4 CHAPTER FOUR: SUMMARY AND CONCLUSIONS
4.1 Summary
The objectives of this study were to describe the cutaneous bacterial microbiota of a population of healthy horses at a farm in Southern Ontario and to evaluate the effect of body site, individual and season on taxonomic distributions as well as alpha and beta diversity. Interestingly, contrary to what has been described in humans1,2 and dogs3,4, we found no significant difference in species diversity, richness or evenness between body sites or individual. In contrast, all alpha diversity parameters trended higher in the Winter and Summer than the Spring and Fall. The reason for these findings is unknown but the author’s hypothesis is that the ground in the paddocks was wetter in the Spring and Fall and may have led to overgrowth of species favouring these conditions. When evaluating community membership and structure (beta diversity) between samples, both skin site and season affected microbial populations, but samples clustered more based on season than site. Thus, our study suggests that for horses living in the same
environment, time of year is the main force driving the skin microbiota composition rather than skin site or individual. This is a significant finding as it highlights the importance of taking seasonality into account when formulating future studies. In this study, I did not specifically look at the effect of haircoat length or blanketing on the cutaneous microbiota but given that bacterial populations in the warmer months (spring and summer) did not cluster together but separate to the colder months (fall and winter), these variables did not appear to play a role.
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In an attempt to further understand disease pathogenesis as well as explore novel therapeutic avenues, many cutaneous microbiota studies within the field of veterinary medicine focus on comparing healthy versus diseased individuals.3,5-8 This study underscores the need for more descriptive studies such as this to identify intrinsic and extrinsic factors affecting natural variations of healthy individuals to truly characterize the cutaneous microbiota prior to being able to compare ‘normal’ versus ‘diseased’. A failure to identify these factors can lead to erroneous conclusions. Studies in
humans,9,10 cats5 and dogs3 have identified a decrease in microbial diversity in inflammatory skin conditions such as atopic dermatitis, when compared to healthy individuals. Our longitudinal study of healthy horses suggests that for any future similar studies evaluating atopic versus healthy horses, sampling date must be homogenous across groups.
Despite the insights provided by this research, it was not without limitations. Ten OTUs were present in all samples regardless of horse, month or site sampled (Psychrobacter, Pseudomonas, Macrococcus, Acinetobacter, Carnobacterium, and Desemzia as well as two OTUs each from the genera Planomicrobium and Arthrobacter). These may
represent core OTUs present in all healthy horses that are stable over time and thus a marker of health. Unfortunately, these same OTUs were also identified in the null
exposure samples and thus could also represent environmental contaminants. The low abundance of microbial DNA present at sites like skin poses a major challenge for studies such as this.11-13 Care must be taken when interpreting results, especially those that did not take any steps to measure or control for contaminant DNA, as it could
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influence results more prominently than in studies where samples contain abundant microbial DNA such as fecal samples. Here, we evaluated sampling and processing contaminants by running negative controls during DNA extraction, amplification and purification. We also processed null exposure swabs (which were obtained by waving the swabs in the air on the farm) in the same manner as skin swabs. From these null exposure swabs, 733 genera were identified. Subtracting lineages present in negative control samples has been described,14 but cannot be recommended in skin samples when evaluating null exposure swabs, as OTUs present in the environment may also be genuine members of the resident microbiota.15 The best way to analyze and control for those contaminants is unknown and is an important area of study to ensure that useful and unbiased data are obtained. Future studies comparing cutaneous to null exposure samples from different farms as well as studies comparing different sampling protocols, such as the inclusion of an ‘environmental decontamination’ step may be helpful to further distinguish between resident, transient and contaminant bacterial DNA.
An additional limitation of this study includes the fact that all horses in this study were sampled from the same farm. This was done intentionally to reduce confounding variables. Whether the findings from this study can be extrapolated to the equine species as a whole, is questionable. Furthermore, the number of anatomical locations sampled in this study was low at only four sites. It is possible that the lack of significant findings in alpha diversity parameters between body sites was due to the low number of sites sampled and the fact that these sites were too similar. Lastly, this study only evaluated one timepoint per season. Sampling the same seasons over multiple years,
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as well as more timepoints within a year would be needed to make any definitive conclusions of seasonal effect on the cutaneous microbiota. Additional limitations, include those that are inherent to 16S rRNA gene sequencing, including a lack of
standardization of protocols, the fact that it cannot distinguish between bacteria that are alive or dead,16 and the fact that it is only able to characterize sequences to the genus level.
If this study were to be repeated, an increase in the number of anatomical sites sampled, including samples from mucocutaneous junctions as well as the ear canal would be warranted to see if alpha diversity is decreased in these areas, similarly to dogs and cats.3-5 In addition, including horses from different farms to see if horses from the same farm are more similar to one another than horses from a different farm would be of value. This would also help identify whether the same core OTUs that were
identified in this cohort of horses are present in horses from different environments, and thus potentially a true marker of health. To further investigate the influence of season, sampling horses at multiple timepoints within the same season and sample individuals across several years would be recommended. By increasing anatomical sites, farms and timepoints sampled within a year as well as over several years, a more complete picture of the healthy cutaneous microbiota of healthy horses could be obtained.
106 4.2 Conclusions
Many studies in humans and companion animals have found associations between cutaneous dysbiosis and inflammatory skin diseases. The prevalence of allergic skin disease in horses is reported to be similar to that in dogs.17 Descriptive studies such as this are critical as it is important to identify non-disease influencing factors first, if we want to further understand disease. This information can then be used to understand how deviations from ‘normal’ may be causing or contributing to disease status. Failure to consider these could lead to erroneous results or interpretations. Considering the findings of this study, it appears that any comparison of horses with skin disease should have null samples as a negative control, and other horses in the same environment as control animals.
This study revealed that the cutaneous microbiota of healthy horses is more diverse than previously described by culture methods,18,19 that time of year and skin site play a role in bacterial species found on equine skin but that the former appears to be a more important driving factor. It also revealed that there may be a core set of bacteria that are present across all sites and individuals that is stable over time. Lastly, this study
highlights the need for more depth of investigation into the effect of design methods and environmental influences of healthy animals to truly characterize the cutaneous
microbiota. Our hope is that this topographical and temporal survey may serve as a baseline for future studies investigating the effect of the cutaneous microbiota of horses in disease pathogenesis.
107 4.3 References
1. Grice EA, Kong HH, Conlan S et al. Topographical and temporal diversity of the human skin microbiome. Science 2009; 324: 1,190–1,192
2. Costello EK, Lauber CL, Hamady M et al. Bacterial community variation in human body habitats across space and time. Science 2009; 326: 1694-1697
3. Rodrigues Hoffmann A, Patterson AP, Diesel A, Lawhon SD, Ly HJ, Elkins Stephenson C, Mansell J, Steiner JM, Dowd SE, Olivry T, Suchodolski JS. The skin microbiome in healthy and allergic dogs. PLoS One 2014; 9: e83197.
4. Cuscó A, Sanchez A, Altet L, Ferrer L, Francino O. Individual signatures define canine skin microbiota composition and variability. Front Vet Sci 2017; 4: 6.
5. Older CE, Diesel A, Patterson AP et al. The feline skin microbiota: The bacteria inhabiting the skin of healthy and allergic cats. PLoS One 2017; 12:e0178555.
6. Korbelik J, Singh A, Rousseau J et al. Characterization of the otic bacterial microbiota in dogs with otitis externa compared to healthy individuals. Vet Dermatol 2019; 30: 228-e70.
7. Ngo J, Taminiau B, Fall PA et al. Ear canal microbiota – a comparison between healthy dogs and atopic dogs without clinical signs of otitis externa. Vet Dermatol 2018; 29:425-e140
8. Kobayashi T, Glatz M, Horiuchi K et al. Dysbiosis and Staphylococcus aureus colonization drives inflammation in atopic dermatitis. Immunity 2015; 42: 756–766.
9. Paller AS, Kong HH, Seed P et al. The microbiome in patients with atopic dermatitis. J Allergy Clin Immunol 2018; 143: 26-35
10. Kong HH, Oh J, Deming C et al. Temporal shifts in the skin microbiome associated with disease flares and treatment in children with atopic dermatitis. Genome Res 2012; 22: 850–859.
11. Grogan MD, Bartow-McKenney C, Flowers L, Knight SAB, Uberoi A, Grice EA. Research Techniques made simple: profiling the skin microbiota. J of Invest Dermatol 2019; 139: 747-752 12. Goodrich JK, Di Rienzi SC, Poole AC, Koren O, Walters WA, Caporaso JG, Knight R, Ley RE.
Conducting a Microbiome Study. Cell 2014; 158 (2): 250-262
13. Salter SJ, Cox MJ, Turek EM, Calus ST, Cookson WO, Moffatt MF, Turner P, Parkhill J, Loman NJ, Walker AW. Reagent and laboratory contamination can critically impact sequence-based microbiome analyses. BMC Biology 2014; 12: 87
14. Jervis-Bardy J, Leong LEX, Marri S, et al. Deriving accurate microbiota profiles from human samples with low bacterial content through post-sequencing processing of Illumina MiSeq data.
Microbiome 2015; 3:19 doi 10.1186/s40168-015-0083-8
15. Kim D, Hofstaedter CE, Zhao C, Mattei L, Tanes C, Clarke E, Lauder A, Sherrill-Mix S, Chehoud C, Kelsen J, Conrad M, Collman RG, Baldassano R, Bushman FD, Bittinger K. Optimizing methods and dodging pitfalls in microbiome research. Microbiome. 2017; 5: 52
16. Rodrigues Hoffmann A. The cutaneous ecosystem: the roles of the skin microbiome in health and its association with inflammatory skin conditions in humans and animals. Vet Dermatol 2017; 28:
60-e15.
17. Marsella R, De Benedetto A. Atopic dermatitis in animals and people: an update and comparative review. Vet Sci 2017; 4(3): 37
18. Scott DW, Miller WH. Equine Dermatology, 2nd edition. Maryland Heights, MO: Saunders Elsevier, 2011; p 17
19. Adams MK, Hendrickson DA, Rao S et al. The bacteria isolated from the skin of 20 horses at a Veterinary Teaching Hospital. J of Eq Vet Sci. 2010; 30: 687-695
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