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Leishmania infantum infection does not affect the main composition of the intestinal microbiome of the Syrian hamster



Visceral leishmaniasis (VL) is the most severe form of all leishmanial infections and is caused by infection with protozoa of Leishmania donovani and Leishmania infantum. This parasitic disease occurs in over 80 countries and its geographic distribution is on the rise. Although the interaction between the intestinal microbiome and the immune response has been established in several pathologies, it has not been widely studied in leishmaniasis. The Syrian hamster is the most advanced laboratory model for developing vaccines and new drugs against VL. In the study reported here, we explored the relationship between the intestinal microbiome and infection with L. infantum in this surrogate host.


Male Syrian hamsters (120–140 g) were inoculated with 108 promastigotes of a canine-derived L. infantum strain or left as uninfected control animals. Infection was maintained for 19 weeks (endpoint) and monitored by an immunoglobulin G (IgG) enyzme-linked immunosorbent assay throughout the experiment. Individual faecal samples, obtained at weeks 16, 18 and 19 post-inoculation, were analysed to determine the 16S metagenomic composition (the operational taxonomic units [OTUs] of the intestinal microbiome and the comparison between groups were FDR (false discovery rate)-adjusted).


Leishmania infantum infection elicited moderate clinical signs and lesions and a steady increase in specific anti-Leishmania serum IgG. The predominant phyla (Firmicutes + Bacteriodetes: > 90%), families (Muribaculaceae + Lachnospiraceae + Ruminococcaceae: 70–80%) and genera found in the uninfected hamsters showed no significant variations throughout the experiment. Leishmania infantum infection provoked a slightly higher—albeit non-significant—value for the Firmicutes/Bacteriodetes ratio but no notable differences were found in the relative abundance or diversity of phyla and families. The microbiome of the infected hamsters was enriched in CAG-352, whereas Lachnospiraceae UCG-004, the [Eubacterium] ventriosum group and Allobaculum were less abundant.


The lack of extensive significant differences between hamsters infected and uninfected with L. infantum in the higher taxa (phyla, families) and the scarce variation found, which was restricted to genera with a low relative abundance, suggest that there is no clear VL infection-intestinal microbiome axis in hamsters. Further studies are needed (chronic infections, co-abundance analyses, intestinal sampling, functional analysis) to confirm these findings and to determine more precisely the possible relationship between microbiome composition and VL infection.

Graphical Abstract


Leishmaniasis refers to a broad group of vector-borne parasitic diseases caused by species of the genus Leishmania. Infection can occur in humans and other mammals and causes a range of clinical presentations (visceral, cutaneous, mucocutaneous, post-kala-azar dermal leishmaniasis) [1]. The most severe disease is visceral leishmaniasis (VL), which is fatal unless treated, and is caused by Leishmania donovani and Leishmania infantum (= L. chagasi) [2, 3]. The geographic distribution of VL is increasing due to human migration and travel, as well as the spread of vector populations to previously unaffected regions [4]. The infection is also frequent in HIV-positive patients in endemic areas [5,6,7] and has been reported in recipients of solid organ transplants [8, 9]. While infections by L. donovani are regarded as anthroponotic, leishmaniasis caused by L. infantum is zoonotic (zoonotic VL), and dogs are considered to be the main reservoir of human disease [10,11,12]. Disease outcome after inoculation depends on the inherent virulence of the Leishmania strain, individual immune response, host health status and intercurrent infections, among other factors [13, 14]. The Syrian hamster is the most advanced surrogate model for Leishmania infections, particularly those caused by L. donovani and L. infantum, and is widely used to study pathogenicity and experimental chemotherapy [15,16,17,18,19,20].

There is a growing awareness of the importance of gut microbiota for general health status [21]. It has been shown that systemic infections can alter the intestinal microbiota [22], and current evidence suggests that there is a two-way interaction where alterations in gut microbiota affect infectious diseases, while infectious diseases in turn regulate the structure and function of gut microbiota [23,24,25,26]. However, there is little information on the intestinal microbiome of the hamster [27, 28], and to the best of our knowledge, the only relationship studied in this host species is between L. infantum infection and Bifidobacterium spp. and Lactobacillus spp. [29]. Thus, in the context of a wider project whose results will be published elsewhere, we have determined the 16S metagenomic composition of the intestinal microbiome of hamsters experimentally infected with L. infantum.


Experimental design

Syrian male hamsters (Mesocricetus auratus) (n = 14), aged 7–8 weeks, each weighing between 85 and 120 g, were purchased from Janvier Labs (Le Genest-Saint-Isle, France) and placed in quarantine. The animals were kept under observation and housed under a controlled temperature regimen and 12:12-h light:dark cycle (Instituto de Investigación Hospital 12 de Octubre, Madrid, Spain). They were provided with commercial pelleted food and water ad libitum. When the animals reached a weight of 120–140 g they were divided in a stratified manner (live weight [lw]) and inoculated with a canine-derived autochthonous strain of L. infantum (MCAN/ES/96/BCN150) (experimental group 3 [G3]; n = 8) at 108 promastigotes/hamster [19, 30] or maintained uninfected (control group 1 [G1]: n = 6). The animals were euthanized at approximately 19 weeks post inoculation (wpi). The size of the uninfected control group was kept to a minimum on ethical grounds and from prior experience with such control groups.

The lw was determined on day 0 (preinfection), 16 wpi and at the endpoint of the experiment. Blood samples were obtained from the cava vein under anaesthesia (isoflurane 2–4%) before infection and 16 wpi; the endpoint sample was obtained by intracardiac puncture. Sera from all the experimental animals were used to determine the infection status of the hamsters by the immunoglobulin G enzyme-linked immunosorbent immunoglobulin (IgG ELISA) using the standard indirect ELISA protocol from our laboratory. The 96-well ELISA plates (Nunc MaxiSorp™, Thermo Fisher Scientific, Waltham, MA, USA) were coated with 50 µl/well of 7.5 µg/ml soluble L. infantum antigen in HCO3/CO3 (4 °C, overnight). Sera samples were added to the wells at 1/100 dilution, 50 µl/well and incubated at 37 °C for 1 h. The secondary antibody was goat anti-hamster IgG (H+L)-HRP (Southern Biotech, Birmingham, AL, USA) (1/2000 dilution; incubated for 30 min at room temperature); o-phenylenediamine (1 mg/ml) (Sigma-Aldrich, St. Louis, MO, USA) + H2O2 (1/1000) solution were added (100 µl/well) and the reaction was stopped with the addition of 50 µl/well of H2SO4 (3 N). Absorbance at 492 nm was determined in a Multiskan™ GO Microplate Spectrophotometer (Thermo Fisher Scientific). ELISA results (optical density [OD]) were expressed as a percentage of the positive control serum. The OD cut-off (±) was established at mean preinfection values of +3 standard deviations (SD) (13.74%). All determinations were performed at least in triplicate.

Genomic DNA extraction, 16S metagenome library construction and next-generation sequencing

Individual faecal samples were obtained at week 16 pi and week 18 pi by temporarily isolating individual animals and collecting faecal pellets immediately after dropping. At the end of the experiment (endpoint: 19 wpi), faecal samples were obtained from the rectum during the necropsy and stored at − 80 °C until processing. For the genomic analyses, five faecal samples from G1 (uninfected control hamsters) and four from G3 (L. infantum-infected hamsters) were randomly selected from each sampling time.

DNA extraction, construction of the next-generation sequencing (NGS) library and sequencing were carried out at the Unidad de Genómica, Complutense University of Madrid (Spain). Total DNA from hamster faecal samples was extracted with the DNeasy PowerLyser PowerSoil DNA Kit (Qiagen, Hilden, Germany) following the manufacturer’s instructions. DNA concentration was estimated using the Qubit 2.0 fluorimeter (Life Technologies™, Thermo Fisher Scientific). DNA libraries from each sample were prepared following the Illumina 16S Metagenomic Sequencing Library Preparation manual (Illumina, San Diego CA, USA). In brief, the V3–V4 region of the prokaryotic 16S ribosomal RNA (rRNA) was amplified for each sample with primers containing the 341F and 805R sequences and Illumina-specific adapters. In a second PCR amplification, two specific 8-nucleotide index and i5/i7 Illumina adapters were added to the previous amplicons. DNA libraries were checked with the Bioanalyzer 2100 platform (Agilent Technologies, Palo Alto, CA, USA). A library pool was prepared for sequencing by mixing equal amounts of the individual sample libraries, and then sequenced in the Illumina MiSeq benchtop sequencer with 2 × 300 reads using the 600 cycle MiSeq Reagent Kit v3 in accordance with the manufacturer’s recommended protocol.

Sequence data analysis

The FASTQ files containing the sequencing reads were analysed using the CLC Genomics Workbench version 20.0.4 (QIAGEN Aarhus A/S, Aarhus, Denmark; Sequence data were trimmed using 0.05 as a limit for quality scores, with 2 as the maximum number of ambiguities. The reads after trimming were analysed using the CLC Microbial Genomics Module version 20.1.1. The optional merge paired reads method was run with default settings (mismatch cost = 1; minimum score = 40; gap cost = 4; maximum unaligned end mismatch = 5). Sequence reads were clustered and chimeric sequences detected using an identity of 97% as the operational taxonomic unit (OTU) threshold. The reference OTU data used in the present study were downloaded from the SILVA database [31] v132 for 16S rRNA. Shannon’s diversity index was calculated considering the assigned species. The raw sequencing data were deposited in the NCBI Sequence Read Archive database (BioProject ID: PRJNA843999) (

Statistical analysis

The experimental groups were included in a larger experiment, and the number of animals was selected to give a Z-power of 0.8 and a 95% level of significance. Unless otherwise stated, the numerical values are presented as the mean ± SD. Statistical analyses included parametric and non-parametric tests (1w- and 2w-analysis of variance [ANOVA], Mann–Whitney test, Student t-test), and the level of significance was set at P ≤ 0.05. The taxonomic comparison between the groups was performed with the differential abundance analysis tool from the CLC Microbial Genomics Module. The table of OTUs generated by the CLC Microbial Genomics Module from each microbiome classified at phylum, family or genus levels was used as the input. Unless otherwise stated, only changes of at least ± twofold (±) in the present taxa, and false discovery rates (FDR) with an adjusted P-value of ≤ 0.05, were considered as significant. The figures were prepared with GraphPad Prism 9.0 (GraphPad Software, San Diego, CA, USA) and Microsoft Excel (Microsoft Corp., Redmond, WA, USA).


Leishmania infantum infection

None of the non-inoculated animals developed any cutaneous lesion throughout the experiment, whereas lesions were observed in five of the eight infected hamsters (62.5%). Lesions included dermatitis, exfoliative dermatitis and crusts in different locations (e.g. back, abdomen, legs, neck, inguinal and axillar regions, ears) and alopecia (chest, face, periorbitary). Inoculation with L. infantum did not cause significant lw loss, and hamsters reached comparable lw at week 16 irrespective of their infection status (161.85 ± 13.19 g [uninfected G1] vs 162.33 ± 11.15 g) [infected G3]).

The control animals (G1) showed no variation in the the serum-specific IgG response throughout the experiment, and their OD values were in all cases below the cut-off (Fig. 1). In the inoculated animals (G3), there was a notable increase in anti-Leishmania IgG levels at week 16 pi and 3 weeks later (i.e. endpoint: 19 weeks) when the inoculated hamsters attained an anti-Leishmania IgG level of 104.9 ± 12.9% of that of the positive control serum compared to the uninfected control group (9.4 ± 0.5%).

Fig. 1
figure 1

Serum-specific anti-Leishmania immunoglobulin G (IgG) response of inoculated (G3) (solid circles) and control (G1) (empty circles) hamsters throughout the experiment. ELISA values correspond to the percentage OD from the positive control sera. Serum IgG levels were determined before infection (preinfection), 16 WPI and at the endpoint of the experiment (19 WPI). Individual values, mean of the group and standard deviation (%) are given. ELISA, Enzyme-linked immunosorbent assay; OD, optical density; WPI, weeks post inoculation

Intestinal microbiome of the control hamsters

The analysis of the predominant taxa (OTU) in the microbiome in the faecal samples obtained at week 16 yielded 17 phyla. The most abundant were Firmicutes and Bacteriodetes, representing an average of ≥ 90% of the OTUs, followed by Verrucomicrobia (1.7–< 0.1%), Proteobacteria (1.9–0.7%), Cyanobacteria (1.3–0.4%), Patescibacteria (1.4–0.1%), Actinobacteria (0.3–< 0.1%), Tenericutes, Deferribacteres and Elusimicrobia. The most represented families were: Muribaculaceae (Bacteroidetes), Lachnospiraceae (Firmicutes) and Ruminococcaceae (Firmicutes), which accounted for 70–80% of the total number (n = 78) of families; Prevotellaceae (Phy. Bacteroidetes), Akkermansiaceae (Phy. Verrucomicrobia) and Lactobacillaceae (Phy. Firmicutes) represented approximately 6–12% of families (Additional file 1: Table S1). A total of 169 genera were found, of which—in addition to the dominant uncultured bacterium (41.1 ± 6%)—the most abundant were the Lachnospiraceae NK4A136 group (Firmicutes, Lachnospiraceae), Ruminococcaceae UCG-014 (Firmicutes, Ruminococcaceae) and Alloprevotella (Bacteroidetes, Prevotellaceae), although there was ample representation of other genera (Fig. 2).

Fig. 2
figure 2

Dominant genera (20 most abundant) identified in the faecal samples obtained from the uninfected control hamsters (G1) at week 16 of the experiment (16 wpi). The values represent the mean relative abundance (%) in the samples analysed (n = 5)

The relative abundance of OTUs in the uninfected animals (G1) did not vary for the most prevalent taxa throughout the experimental period, and Shannon’s index suggested a similar diversity (16 weeks: 3.3; 18 weeks: 3.1; 19 weeks: 3.3). The microbiome analysis of the uninfected hamsters was therefore homogeneous in terms of the most represented phyla, most abundant families (Fig. 3) and the genera. The statistical analysis revealed no significant difference throughout the experiment.

Fig. 3
figure 3

Relative abundance (% and colour scale) of the most represented phyla and families in the faecal samples from the uninfected hamsters (G1) throughout the experimental period, and from the infected hamsters (G3) at the endpoint. Values represent the mean relative abundance (%) in the samples (n = 5 for G1 samples; n = 4 for G3 samples). The colour scale (green to red) shows a graphic representation of the relative abundance within each group and sampling time. 16w, 16 weeks post inoculation; 18w, 18 weeks post inoculation; 19w (EP), 19 weeks post inoculation (endpoint)

Effect of L. infantum infection on microbiome composition

No significant differences were found in the phyla at 16 wpi between the uninfected control hamsters (G1) and hamsters infected with L. infantum (G3) (G1 vs G3, P = 0.414), with OTUs of > 99.9% of all reads. This lack of significant differences was also observed 2 weeks later (G1 vs G3, FDR P-value = 0.196) and at the endpoint of the experiment (G1 vs G3, FDR P-value = 0.99) (Fig. 3). Despite the slightly higher value of the Firmicutes/Bacteroidetes (F/B) ratio found in infected animals (F/B = 2.47), the difference was not significant. Similar results were obtained for the families, considering the 30 most abundant (G1 vs G3, FDR P-value ≥ 0.99) for all the sampling times (16 weeks, 18 weeks and endpoint).

No significant differences were found in any of the time-matched samplings when the values of the 30 most abundant genera were compared (> 94% reads). Moreover, neither the phyla, families nor genera in the control group showed any significant variation between the three sampling times (G1: uninfected: 16 vs 18 weeks vs endpoint; G3: 16 vs 18 weeks vs endpoint).

The taxonomic comparison (FDR analysis, FDR P-value) of the hamster groups showed no significant differences in phyla between the uninfected control animals (G1) and the L. infantum-infected hamsters at 16 weeks (G3); this lack of significance was maintained for the entire experiment (minimum FDR value in all comparisons = 0.0975). Similarly, there were no significant differences in the relative abundance of families in both groups (minimum FDR P-value in all comparisons = 0.1237) (Additional file 2: Table S2). However, there were variations in the relative abundance of the genera identified: CAG-352 (Clostridia, Firmicutes) significantly increased its presence at week 16 (FDR P-value = 0.0078), week 18 and at the endpoint of the experiment (week 19, FDR P-value = 0.0309), while in the final sampling, Lachnospiraceae UCG-004, [Eubacterium] ventriosum and the Allobaculum group showed a significantly lower abundance (FDR P-value = 0.0059, 0.0015 and 0.0314, respectively). Significant differences (FDR P-value = 3.51 × 10–6 in the relative abundance of Ruminococcus 2 were found between groups at week 16 but these progressively disappeared, and no differences were observed by the end of the experiment (FDR P-value > 0.05) (Fig. 4).

Fig. 4
figure 4

Comparison of genera abundance (log2) in the intestinal microbiome of hamsters infected with L. infantum (G3) and uninfected (G1) throughout the experiment. Asterisk indicates a statistically significant difference (false discovery rate [FDR] P-value < 0.05)


The course of leishmaniasis is dependent on the imbalance in the host’s immune response [13, 14], which in turn is related to the virulence of Leishmania and the presence of intercurrent infections, among other factors. In experimental infections in surrogate models, the dose administered has been shown to affect the infection outcome with different Leishmania spp. [32,33,34]. Experimental infections of VL in hamsters have been carried out with a variety of infective doses and inoculation procedures: intracardiac [15, 18, 35,36,37], intraperitoneal [36, 38] and intradermic) [36]. In our experiment, hamsters were inoculated by the retro-orbitary route with 108 stationary phase promastigotes of L. infantum, since both infective dose and inoculation procedure elicit consistent infections in hamsters with the same L. infantum strain [39].

Under our experimental conditions, the inoculation of Syrian hamsters with L. infantum promastigotes provoked a steady rise in specific antibodies in all inoculated animals throughout the experiment, confirming the infectivity of the parasite strain and the efficiency of the inoculation procedure. The absence of weight loss and the moderate cutaneous alterations in the infected hamsters were comparable to previous findings under similar experimental conditions (age of animals, infective dose, Leishmania strain) [19]. Although most exploratory research on VL (e.g. immune response, pathology, vaccine candidates, chemotherapeutic lead compounds) is done in mice, hamster infections are long-lasting [15] and mimic the infections in the natural hosts (humans, dogs) more closely since they lack the nitric oxide response against Leishmania [16] and are considered to be a better model for VL [17,18,19,20]. The relationship between the intestinal microbiome and health is considered to be of paramount importance and has fuelled the exploration of its composition both in humans [21,22,23,24, 26, 40] and in dogs and cats [41,42,43]. While there is abundant information on the intestinal microbiome of mice [26, 44,45,46], very few studies have been conducted and published on hamsters. We found a higher diversity in the microbiome than previously reported for Syrian hamsters [47,48,49]. The most abundant phyla (Firmicutes, Bacteriodetes, Verrucomicrobia, Proteobacteria, Saccharibacteria) had previously been reported in this animal species, with the exception of a nutrition-focused study [48]. The relative abundance was variable, although the values for the main phyla were comparable to those of the most recent study [49]; the predominance of Firmicutes (94%) reported by Martínez et al. [47] could not be confirmed. In fact, the relative abundance of OTUs in our study in the case of phyla is in line with the findings reported in most animal species, including mice [46, 48], dogs and cats [41, 42, 50, 51] and humans [52, 53]. The high variability reported could be related to the strong influence of diet, genotype and environmental factors [44, 54], including housing conditions, which—together with the different methodologies used—would affect their inter-laboratory reproducibility. This is relevant since the results cannot be compared unless standardized designs and methods are used.

The interaction between several pathological conditions and the intestinal microbiome composition of humans [21, 24,25,26, 53] and other animals, including dogs and cats [41, 42, 51], has been explored. In the only study carried out in humans with VL, despite the reduction of Ruminococcaceae UCG-014 and Gastranaerophilales uncultured bacterium in VL patients (n = 23), the overall comparison (alpha and beta analyses) showed no variation in infected individuals [55]. Cross-sectional studies are hampered by individual variations (diet, habits), and data derived from such studies are thus difficult to interpret and extrapolate. In our study the hamsters were obtained from a standardized supplier and maintained throughout the experiment in the same animal facilities and under the same conditions; thus, the intergroup differences (uninfected vs L. infantum-infected animals) could be considered to be factual. In the only study carried out so far with Leishmania-infected hamsters under controlled conditions, no relationship was found between the abundance of intestinal Bifidobacterium spp. and Lactobacillus and infection with L. infantum despite the duration of the infection (up to 8 months) [29]. In the present experiment, we used a powerful analytical technique (16S) followed by a robust statistical analysis (FDR filtered P-value < 0.05), and found no major differences in the relative abundance of phyla and families in the intestinal microbiome; the variation observed was restricted to certain non-abundant genera (CAG-352, Lachnospiraceae UCG-004, [Eubacterium] ventriosum group and Allobaculum). CAG-352 has been linked to human prostate cancer [56], and the role of the Lachnospiraceae family is far from clear [52, 57]. No mechanistic explanation is therefore available on the variation of these Clostridia (Firmicutes) in L. infantum-infected hamsters. Increases in butyrate-producing isolates of Eubacterium spp. have been related to higher body mass, which implies more efficient energy utilization [58]. However, in our case no significant differences were observed in the lw of the experimental animals. More recently this genus has also been implicated in the modulation of inflammation and the regulation of immune responses [59], and leishmaniasis outcomes are linked to an unbalanced host-parasite immune response [13, 14]. In our experiment the infected hamsters showed a 165-fold reduction in the [Eubacterium] ventriosum group compared to the uninfected animals (endpoint, week 19), accompanied by high levels of anti-Leishmania IgG antibodies. Although this relationship is suggestive, the number of experimental animals, duration of the infection and low relative abundance of the taxon (0.2% in healthy hamsters) do not support a causality relationship. More research is therefore required.


In this study, Syrian hamsters were inoculated with a canine-derived L infantum strain and maintained for 19 weeks. The infection status of the animals was assessed by the presence of clinical signs and lesions and indirect IgG ELISA. The relationship between the intestinal microbiome (16S) and L. infantum infection in hamsters was studied for the first time. No major differences were found in higher taxa, and the actual significance of the slight variations found in non-abundant genera is unknown. Although we are aware of the limitations of the study (e.g. study duration, number of animals, co-abundance analysis and additional parasitological determinations) and the probable scarce translatability of microbiome results in rodents to the target hosts [54], the main conclusion is that no clear VL infection-intestinal microbiome axis has been identified. This finding should be confirmed in the future with chronically infected animals and by precisely determining the possible role of specific bacterial differences in the pathogenesis of VL.

Availability of data and materials

All relevant data are given in the manuscript and Supplementary information. Materials, when available, can be requested to the authors.



False discovery rate


Immunoglobulin G enzyme-linked immunosorbent assay


Live weight


Next-generation sequencing


Operational taxonomic unit


Visceral leishmaniasis


weeks post inoculation


  1. WHO/Expert Committee on the Control of the Leishmaniases. Control of the leishmaniases: report of a meeting of the WHO Expert Committee on the Control of Leishmaniases, Geneva, 22–26 March 2010. 2010. Accessed 24 June 2022.

  2. Alvar J, Vélez ID, Bern C, Bern C, Herrero M, Desjeux P, et al. Leishmaniasis worldwide and global estimates of its incidence. PLoS ONE. 2012;7:35671.

    Article  CAS  Google Scholar 

  3. Pace D. Leishmaniasis. J Infect. 2014;69:510–8.

    Article  Google Scholar 

  4. Naucke TJ, Amelung S, Lorentz S. First report of transmission of canine leishmaniosis through bite wounds from a naturally infected dog in Germany. Parasit Vectors. 2016;9:256.

    Article  CAS  Google Scholar 

  5. Pintado V, López-Vélez R. HIV-associated visceral leishmaniasis. Clin Microbiol Infect. 2001;7:291–300.

    Article  CAS  Google Scholar 

  6. Monge-Maillo B, Norman FF, Cruz I, Alvar J, López-Vélez R. Visceral leishmaniasis and HIV coinfection in the Mediterranean Region. PLoS Negl Trop Dis. 2014;8:e3021.

    Article  Google Scholar 

  7. Van Griensven J, Carrillo E, López-Vélez R, Moreno J. Leishmaniasis in immunosuppressed individuals. Clin Microbiol Infect. 2014;20:286–99.

    Article  Google Scholar 

  8. Antinori S, Cascio A, Parravicini C, Bianchi R, Corbellino M. Leishmaniasis among organ transplant recipients. Lancet Infect Dis. 2008;8:191–9.

    Article  Google Scholar 

  9. Gajurel K, Dhakal R, Deresinski S. Leishmaniasis in solid organ and hematopoietic stem cell transplant recipients. Clin Transplant. 2017;31:e12867.

    Article  Google Scholar 

  10. Alvar J, Cañavate C, Molina R, Moreno J, Nieto J. Canine leishmaniasis. Adv Parasitol. 2004;57:1–88.

    Article  Google Scholar 

  11. Burza S, Croft SL, Boelaert M. Leishmaniasis. Lancet. 2018;392:951–70.

    Article  Google Scholar 

  12. Morillas F, Sánchez Rabasco F, Ocaña J, Martín-Sánchez J, Ocaña-Wihelmi J, Acedo C, et al. Leishmaniosis in the focus of the Axarquia region, Málaga province, southern Spain: a survey of the human, dog, and vector. Parasitol Res. 1996;82:569–70.

    Article  CAS  Google Scholar 

  13. Kumar R, Nylén S. Immunobiology of visceral leishmaniasis. Front Immunol. 2012;3:251.

    Article  CAS  Google Scholar 

  14. McCall LI, Zhang WW, Matlashewski G. Determinants for the development of visceral leishmaniasis disease. PLoS Pathog. 2013;9:e1003053.

    Article  CAS  Google Scholar 

  15. Requena JM, Soto M, Doria MD, Alonso C. Immune and clinical parameters associated with Leishmania infantum infection in the golden hamster model. Vet Immunol Immunopathol. 2000;76:269–81.

    Article  CAS  Google Scholar 

  16. Melby PC, Chandrasekar B, Zhao W, Coe JE. The hamster as a model of human visceral leishmaniasis: progressive disease and impaired generation of nitric oxide in the face of a prominent Th1-like cytokine response. J Immunol. 2001;166:1912–20.

    Article  CAS  Google Scholar 

  17. Garg R, Dube A. Animal models for vaccine studies for visceral leishmaniasis. Indian J Med Res. 2006;123:439–54.

    Google Scholar 

  18. Dea-Ayuela MA, Rama-Íñiguez S, Alunda JM, Bolás-Fernández F. Setting new immunobiological parameters in the hamster model of visceral leishmaniasis for in vivo testing of antileishmanial compounds. Vet Res Commun. 2007;31:703–17.

    Article  CAS  Google Scholar 

  19. Jiménez-Antón MD, Grau M, Olías-Molero AI, Alunda JM. Syrian hamster as an advanced experimental model for visceral Leishmaniasis. Methods Mol Biol. 2019;1971:303–14.

    Article  CAS  Google Scholar 

  20. Saini S, Rai AK. Hamster, a close model for visceral leishmaniasis: opportunities and challenges. Parasite Immunol. 2020;42:e12768.

    Article  Google Scholar 

  21. Heintz-Buschart A, Wilmes P. Human gut microbiome: function matters. Trends Microbiol. 2017;26:563–74.

    Article  CAS  Google Scholar 

  22. Woodall CA, McGeoch LJ, Hay AD, Hammond A. Respiratory tract infections and gut microbiome modifications: a systematic review. PLoS ONE. 2022;17:e0262057.

    Article  CAS  Google Scholar 

  23. Belkaid Y, Hand T. Role of the microbiota in immunity and inflammation. Cell. 2014;157:121–41.

    Article  CAS  Google Scholar 

  24. Ho JT, Chan GC, Li JC. Systemic effects of gut microbiota and its relationship with disease and modulation. BMC Immunol. 2015;16:21.

    Article  CAS  Google Scholar 

  25. Lv L-X, Jiang H-Y, Yan RY, Li L. Interactions between gut microbiota and hosts and their role in infectious diseases. Infect Microbes Dis. 2019;1:3–9.

    Article  CAS  Google Scholar 

  26. Zheng D, Liwinski T, Elinav E. Interaction between microbiota and immunity in health and disease. Cell Res. 2020;30:492–506.

    Article  Google Scholar 

  27. Philippe-Taine G, Coroler L, Levy RH, Gillardin JM. Dose-dependent preventive effect of Saccharomyces boulardii on clindamycin-induced alterations in intestinal aerobic flora of the hamster. Microb Ecol Health Dis. 2003;15:126–30.

    Article  CAS  Google Scholar 

  28. Miezeiewski M, Schnaufer T, Muravsky M, Wang S, Caro-Aguilar I, Secore S, et al. An in vitro culture model to study the dynamics of colonic microbiota in Syrian golden hamsters and their susceptibility to infection with Clostridium difficile. ISME J. 2015;9:321–32.

    Article  CAS  Google Scholar 

  29. Correia Passos F, BiondaroGois M, Duranes Sousa A, Lima de Marinho AI, Corvo L, Soto M, et al. Investigating associations between intestinal alterations and parasite load according to Bifidobacterium spp. and Lactobacillus spp. abundance in the gut microbiota of hamsters infected by Leishmania infantum. Mem Inst Oswaldo Cruz Rio de Janeiro. 2020;115:e200377.

    Article  CAS  Google Scholar 

  30. Jiménez-Antón MD, Grau M, Corral MJ, Olías-Molero AI, Alunda JM. Efficient infection of hamster with Leishmania donovani by retro-orbital inoculation. Virulence. 2019;10:711–8.

    Article  CAS  Google Scholar 

  31. Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2013;41:D590–6.

    Article  CAS  Google Scholar 

  32. Rostamian M, Jafari D, Abolghazi M, Farahani H, Niknam HM. Leishmania tropica: suggestive evidences for the effect of infectious dose on pathogenicity and immunogenicity in an experimental model. Parasitol Res. 2018;117:2949–56. (Epub 2018 Jul 5).

    Article  Google Scholar 

  33. Doherty TM, Coffman RL. Leishmania major: effect of infectious dose on T cell subset development in BALB/c mice. Exp Parasitol. 1996;84:124–35.

    Article  CAS  Google Scholar 

  34. Loeuillet C, Bañuls AL, Hide M. Study of Leishmania pathogenesis in mice: experimental considerations. Parasites Vectors. 2016;9:144.

    Article  CAS  Google Scholar 

  35. Fernández L, Solana JC, Sánchez C, Jiménez MÁ, Requena JM, Coler R, et al. Protective efficacy in a hamster model of a multivalent vaccine for human visceral leishmaniasis (MuLeVaClin) consisting of the KMP11, LEISH-F3+, and LJL143 antigens in virosomes, plus GLA-SE adjuvant. Microorganisms. 2021;9:2253.

    Article  CAS  Google Scholar 

  36. Moreira NDD, Vitoriano-Souza J, Roatt BM, Vieira PMDA, Ker HG, de Oliveira Cardoso JM, et al. Parasite burden in hamsters infected with two different strains of Leishmania (Leishmaniainfantum: “leishman Donovan units” versus real-time PCR. PLoS ONE. 2012;7:e47907.

    Article  CAS  Google Scholar 

  37. Fortin A, Dorlo TPC, Hendrickx S, Maes L. Pharmacokinetics and pharmacodynamics of oleylphosphocholine in a hamster model of visceral leishmaniasis. J Antimicrob Chemother. 2016;71:1892–8.

    Article  CAS  Google Scholar 

  38. Corral MJ, Serrano DR, Moreno I, Torrado JJ, Domínguez M, Alunda JM. Efficacy of low doses of amphotericin B plus allicin against experimental visceral leishmaniasis. J Antimicrob Chemother. 2014;69:3268–74. (Epub 2014 Aug 4).

    Article  CAS  Google Scholar 

  39. Jiménez-Antón MD, García-Calvo E, Gutiérrez C, Escribano MD, Kayali N, Luque-García JL, et al. Pharmacokinetics and disposition of miltefosine in healthy mice and hamsters experimentally infected with Leishmania infantum. Eur J Pharm Sci. 2018;121:281–6.

    Article  CAS  Google Scholar 

  40. Hooper LV. Bacterial contributions to mammalian gut development. Trends Microbiol. 2013;12:129–34.

    Article  CAS  Google Scholar 

  41. Coelho LP, Kultima JR, Costea PI, Fournier C, Pan Y, Czarnecki-Maulden G, et al. Similarity of the dog and human gut microbiomes in gene content and response to diet. Microbiome. 2018.

    Article  Google Scholar 

  42. Pilla R, Suchodolski JS. The role of canine gut microbiome and metabolome in health and gastrointestinal disease. Front Vet Sci. 2020;6:498.

    Article  Google Scholar 

  43. Wernimont SM, Radosevich J, Jackson MI, Ephraim E, Badri DV, MacLeay JM, et al. The effects of nutrition on the gastrointestinal microbiome of cats and dogs: impact on health and disease. Front Microbiol. 2020;11:1266.

    Article  Google Scholar 

  44. Parker KD, Albeke SE, Gigley JP, Goldstein AM, Ward NL. Microbiome composition in both wild-type and disease model mice is heavily influenced by mouse facility. Front Microbiol. 2018;9:1598.

    Article  Google Scholar 

  45. Wang J, Lang T, Shen J, Dai J, Tian L, Wang X. Core gut bacteria analysis of healthy mice. Front Microbiol. 2019;10:887.

    Article  Google Scholar 

  46. Chung YW, Gwak H-J, Moon S, Rho M, Ryu J-H. Functional dynamics of bacterial species in the mouse gut microbiome revealed by metagenomic and metatranscriptomic analyses. PLoS ONE. 2020;15:e0227886.

    Article  CAS  Google Scholar 

  47. Martínez I, Wallace G, Zhang C, Legge R, Benson AK, Carr TP, et al. Diet-induced metabolic improvements in a hamster model of hypercholesterolemia are strongly linked to alterations of the gut microbiota. Appl Environ Microbiol. 2009;75:4175–84.

    Article  CAS  Google Scholar 

  48. Kim H, Kim D-H, Seo K-H, Chon Y-W, Nah S-Y, Bartley GE, et al. Modulation of the intestinal microbiota is associated with lower plasma cholesterol and weight gain in hamsters fed Chardonnay grape seed flour. J Agric Food Chem. 2015;63:1460–7.

    Article  CAS  Google Scholar 

  49. Li X, Zhang Z, Cheng J, Diao C, Yan Y, Liu D, et al. Dietary supplementation of soybean-derived sterols regulates cholesterol metabolism and intestinal microbiota in hamsters. J Funct Foods. 2019;59:242–50.

    Article  CAS  Google Scholar 

  50. You I, Kim MJ. Comparison of gut microbiota of 96 healthy dogs by individual traits: breed, age, and body condition score. Animals. 2021;11:2432.

    Article  Google Scholar 

  51. Suchodolski JS. Analysis of the gut microbiome in dogs and cats. Vet Clin Pathol. 2022;50:6–17.

    Article  Google Scholar 

  52. Vacca M, Celano G, Calabrese FM, Portincasa P, Gobbetti M, De Angelis M. The controversial role of human gut Lachnospiraceae. Microorganisms. 2020;8:573.

    Article  CAS  Google Scholar 

  53. Raethong N, Nakphaichit M, Suratannon N, Sathitkowitchai W, Weerapakorn W, Keawsompong S, et al. Analysis of human gut microbiome: taxonomy and metabolic functions in Thai adults. Genes. 2021;12:331.

    Article  CAS  Google Scholar 

  54. Hugenholtz F, de Vos WM. Mouse models for human intestinal microbiota research: a critical evaluation. Cell Mol Life Sci. 2018;75:149–60.

    Article  CAS  Google Scholar 

  55. Lappan R, Classon C, Kumar S, Singh OP, de Almeida RV, Chakravarty J, et al. Meta-taxonomic analysis of prokaryotic and eukaryotic gut flora in stool samples from visceral leishmaniasis cases and endemic controls in Bihar State India. PLoS Negl Trop Dis. 2019;13:e0007444.

    Article  CAS  Google Scholar 

  56. Kalinen S, Kallonen T, Gunell M, Ettala O, Jambor I, Knaapila J, et al. Gut microbiota signatures associate with prostate cancer risk. MedRxiv. 2021.

    Article  Google Scholar 

  57. Matsumoto N, Park J, Tomizawa R, Kawashima H, Hosomi K, Mizuguchi K, et al. Relationship between nutrient intake and human gut microbiota in monozygotic twins. Medicina. 2021;57:275.

    Article  Google Scholar 

  58. Tims S, Derom C, Jonkers DM, Vlietinck R, Saris WH, Kleerebezem M, et al. Microbiota conservation and BMI signatures in adult monozygotic twins. ISME J. 2013;7:707–17.

    Article  CAS  Google Scholar 

  59. Mukherjee A, Lordan C, Ross RP, Cotter PD. Gut microbes from the phylogenetically diverse genus Eubacterium and their various contributions to gut health. Gut Microbes. 2020;12:1802866.

    Article  CAS  Google Scholar 

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We are very grateful to Montserrat Grau DVM, the veterinarian responsible for the animal housing, Instituto de Investigación Hospital 12 de Octubre, Madrid, whose commitment, technical skills and assistance have made the animal experiment possible. We also much appreciate the collaboration of the animal caretakers. We would like to thank Sara Álvarez (Unidad de Genómica—UCM, Madrid) for her expert work on DNA extractions.


The research was partially funded by project PR87/19-22646 (Proyectos Santander/Complutense).

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Authors and Affiliations



Conceptualization: AIOM, JJT, JMA. Funding acquisition: JJT, JMA. Investigation: AIOM, MC, ST, EB, PB, JGC, JJT, JMA. Methodology: PB, JGC, AIOM, JMA. Software: PB, JGC. JJT and JMA coordinated the research. All authors read and approved the final manuscript.

Corresponding author

Correspondence to José María Alunda.

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The animal experiments were approved by the Committee of Animal Experimentation (Universidad Complutense de Madrid, UCM) and the regional authorities (Madrid Region) (PROEX 169/15). The 3R principles were followed and all procedures involving experimental animals were performed by qualified personnel supervised by a veterinary doctor. All efforts were made to minimize suffering.

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Supplementary Information

Additional file 1: Table S1.

Identification and relative quantification of phyla, families and genera detected in individual microbiomes of G1-16w samples.

Additional file 2: Table S2.

Changes in microbiota composition between G1 and G3 groups, expressed as log2 ratio, at the level of phylum, family and genus. The FDR P-value is indicated. 16w: 16 weeks post inoculation; 18w: 18 weeks post inoculation; EP: end point.

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Olías-Molero, A.I., Botías, P., Cuquerella, M. et al. Leishmania infantum infection does not affect the main composition of the intestinal microbiome of the Syrian hamster. Parasites Vectors 15, 468 (2022).

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