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. 2023 Oct 30;89(11):e00977-23. doi: 10.1128/aem.00977-23

Tracking the fecal mycobiome through the lifespan of production pigs and a comparison to the feral pig

Tausha L Prisnee 1, Rajibur Rahman 1, Janelle M Fouhse 1, Andrew G Van Kessel 2, Ryan K Brook 3, Benjamin P Willing 1,✉
Editor: Irina S Druzhinina4
PMCID: PMC10686082  PMID: 37902410

ABSTRACT

Fungi in the pig gastrointestinal tract (the mycobiome) are believed to be important for host health and development. However, studies profiling the mycobiome over a production cycle and of the feral pig are lacking. The objectives of this study were to profile the pig mycobiome over one production cycle and profile the mycobiome of feral pigs. A total of 24 pigs from 12 litters (n = 2 per litter) had fecal swabs taken at 11 days of age (D11), the day before weaning (W − 1), 7 days post-weaning (W + 7), and 119 days post-weaning (W + 119) in a research facility. Their dams (n = 12) and an additional eight sows from a commercial barn had fecal swabs taken 3 days after farrowing. A total of 20 feral pigs had intestinal contents collected from the ileum and cecum for a total of 24 samples. Samples were analyzed via amplicon sequencing of the internal transcribed spacer 2 rRNA gene. Piglets tended to cluster based on their mother’s Kazachstania slooffiae levels on D11 (P = 0.087) and significantly clustered on W + 119 (P = 0.046). Piglets clustered with their littermate on D11 (P = 0.006) and on W + 119 (P = 0.007). In summary, we found that pigs maintain the same K. slooffiae status as their mothers until the end of the production cycle and show that the feral pig has a complex and unique mycobiome largely represented by ingested fungal species.

IMPORTANCE

This work provides evidence that early-life fungal community composition, or host genetics, influences long-term mycobiome composition. In addition, this work provides the first comparison of the feral pig mycobiome to the mycobiome of intensively raised pigs.

KEYWORDS: Kazachstania slooffiae, mycobiome, wild boar, feral pig, porcine

INTRODUCTION

The gastrointestinal tract (GIT) is home to a vast array of microbes including bacteria, archaea, viruses, protozoa, and fungi (1). While fungi make up less than 0.1% of total microbial reads in the human GIT (1), fungi contribute to host health and development and are immunomodulatory (2, 3). The collection of fungi in the GIT is termed the mycobiome and although it has gained increased attention in recent years is largely understudied.

In both humans and pigs, yeast is the most common fungi in the GIT (4, 5). The mycobiome has been well-profiled in pigs up to 35 days of age (5 – 7). However, studies tracking the fungal community of a pig through the production cycle have not been completed. We have previously found that the maternal mycobiome drives piglet mycobiome assembly (8). Specifically, we found that differences in fungal community structure between litters were driven by sow Kazachstania slooffiae abundance (8). K. slooffiae is a commensal yeast found in pigs across geographic locations and under different types of production systems (9 – 12). Previous studies have found that K. slooffiae is most abundant in post-weaning pigs (5, 7, 13), although we have previously noted the presence of K. slooffiae in pigs as young as 3 days of age (8). K. slooffiae may contribute to pig health by altering intestinal bacteria, increasing short-chain fatty acids in feces, and acting as a source of amino acids (especially lysine) and vitamin C (14, 15).

There is also a recent growing interest in “wild” microbiomes as a result of their potential to protect against disease. The feral pig therefore represents an interesting comparison for the identification of organisms that may have been lost as a result of high-level sanitation in conventional pig production. Domestic wild boar were originally imported to Canada in the 1980s for livestock diversification where most animals were cross-bred with domestic pigs and in some cases with mini-pigs and pot-bellied pigs (16). Since then, they have escaped or have been released from the farms and these feral pigs are now free-ranging across over one million km2 of Canada (17, 18). Feral pigs now roam the Canadian prairies where they wreak havoc on cropland and pose a very concerning disease risk to domestic pigs (19 – 21). While sanitation practices help to prevent disease among pigs, there may also be alterations in the bacterial and fungal communities. Indeed, one study has shown that exposure to soil can accelerate the maturation of the bacterial community (22). It is not currently known how domestication affects the pig mycobiome.

Drawing on previous research, we can assume that K. slooffiae provides at least some benefit to the pig. While K. slooffiae is an important member of the mycobiome, there are other fungi that may also contribute to host health. However, it is unclear how the mycobiome changes over a production cycle, nor is it clear how the domestic pig may differ when compared to feral pigs in terms of the mycobiome. In this study, we seek to understand these questions by (i) tracking the pig mycobiome through the lifespan (production cycle), including the sows; (ii) profiling the mycobiome of feral pigs; and (iii) profiling the mycobiome of sows raised in a commercial facility.

MATERIALS AND METHODS

Animal use and care

This animal study was approved by the Animal Care and Use Committee of the University of Alberta (Edmonton, AB, CAN) and conducted in accordance with the guidelines of the Canadian Council on Animal Care under AUP00002214. Feral pig capture and handling protocols were approved by the University of Saskatchewan Animal Research Ethics Board under AUP20150024 and the Saskatchewan Ministry of Environment under permit 17FW027. A total of 36 pigs were used from the Swine Research and Technology Center (SRTC) at the University of Alberta (Edmonton, AB, CAN). The pigs at SRTC represent pigs raised under experimental farm conditions, although they were raised with commercial production practices. Sows (n = 12; Large White/Landrace) had fecal samples taken 3 days after farrowing. Piglets (n = 24; Large White/Landrace x Duroc) had fecal swabs taken at 11 days of age (D11), 1 day prior to weaning (W − 1), 7 days post-weaning (W + 7), and 119 days post-weaning (W + 119). W + 119 represents a pig at market weight and the end of a production cycle. As we previously documented the pig mycobiome up to 8 days of age (8), we opted to focus on time points after 8 days of age. Two piglets, whose weights were close to the median litter weight, were selected per sow to follow through the trial. Piglets were housed with their mother and littermates until 21 days of age when weaning occurred. Following weaning, piglets were housed with only their litter mates until 4 weeks following weaning when they were housed in groups separate from any of their litter mates. Five pigs in the study received antibiotics (penicillin) prior to D11 due to an outbreak of scours in the farrowing room. Piglets were introduced to creep feed at 14 days of age. Fecal samples were taken from additional sows (n = 8; Large White) from a commercial farm 3 days after farrowing for mycobiome analysis. Feral pigs were captured from two locations: Moose Mountain (approximately 49.8338° N, 102.2922° W) and Melfort (approximately 52.8608° N, 104.6143° W), Saskatchewan, Canada. Pigs were located via helicopter and captured with a net gun. Following capture, the pigs were euthanized via a captive bolt, and intestinal contents were collected. Ileum content was collected 5 cm from the ileocecal junction and cecal content was collected from the tip of the cecum. A total of 12 ileum samples and 12 cecal samples were selected for mycobiome analysis from 20 pigs (4 of the pigs had samples available from the ileum and cecum for sequencing).

DNA extraction

Total genomic DNA was extracted using the DNeasy PowerSoil Pro kit (Qiagen, CA, USA) with no modifications. Bead beating was performed on a FastPrep-24 (MP Biomedicals, OH, USA) homogenizer at 5 m/s for 45 seconds.

Fungal sequencing

Internal transcribed spacer (ITS) 2 sequencing was performed at Microbiome Insights (Richmond, BC, CAN). Sequencing was done using the following primers: forward (ITSF) 5′-CCTCCGCTTATTGATATGC-3′ and reverse (ITSR) 5′-CCGTGARTCATCGAATCTTTG-3 (23). Paired-end sequencing was done on the Illumina MiSeq platform (Illumina, CA, USA) using 2 × 300 cycles and PCR parameters have been previously described (24).

Sequencing analysis was performed using Quantitative Insight into Microbial Ecology (QIIME) 2 (v2022.11) (25). Only forward reads were used and reads were truncated at 240 base pairs. The Divisive Amplicon Denoising Algorithm (v2) plugin was used to perform demultiplexing, quality filtering, denoising, and filtering out chimeras (26). Mafft was used to align Amplicon Sequence Variants (ASVs) (27). Taxonomy was assigned to ASVs using the classify-sklearn naïve Bayes taxonomic classifier (via the q2-feature-classifier plugin) (28), and the UNITE database (v8.4) was used (29).

Statistical analysis

K. slooffiae level in sows was broken into low, medium, and high by assigning levels at natural breaks, with low being ≤10% total reads (n = 7), medium being the lone sow at 35%, and high being ≥70% (n = 4). The random sow was found using the randomize function in Excel. Differences in the percentage of K. slooffiae and the distance to the maternal sow versus a random sow were calculated using a Mann-Whitney U test in GraphPad Prism 9.5.1 based on Bray-Curtis distance. Phyloseq (v1.34.0) was used in R to analyze microbial community structure and diversity (30). Differences in fungal community composition were measured using Bray-Curtis dissimilarity and Permutational Multivariate Analysis of Variance (PERMANOVA) and were visualized using principal coordinates analysis (PCoA) (R, v4.0.5). Homogeneity of dispersion was measured using the Betadisper function in phyloseq (30). α-diversity was calculated in QIIME (v2022.11) and was statistically analyzed using Kruskal-Wallis in GraphPad Prism 9.5.1 with Dunn’s multiple comparisons. Prior to α-diversity analysis, reads were rarefied to 1,700 reads. Differential abundance was calculated using Analysis of Composition of Microbiomes (ANCOM) in QIIME (v2022.11) and was compared using a Kruskal-Wallis test with Dunn’s multiple comparisons in GraphPad Prism 9.5.1. Differences in average daily gain (ADG) were compared using a Kruskal-Wallis test with Dunn’s multiple comparisons in GraphPad Prism 9.5.1. The correlation between pig K. slooffiae and weight was computed using Spearman’s correlation in GraphPad Prism 9.5.1.

RESULTS

Sows are colonized with varying levels of K. slooffiae

Levels of K. slooffiae varied among sows. All but one of the sows fell into one of two categories: ≤10% K. slooffiae (n = 7) or ≥70% K. slooffiae (n = 4). One sow had 35% K. slooffiae.

Sow K. slooffiae colonization level shapes piglet mycobiome during early and later life, but not around the weaning transition

On D11, piglets tended to cluster depending on whether the sows were colonized with high, medium, or low levels of K. slooffiae (Fig. 1a, P = 0.087, β-dispersion P = 0.181). By W − 1, piglets no longer clustered based on their sow’s K. slooffiae status (Fig. 1b, P = 0.297, β-dispersion P = 0.726). On W + 7, sow K. slooffiae status also made no difference to the mycobiome (Fig. 1c, P = 0.859, β-dispersion P = 0.509). However, by W + 119, sow K. slooffiae status influenced pig mycobiome composition (Fig. 1d, P = 0.046, β-dispersion P = 0.278). There was an effect of day on the percentage of K. slooffiae present in the mycobiome (P < 0.001). On day 11, piglets had higher mean K. slooffiae than on W + 7 (Fig. 2, P < 0.001) and sows had more K. slooffiae than piglets on W + 7 (Fig. 2, P = 0.013).

Fig 1.

Fig 1

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β-diversity based on Bray-Curtis dissimilarity of the mycobiome based on sow Kazachstania slooffiae level. (a) Piglets 11 days of age (D11) showed a trend in clustering by sows K. slooffiae status (P = 0.087, β-dispersion P = 0.181). (b) Piglets on the day before weaning (W − 1) did not cluster by sows K. slooffiae status (P = 0.297, β-dispersion P = 0.726). (c) Pigs 7 days following weaning (W + 7) did not cluster by sow K. slooffiae status (P = 0.859, β-dispersion P = 0.509). (d) At 119 days following weaning, that is, the end of a production cycle, pigs clustered by their mother’s K. slooffiae status (P = 0.046, β-dispersion P = 0.278). Trends were defined as P < 0.1, and significance was defined as P ≤ 0.05.

Fig 2.

Fig 2

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Percentage of K. slooffiae at different time points across a production cycle presented as the mean at each time point ± SEM. There was a significant impact of time on the percentage of K. slooffiae (P < 0.001). * indicates P ≤ 0.05 and ** indicates P < 0.001. Significance was defined as P ≤ 0.05.

The amount of K. slooffiae a sow was colonized with did not correlate with the weight of the piglets on D11 (P = 0.903, r = −0.0263), W − 1 (P = 0.720, r = −0.0852), W + 7 (P = 0.783, r = −0.0606), or W + 119 (P = 0.945, r = −0.0148). The amount of K. slooffiae a pig was colonized with did not associate with pig weight on D11 (P = 0.703, r = −0.0821), W − 1 (P = 0.163, r = −0.3149), W + 7 (P = 0.140, r =−0.3171), or W + 119 (P = 0.574, r = −0.121).

Piglets cluster by litter except on the day before weaning

Piglets clustered by litter on D11 (Fig. 3a, P = 0.006). Around the weaning transition (W − 1), piglets are no longer clustered by litter (Fig. 3b, P = 0.184). By W + 7, piglets tended to cluster by litter (Fig. 3c, P = 0.087). On W + 119, piglets clustered by litter (Fig. 3d, P = 0.007). On D11 piglets were closer to their maternal sow than a random sow (Fig. 4a, P < 0.001), however, on W − 1, W + 7, and W + 119, there was no difference between the distance to the maternal sow and a random sow (Fig. 4b, P = 0.418; Fig. 4c, P = 0.116; Fig. 4d, P = 0.511).

Fig 3.

Fig 3

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β-diversity of pigs by sow based on Bray-Curtis dissimilarity. (a) Piglets at 11 days of age (D11) clustered with their littermate (P = 0.006, β-dispersion P < 0.001). (b) Pigs on the day prior to weaning (W − 1) no longer clustered by sow ID (P = 0.184, β-dispersion P = 1.0). (c) Pigs at 7 days following weaning (W + 7) tended to cluster by sow ID (P = 0.087, β-dispersion P < 0.001). Pigs 119 days following weaning (W + 119) clustered by sow ID (P = 0.007, β-dispersion P < 0.001).

Fig 4.

Fig 4

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Distance of piglets to their maternal sow versus the distance to a randomly selected sow. (a) On D11, piglets were closer to their maternal sow than they were to their randomly selected sow (P < 0.001). (b) On the day before weaning, piglets were no closer to their maternal sow than to a randomly selected sow (P = 0.418). (c) 7 days after weaning, pigs were no closer to their maternal sow than to a randomly selected sow (P = 0.116). (d) At 119 days following weaning, pigs were no closer to their maternal sow than to a randomly selected sow (P = 0.511). Data are presented as mean ± SEM. Significance was defined as P ≤ 0.05. * indicates P < 0.001.

Both α-diversity and β-diversity were altered by pig age

Mycobiomes clustered based on time point (Fig. 5a, P = 0.001, β-dispersion P = 0.253); however, there was no clear distinction based on younger or older pigs, with no clear break at weaning. α-diversity decreased with time (Fig. 5b, P < 0.001). For the five pigs that received antibiotics, no apparent effect on mycobiome β-diversity at any timepoint was observed (P = 0.359).

Fig 5.

Fig 5

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(a) β-diversity of pigs at all time points as measured by Bray-Curtis dissimilarity. Pigs clustered based on time point (P = 0.001, β-dispersion P = 0.253). Pairwise comparisons between different time points are shown. (b) α-diversity of pigs at different timepoints as measured by the Shannon diversity index (P < 0.001). * indicates P ≤ 0.05 and ** indicates P < 0.001. Box and whisker plot shows the mean with a 95% confidence interval. Significance was defined as P ≤ 0.05.

K. slooffiae is more prevalent in intensively raised pigs than in feral pigs

K. slooffiae was among the 10 most dominant fungi at all time points in the experimental piglets and in experimental sows (Fig. 6a through d and 7a). In commercial sows, K. slooffiae was the most dominant fungi (Fig. 7b). While K. slooffiae was present in all feral pigs in both gut sections, it was not among the 10 most dominant fungi (Fig. 7c and d). The mycobiome of feral pigs was more complex than that of both experimental and domestic pigs. Following singleton removal, there were a total of 224 genera in experimental pigs, 123 genera in commercial pigs, and 280 genera in feral pigs. In addition, feral pigs clustered separately from commercial and experimental pigs (Fig. 8c, P = 0.002, β-dispersion P = 0.01). ANCOM revealed several differential genera between experimental, commercial, and feral pigs: Kazachstania, Saccharomyces, Aspergillus, Monilia, Kalmanozyma, Xeromyces, Naganishia, Hyphopichia, and Diutina. Kazachstania and Saccharomyces were more abundant in both commercial and experimental pigs than feral pigs (Fig. 8a, P < 0.01) and there was no difference between experimental and commercial pigs (Fig. 8a, P = 0.90). Aspergillus was more abundant in commercial pigs than in feral pigs (Fig. 8b, P < 0.001) and not in commercial versus experimental or in experimental versus feral (Fig. 8b, P = 0.052 and P = 0.08). Monilia was only present in commercial pigs and was therefore significantly higher in this group than in experimental or feral pigs (Fig. 8b, P < 0.001). Kalmanozyma and Xeromyces were only found in commercial and experimental pigs and not in feral pigs and were therefore more abundant in these groups (Fig. 8b, P < 0.001). Naganishia was more abundant in commercial pigs than in feral pigs and in commercial versus experimental pigs (Fig. 8b, P < 0.01) but not in experimental versus feral pigs (Fig. 8b, P = 0.327). Hyphopichia was more abundant in commercial pigs than in experimental or feral pigs (Fig. 8b, P < 0.001) but not in experimental versus feral (Fig. 8b, P > 0.999). Finally, Diutina was more abundant in commercial pigs than in experimental or feral pigs (Fig. 8b, P < 0.001 and P = 0.01) but not in experimental versus feral pigs (Fig. 8b, P = 0.222).

Fig 6.

Fig 6

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(a) Top 10 genera in experimental pigs at 11 days of age. (b) Top 10 genera on the day prior to weaning. (c) Top 10 genera at 7 days post-weaning. (d) Top 10 genera at 119 days post-weaning.

Fig 7.

Fig 7

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(a) Top 10 most abundant genera in experimental sows. (b) Top 10 most abundant genera in commercial sows. (c) Top 10 most abundant taxa in the ileum feral pigs. All taxa are genera unless otherwise noted. The prefix f_ indicates a fungal family. Feral pigs are from two different locations—Melfort and Moose Mountain Saskatchewan Canada. (d) Top 10 most abundant taxa in the cecum of feral pigs. All taxa are genera unless otherwise noted. The prefix f_ indicates a fungal family and the prefix o_ indicates a fungal order.

Fig 8.

Fig 8

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(a) Kazachstania and Saccharomyces as identified by ANCOM as being differential. (b) All other differential taxa as identified by ANCOM. (c) β-diversity based on Bray-Curtis dissimilarity of the mycobiome of commercial, experimental, and feral pigs. (d) β-diversity based on Bray-Curtis dissimilarity of the mycobiome of feral pigs by location and intestinal section. Significance was defined as P ≤ 0.05.

The feral pig mycobiome differed based on the location of the pig

Feral pigs clustered based on their geographic location (Fig. 8d, P = 0.047, β-dispersion P = 0.059) but not based on the ileum versus the cecum (Fig. 8d, P = 0.114, β-dispersion P = 0.379). ANCOM analysis revealed the genus Gibellulopsis as the only differential taxa between the two locations.

DISCUSSION

This study brings new advances to our understanding of the pig mycobiome throughout life. We found that early-life fungal community composition coincided with fungal composition later in life, a finding that may have consequences for production performance and disease risk. In addition, we showed that the feral pig mycobiome is more complex than the mycobiome of intensively raised pigs and that the feral pig mycobiome contains the key taxa K. slooffiae, albeit at a lower level. This provides further evidence that K. slooffiae is a core microbe of the pig.

We found that pigs had differences in their mycobiomes based on their age. This trend is similar to what has previously been described for bacterial populations (5, 7). However, unlike the aforementioned studies, which saw a clear break in pig mycobiomes surrounding the weaning transition, this break was not present in our study; likely due to K. slooffiae colonization in pre-weaning piglets. In the present study, K. slooffiae made up a larger percentage of the young pig mycobiome, converse to studies indicating that it becomes detectable in large numbers after weaning (5 – 7, 12, 13).

The K. slooffiae levels of the maternal sow drove piglet K. slooffiae levels early and late in life but did not impact the level of K. slooffiae around the weaning transition. Indeed, W + 7 was the time point with the lowest mean K. slooffiae levels suggesting perturbation of the mycobiome around the weaning transition. In addition, piglets clustered by litter and were, therefore, more similar to their littermate on D11 and W + 119 with disruption right before and after weaning. It remains unclear as to whether these findings are due to early life exposure or due to other factors such as genetic susceptibility to fungal colonization. Early-life bacterial exposure has been shown to impact the bacteriome later in life (31), and therefore there may also be an impact of early-life fungal exposure. This is particularly important because in humans composition of the mycobiome has been shown to contribute to several different diseases including inflammatory bowel disease, multiple sclerosis, colon cancer, and asthma (32 – 35). While it is not clear how the mycobiome affects pig health and performance long term, it is possible that there is some impact, as the bacterial community has been previously implicated in pig performance (36, 37). It also remains unclear why some pigs have large amounts of K. slooffiae and others do not. The overgrowth of other fungi such as Candida has been shown to have a genetic component (38) as well as a host metabolome component (39). Therefore, there may be a number of host factors that may help determine the level to which K. slooffiae can colonize and that dissimilarity between mycobiomes may have a genetic component. If fungal exposure during early life is the driving force behind piglet mycobiome development, it may be possible to develop supplementation strategies to alter long-term mycobiome composition, especially with respect to K. slooffiae abundance. If genetics are the driving force behind the fungal composition, it may be possible to promote favorable mycobiomes through genetic selection. However, it remains unclear what the composition of a favorable mycobiome is. To determine whether piglet exposure or genetics is the driving factor in piglet mycobiome development, a cross-fostering study should be performed.

We have previously shown that piglets have mycobiomes that are more similar to their maternal sows than to that of other sows at 8 days of age (8). In the present study, we show that this result is consistent, with piglets having mycobiomes that are more similar to their maternal sow than to that of a random sow at 11 days of age. However, by the day prior to weaning, they no longer have mycobiomes that are closer to their maternal sow than to a random sow, despite still being housed in a pen with their mother. This is contrary to what is seen in humans, where the infant fecal mycobiome is no more similar to their own mother’s mycobiome than that of a randomly selected mother (40). The reason for this difference likely comes down to the environment in which piglets live, where they are in contact with the maternal feces, and are therefore able to obtain microbes from their mothers through repeated exposure. However, as piglets age and begin to consume solid feed, their mycobiomes become less like their mothers, suggesting that the diet contributes to mycobiome composition, a finding that has been previously noted (5).

The weaning transition is a health-challenging time for piglets (41). During the weaning transition, piglets will transition to solid feed, encounter stress from social mixing, and develop transient intestinal malabsorption (42 – 44). We found that piglets on W − 1 and W + 7 showed the most variability in that they did not cluster with their litter mate and did not cluster by sow K. slooffiae level. Microbial dysbiosis during the weaning transition has previously been noted in bacterial communities as piglets begin to transition to a solid diet high in cereal grain (45). Given that piglets return to clustering with their litter mate and by sow K. slooffiae level later in life, we can assume that something similar is happening to the mycobiome. In our model, piglets were introduced to creep feed at 14 days of age. The amount of creep feed consumed is variable among piglets (46); therefore, differences in creep feed consumption may explain the variability in mycobiome compassion among littermates at W − 1.

Sows from commercial barns displayed a similar pattern of K. slooffiae colonization to experimental sows, with some sows having a high abundance and others having a low abundance. This suggests that in intensively raised pigs, K. slooffiae is a commonly abundant microbe, which multiple studies have previously noted (5 – 7, 12, 13, 47).

The mycobiome of feral pigs differed from that of domestic pigs. Perhaps the most glaring difference was the presence of Vishniacozyma as the most dominant genera of fungi in the ileum and the second most dominant in the cecum. While this microbe is also present in the experimental pigs and commercial sows, Kazachstania is instead the most dominant genera in intensively raised pigs. This is likely because feral pigs are in contact with more soil as they are free-roaming and feed on crops and crop stubble (48). Indeed, many members of the feral pig mycobiome are microbes regularly found on plants or in soil. This is similar to what has been described in Tibetan pigs fed a high forage diet, where the most prominent genus was Russula (49), which is often found in soil (50). While Russula was not a common finding in the feral pigs in the present study, it was present in two individuals in small amounts. Vishniacozyma may not actually colonize the intestine but instead may be a passenger that simply travels through the GIT, as is potentially the case for the Cladosporium found in another study (7, 51). Vishniacozyma victoriae, the species of Vishniacozyma found in the present study, was originally found in Antarctica, although it has also been isolated from living environments in other locations, and was found to grow between 4°C and 20°C (52). Likewise, it has been found that Vishniacozyma victoriae is incapable of growing at human body temperature (37°C) (53). It remains unclear what impact Vishniacozyma has on the pig GIT.

Not all the fungi present in the feral pig are unable to colonize. Candida was the most abundant genera in the cecum of feral pigs, and Candida species have been previously identified in the feces of pigs (12, 47, 49). Candida species are able to grow at body temperature (54) and are usually considered opportunistic pathogens (55). In healthy humans, Candida coexists with commensal bacteria, whereas Lactobacillus has an antagonistic relationship and causes the loss of pathogenicity factors (56). However, Candida is also able to overgrow and cause sepsis, typically in hosts who are immunocompromised (57). In pigs, Candida tropicalis has been shown to cause mucosal disease of the GIT (58) and invasion of the oral cavity and stomach (59), although the vast majority of pigs appear unaffected. Interestingly, the commercial pigs as a whole have much less Candida than the feral pigs; however, most pigs have at least some Candida, and a couple of individuals have up to 96.7% of their mycobiomes made up by Nakaseomyces, a clade made up of pathogenic Candida species (60). This suggests that there is some individual susceptibility to high levels of Candida colonization, which has previously been shown in humans and is influenced by genetics, the host metabolite pool, and antibiotic use (38, 61).

Conclusion

We found that the amount of K. slooffiae the maternal sow had coincided with piglet K. slooffiae status up to 119 days post-weaning. In addition, we found that piglets had more comparable mycobiome compositions to their litter mates in early life as well as at the end of one production cycle. Data from this work indicate a potential transient disruption to the mycobiome during the weaning, where littermate groupings no longer exist, paralleling the dysbiosis that has been documented in bacterial taxa. Overall, this study provides evidence of the importance of early-life mycobiome composition on long-term mycobiome composition in pigs and how conventionally raised pigs’ mycobiomes differ significantly in composition and complexity from their feral counterparts.

ACKNOWLEDGMENTS

This work was supported by a Swine Innovation Porc grant held by B.P.W. and a Discovery grant from the Natural Sciences and Engineering Research Council also held by B.P.W. T.L.P. was supported by the Frank Aherne Graduate Scholarship in Swine Research and the Alberta Graduate Excellence Scholarship. B.P.W. is supported by the Canada Research Chairs Program.

Contributor Information

Benjamin P. Willing, Email: willing@ualberta.ca.

Irina S. Druzhinina, Royal Botanic Gardens, Surrey, United Kingdom

DATA AVAILABILITY

ITS sequence data is located in the National Center for Biotechnology Information Sequence Read Archive and is available under BioProject accession number PRJNA1008685.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Data Availability Statement

ITS sequence data is located in the National Center for Biotechnology Information Sequence Read Archive and is available under BioProject accession number PRJNA1008685.

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