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Published in final edited form as: Parasitology. 2019 Sep 26;146(14):1707–1713. doi: 10.1017/S0031182019001252

Leptospira in livestock in Madagascar: uncultured strains, mixed infections and small mammal-livestock transmission highlight challenges in controlling and diagnosing leptospirosis in the developing world

Soanandrasana Rahelinirina 1,†, Mark H Moseley 2,†, Kathryn J Allan 3, Emmanuel Ramanohizakandrainy 4, Sati Ravaoarinoro 4, Minoarisoa Rajerison 1, Vincent Rakotoharinome 5, Sandra Telfer 2
PMCID: PMC6935375  EMSID: EMS84709  PMID: 31554531

Abstract

In developing countries, estimates of the prevalence and diversity of Leptospira infections in livestock, an important but neglected zoonotic pathogen and cause of livestock productivity loss, are lacking. In Madagascar, abattoir sampling of cattle and pigs demonstrated a prevalence of infection of 20% in cattle and 5% in pigs by real-time PCR. In cattle, amplification and sequencing of the Leptospira-specific lfb1 gene revealed novel genotypes, mixed infections of two or more Leptospira species and evidence for potential transmission between small mammals and cattle. Sequencing of the secY gene demonstrated genetic similarities between Leptospira detected in Madagascar and, as yet, uncultured Leptospira strains identified in Tanzania, Reunion and Brazil. Detection of Leptospira DNA in the same animal was more likely in urine samples or pooled samples from four kidney lobes relative to samples collected from a single kidney lobe, suggesting an effect of sampling method on detection. In pigs, no molecular typing of positive samples was possible. Further research into the epidemiology of livestock leptospirosis in developing countries is needed to inform efforts to reduce human infections and to improve livestock productivity.

Keywords: Zoonosis, pathogen, veterinary, Madagascar, Tanzania, spirochaete, spillover, Africa

Introduction

Zoonoses account for 61% of infectious diseases of humans and 75% of emerging infectious diseases (Taylor et al., 2001; Woolhouse and Gowtage-Sequeria, 2005). Endemic zoonoses, characterised by a widespread distribution and frequent transmission between animals and humans, include some of the most important diseases of poverty (Maudlin et al., 2009; ILRI, 2012). However, in the poorest communities in the developing world, a lack of surveillance and control (Halliday et al., 2012) and misdiagnosis (Maudlin et al., 2009) result in these diseases being amongst the most neglected in the world (ILRI, 2012). It has been suggested that, in terms of human health impacts, livestock productivity loss and amenity to agricultural intervention, leptospirosis, a neglected endemic zoonosis causing an estimated 1 million cases of human disease annually (Costa et al., 2015), is the second most important zoonosis globally (ILRI, 2012). Although rodents are frequently implicated as the key reservoir hosts of Leptospira (Picardeau, 2017), recent evidence suggests that, in rural Africa, livestock may also be significant reservoir hosts (Allan et al., 2018).

Leptospira are phylogenetically delineated into 22 species, 10 of which are pathogenic, and further divided into more than 300 serovars which may demonstrate specific, but not absolute, host preferences (Picardeau, 2017). In Madagascar, four Leptospira species have been identified in terrestrial small mammals (Rahelinirina et al., 2010; Dietrich et al., 2014; Moseley et al., 2018) and a molecular link demonstrated between a recent acute, severe case of human leptospirosis (Pagès et al., 2015) and small mammals (Moseley et al., 2018). However, a recent household cross-sectional serosurvey in Madagascar identified contact with cattle as the only significant risk factor (OR=3, 95% CI [1.03–10.03]) for human exposure despite serological typing based on Microscopic Agglutination Testing (MAT) identifying Icterohaemorrhagiae, a serogroup traditionally associated with rodents, as the predominant serogroup (Ratsitorahina et al., 2015).

Therefore, understanding the role of livestock in the epidemiology of leptospirosis is critical to understanding public health risks and informing control strategies. Vaccination of livestock is used to both improve productivity and to reduce zoonotic risk from livestock (Marshall and Chereshsky, 1996). However, recent studies have demonstrated that commercial leptospirosis vaccines may lack efficacy when challenged by autochthonous livestock strains in Brazil (Sonada et al., 2018) and will not reduce human infections from serovars associated with wildlife (Thornley et al., 2002). Therefore, to inform control measures, we used abattoir sampling and molecular methods to determine the prevalence and diversity of Leptospira in livestock in Madagascar and their phylogenetic relationship to Leptospira identified in livestock from similar studies in other developing countries.

Materials and methods

Abattoir sampling

In 2015, we estimated Leptospira prevalence by sampling a total of 205 livestock: 25 cattle and 25 pigs sampled at each of 3 abattoirs around the capital, Antananarivo, and 30 cattle and 25 pigs sampled at the abattoir in the town of Moramanga. Livestock presented at abattoirs were sourced from a wide geographic area (Supplementary - Figure S1). Whole kidneys and urine aspirates were collected and cooled for transportation to a laboratory.

Sample preparation

As Leptospira may not be homogeneously distributed within kidney tissue and urinary shedding is intermittent, to advise future studies/surveillance we compared results from three sample preparations: a single kidney excision, a pool of four kidney excisions from different lobes (cattle) or anatomical locations (pigs) and urine. Kidney samples were stored in 95% ethanol and urine samples at -80°C.

i. Kidney samples

Extraction of the kidney samples was performed on 40mg of tissue using the DNeasy Blood and Tissue Kit (Qiagen) as per the manufacturer’s instructions with the volumes of buffers ATL and AL increased to account for the increased amount of tissue extracted and the volume of elution buffer reduced to 100μl to concentrate the DNA.

For preparations from a single lobe/location, 40mg of tissue from the cortico-medullary junction was extracted. For pooled samples of four kidney lobes, 40mg of tissue from each lobe was placed in a sterile 1.5ml Eppendorf tube containing 400μl of the ATL lysis buffer and ground with a glass pestle to obtain a homogenate. One hundred microliters of the homogenate was then placed in a new sterile 1.5ml Eppendorf and extracted as outlined above. To monitor inhibition, 1.5μl of DNA extraction control 560 (Bioline) was added to each extraction.

ii. Urine samples

After slaughter urine samples were obtained by aspiration of 2ml of urine from the bladder using sterile needles and 2ml syringes. If samples arrived at the laboratory within 24 hours they were transferred to a clean cryotube and stored at -80°C. For the Moramanga abattoir, this was not possible. Consequently, samples were centrifuged at 7000 rpm for 20 minutes before discarding the supernatant and adding 200μl of TE buffer (Sigma-Aldrich). The samples were then kept at +4°C, before transferring to the laboratory in a cool box with ice packs.

Extraction of the urine samples was performed using the same protocol as for the kidney samples with the exception of the following pre-extraction step. Samples frozen at -80°C were incubated at 67°C for one hour before 1ml was transferred to a 2ml Eppendorf. Samples were then centrifuged at 7000 rpm for 10 minutes. After discarding the supernatant, the pellet was suspended in 200μl of TE buffer. The re-suspended pellet was centrifuged again at 7000 rpm for 10 minutes and the supernatant discarded. The pellet was then re-suspended again with 100μl of TE buffer, 50μl of lysozyme (10μg/ml), 50μl of mutanolysin (4KU/ml), 4μl of lysostaphin and incubated at 37°C for one hour. After addition of 20μl of proteinase K and 180μl of AL buffer (Qiagen) and vortexing, samples were incubated at 56°C for 10 minutes. The final step involved the addition of 200μl of 95% ethanol and vortexing before following the extraction protocol used for kidney samples.

Leptospira detection and DNA sequencing

Prevalence estimates were obtained using a 16s (rrs) qPCR (Smythe et al., 2002) as described previously (Moseley et al., 2018). Each sample preparation (single lobe/location, four lobes/location and urine sample) was tested separately. An individual animal was identified as infected if any of the sample preparations tested positive. Chi-square tests were used to test for differences in prevalence estimates between cattle and pigs and between prevalence estimates from different sample preparations within the same host. Mann-Whitney-Wilcoxon tests were used to determine whether parasite load, as measured by 16s qPCR amplification threshold (Ct), affected typing success. Statistical analyses were performed using R version 3.4.1 software (R Core Team).

Amplification of 200-300bp of the lfb1 gene (Merien et al., 2005; Moseley et al., 2018) was then performed on positive kidney and urine sample from each animal and on positive livestock samples previously identified in Tanzania (Allan et al., 2018). Initially, samples were tested using lfb1 primers (Merien et al., 2005) designed to detect all pathogenic Leptospira. Previous molecular studies in livestock in Africa have identified L. borgpetersenii, L. kirschneri and a L. kirschneri-like species (Allan et al., 2018) and previous studies suggest that existing lfb1 primers fail to amplify L. borgpetersenii in some cases (Moseley et al., 2018). Therefore, redesigned lfb1 forward primers targeting L. kirschneri and L. borgpetersenii (Moseley et al., 2018) were subsequently used on samples that tested negative using the standard lfb1 qPCR. In addition, to test for mixed infections, samples infected with L. borgpetersenii were tested using L. kirschneri targeting primers and vice versa. All lfb1 assays were performed using reaction conditions as previously described (Moseley et al., 2018) and all amplicons were sequenced.

To facilitate comparisons with other studies, amplification and sequencing of the secY gene, a widely used target for phylogenetic analysis of Leptospira (Victoria et al., 2008), was undertaken. Initial amplification of a ~450bp fragment was performed using reaction conditions previously described (Allan et al., 2018). Where initial amplification failed to amplify sufficient material for sequencing, a second round of amplification of a ~350bp fragment was performed in a 25μl reaction volume was performed using a nested forward primer [5’-AATCCATTYTCYCARATYTGGTA-3’] and the first round reverse primer at concentrations of 0.5μM, 12.5μl of MyTaq Red mix 2x (Bioline Reagents Ltd), 9.5μl of molecular grade water and 1μl of first round product. The thermal profile comprised initial denaturation at 95°C for 3min, followed by 20 cycles of denaturation at 95°C for 30s, annealing at 50°C for 30s and extension at 72°C for 1min, with a final extension at 72°C for 7min.

DNA sequence analysis

Phylogenetic analysis of lfb1 sequences was supplemented with lfb1 sequences obtained from small mammals in Madagascar (Moseley et al., 2018), a goat opportunistically sampled during village-based rodent surveys and four cattle sampled during a pilot study in Antananarivo. In addition, to provide additional lfb1 sequences from the region and to test for mixed infections, the same lfb1 assays were used to obtain sequences from 28 livestock sampled in a similar abattoir study in northern Tanzania (Allan et al., 2018) and typed using secY sequencing. To identify the serovars to which these strains are most closely related, lfb1 sequences were queried against the NCBI refseq_genomes and nr/nt database using the blastn algorithm and identical or closely related (>99% identity) records with associated serovar information reported.

To place the strains identified in this study in a global context, secY sequences were analysed alongside sequences from similar livestock studies from Tanzania (Allan et al., 2018), Reunion (Guernier et al., 2016) and Brazil (Hamond et al., 2015; Guedes et al., 2019) (Supplementary - Table S1). Reference secY sequences (Victoria et al., 2008) with 100% identity to any of the secY genotypes were included in the phylogenetic analysis. Sequence alignment and phylogenetic analysis was performed using MEGA7 (Kumar et al., 2016).

Results

Prevalence estimates and effect of sample preparation

Cattle had a significantly higher overall prevalence than pigs (19% vs 5%, χ2=8.17, p<0.01) (Table 1). Cattle had 13 infections detected in kidney samples and 11 in urine samples, with only 4 individuals testing positive for both sample types. Pooled kidney excisions detected more infections than single excisions (12% vs 6%) (Table 1), although the difference was not significant (χ2=1.11, p=0.29), and only one individual tested positive by the single excision but negative by the pooled excision. Of the positive pigs, only one infection was detected in kidney (single excision), compared to four infections detected in urine.

Table 1.

Prevalence of Leptospira infection using different sample preparations from cattle and pigs. Cattle kidneys were sampled at four lobes and pig kidneys were sampled at four locations (pig kidneys are not lobed). Infection prevalence and 95% confidence intervals (logit method) were determined using the binom package (https://cran.r-project.org/ package=binom). Samples with evidence of inhibition were excluded from prevalence estimate calculations for each sample preparation but each individual animal had at least one sample preparation without evidence of inhibition so every animal was represented in overall prevalence estimates

Host Sample preparations Overall prevalence (P/n, 95% CI)
Sample type Inhibitors Prevalence (P/n, 95% CI)
Cattle (n=105) Kidney (1 lobe) (n=105) 11 6% (6/94, 3-13%) 19% (20/105, 13-28%)
Kidney (4 lobes) (n=105) 3 12% (12/102, 7-20%)
Urine (n=99) 3 11% (11/96, 6-20%)
Pigs (n=100) Kidney (1 location) (n=100) 1 1% (1/99, 0-6%) 5% (5/100, 2-11%)
Kidney (4 locations) (n=100) 7 0% (0/93, 0-4%)
Urine (n=95) 0 4% (4/95, 2-11%)
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Typing success

In Madagascar, interpretable lfb1 and secY sequences were obtained from 50% (10/20) and 40% (8/20) of 16s qPCR positive cattle samples respectively. All secY sequences were obtained from samples in which an lfb1 sequence had previously been amplified. In kidney samples, parasite load, as measured by 16s qPCR Ct, had an effect on sequencing success with samples from which sequencing data was obtained having a higher parasite load (median 16s Ct=33.31, range 32.38-33.99) than samples in which sequencing was unsuccessful (median 16s Ct=37.85, range 33.95-39.51) (W=2, p=0.002). No such effect was noted in urine samples (median 16s Ct=35, range 34-37 vs median 16s Ct=37.4, range 33-39) (W=13.5, p=0.18). None of the 16s qPCR positive pig (n=5, 16s Ct range 37-39) samples yielded interpretable lfb1 or secY sequences. Samples from Tanzania identified to be infected with an unknown L. kirschneri-like Leptospira species based on secY sequences (Allan et al., 2018), failed to satisfactorily amplify using any of the lfb1 primer combinations used.

Genetic diversity and mixed infections

Based on lfb1 sequences, we identified L. borgpetersenii and L. kirschneri in cattle and L. interrogans in a goat sample in Madagascar (Figure 1). Including lfb1 sequences from both Madagascan and Tanzanian samples, five L. borgpetersenii lfb1 clades (A-E) were identified in cattle, two of which (clade A and clade D) were shared between both countries. L. borgpetersenii clade A sequences were identical to serovar Hardjo-bovis strain L550, a livestock associated strain (Bulach et al., 2006). However, L. borgpetersenii clade D, had no lfb1 homologue. Of the remaining three L. borgpetersenii clades, two (clade C and clade E) were present in Madagascar, clade E sequences were identical to lfb1 sequences previously obtained from small mammals (Moseley et al., 2018) and the clade C sequence was identical to a strain previously identified as serovar Tarassovi. The remaining L. borgpetersenii clade, detected only in Tanzania, was identical to serovar Hardjo-bovis strain JB197 (Bulach et al., 2006).

Figure 1.

Figure 1

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Maximum likelihood phylogenetic tree of 171bp lfb1 sequences (accession numbers MK244296-MK244341) using the Kimura 2-Parameter model with a uniform distribution. Samples from Madagascar and Tanzania are highlighted in bold and the number of animals from which sequences were obtained are indicated. Reference sequences are labelled by Leptospira species, serovar and accession number. Nodes are labelled with bootstrap support.

L. kirschneri lfb1 sequences, obtained from the kidney and urine samples of a single animal from the Moramanga region of Madagascar, demonstrated 99% identity to sequences previously obtained from endemic small mammals (Hemicentetes semispinosus) (Moseley et al., 2018) in the same region and the L. interrogans sequence obtained from a goat was identical to a genotype previously identified predominantly in black rats (Rattus rattus) (Moseley et al., 2018) and to that obtained from an acute human case of leptospirosis (Pagès et al., 2015).

Using the lfb1 primers targeting different Leptospira species, mixed infections were detected in two cattle in Madagascar and one in Tanzania. In Madagascar, L. kirschneri and L. borgpetersenii clade E were detected in a urine sample from one animal and L.borgpetersenii clade A was detected in the urine sample and L. borgpetersenii clade E in the kidney sample from another animal. In Tanzania, L. kirschneri and L. borgpetersenii clade A were detected in a single kidney sample.

Phylogenetic relationship to Leptospira from other livestock studies in the developing world

Analysis of secY sequences from this study and other studies in northern Tanzania (Allan et al., 2018), Reunion island (Guernier et al., 2016) and Brazil (Hamond et al., 2015; Guedes et al., 2019) that used similar molecular approaches identified five recognised pathogenic Leptospira species (L. borgpetersenii, L. santarosai, L. interrogans, L. noguchii, L. kirschneri) and one L. kirschneri-like species infecting cattle (Figure 2). In Madagascar, four L. borgpetersenii secY clades (A-D) were identified. With the exception of clade B, all remaining clades contained secY sequences from cattle from other regions of the world. Both clade A and D were detected in cattle from Tanzania (Allan et al., 2018), Reunion (Guernier et al., 2016) and Brazil (Hamond et al., 2015; Guedes et al., 2019). Clade A sequences were identical to L. borgpetersenii serovar Hardjo-bovis strains JB197 and L550, strains associated with cattle (Bulach et al., 2006), which lfb1 sequencing had been able to differentiate in Tanzanian samples (Figure 1: L. borgpetersenii clade A and B). However, clade D sequences matched no reference strains, with sequences in clade B providing the closest match (98% identity). Sequences in clade C were detected in a single animal in this study and a single animal in Reunion (Guernier et al., 2016) and were identical to reference sequences from strains previously associated with small mammals (Victoria et al., 2008). Moreover, the corresponding lfb1 sequence from Madagascan cattle (Figure 1, L. borgpetersenii clade E) was identical to numerous lfb1 sequences obtained from Madagascan small mammals (Moseley et al., 2018). Although the single L. kirschneri sequence from this study was novel, it was closely related to sequences obtained from cattle in the Brazilian Amazon (Guedes et al., 2019) and to L. kirschneri serovar Galtoni, which was obtained from a cattle kidney sample in Argentina (Victoria et al., 2008), differing by a single synonymous polymorphism. L. santarosai and L. noguchii, which have been identified in Brazil (Hamond et al., 2015; Guedes et al., 2019) and a L. kirschneri-like species recently identified in both Brazil (Guedes et al., 2019) and Tanzania (Allan et al., 2018) were not identified in Madagascan cattle.

Figure 2.

Figure 2

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Maximum likelihood phylogenetic tree of 329bp secY sequences (accession numbers MK244342-MK244344 and MK610269-MKMK610274) using the Tamura 3-Parameter model with a discrete Gamma distribution. Samples from Madagascar are highlighted in bold with the number of animals from which sequences were obtained and a representative accession number indicated. Sequences from similar livestock studies are labelled by country of origin, host, number of sequences and accession number of a representative sequence. Reference sequences (Victoria et al., 2008) are labelled by Leptospira species, serovar and accession number. Nodes are labelled with bootstrap support.

Discussion

We show that cattle have a higher prevalence of Leptospira infection than pigs in Madagascar and are potential reservoir hosts for a diversity of Leptospira with genetic similarities to Leptospira strains identified in cattle in other tropical regions of the world, some of which are genetically distinct from reference strains. Increased probability of detection of Leptospira infections in sample preparations that included multiple kidney lobes or urine rather than single kidney lobes, suggest a localised distribution for Leptospira infections in cattle kidneys or low Leptospira loads in single samples. These findings support previous studies (Guedes et al., 2019), where prevalence estimates from urine samples (14.9%, 31/208) were higher than those obtained from kidney samples (5.8%, 12/208), and highlight the importance of evaluating sample collection methods when evaluating prevalence estimates. Leptospira sequencing success was dependent on parasite load and the inability to obtain Leptospira sequence data from pigs was likely due to the low parasite load in these samples.

The identification of strains in Madagascar, Tanzania, Reunion and Brazil, such as L. borgpetersenii clade D (Figure 2), genetically distinct from any reference strains, has implications for the control and diagnosis of leptospirosis in the developing world. In Brazil, it has been demonstrated that the efficacy of commercial vaccines is poor in the face of autochthonous strain challenge (Sonada et al., 2018) and it is likely that commercial vaccines are likely to perform as poorly in Madagascar and other developing countries. Moreover, the inclusion of local isolates in antigen panels for serological assays, such as MAT, can dramatically increase the serological detection of infections (Mgode et al., 2015). Therefore, the identification of strains with no associated reference strains suggest that existing antigen panels might underestimate the prevalence of human infections.

In addition to strains associated with livestock, we also identified infection of cattle with strains previously detected in small mammals. For example, in Madagascar, a L. interrogans strain common in small mammals (Moseley et al., 2018) and identified in an acute human case of leptospirosis (Pagès et al., 2015) was also identified in a goat. Moreover, secY sequencing identified a L. borgpetersenii strain previously detected in small mammals (Victoria et al., 2008) in cattle in this study and in Reunion (Guernier et al., 2016) and lfb1 sequencing confirmed this strain as identical to L. borgpetersenii previously identified in small mammals in Madagascar (Moseley et al., 2018). Thus, our results suggest transmission of at least some Leptospira strains between small mammals and livestock, and raise the potential that livestock may play a role in the transmission of these strains. For example, due to their large size, livestock could act as amplification hosts for small mammal associated strains by increasing environmental contamination through urinary shedding after incidental infection or could act as reservoir hosts in their own right, possibly explaining why contact with cattle has been identified as a risk factor for human Leptospira infection in Madagascar despite serological typing identifying serogroup Icterohaemorrhagiae, a serogroup associated with rodents, as the predominant serogroup (Ratsitorahina et al., 2015). In contrast, in Tanzania, where small mammals sampled tested negative for Leptospira infection (Allan et al., 2018), no evidence was found for infection of livestock with strains previously associated with small mammals.

We also confirm the presence of mixed infections in livestock in both Madagascar and Tanzania, supporting previous evidence for mixed infections in small mammals (Moseley et al., 2018). Horizontal genetic transfer plays an important role in the evolution and serological classification of Leptospira (Llanes et al., 2016), and mixed infections within the same host provide the ideal environment for this to take place. Moreover, for Borrelia, it has been proposed that mixed infections may facilitate maintenance of infection in reservoir hosts (Andersson et al., 2013). Where serological diagnostic assays, such as MAT, rely on evaluating serological response to specific antigens, mixed infections with strains which may represent different serovars could complicate interpretation. Further research is needed to clarify the role of mixed infections in the evolution and epidemiology of Leptospira.

Our results emphasise that Leptospira epidemiology in tropical, developing country contexts, where close human contact with livestock is more likely and farming systems may promote contact between small mammals and livestock, may be very different to developed country settings. In addition, considering sampling methodology is important when comparing studies and planning surveillance and further work is needed to optimise abattoir sampling strategies. In Madagascar and other developing countries, evidence that livestock are infected with potentially novel Leptospira strains highlights the need for understanding the diversity of Leptospira circulating in livestock to inform diagnostic antigen panels and vaccine development in these regions.

Supplementary Material

Supplementary information

Acknowledgements

We thank the staff of the Plague Central Laboratory, Institut Pasteur de Madagascar, for technical assistance, especially Fehivola Andriamiarimanana, as well as staff from the abattoirs for their assistance during sampling.

Financial Support

This work was supported the Wellcome Trust (Senior Fellowship no. 095171 to S.T., Veterinary Training Fellowship no. 096400/Z/11/Z to K.A., Institutional Strategic Support Fund to the University of Aberdeen no. 2014815/Z/16/Z), the Biotechnology and Biological Sciences Research Council (no. BB/M010996/1), the University of Aberdeen Environment and Food Security theme and Institut Pasteur Madagascar.

Footnotes

Conflicts of Interest

None

Ethical Standards

Not applicable

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