thomas F, Guégan JF. Evidence for phylogenetic inheritance in pathogenicity of Mycobacterium. Antonie van Leeuwenhoek 2003;83:265–74. pmid:12776922
View Article
PubMed/NCBI
Google Scholar

[5] View Article

[6] PubMed/NCBI

[7] Google Scholar

[ref3] 3. Cook JL. Nontuberculous Mycobacteria: Opportunistic environmental pathogens for predisposed hosts. Brit Med Bull. 2010; 96:45–59. pmid:20977990
View Article
PubMed/NCBI
Google Scholar

[9] View Article

[10] PubMed/NCBI

[11] Google Scholar

[ref4] 4. Falkinham JO. Epidemiology of infection by non-tuberculous mycobacteria. Clin Microbiol Rev 1996;9:177–215. pmid:8964035
View Article
PubMed/NCBI
Google Scholar

[13] View Article

[14] PubMed/NCBI

[15] Google Scholar

[ref5] 5. Howard ST, Byrd TF. The rapidly growing mycobacteria: saprophytes and parasites. Microbes Infect. 2000;2:1845–53. pmid:11165929
View Article
PubMed/NCBI
Google Scholar

[17] View Article

[18] PubMed/NCBI

[19] Google Scholar

[ref6] 6. Primm TP, Lucero CA, Falkinham JO. Health impacts of environmental mycobacteria. Clin Microbiol Rev 2004;17:98–106. pmid:14726457
View Article
PubMed/NCBI
Google Scholar

[21] View Article

[22] PubMed/NCBI

[23] Google Scholar

[ref7] 7. Adrados B, Julian E, Codony F, Torrents E, Luquin M, Morato J. Prevalence and concentration of non-tuberculous mycobacteria in cooling towers by means of quantitative PCR: a prospective study. Curr Microbiol 2011;62:313–9. pmid:20640853
View Article
PubMed/NCBI
Google Scholar

[25] View Article

[26] PubMed/NCBI

[27] Google Scholar

[ref8] 8. Brooks RW, Parker BC, Gruft H, Falkinham JO. Epidemiology of infection by nontuberculous mycobacteria. Am rev respir dis. 1984;130:630–633 pmid:6486562
View Article
PubMed/NCBI
Google Scholar

[29] View Article

[30] PubMed/NCBI

[31] Google Scholar

[ref9] 9. Dantec C, Duguet JP, Montiel A, Dumoutier N, Dubrou S, Vincent V. Occurrence of mycobacteria in water treatment lines and in water distribution systems. Appl environ microb. 2002;68:5318–5325.
View Article
Google Scholar

[33] View Article

[34] Google Scholar

[ref10] 10. Kirschner R. Epidemiology of infection by nontuberculous mycobacteria—Mycobacterium avium, Mycobacterium intracellulare and Mycobacterium scrofulaceum in acid brown water swamps of the South-Eastern United States and their association with environmental variables. Am rev respir dis. 1992;145:271–275. pmid:1736730
View Article
PubMed/NCBI
Google Scholar

[36] View Article

[37] PubMed/NCBI

[38] Google Scholar

[ref11] 11. Leys NM,Ryngaert A, Bastianens L, Wattiau P, Top EM, Verstraete W, Springael D. Occurrence and community composition of fast-growing Mycobacterium in soils contaminated with polycyclic aromatic hydrocarbons. FEMS microbiol ecol. 2005;51:375–88. pmid:16329885
View Article
PubMed/NCBI
Google Scholar

[40] View Article

[41] PubMed/NCBI

[42] Google Scholar

[ref12] 12. Niva M, Hernesmaa A, Haahtela K, Salkinoja-salonen M, Haukka K. Actinobacterial communities of boreal forest soil and lake water are rich in mycobacteria. Boreal environ res. 2006;11:45–53.
View Article
Google Scholar

[44] View Article

[45] Google Scholar

[ref13] 13. Thorel MF, Falkinham JO, Moreau RG. Environmental mycobacteria from alpine and subalpine habitats. FEMS microbiol ecol. 2004;49:343–7. pmid:19712284
View Article
PubMed/NCBI
Google Scholar

[47] View Article

[48] PubMed/NCBI

[49] Google Scholar

[ref14] 14. Dailloux M, Laurain C, Webber M, Harterman P. Water and nontuberculous Mycobacteria. Water Res. 1999;33:2219–2228
View Article
Google Scholar

[51] View Article

[52] Google Scholar

[ref15] 15. Falkinham J. O. The Biology of environmental mycobacteria. Environ microbial reports. 2009;1:477–487
View Article
Google Scholar

[54] View Article

[55] Google Scholar

[ref16] 16. Ingen J, Bendien SA, de Lange WCM, Hoefsloot W, Dekhuijzen , Boeree MJ, Soolingen D. Clinical relevance of non-tuberculous mycobacteria isolated in the Nijmegen-Arnhem region, The Netherlands. Thorax. 2009;64:502–504. pmid:19213773
View Article
PubMed/NCBI
Google Scholar

[57] View Article

[58] PubMed/NCBI

[59] Google Scholar

[ref17] 17. Vaereqijck MJM, Huys G, Palomino JC, Swings J, Porteals F. Mycobacteria in drinking water distribution systems: ecology and significance for human health. FEMS microbial rev. 2005; 29:911–934
View Article
Google Scholar

[61] View Article

[62] Google Scholar

[ref18] 18. Black GF, Weir RE, Floyd S, Bliss L, Warndorff DK, Crampin AC, Ngwira B, Dockrell HM. BCG-induced increase in interferon gamma response to Mycobacterial antigens and efficacy of BCG vaccination in Malawi and the UK: two randomised control studies. Lancet. 2002;359:1396–1401
View Article
Google Scholar

[64] View Article

[65] Google Scholar

[ref19] 19. Brandt L, Cunha JF, Olsen AW, Chilima BZ, Hirsch P, Appelberg R, Anderson P. Failure of the Mycobacterium bovis BCG vaccine: some species of environmental mycobacteria block multiplication of BCG and induction of protective immunity to tuberculosis. Infect and immun. 2002;70:672–678.
View Article
Google Scholar

[67] View Article

[68] Google Scholar

[ref20] 20. Fine PEM. Variation in protection by BCG: implications of and for heterologous immunity. Lancet. 1995;346:1339–1345 pmid:7475776
View Article
PubMed/NCBI
Google Scholar

[70] View Article

[71] PubMed/NCBI

[72] Google Scholar

[ref21] 21. Flaherty DK, Vesosky B, Beamer GL, Stromberg P, Turner J. Exposure to Mycobacterium avium can modulate established immunity against Mycobacterium tuberculosis infection generated by Mycobacterium bovis BCG vaccination. Journ leukocyte biol. 2006; 80: 1262–1271
View Article
Google Scholar

[74] View Article

[75] Google Scholar

[ref22] 22. Weir RE, Fine PE, Floyd S, Stenson S, Stanley C, Branson K, Dockerell HM. The influence of previous exposure to environmental mycobacteria on the interferon gamma response to Bacilli Calmette-Guerin vaccination in southern England and northern Malawi. Clin exp immunol. 2006;146:390–399 pmid:17100757
View Article
PubMed/NCBI
Google Scholar

[77] View Article

[78] PubMed/NCBI

[79] Google Scholar

[ref23] 23. Kopecky J, Kyselkova M, Omelka M, Cermak L, Novotna J, Grundmann G, Moenne-Loccoz Y, Sagova-Mareckova M. Environmental mycobacteria closely related to the pathogenic species evidenced in an acidic forest wetland. Soil Biol Biochem. 2011;43:697–700
View Article
Google Scholar

[81] View Article

[82] Google Scholar

[ref24] 24. Norby B, Fosgate GT, Manning EJB, Collins MT, Roussel AJ. Environmental mycobacteria in soil and water on beef ranches: association between presence of cultivable mycobacteria and soil and water physicochemical characteristics. Vet microbiol. 2007;124:153–9. pmid:17512144
View Article
PubMed/NCBI
Google Scholar

[84] View Article

[85] PubMed/NCBI

[86] Google Scholar

[ref25] 25. Donoghue HD, Overend E, Stanford JL. A longitudinal study of environmental mycobacteria on a farm in south-west England. Journ appl microbial. 1997;82:57–67
View Article
Google Scholar

[88] View Article

[89] Google Scholar

[ref26] 26. Roth A, Fischer M, Hamid ME, Michalke S, Ludwig W, Mauch H. Differentiation of phylogenetically related slow rowing mycobacteria based on 16S rRNA gene internal transcribed spacer sequences. Journ clin micro. 1998;36:139–147
View Article
Google Scholar

[91] View Article

[92] Google Scholar

[ref27] 27. Eren AM, Maignien L, Sul WJ, Murphy LG, Grim SL, Morrison HG, Sogin ML. Oligotyping: differentiating between closely related microbial taxa using 16S rRNA gene data. Meth ecol evo. 2013.
View Article
Google Scholar

[94] View Article

[95] Google Scholar

[ref28] 28. Jacobs J, Rhodes M, Sturgis B, Wood B. Influence of environmental gradients on the abundance and distribution of Mycobacterium spp. in a coastal lagoon estuary. Appl envirol microbial. 2009;75:7378–84.
View Article
Google Scholar

[97] View Article

[98] Google Scholar

[ref29] 29. Global tuberculosis Report 2013, World Health Organisation (WHO), http://www.who.int/tb/country/en/

[ref30] 30. Berg S, Firdessa R, Habatmu M, Gadisa E, Mangistu A, Yamuah L, Ameni G. The burden of mycobacterial disease in Ethiopian cattle: implications for public health. PloS One. 2009;4:1–8
View Article
Google Scholar

[101] View Article

[102] Google Scholar

[ref31] 31. Chilima BZ, Clark IM, Floyd S, Fine PEM, Hirsch PR. Distribution of environmental mycobacteria in Karonga District, northern Malawi. Appl enviro microbiol 2006;72:2343–2350
View Article
Google Scholar

[104] View Article

[105] Google Scholar

[ref32] 32. Kamala T, Paramasivan CN, Herbert D, Venkatesan P, Prabhakar R. Isolation and Identification of Environmental Mycobacteria in the Mycobacterium bovis BCG Trial Area of South India. Appl enviro microbiol 1994;60:2180–3.
View Article
Google Scholar

[107] View Article

[108] Google Scholar

[ref33] 33. Torvinen E, miklin T, Torkko P, Suomalainen S, Reiman M, Katila ML, Paulin L, Nevalainen A. Mycobacteria and fungi in moisture-damaged building materials. Appl envirl microbial. 2006;72:6822–4.
View Article
Google Scholar

[110] View Article

[111] Google Scholar

[ref34] 34. Shitaye JE, Tsegaye W, Pavlik I. Bovine tuberculosis infection in animal and human populations in Ethipia: A Review. Veterinarni Medicina. 2009;8:317–332.
View Article
Google Scholar

[113] View Article

[114] Google Scholar

[ref35] 35. Shargie EB, Lindtjørn B. Determinants of treatment adherence among smear-positive pulmonary tuberculosis patients in Southern Ethiopia. PLoS Med 2007;4: e37. pmid:17298164
View Article
PubMed/NCBI
Google Scholar

[116] View Article

[117] PubMed/NCBI

[118] Google Scholar

[ref36] 36. Cowie CE, Hutchings MR, Barasona JA, Gortazar C, Vicente J, White PCL. Interactions between four species in a complex wildlife: livestock disease community: implications for Mycobacterium bovis maintenance and transmission. 2016; 62: 51–64
View Article
Google Scholar

[120] View Article

[121] Google Scholar

[ref37] 37. Barasona JA, Vicente J, Diez-Delgado I, Aznar J, Gortazar C, Torres MJ. Environmental presence of Mycobacterium tuberculosis complex in aggregation points at the Wildlife/livestock interface. Transbound emerg disease. 2016;
View Article
Google Scholar

[123] View Article

[124] Google Scholar

[ref38] 38. Ghodbane R, Mba Medie F, Lepidi H, Nappez C, Drancourt M. Long-term survival of tuberculosis complex mycobacteria in soil. Microbuil 2014;160:496–501.
View Article
Google Scholar

[126] View Article

[127] Google Scholar

[ref39] 39. Reed C, Fordham von Reyn C, Chamblee S, Ellerbrock TV, Johnson JW, Marsh BJ, Johnson LS, Trenschel RJ, Hornsburugh CR. Environmental Risk Factors for Infection with Mycobacterium avium Complex. Am. J. Epidemiol. 2006; 164:32–40 pmid:16675537
View Article
PubMed/NCBI
Google Scholar

[129] View Article

[130] PubMed/NCBI

[131] Google Scholar

[ref40] 40. Horsburg CR. Epidemiology of Mycobacterium avium complex. In Mycobacterium avium complex infection: Progress in Research and Treatment. 1^st edition p1–22. Edited by Korvick J. and Benson C.. New York: Mark Dekker 1996

[ref41] 41. Falkinham JO. Nontuberculous mycobacteria from household plumbing of patients with nontuberculous mycobacteria disease. Emerg infect diseases. 2011;17:419–424
View Article
Google Scholar

[134] View Article

[135] Google Scholar

[ref42] 42. Pickup RW, Rhodes G, Arnott S, Sidi-Boumedine K, Bull TJ, Weightman A, Hurley M, Hermon-Taylor J. Mycobacterium avium subsp. Paratuberculosis in the Catchment Area and Water of the River Taff in South Wales, United Kingdom, And its potential relationship to clustering of chrone’s disease cases in the city of Cardiff. Appl environmen microbial. 2005;71:2130–2139
View Article
Google Scholar

[137] View Article

[138] Google Scholar

[ref43] 43. Tortoli E. Phylogeny of the genus Mycobacterium: Many doubts, few certainties. Infect Gen Evol. 2012; 12:827–831
View Article
Google Scholar

[140] View Article

[141] Google Scholar

Figures

Abstract

Introduction

Results

Processed sequences results

Core mycobacterial communities and pathogenic groups in soil and water

Biogeographical variation in mycobacterium community composition and diversity

Environmental predictors of mycobacteria communities in water

Environmental predictors of slow-growing mycobacteria communities in water

Environmental predictors of mycobacteria communities in soil

Environmental predictors of slow growing mycobacteria communities in soil

Discussion

Materials and methods

Environmental sampling sites

Elevation transects

Sample collection and environmental readings

DNA extraction

Quantitative PCR assays

Primers targeting mycobacterium 16S rRNA gene for pyrosequencing

Bacterial tag-encoded FLX amplicon pyrosequencing

Pyrosequencing quality control

Oligotype generation

BLAST assigned taxonomy

Statistical analysis

Acknowledgments

Author Contributions

References