123 COMPARING MICROBIAL COMMUNITIES ACROSS THE GASTROINTESTINAL TRACT OF MOOSE Katie Lynn Anderson and Lisa Shipley Washington State University, Pullman, WA 99164, USA Correspondence author: Katie Lynn Anderson, misskate.anderson3@gmail.com ABSTRACT: Coevolution of microbial communities and their hosts has allowed herbivores to extract energy from recalcitrant complex carbohydrates in plants, and benefit from synthesized protein and vitamins. Ruminants have further evolved 3 locations within their gastrointestinal (GI) tract that house and that facilitate fermentation of plant fiber, the rumen/reticulum, cecum, and colon, but little is known about how the microbiome varies across these organs, especially in free-ranging browsing herbivores like moose (Alces alces). Therefore, we took advantage of a unique opportunity to sample each of these organs from vehicle-struck and hunter-harvested moose in south-central Alaska and eastern Washington/northwestern Idaho, USA. Using culture-free microbial techniques to extract, identify, and quantify microbes in the samples, we found few differences in microbial composition, richness, and diversity among fermentation organs, but large differences between geographic regions. Our results suggest that differences in the macroenvironment and diets across regions play a greater role in microbial communities than different microenvironmental conditions within organs of the GI tract. Furthermore, our findings support microbiome measurements from fresh fecal samples as a surrogate for microbial communities in the rumen and cecum. Understanding the function and com- position of microbial communities within the fermentation organs is increasingly important for quan- tifying how ruminants such as moose will respond to changes in diet and environment, and how these changes may influence their interactions with plant resources. ALCES VOL. 60: 123–145 (2024) Key Words: Alaska, Alces alces, cecum, colon, Idaho, microbial communities, Washington Ruminants have evolved a complex, anoxic foregut portion of their gastrointestinal (GI) tract that houses a large and diverse microbial community (i.e., 1010 to 1011 organisms/ml, (Newbold and Ramos-Morales 2020). This adaptation allows them to convert cellulose (the most abundant biomolecule on Earth), other recalcitrant structural carbohydrates, and elemental nutrients of plants to microbial cell protein and short chain fatty acids that fuel metabolism and build body tissue (Hofmann 1989, Solden et al. 2018). Approximately 80% of the energy derived by the ruminant host is produced from fermentation by the rumen microbiota (Van Soest 1994). Through anaerobic fermentation, microbes in the ruminant foregut not only extract energy and nutrients from plants ingested by their host for their own use, but also synthesize vita- mins, and aid in detoxifying plant secondary metabolites (Kartzinel et al. 2015), which provides critical benefits to their host in the ultimate symbiotic relationship. The evolutionary benefits of microbial fermentation are enhanced when herbivores have multiple fermentation chambers that can increase digestion and absorption of nutrients from the plants they consume and improve detoxification efficiency. Ruminants (Bovidae, Cervidae, and Antilocapridae) mailto:misskate.anderson3@gmail.com ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 124 have 3 sites in their GI tract that house most of the microbes important to nutrition: the rumen/reticulum in the foregut, and the cecum and colon that are collectively referred to as the hindgut. The rumen is the largest fermentation organ, accounting for approximately 16% of a ruminant’s total weight and 84% of the total GI volume (Van Soest 1994, Niehaus et al. 2019). The rumen is the primary location for fermentation and has large papillae and many folds to increase surface area for absorption of energy-rich byproducts of microbial fermentation. The papillae develop as the animal ages, stimu- lated by the presence of ingested food avail- able for fermentation (Robbins 1994). Papillary structure and density are partly influenced by the animal’s diet but are gen- erally larger and denser in the ventral regions of the rumen where nutrient absorption is most pronounced (Van Soest 1994). Fine food particles wash out of the rumen into the reticulum, which is tasked with sorting and handling these particles, aided by its unique honeycomb texture. Like the rumen, the reticulum also hosts fermenting microbes, but its main role is to ensure larger particles stay in the rumen to be further diminished by rumination and fermentation while finer par- ticles and microbial cells move on to the next chamber (Van Soest 1994)(Van Soest 1994). Further along the GI tract beyond the gastric stomach (i.e., abomasum), fermenta- tion also occurs in the cecum and colon. The cecum and colon are often understudied in ruminants because they are smaller than the foregut and are more difficult to access for sampling in live animals. The ruminant cecum, situated between the small intestine and colon, composes approximately 2% of the GI volume (Hoover 1978, Russell 2002). The cecum is a secondary site of fermenta- tion in ruminants, containing a diverse microbial community. The rate and extent of cecal fermentation depends on the amount and type of fermentable digesta that escapes foregut fermentation and gastric catabolism. During periods of heightened food intake (e.g., summer, lactation), passage rate through the GI tract is increased to avoid overfilling the rumen by reducing rumina- tion time and drinking more water (Asano et al. 2007). Under these conditions, larger particles of food are more likely to escape the rumen before they can be fermented fully, at which point the cecum can provide a second chance to capture energy and nutri- ents from hemicellulose and cellulose (Allen et al. 1986, Van Soest 1994). The most distal fermentation site of ruminants, the colon, composes approxi- mately 12.5% of the total GI volume (Hoover 1978), and is the primary location of water, sodium, and copper absorption. However, the colon also houses microbial communi- ties capable of fermentation. As in the cecum, colon communities specialize on the fermentable substances that by-pass the foregut or get washed out before being fully fermented (Hoover 1978, Van Soest 1994). The colon also plays a major role in the recovery and transfer of nitrogen from the microbes to the blood, where it can then be returned to the rumen and used for micro- bial protein synthesis (Hoover 1978). In most mammals, the GI microbi- ome is dominated by bacterial phyla: Firmicutes, Bacteroidota, Actinobacteria, and Proteobacteria (Ley et al. 2008, Manichanh et al. 2010). Because Firmicutes and Bacteroidota are the most abundant phyla in the gut of mammals, including ruminants (Magne et al. 2020, Pinnell et al. 2022, Bensch et al. 2023), and they have different functions, the ratio of the 2 has often been used to evaluate proper gut functioning, such as identifying various health conditions, hor- mone shifts, and seasonal changes in the diet (Springer et al. 2017). Although Firmicutes ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 125 and Bacteroidetes are both important for car- bohydrate metabolism, Firmicutes primarily ferment nonfibrous carbohydrates and pro- teins whereas Bacteroidetes digest and uses cellulose and exogenous peptides as a food source (Hayes and Bleakley 2018, Wu et al. 2022, Fu et al. 2024). The ruminant’s micro- biome also includes bacteria species from Acidobacteria, the Chloroflexi phyla, and some fungal and protozoan species (Ishaq and Wright 2014, Ishaq et al. 2015, Solden et al. 2017). Three bacteria species, Fibrobacter succinogenes within the Fibrobacterota phylum and Ruminococcus albus and Ruminococcus flavefaciens within the Bacillota phylum, have been considered the predominant agents of cellulolysis in the rumen, but the full suite of microbial taxa that are key to cellulose degradation in the rumen has yet to be identified (Weimer 2022). Microbes within the same GI organ spatially segregate, depending on their roles or preferred food sources (Van Soest 1994). Microorganisms that ferment cellu- lose in plant fiber will often etch pits into the available surfaces of the fibrous mate- rial to anchor themselves to the fiber and remain in the rumen longer, but microbes in the rumen liquid are typically non-cellulo- lytic (Kong et al. 2010). Some bacteria attach to the rumen wall, and they are often ureolytic and aid in urea and oxygen trans- fer. Bacteria that are not attached to a sub- strate as well as free floating protozoans and fungi tend to wash out of the rumen at the liquid passage rate, and are later digested by the animal in the abomasum (gastric stomach, Van Soest 1994, Kong et al. 2010, Weimer 2022). Describing and comparing the microbial communities spatially sepa- rated in the digestive tract is in its early stages, and has focused solely on hindgut fermenters or ruminant livestock that con- sume more grasses (Kong et al. 2010, Mao et al. 2015, Holman and Gzyl 2019). Therefore, little is known about these com- munities for browsing ruminants that con- sume plants with high levels of insoluble lignin and often toxic plant secondary metabolites (PSMs, Godoy-Vitorino et al. 2012, Svartström et al. 2017). Not only is the microbial community expected to vary across the GI tract based on the morphology and function of the fermen- tation organ, but also with the composition of diets consumed, and thus the habitats in which herbivores forage. Because of their circumpolar distribution, moose reside in areas that vary greatly in available forage resources. Moose are large, near-obligate dietary specialists on boreal woody plants, primarily deciduous leaves during the grow- ing season and stems and evergreens during the dormant period from mid-autumn through mid-spring. Their diets usually include willows (Salix spp.) and birch (Betula spp.) in Alaska and the Rocky Mountains, and conifers in Maine and Sweden (Shipley 2010). Dormant deciduous and evergreen shrubs not only contain high amounts of cellulose, but also high amounts of lignin and cutin (e.g., 15% - 20%) that cannot be fermented by GI microbes to extract energy (Spaeth et al. 2002, Spalinger et al. 2010). In addition, leaves of both deciduous and evergreen shrubs are chemi- cally defended by PSMs such as condensed tannins and monoterpenes (Palo 1984, McArt et al. 2009, Tremblay et al. 2019). Therefore, moose rely on adaptations to help consume and digest these diets, including specific mouth morphology (e.g., prehensile tongue and lips, a wide mouth, large salivary glands (Hofmann and Nygren 1992, Van Soest 1996, Clifford and Witmer 2004), pro- line-rich saliva that binds tannins (Hagerman and Robbins 1993), and, most importantly, their GI microbiota (Ishaq and Wright 2012, Ishaq et al. 2015, Solden et al. 2017). ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 126 Our objectives were to describe and compare the microbial communities in the rumen, cecum, and colon of free-ranging moose, and compare microbial communities of moose in 2 different study areas in the United States, southcentral Alaska (Alaska, hereafter) and eastern Washington and north- western Idaho (Inland Northwest, hereafter). We hypothesized that the diversity of micro- bial communities would decrease along the GI tract, thus the highest diversity in the rumen and the lowest in the colon. This hypothesis is supported by research with domestic cattle consuming traditional live- stock rations that found lower microbial diversity in the hindgut (Holman and Gzyl 2019), likely because the gastric stomach removes much of the dietary substrates that provide food for microbiota in the hindgut. In addition, the cecum and colon have shorter retention times and less spatial sepa- ration than the rumen, so the microbes also have less time to fully establish into unique subcommunities (Dehority 2002). Because the rumen is the first major fermentation organ chamber in the GI tract, we also hypothesized that its microbial community would be dominated by organisms best able to ferment and transform cellulose and detoxify PSMs that reduce the animal’s bur- den for detoxification later in the GI tract. Therefore, we predicted that the rumen would have a higher abundance of cellulo- lytic bacteria whereas the cecum would have a higher relative abundance of hemicellulose digestors (Kohl et al. 2014, Cholewińska et al. 2021, Pinnell et al. 2022). Finally, we hypothesized that the ratio of Firmicutes to Bacteroidetes would be lowest in the rumen and increase down the GI tract from the cecum to colon, because Bacteroidota would be most beneficial in fermenting complex carbohydrates, especially cellulose, and Firmicutes most beneficial in fermenting starches and proteins that escape the rumen. METHODS Study area We opportunistically collected samples of the GI microbiome of moose in 2 study areas. The samples from Alaska came from 5 moose struck by vehicles in Game Management Units (GMU) 14A and 14C, which included the cities of Anchorage (61°13′ N 149°54’ W), Wasilla (61°34′ N 149°27′ W), and Palmer (61°36′ N 149°07′ W, Table A1). This area houses approxi- mately 40% of the human residents of Alaska, however, most of the area (66% or 19,900 ha) is woodland (mixed, deciduous, coniferous, and shrub). The woodlands con- sisted of paper birch (Betula papyrifera), white spruce (Picea glauca), black spruce (Picea mariana), quaking aspen (Populus tremuloides), balsam poplar (Populus bal- samifera), and black cottonwood (Populus trichocarpa). Shrubs were primarily willow, alder (Alnus spp.), and high-bush cranberry (Viburnum edule). Vehicle collisions were a significant source of mortality for the local moose population, accounting for 200 to 300 moose killed annually depending on snow- fall (Peltier 2015). In 2022, this area accu- mulated approximately 178 cm of snowfall (Alaska Climate Research Center 2022), had an average minimum temperature of -13 °C (U.S. Climate Normals 2021). Vehicle- struck moose provided easily accessible and less invasive samples than wild-caught or captive animals. In the Inland NW study area, we col- lected microbiome samples from both vehi- cle-struck and hunter-harvested animals, including 2 from northwestern Idaho, GMUs 8 and 11, near the cities of Moscow (46°44′ N 117°00 W) and Troy (46°44′ N 116°46′ W), and 5 from eastern Washington, GMU 124, near the city of Spokane (47°40′ N 117°25′ W, Table A1). This area is charac- terized by rolling hills and an open landscape ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 127 that supports plant communities dominated by big sagebrush (Artemesia tridentata), Idaho fescue (Festuca idahoensis), and blue- bunch wheatgrass (Pseudoroegneria spi- cata; Looney and Eigenbrode 2012). Snowberry (Symphoricarpos albus), quak- ing aspen (Populus tremuloides), and willow (Salix spp.) grow along riparian areas, and scattered ponderosa pines (Pinus ponderosa) are distributed along the foothills. There is also extensive farming in the area, producing primarily wheat and legumes. The mean minimum temperature in winter across all years was −4.3° C, and mean annual precip- itation in this area between 2018–2022 was 52 cm, falling mostly as snow in the winter (U.S. Climate Normals 2021). To maintain sample quality, we only collected samples from animals that could be accessed within 3 hours of death as reported by the motorist or hunter. . This ensured minimal microbial degradation as the animal tissues begin to break down. Sampling methods We used sterile instruments to open the animal’s abdomen, rumen, cecum, and fecal samples from the colon (hereafter referred to as colon samples), collecting a mixture of fluid and solid digesta in equal proportion from each organ. We homogenized the solid and fluid contents to minimize potential bias caused by the animal’s body position that could cause the pooling of fluids. Samples were placed on ice immediately after collec- tion, and frozen at -20° C until extraction, which was accomplished no later than 6 months after collection. We analyzed each sample from the rumen, cecum, and colon for microbial composition and diversity using a 16S rRNA approach. This gene sequencing method is commonly used for identifying and quantifying microbes found in samples that could have thousands of dif- ferent bacterial species. This method targets the 16S rRNA genes, a highly conserved region of the transcriptional machinery found in all DNA-based life forms, making it an ideal target for sequencing DNA (Combrink et al. 2023). We extracted samples using the Qiagen RNEasy PowerMicrobiome Kit with manufacturer recommendations. The amount of DNA in each sample was measured using a Qubit (Invitrogen, Inc., Carlsbad, CA), and 5 μl of each DNA suspension was amplified by polymerase chain reaction (PCR) using a MasterCycler thermocycler (Eppendorf, Westbury, NY). Samples were multiplexed and sequenced using Illumina MiSeq Platform using the 300 cycle MiSeq Reagent kit V2 at the Advanced Instrumentation for Microbiome Studies (AIMS) core facility (Brauner and Briggs 2023). We sequenced samples using a 600-cycle MiSeq Reagent Kit v3 (Illumina®) in the presence of 25 % PhiX DNA. Following sequencing, samples were processed using the software Quantitative Insights Into Microbial Ecology (QIIME2), beginning with demultiplexing and removing samples that did not amplify, removing sequences that had less than 300 bp. We then denoised and dereplicated sam- ples using the Divisive Amplicon Denoising Algorithm (DADA2) module within QIIME2 and removed sequences that had less than 300 bp. Unique amplicon sequence variants (ASVs) were generated using DADA2 and assigned a taxonomy using the SILVA 138 ribosomal RNA database. ASVs that could not be assigned to phylum were removed. Downstream analysis was done in R v 4.0.735. We imported QIIME2 readable files (*.qza) into R using the package qiime2R v0.99 (Bisanz 2024). We then used the phyloseq package (McMurdie and Holmes 2013) to remove non-bacterial, mitochondrial, and chloroplast ASVs, we then removed the negative controls (i.e., DNA-free water). ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 128 Statistical analysis We examined the effects of GI organ (rumen, cecum, colon) and geographic region (Alaska, Inland NW) on taxonomic alpha diversity, beta diversity, and relative abun- dance of the most abundant microbial taxa. Alpha diversity quantified diversity within individual samples, allowing comparison across sample groups using 2 metrics-over- all species richness and the Shannon-Weiner Diversity Index. We characterized richness as the number of unique microbial sequences per sample. We calculated the Shannon Index, which is the number of unique reads and their proportional abundance in the sequence, using the phyloseq package (McMurdie and Holmes 2013). We exam- ined the effects of GI organ and geographic region starting with a linear mixed effects model with the individual moose as the grouping variables using the lme4 and lmerTest packages (Bates et al. 2017, Kuznetsova et al. 2013). We calculated esti- mated marginal means (emmeans) and lower and upper limits of the 95% confidence interval using the emmeans package (Lenth et al. 2024). Lastly, we performed a post-hoc Levene’s test to compare variance in alpha diversity among GI organ and study loca- tions (Derrick et al. 2018). We calculated beta diversity to com- pare the dissimilarity between samples (moose location and organ), creating a distance matrix between all pairs of sam- ples. To quantify the beta diversity of the microbiome, we first filtered and clus- tered sequences into amplicon sequence variants (ASVs). ASVs were used to reconstruct phylogenies, allowing us to calculate microbial beta diversity and rel- ative abundances (Combrink et al. 2023). We then transformed the read counts of our non-rarefied data to relative abun- dance by dividing the number of reads for each taxon within a sample by the total number of reads for that sample. We per- formed a Permutational Multivariate Analysis of Variance (PERMANOVA) to compare Bray-Curtis distances among GI organs and geographic regions, using the adonis2 function in the vegan package v2.6-4 (Oksanen et al. 2024). To visualize differences among independent variables, we used a Principal Coordinates Analysis with the Bray-Curtis distance that consid- ers presence/absence of different ASVs and their abundance and an Analysis of Similarity (ANOSIM). We compared the response of the relative abundance of selected microbial families or phyla, including differences in the abundances of Firmicutes and Bacteroidota phylum, to GI organs and geographic regions using ANOVAs, and means were separated using the Tukey test. For our analyses, we set the statistical significance at α = 0.10. We used this α (higher than the conventional (α = 0.05) to better align with our exploratory research objectives (McDonald 2009). RESULTS Of the 30 samples collected from 3 GI organs of 5 moose in Alaska and 7 moose in the Inland NW, we removed 4 samples because they did not amplify during PCR. Two of the Alaska moose samples had with organs that had burst and therefore we were unable to collect a sample. In our linear models that contained both geographic region and GI organ, microbial richness did not vary with region (Alaska: emmeans = 551, SE = 119, Inland NW: emmeans = 481, SE = 96, P = 0.66) or GI organ (all P > 0.1). However, the Shannon Index was lower in Alaska (emmeans = 3.08, SE = 0.56) than the Inland NW (emmeans = 4.09, SE = 0.46, F = 6.30, P = 0.03) but did not differ among organs (all P > 0.1, Fig. 1). The variance of micro- bial richness was greater in the Alaska ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 129 samples (var = 261,746) than the Inland NW samples (var = 22,640, F = 9.29, P = 0.005), and for both areas combined higher in the rumen (var = 155,147) and cecum (var = 154,995) than the colon (var = 11,613, F = 2.52, P = 0.1; Fig. 1). The variance of the Fig. 1. Microbial richness (A) and Shannon Index (B) of the microbial community found in the rumen, cecum, and colon of moose in Alaska and the Inland NW (eastern Washington and northwestern Idaho), United States. Samples were collected from vehicle collisions and hunter harvests during the fall and winter seasons, 2022–2024. Asterisk denotes a difference between regions for the Shannon Index Values. ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 130 Shannon Index was also greater in the Alaska samples (var = 4.55) than the Inland NW samples (var = 0.20, F = 26.32, P < 0.0001). However, the variances did not differ across GI organs (F = 0.034, P = 0.96; Fig. 1). Much like alpha diversity, we found that geographic region (F = 7.14, R2 = 0.21, P = 0.001), influenced beta diversity but GI organ did not (F = 0.63, P = 0.96; Fig. 2). The ANOSIM also showed that regions dif- fered significantly (F = 7.14, R2 = 0.24, P = 0.001) but there were no differences among GI organs. The first two axes of the PCoA using the Bray-Curtis distance for ordinations accounted for 30% of the varia- tion in beta diversity for moose in both regions, clustering by geographic region but not GI organ (Fig. 2). Fig. 2. Ordinations of microbial beta diversity obtained from the rumen, cecum, and colon of moose in Alaska and Inland NW (northwestern Idaho and eastern Washington), United States. Samples were collected from vehicle collisions and hunter harvests in the fall and winter seasons, 2022–2024. Beta diversity ordinations based on a PCoA with the Bray-Curtis distance metric (A), ordinations were then replotted using just the centroids with the arrows indicating the direction of change (B). ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 131 In addition to differences in microbial alpha and beta diversity, the 2 regions had distinct microbial community compositions (Fig. 3, Table A2). We identified 7 families that were present only in the Alaska samples and 8 present only in the Inland NW. Furthermore, families that were identified in both regions differed in relative abundance. The top family identified in the Alaska sam- ples, Enterobacteriaceae, had a higher abun- dance in the Alaska samples (x̄ = 28.49, SE = 31.39) than the Inland NW (x̄ = 3.39, SE = 3.32, F = 15.08, P = 0.002, Fig. 3). The top 3 identified families for Inland NW moose, Oscillospiraceae (NW x̄ = 20.95, SE = 2.03 vs. AK = x̄ 15.49, SE = 8.57, F = 17.17, P = 0.01), Lachnospiraceae (NW x̄ = 17.06, SE = 3.85 vs. AK x̄ = 10.88, SE = 7.46, F = 38.60, P = 0.002), and Christensenellaceae (NW x̄ = 13.12, SE = 2.66 vs. AK x̄ = 8.31, SE = 5.73, F = 43.51, P = 0.002) all had greater relative abundances than in Alaska (Fig. 3, Table A2). However, across GI organs within regions, we found that each organ had the same top 5 families except Oscillospiraceae in the Alaska samples (F = 3.40, P = 0.08), where relative abundance for Oscillospiraceae was highest in the cecum (x̄ = 20.83, SE = 7.01), followed by the rumen (x̄ = 2.57, SE = 3.96) and finally the colon (x̄ = 6.93, SE = 1.01, Fig. 3). In addition, the relative abundance of Firmicutes was lower in the colon (x̄ = 1.76, SE = 0.99) than in the rumen (x̄ = 5.38, SE = 2.08, P = 0.09) and cecum for Alaskan moose (x̄ = 4.93, SE = 1.66, P = 0.1, Table 1). We find no differences between the 2 regions or within the Inland NW samples for either Firmicutes, Bacteroidota, or the F:B ratio between geographic location or among GI organs (all P > 0.1, Fig. 4). DISCUSSION Although we expected to find differences in microbial diversity and composition across the 3 fermentation organs of moose GI tracts (i.e., rumen, cecum, and colon), these differences were minimal in both geographic regions we sampled. On the other hand, we found differences in micro- bial diversity and composition between the 2 geographic regions. Taken together, our results suggest that regional environmental characteristics, such as diet composition and seasonal availability of forages, are more important drivers of microbial com- munity diversity and composition than the internal microenvironment caused by phys- iological and functional differences along the GI tract. Because of the minimal differ- ences in the microbiome from the rumen to the colon, our findings support sampling the microbiome from fresh fecal samples as a reasonable surrogate for microbial com- munities in the rumen and cecum. We expected that microbial diversity would decrease along the GI tract because of the decrease in the amount of substrate for the microbes and the faster passage rate after the gastric stomach (Van Soest 1996, Stevens and Hume 1998, Clauss and Hummel 2005). However, microbial diversity and richness did not vary among the GI fermentation organs of moose in either region. These sim- ilar levels of microbial diversity may reflect the need for functional redundancy in GI microbial community structure that addresses the most important task of break- ing down the complex fibers in their diet and adds resilience as diet changes across sea- sons and locations (Ribas et al. 2023). On the other hand, diets consumed by moose during the non-growing season (e.g., decidu- ous twigs and conifers) are relatively less diverse than in the growing season, which might have resulted in a relatively low diver- sity overall (Solden et al. 2017). For exam- ple, the mean value of the Shannon index across the GI tract of moose in our study was lower (3.08 in Alaska and 4.09 in Inland ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 132 Fig. 3. Top 20 bacterial families identified in the rumen, cecum, and colon of moose in Alaska (A) and Inland NW (B, northwestern Idaho and eastern Washington), United States. Samples were collected from vehicle collisions and hunter harvests in the fall and winter seasons between 2022 and 2024. Families are ordered from most to least abundant from bottom to top. Asterisks denote significant differences between the two geographic regions, α = 0.10. ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 133 Northwest) than the value (i.e., 6) recorded for the American bison (Bison bison) during the fall in Colorado (Bergmann 2017), but were similar to that of pronghorn (Antilocapra americana) fecal samples (i.e., 4.3) in fall and winter in Wyoming (Buchanan et al. 2024). We also expected the rumen to have the highest diversity of microbes and contain primarily cellulolytic bacteria compared to the colon because of the change in available substrates along the GI tract. Contrary to our expectations, however, microbial taxa and the Firmicutes: Bacteroidetes ratio were Fig. 4. Relative Abundances of the Firmicutes and Bacteroidetes phyla identified in the rumen, cecum, and colon of moose in Alaska and Inland NW (northwestern Idaho and eastern Washington), United States. Samples were collected from vehicle collisions and hunter harvests in the fall and winter seasons, 2022–2024. Different capital letters indicate differences among means for Firmicutes from the Tukey means separation tests (α = 0.05). The same lowercase letters indicate that the relative abundance of Bacteroidota did not differ among organs within regions. Table 1. Mean relative abundance (SE) for the Firmicutes and Bacteroidota phyla for moose by geographic region (Alaska vs. Inland NW [eastern Washington and western Idaho], USA) and gastrointestinal (GI) organ during the fall and winter seasons of 2022–2024. Region GI organ Firmicutes Bacteroidota Ratio Alaska Rumen 5.38 (2.08) 2.49 (0.93) 2.16 Cecum 4.93 (1.66) 1.45 (0.53) 3.41 Colon 1.76 (0.99) 0.19 (0.09) 9.08 Inland NW Rumen 6.70 (1.94) 3.44 (1.12) 1.94 Cecum 6.36 (1.95) 4.45 (1.36) 1.43 Colon 6.28 (1.91) 3.94 (1.32) 1.59 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 134 generally similar among the fermentation organs, but we did note 1 microbial phylum and 1 family that differed in Alaskan moose. Firmicutes were more abundant in the rumen than the cecum and colon, whereas Oscillospiraceae, an epithelial group, were more abundant in the cecum than the rumen, and more abundant in the rumen than the colon. The higher abundance of Firmicutes in the rumen may be due to the energetic requirements of the season when the animals were sampled. Firmicutes can act as an effi- cient energy source, leading to more calorie absorption which is important during winter when food is scarce (Söllinger et al. 2018). Likewise, domestic cattle (Bos taurus) fed a high forage diets had higher abundances of Firmicutes, but when fed a high concentrate diet, they had higher abundances of Bacteroidota (Clemmons et al. 2019). The higher abundance of Oscillospiraceae in the hindgut is unsurprising based on the func- tion of the cecum in foregut fermenters, which serves as a site for absorption of water and nutrients from bacterial fermentation. As an epithelial group, Oscillospiraceae would have the ability to absorb the micro- bial proteins, the nutrients that were partially digested in the acid stomach, and the meta- bolic products from other microbes that can then be used as a substrate by other microbes (Grond et al. 2021). Microbes engage in cross-feeding, whereby they produce SCFAs that are not only useful to the host and other microbes, but can also digest those microbes themselves (Solden et al. 2018). Cecal fer- mentation can be important to a ruminant because both the microbes and the animal can take advantage of nutrients that require prior breakdown, either through digestion or fermentation in the rumen and small intes- tine (Hoover 1978). Although none of our 3 hypotheses were strongly supported, our results align with some studies with other species. For example, research with ruminant livestock (Mao et al. 2015), captive-raised bison (Bergmann 2017), and moose in Vermont (Ishaq and Wright 2012) found differences in the relative abundances of different bacterial orders among GI organs, but did not find a consistent decrease in the abundances along the GI tract. Like Ishaq and Wright (2014), our samples also con- tained a relatively high abundance of the Lachnospiraceae family. Lachnospiraceae have been found in the human large intes- tine (Flint et al. 2012) and are fiber degrad- ers often found in the solid digesta fractions (Deusch et al. 2017). Because of its ability to produce butyrate, this family can help reduce the side effects of gastrointestinal inflammation and can help stimulate growth of papillae (Ishaq and Wright 2014). One potential reason for the amount of overlap in the microbial species between the GI organs in moose is that the general purpose of each organ is essentially the same, fer- menting cellulose, hemicellulose, and other nutrients to capture energy contained in SCFAs, break down food particles, and increase the digestive efficiency of the ani- mal (Van Soest 1996, Kohl et al. 2014). This functional redundancy across the GI tract ensures that the animal can extract the most energy and nutrients from a complex diet, well-defended by insoluble fiber. In contrast to our moose, woodrats (Neotoma spp.), which have a small foregut fermenta- tion portion of their bilocular stomach and a larger cecum, were found to have a foregut community that was better suited for detox- ification of PSMs and a hindgut community that was bettered suited for SCFA produc- tion (Kohl et al. 2014). A second potential reason that microbial communities did not differ across the GI organs is our relatively small sample size and high variance among individuals in our study, even though we controlled for ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 135 individual variation in our models using a grouping variable. The variance for both microbial richness and Shannon Index was particularly high in the rumen and cecum of the Alaska samples. Experiments with cap- tive ruminants in controlled conditions have shown substantial individual variation in diet selection, thus differences in microbial communities might be expected to be ampli- fied in free-ranging animals in natural land- scapes (Provenza and Balph 1987, Anderson 2020). Dietary differences might have the greatest influence on microbes in the rumen because the food entering the rumen has undergone only minimal transformation from saliva. The rumen provides a stable environment for microbes to establish larger communities, with perturbations that contin- ually expose new substrates to the microbes (Troyer 1984, Weimer 2015). Rumen microbial diversity also depends in part on the chemical complexity of the foods consumed, specialization of subcom- munities for maximal biochemical effi- ciency, and the possibility that the best-suited microbes will displace others that are under-suited and create opportuni- ties for new microbes (Russell 2002). Therefore, the rumen microbial community would be expected to vary the most among individuals based on the animal’s early life experiences, health status, physiology, behavior, diet, and other members of the microbial community (e.g. cross feeding and the presence of protozoa [Troyer 1984, Ley et al. 2008, Rosenberg and Zilber- Rosenberg 2016, Clemmons et al. 2019, Fountain-Jones et al. 2020, Dearing and Weinstein 2022]). Variation in microbial communities in moose likely also reflects the substantial chemical diversity con- sumed in the form of PSMs. Deciduous and coniferous shrubs and trees are heavily defended by antimicrobial chemicals (terpenes and phenolic resins), complex polyphenolics that are known to interfere with protein digestion (e.g., tannins), as well as a suite of other primarily small or intermediate-sized phenolics that can act as metabolic toxins to the animals (Richards et al. 2016). The microbial community within the GI tract can enhance the herbivore’s innate detoxification system while also detoxifying PSMs themselves, lessening the overall bur- den on the host (Dearing et al. 2005). Although microbial communities did not vary greatly among fermentation organs, microbial diversity and composition differed substantially between our study areas. These differences most likely reflect differences in diet and other macro-environmental condi- tions. Similarly, Ishaq and Wright (2014) found that the differences in region out- weighed the differences in the GI organ sam- pled for moose in Vermont, Norway, and Alaska. In addition, microbial communities could in part reflect small differences in the GI tract of moose based on their phylogeny between our study regions (i.e., A. a. gigas in Alaska vs. A. a. shirasi in the continental U.S., Chong et al. 2018, Arshad et al. 2021). We sampled from 2 study regions to increase our sample size and to determine the univer- sality of patterns in the GI microbiome but did not have a priori expectations about which geographic location would support the most diverse GI microbiome, in part because we lacked adequate information about their diets during fall and winter. Previous studies have demonstrated that moose in Alaska consume a large variety of willow species in both summer and winter, but the only published data on diets of Inland NW moose suggests that they might con- sume a greater variety of plant functional groups such as grass and forbs, at least in summer (Schrempp et al. 2019). Because our samples were collected in fall and win- ter, snow cover and reduced abundance of nutritious forages likely restricted diet ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 136 choices, especially for the Alaska moose (Hamilton et al. 1980, Hoy et al. 2022). For example, Cook et al. (2021) found that approximately 80% of the moose sampled in Northwestern Washington showed some level of starvation and starvation has also been the cause of winter mortality for many Alaskan moose (Testa 2004, Boertje et al. 2007, 2019). GI microbial communities respond to declining abundance and nutri- tional quality and diversity of diets from the summer to the winter (Solden et al. 2018, Zou et al. 2019). To gain insight into causes of regional differences in the GI microbiome of moose, future studies could directly link diet to microbiome by 1) collecting simulta- neous rumen or fecal samples and identify- ing diet composition using molecular techniques similar to those of the microbi- ome (Kartzinel et al. 2015, Sonsthagen et al. 2020), 2) minimizing variation of sample collection dates or collect samples through- out each season, or 3) using controlled feed- ing studies with specific diet items of interest. Not only did microbial diversity differ between Alaska and Inland NW, but the rel- ative abundance of microbial families did also. In fact, almost half of the 20 families were not found in samples from the other region. The top family identified in the Alaska samples, Enterobacteriaceae, a member of the Proteobacteria phylum, is a facultative anaerobic family that ferments lactose and produces acid from glucose. This family is more abundant in hosts that live in high-elevation and/or cold environments, which would be consistent with the Alaskan region (Wu et al. 2022). The second most abundant was unclassified Clostridia_UCG- 014, which is an order found in the Firmicutes phylum that could not be further refined to family or genus. This order has been cor- related with tryptophan metabolism, helps maintain intestinal barrier function, and has been found in the cecum of several species (e.g., chickens and geese, Jiang et al. 2023; Zhao et al. 2025). The third most abundant family was Oscillospiraceae, which was also the top identified family for the Inland NW samples. Oscillospiraceae has been positively associated with growth and lacta- tion of ruminants, and can enhance growth and production of other microbes by mediat- ing SCFA transport (Tong et al. 2018; Pinnell et al. 2022; Chen et al. 2024). The top three identified families for the Inland NW, Oscillospiraceae, Lachnospiraceae, and Christensenellaceae, are all families within the Firmicutes phylum. Using metagenomic analyses, future studies could further deter- mine the function of various microbial taxa to better understand the relationship between microbial communities and the internal (GI tract) and external (regional) environments. Elucidating the function and composi- tion of microbial communities within fer- mentation organs is key to understanding how ruminants like moose respond to changes in diet and environment, and how these changes might influence their inter- actions with their plant resources. However, the logistics of sampling wild ruminants can often be a barrier to quanti- fying the complex interactions between the microbes and their hosts, especially considering the diversity of microbes within spatially segregated portions of the GI tract. Sampling recently killed vehi- cle-struck or hunter-harvested animals in our study allowed for relatively easy and precise sampling of potentially unique microbial communities (Kohl et al. 2014) within each fermentation chamber, with- out the ethical concerns related to surgical access to the GI in live animals or harvest- ing animals specifically to sample their microbiome. Although we were able to bypass some logistic limitations by oppor- tunistically sampling vehicle-struck or ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 137 hunter-harvested animals, this approach cannot completely control for variation caused by postmortem physiological pro- cesses. Microbial communities change after the host dies as the stability of the environment and the substrates they rely on for food are no longer available. Although changes in the microbial com- munities after host death are used as a forensic technique on human cadavers (DeBruyn and Hauther 2017), to our knowledge there has not been a study on a ruminant that quantifies how the commu- nity changes postmortem. Microbes capable of fermentation pro- vide a quid pro quo relationship with their host. The microbes providing the host SCFAs, essential amino acids, vitamins, and microbial protein; whereas the herbivores provide a consistent anoxic environment and substrate (Troyer 1984, Söllinger et al. 2018). However, the characteristics of the GI microbiome have implications for more than just extracting nutrients from plants. Recent research suggests that the microbiome may have a role in mental and physical health, hormone balance, and immune response within humans, livestock, and wildlife (Delzenne et al. 2019, Anand and Mande 2022, Vos et al. 2022). Because climate change is expected to influence allocation of resources within plants, which in turn affects how ruminants and their symbiotic micro- bial communities can sustain themselves and interact with their environment, this research provides a baseline for future comparisons (Herms and Mattson 1992, Moore et al. 2015, Chen et al. 2022, Beale et al. 2023). ACKNOWLEDGEMENTS We would like to thank B. Briggs and the AIMS Core Lab at the University of Alaska Anchorage for running the 16S samples. We also appreciate the assistance collecting moose samples provided by Alaska Department of Fish and Game, Washington Department of Fish and Wildlife, and Idaho Department of Fish and Game, especially M. DeVivo and I. Hull. LITERATURE CITED AlAskA ClimAte ReseARCh CenteR. 2022. Snow Summary. Snow Summary. https://akclimate.org/snow-summary/. Accessed 8 Aug 2025. Allen, M. S., C. J. sniffen, And P. J. V. soest. 1986. 2. Rumen Dynamics. Pages 21–42 in M. J. Dobson, editor. Aspects of Digestive Physiology in Ruminants: Proceedings of a Satellite Symposium of the 30th International Congress of the International Union of Physiological Sciences, Held at Cornell University, Ithaca, New York, July 21-23, 1986. Cornell University Press. AnAnd, S., And S. S. mAnde. 2022. Host- microbiome interactions: Gut-Liver axis and its connection with other organs. Biofilms and Microbiomes 8:1–89. AndeRson, K. L. 2020. Effects of Fire on Diet Composition, Foraging Behavior, and Nutritional Status of Moose in South-Central and Interior Alaska. Master’s of Science, University of Alaska Anchorage. ARshAd, M. A., F. hAssAn, M. S. RehmAn, S. A. huws, Y. Cheng, And A. U. din. 2021. Gut microbiome colonization and development in neonatal ruminants: Strategies, prospects, and opportunities. Animal Nutrition 7:883–895. AsAno, S., S. ikedA, Y. kuRokAwA, S. kAndA, And H. itAbAshi. 2007. Seasonal changes in digestibility, passage rate and rumen fermentation of alfalfa hay in sika deer (Cervus nippon) under restricted feed- ing. Animal Science Journal 78:28–33. bAtes, D., M. mAeChleR, B. bolkeR, S. wAlkeR, R. H. B. ChRistensen, H. singmAnn, B. dAi, F. sCheipl, G. https://akclimate.org/snow-summary/ ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 138 gRothendieCk, And P. gReen. 2017. lme4: Linear Mixed-Effects Models using “Eigen” and S4. https://CRAN.R- project.org/package=lme4. Accessed 3 Dec 2024. beAle, P. K., W. J. foley, B. D. mooRe, And K. J. mARsh. 2023. Warmer ambi- ent temperatures reduce protein intake by a mammalian folivore. Philosophical Transactions of the Royal Society B: Biological Sciences 378:1–9. bensCh, H. M., C. tolf, J. wAldenstRöm, D. lundin, And M. Zöttl. 2023. Bacteroidetes to Firmicutes: captivity changes the gut microbiota composition and diversity in a social subterranean rodent. Animal Microbiome 5:1–11. beRgmAnn, G. T. 2017. Microbial commu- nity composition along the digestive tract in forage- and grain-fed bison. BMC Veterinary Research 13:1–9. bisAnZ, J. 2024. jbisanz/qiime2R. HTML. https://github.com/jbisanz/qiime2R. Accessed 21 Nov 2024. boeRtje, R. D., G. G. fRye, And D. D. young. 2019. Lifetime, known-age moose reproduction in a nutritionally stressed population. The Journal of Wildlife Management 83:610–626. boeRtje, R. D., K. A. kellie, C. T. seAton, M. A. keeCh, D. D. young, B. W. dAle, L. G. AdAms, And A. R. AdeRmAn. 2007. Ranking Alaska moose nutrition: Signals to begin liberal antlerless harvests. The Journal of Wildlife Management 71:1494–1506. bRAuneR, M., And B. R. bRiggs. 2023. Microbial iron acquisition is influenced by spatial and temporal conditions in a glacial influenced river and estuary system. Environmental Microbiology 25:3450–3465. buChAnAn, C. E., S. J. gAllA, M. E. musCARellA, J. S. foRbey, A. K. Reinking, And J. L. beCk. 2024. Relating gut microbiome composition and life his- tory metrics for pronghorn (Antilocapra americana) in the Red Desert, Wyoming. PLOS ONE 19:1–27. Chen, J., X. ZhAng, X. ChAng, B. wei, Y. fAng, S. song, D. gong, D. huAng, Y. sun, X. dong, Y. ZhAo, And Z. ZhAo. 2024. Multi-omics analysis reveals the effects of host-rumen microbiota inter- actions on growth performance in a goat model. Frontiers in Microbiology 15:1–13. Chen, S., M. holyoAk, H. liu, H. bAo, Y. mA, H. dou, G. Li, N. J. RobeRts, And G. jiAng. 2022. Global warming responses of gut microbiota in moose (Alces alces) populations with different dispersal patterns. Journal of Zoology 318:63–73. Cholewińska, P., M. wołoszyńska, M. miChAlAk, K. Czyż, W. RAnt, J. smoliński, A. wyRostek, And K. wojnARowski. 2021. Influence of selected factors on the Firmicutes, Bacteroidetes phyla and the Lactobacillaceae family in the digestive tract of sheep. Scientific Reports 11:1–18. Chong, C. Y. L., F. H. bloomfield, And J. M. o’sullivAn. 2018. Factors affecting gas- trointestinal microbiome development in neonates. Nutrients 10:1–17. ClAuss, M., And J. hummel. 2005. The diges- tive performance of mammalian herbi- vores: why big may not be that much better. Mammal Review 35:174–187. Clemmons, B. A., B. H. voy, And P. R. myeR. 2019. Altering the gut microbiome of cattle: Considerations of host-microbi- ome interactions for persistent microbi- ome manipulation. Microbial Ecology 77:523–536. CliffoRd, A. B., And L. M. witmeR. 2004. Case studies in novel narial anatomy: 2. The enigmatic nose of moose (Artiodactyla: Cervidae: Alces alces). Journal of Zoology 262:339–360. CombRink, L., I. R. humphReys, Q. wAshbuRn, H. K. ARnold, K. stAgAmAn, K. D. kAssChAu, A. E. jolles, B. R. beeChleR, And T. J. shARpton. 2023. Best practice for wildlife gut microbiome research: A comprehensive review of methodology for 16S rRNA gene investigations. Frontiers in Microbiology 14:1–18. https://CRAN.R-project.org/package=lme4 https://CRAN.R-project.org/package=lme4 https://github.com/jbisanz/qiime2R ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 139 Cook, R. C., J. Oyster, K. Mansfield, And R. B. Harris. 2021. Evidence of summer nutritional limitations in a Northeastern Washington moose population. Alces 57:23–46. deARing, M. D., W. J. foley, And S. mCleAn. 2005. The influence of plant secondary metabolites on the nutritional ecology of herbivorous terrestrial vertebrates. Annual Review of Ecology, Evolution, and Systematics 36:169–189. deARing, M. D., And S. B. weinstein. 2022. Metabolic enabling and detoxification by mammalian gut microbes. Annual Review of Microbiology 76:579–596. debRuyn, J. M., And K. A. hAutheR. 2017. Postmortem succession of gut microbial communities in deceased human sub- jects. PeerJ 5:1–14. dehoRity, B. A. 2002. Gastrointestinal tracts of herbivores, particularly the ruminant: anatomy, physiology and microbial digestion of plants. Journal of Applied Animal Research 21:145–160. delZenne, N. M., C. knudsen, M. beAumont, J. RodRigueZ, A. M. neyRinCk, And L. B. bindels. 2019. Contribution of the gut microbiota to the regulation of host metabolism and energy balance: a focus on the gut–liver axis. Proceedings of the Nutrition Society 78:319–328. deRRiCk, B., A. RuCk, D. toheR, And P. white. 2018. Tests for equality of vari- ances between two samples which con- tain both paired observations and independent observations. Journal of Applied Quantitative Methods 13:36–47. deusCh, S., A. CAmARinhA-silvA, J. ConRAd, U. beifuss, M. RodehutsCoRd, And J. seifeRt. 2017. A structural and functional elucidation of the rumen microbiome influenced by various diets and microen- vironments. Frontiers in Microbiology 8:1–21. flint, H. J., K. P. sCott, S. H. dunCAn, P. louis, And E. foRAno. 2012. Microbial degradation of complex carbohydrates in the gut. Gut Microbes 3:289–306. fountAin-jones, N. M., N. J. ClARk, A. C. kinsley, M. CARstensen, J. foResteR, T. J. johnson, E. A. milleR, S. mooRe, T. M. wolf, And M. E. CRAft. 2020. Microbial associations and spatial prox- imity predict North American moose (Alces alces) gastrointestinal commu- nity composition. Journal of Animal Ecology 89:817–828. fu, L., L. liu, L. ZhAng, Y. hu, Y. Zeng, Q. RAn, Y. Zhou, P. Zhou, J. Chen, J. J. looR, G. wAng, And X. dong. 2024. Inoculation of newborn lambs with ruminal solids derived from adult goats reprograms the development of gut microbiota and serum metabolome and favors growth performance. Journal of Agricultural and Food Chemistry 72:983–998. godoy-vitoRino, F., K. C. goldfARb, U. kARAoZ, S. leAl, M. A. gARCiA-AmAdo, P. hugenholtZ, S. G. tRinge, E. L. bRodie, And M. G. domingueZ-bello. 2012. Comparative analyses of foregut and hindgut bacterial communities in hoatzins and cows. The ISME Journal 6:531–541. gRond, K., C. C. kuRtZ, J. hAtton, M. M. sonsAllA, And K. N. duddleston. 2021. Gut microbiome is affected by gut region but robust to host physio- logical changes in captive active- season ground squirrels. Animal Microbiome 3:1-11. hAgeRmAn, A. E., And C. T. Robbins. 1993. Specificity of tannin-binding salivary proteins relative to diet selection by mammals. Canadian Journal of Zoology 71:628–633. hAmilton, G. D., P. D. dRysdAle, And D. L. euleR. 1980. Moose winter browsing patterns on clear-cuttings in northern Ontario. Canadian Journal of Zoology 58:1412–1416. hAyes, M., And S. bleAkley. 2018. 21 - Peptides from plants and their applica- tions. Pages 603–622 Peptide Applications in Biomedicine, Biotechnology and ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 140 Bioengineering. in S. Koutsopoulos, editor. Woodhead Publishing. heRms, D. A., And W. J. mAttson. 1992. The dilemma of plants: To grow or defend. The Quarterly Review of Biology 67:283–335. hofmAnn, R. R. 1989. Evolutionary steps of ecophysiological adaptation and diversi- fication of ruminants: a comparative view of their digestive system. Oecologia 78:443–457. hofmAnn, R. R., And K. nygRen. 1992. Morphophysiological specialization and adaptation of the moose digestive sys- tem. Alces 1:91–100. holmAn, D. B., And K. E. gZyl. 2019. A meta-analysis of the bovine gastrointesti- nal tract microbiota. FEMS Microbiology Ecology 95:1–9. hooveR, W. H. 1978. Digestion and Absorption in the Hindgut of Ruminants. Journal of Animal Science 46:1789–1799. hoy, S., J. S. foRbey, D. P. melody, L. M. vuCetiCh, R. O. peteRson, K. B. koitZsCh, L. O. koitZsCh, A. L. von duyke, J. J. hendeRson, G. L. pARikh, And J. A. vuCetiCh. 2022. The nutri- tional condition of moose co-varies with climate, but not with density, pre- dation risk or diet composition. Oikos 2022:1-13. ishAq, S. L., C. J. kim, D. Reis, And A.-D. G. wRight. 2015. Fibrolytic bacteria iso- lated from the rumen of North American moose (Alces alces) and their use as a probiotic in neonatal Lambs. PLOS ONE 10:1–25. ishAq, S. L., And A.-D. wRight. 2014. High- Throughput DNA sequencing of the ruminal bacteria from moose (Alces alces) in Vermont, Alaska, and Norway. Microbial Ecology 68:185–195. ishAq, S. L., And A.-D. G. wRight. 2012. Insight into the bacterial gut microbiome of the North American moose (Alces alces). BMC Microbiology 12:1–12. jiAng, X., B. ZhAng, F. lAn, C. Zhong, J. jin, X. li, Q. Zhou, J. li, N. yAng, C. wen, And C. sun. 2023. Host genetics and gut microbiota jointly regulate blood bio- chemical indicators in chickens. Applied Microbiology and Biotechnology 107:7601–7620. kARtZinel, T. R., P. A. Chen, T. C. Coverdale, D. L. Erickson, W. J. Kress, M. L. Kuzmina, D. I. Rubenstein, W. Wang, And R. M. Pringle. 2015. DNA metabar- coding illuminates dietary niche parti- tioning by African large herbivores. Proceedings of the National Academy of Sciences 112:8019–8024. kohl, K. D., A. W. milleR, J. E. mARvin, R. mACkie, And M. D. deARing. 2014. Herbivorous rodents (Neotoma spp.) har- bour abundant and active foregut micro- biota. Environmental Microbiology 16:2869–2878. kong, Y., R. teAtheR, And R. foRsteR. 2010. Composition, spatial distribution, and diversity of the bacterial communities in the rumen of cows fed different forages: Effect of diet on the rumen microbial composition. FEMS Microbiology Ecology 74:612–622. kuZnetsovA, A., P. bRuun bRoCkhoff, And R. hAubo bojesen ChRistensen. 2020. lmerTest: Tests in Linear Mixed Effects Models. https://cran.r-project.org/web/ packages/lmerTest/index.html Accessed 21 Nov 2024. Lenth, R. V., B. Banfai, B. Bolker, P. Buerkner, I. Giné-Vázquez, M. Herve, M. Jung, J. Love, F. Miguez, J. Piaskowski, H. Riebl, And H. Singmann. 2024. emmeans: Estimated Marginal Means, aka Least- Squares Means. https://cran.r-project.org/ web/packages/emmeans/index.html. Accessed 21 Nov 2024. ley, R. E., M. hAmAdy, C. loZupone, P. J. tuRnbAugh, R. R. RAmey, J. S. biRCheR, M. L. sChlegel, T. A. tuCkeR, M. D. sChRenZel, R. knight, And J. I. goRdon. 2008. Evolution of mammals and their gut microbes. Science 320:1647–1651. looney, C., And S. D. eigenbRode. 2012. Characteristics and distribution of https://cran.r-project.org/web/packages/lmerTest/index.html https://cran.r-project.org/web/packages/lmerTest/index.html https://cran.r-project.org/web/packages/emmeans/index.html https://cran.r-project.org/web/packages/emmeans/index.html ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 141 Palouse Prairie remnants: implications for conservation planning. Natural Areas Journal 32:75–85. mAgne, F., M. gottelAnd, L. gAuthieR, A. ZAZuetA, S. pesoA, P. nAvARRete, And R. bAlAmuRugAn. 2020. The Firmicutes/ Bacteroidetes ratio: a relevant marker of gut dysbiosis in obese patients? Nutrients 12:1–17. mAniChAnh, C., J. ReedeR, P. gibeRt, E. vARelA, M. llopis, M. Antolin, R. guigo, R. knight, And F. guARneR. 2010. Reshaping the gut microbiome with bacterial transplantation and anti- biotic intake. Genome Research 20:1411–1419. mAo, S., M. ZhAng, J. liu, And W. Zhu. 2015. Characterizing the bacterial microbiota across the gastrointestinal tracts of dairy cattle: membership and potential func- tion. Scientific Reports 5:1–14. mCARt, S. H., D. E. spAlingeR, W. B. Collins, E. R. sChoen, T. stevenson, And M. buCho. 2009. Summer dietary nitrogen availability as a potential bottom-up con- straint on moose in south-central Alaska. Ecology 90:1400–1411. mCdonAld, J. H. 2009. Handbook of Biological Statistics. 2nd edition. Sparky House Publishing, Baltimore, Maryland, U.S.A. mCmuRdie, P. J., And S. holmes. 2013. phy- loseq: An R package for reproducible interactive analysis and graphics of microbiome census data. PLOS ONE 8:1–11. mooRe, B. D., N. L. wiggins, K. J. mARsh, M. D. deARing, And W. J. foley. 2015. Translating physiological signals to changes in feeding behaviour in mam- mals and the future effects of global climate change. Animal Production Science 55:272–283. newbold, C. J., And E. RAmos-moRAles. 2020. Review: Ruminal microbiome and microbial metabolome: effects of diet and ruminant host. Animal 14:s78–s86. niehAus, A. 2009. Rumenotomy and Rumenostomy. Pages 27–29 in. Food Animal Practices. 5th edition. oksAnen, J., G. L. simpson, F. G. blAnChet, R. kindt, P. legendRe, P. R. minChin, R. B. o’hARA, P. solymos, M. H. H. stevens, E. sZoeCs, H. wAgneR, M. bARbouR, M. bedwARd, B. bolkeR, D. boRCARd, G. CARvAlho, M. ChiRiCo, M. D. CACeRes, S. duRAnd, H. B. A. evAngelistA, R. fitZjohn, M. fRiendly, B. fuRneAux, G. hAnnigAn, M. O. hill, L. lAhti, D. mCglinn, M.-H. ouellette, E. R. CunhA, T. smith, A. stieR, C. J. F. T. bRAAk, And J. weedon. 2024. vegan: Community Ecology Package. https:// cran.r-project.org/web/packages/vegan/ index.html. Accessed 15 May 2024. pAlo, R. T. 1984. Distribution of birch (Betula spp.), willow (Salix spp.), and poplar (Populus spp.) secondary metab- olites and their potential role as chemi- cal defense against herbivores. Journal of Chemical Ecology 10:499–520. peltieR, T. C. 2015. Moose management report and plan, Game Management Units 16A and 16B: Report period 1 July 2010-30 June 2015, and plan period 1 July 2015-30 June 2020. pinnell, L. J., A. A. Reyes, C. A. wolfe, M. D. weinRoth, J. L. metCAlf, R. J. delmoRe, K. E. belk, P. S. moRley, And T. E. engle. 2022. Bacteroidetes and fir- micutes drive differing microbial diver- sity and community composition among micro-environments in the bovine rumen. Frontiers in Veterinary Science 9:1–12. pRovenZA, F. D., And D. F. bAlph. 1987. Diet learning by domestic ruminants: Theory, evidence and practical implica- tions. Applied Animal Behaviour Science 18:211–232. RibAs, M. P., M. gARCíA-ulloA, J. espunyes, And O. CAbeZón. 2023. Improving the assessment of ecosystem and wildlife health: microbiome as an early indicator. Current Opinion in Biotechnology 81:1–8. https://cran.r-project.org/web/packages/vegan/index.html https://cran.r-project.org/web/packages/vegan/index.html https://cran.r-project.org/web/packages/vegan/index.html ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 142 RiChARds, L. A., A. E. glAssmiRe, K. M. oChsenRideR, A. M. smilAniCh, C. D. dodson, C. S. jeffRey, And L. A. dyeR. 2016. Phytochemical diversity and syner- gistic effects on herbivores. Phytochemistry Reviews 15:1153–1166. Robbins, C. T. 1994. Wildlife Feeding and Nutrition. 2nd edition. Academic Press. RosenbeRg, E., And I. ZilbeR-RosenbeRg. 2016. Microbes Drive Evolution of Animals and Plants: the Hologenome Concept. R. J. Collier, editor. mBio 7:5–15. Russell, J. B. 2002. Rumen Microbiology and Its Role in Ruminant Nutrition. James B. Russell, Ithaca, NY. sChRempp, T. V., J. L. RAChlow, T. R. johnson, L. A. shipley, R. A. long, J. L. AyCRigg, And M. A. huRley. 2019. Linking forest management to moose population trends: The role of the nutritional landscape. PLOS ONE 14:e0219128. shipley, L. A. 2010. Fifty years of food and foraging in moose: Lessons in ecology from a model herbivore. Alces 46:1–13. solden, L. M., D. W. hoyt, W. B. Collins, J. E. plAnk, R. A. dAly, E. hildebRAnd, T. J. beAveRs, R. wolfe, C. D. niCoRA, S. O. puRvine, M. Carstensen, M. S. lipton, D. E. spAlingeR, J. L. fiRkins, B. A. wolfe, And K. C. wRighton. 2017. New roles in hemicellulosic sugar fermentation for the uncultivated Bacteroidetes family BS11. The ISME Journal 11:691–703. solden, L. M., A. E. nAAs, S. Roux, R. A. dAly, W. B. Collins, C. D. niCoRA, S. O. puRvine, D. W. hoyt, J. sChüCkel, B. jøRgensen, W. willAts, D. E. spAlingeR, J. L. fiRkins, M. S. lipton, M. B. sullivAn, P. B. Pope, And K. C. Wrighton. 2018. Interspecies cross-feeding orchestrates carbon degradation in the rumen ecosys- tem. Nature Microbiology 3:1274–1284. söllingeR, A., A. T. tveit, M. poulsen, S. J. noel, M. bengtsson, J. beRnhARdt, A. L. fRydendAhl hellwing, P. lund, K. Riedel, C. sChlepeR, O. højbeRg, And T. uRiCh. 2018. Holistic assessment of rumen micro- biome dynamics through quantitative metatranscriptomics reveals multifunc- tional redundancy during key steps of anaerobic feed degradation. mSystems 3:10.1128/msystems.00038-18. sonsthAgen, S. A., C. V. jAy, R. S. CoRnmAn, A. S. fisChbACh, J. M. gRebmeieR, And S. L. tAlbot. 2020. DNA metabarcod- ing of feces to infer summer diet of Pacific walruses. Marine Mammal Science 36:1196–1211. spAeth, D. F., R. T. bowyeR, T. R. stephenson, P. S. bARboZA, And V. vAn bAllenbeRghe. 2002. Nutritional quality of willows for moose: effects of twig age and diameter. Alces 38:143–154. spAlingeR, D. E., W. B. Collins, T. A. Hanley, N. E. Cassara, And A. M. Carnahan. 2010. The impact of tannins on protein, dry matter, and energy diges- tion in moose (Alces alces). Canadian Journal of Zoology 88:977–987. spRingeR, A., C. fiChtel, G. A. Al-ghAlith, F. koCh, K. R. AmAto, J. B. ClAyton, D. knights, And P. M. kAppeleR. 2017. Patterns of seasonality and group mem- bership characterize the gut microbiota in a longitudinal study of wild Verreaux’s sifakas (Propithecus verreauxi). Ecology and Evolution 7:5732–5745. stevens, C. E., And I. D. hume. 1998. Contributions of microbes in vertebrate gastrointestinal tract to production and conservation of nutrients. Physiological Reviews 78:393–427. svARtstRöm, O., J. AlnebeRg, N. teRRApon, V. lombARd, I. de bRuijn, J. mAlmsten, A.-M. dAlin, E. EL mulleR, P. shAh, P. wilmes, B. henRissAt, H. AspeboRg, And A. F. AndeRsson. 2017. Ninety-nine de novo assembled genomes from the moose (Alces alces) rumen microbiome provide new insights into microbial plant biomass degradation. The ISME Journal 11:2538–2551. testA, J. W. 2004. Population dynamics and life history trade-offs of moose (Alces alces) in south-central Alaska. Ecology 85:1439–1452. ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 143 tong, J., H. ZhAng, D. yAng, Y. ZhAng, B. xiong, And L. jiAng. 2018. Illumina sequencing analysis of the ruminal micro- biota in high-yield and low-yield lactat- ing dairy cows. PLOS ONE 13:e0198225. tRemblAy, J.-P., A. A. Royo, É. ChAmpAgne, And P. RAymond. 2019. Phytochemicals involved in plant resistance to leporids and cervids : a systematic review. Journal of Chemical Ecology 46:84–98 tRoyeR, K. 1984. Microbes, herbivory and the evolution of social behavior. Journal of Theoretical Biology 106:157–169. u.s. ClimAte noRmAls. 2021. National Centers for Environmental Information (NCEI). https://www.ncei.noaa.gov/prod- ucts/land-based-station/us-climate- normals. Accessed 8 Aug 2025. vAn soest, P. 1994. Nutritional Ecology of the Ruminant. Second edition. Cornell University Press. vAn soest, P. J. 1996. Allometry and ecol- ogy of feeding behavior and digestive capacity in herbivores: A review. Zoo Biology 15:455–479. vos, W. M. de, H. tilg, M. V. hul, And P. D. CAni. 2022. Gut microbiome and health: mechanistic insights. Gut 71:1020–1032. weimeR, P. J. 2015. Redundancy, resilience, and host specificity of the ruminal microbiota: implications for engineering improved ruminal fermentations. Frontiers in Microbiology 6:1–16. WeimeR, P. J. 2022. Degradation of cellulose And hemicellulose by ruminal microor- ganisms. Microorganisms 10:1–30. Wu, X., Q. Wei, X. WAng, Y. ShAng, And H. ZhAng. 2022. Evolutionary and dietary relationships of wild mammals based on the gut microbiome. Gene 808:0378–1119. ZhAo, T., H. Y. HuAng, R. Q. Mu, M. D. Lu, Z. M. Geng, S. M. Hu, W. W. MA, And C. Zhou. 2025. Clostridia enhances intestinal immunity to alle- viate ovalbumin-induced allergy. International Archives of Allergy and Immunology. Zou, H., R. Hu, Z. WAng, A. M. ShAh, S. Zeng, Q. Peng, B. Xue, L. WAng, X. ZhAng, X. WAng, J. Shi, F. Li, And L. Zeng. 2019. Effects of nutritional deprivation and re-alimentation on the feed efficiency, blood biochemis- try, and rumen microflora in yaks (Bos grunniens). Animals 9:807. https://www.ncei.noaa.gov/products/land-based-station/us-climate-normals https://www.ncei.noaa.gov/products/land-based-station/us-climate-normals https://www.ncei.noaa.gov/products/land-based-station/us-climate-normals ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE ALCES VOL. 60, 2024 144 APPENDIX Table A1. Sample information moose sampled from the rumen, cecum, and colon in Alaska (n = 5) and the Inland NW (n = 7, eastern Washington and western Idaho), United States, from the fall and winter seasons of 2022–2024. Geographic Region Sample ID Closest City Date Collected Sex Age GI Organ Sampled Alaska AK1 Anchorage 1/24/2022 Female Adult Rumen Alaska AK1 Anchorage 1/24/2022 Female Adult Cecum Alaska AK2 Palmer 1/14/2022 Female Calf Rumen Alaska AK2 Palmer 1/14/2022 Female Calf Cecum Alaska AK3 Wasilla 12/8/2022 Female Calf Rumen Alaska AK3 Wasilla 12/8/2022 Female Calf Cecum Alaska AK3 Wasilla 12/8/2022 Female Calf Fecal Alaska AK4 Wasilla 1/28/2022 Female Adult Rumen Alaska AK4 Wasilla 1/28/2022 Female Adult Cecum Alaska AK4 Wasilla 1/28/2022 Female Adult Fecal Alaska AK5 Palmer 2/19/2022 Female Adult Rumen Inland NW INW1 Palmer 2/19/2022 Female Adult Fecal Inland NW INW1 Moscow 9/23/2022 Female Adult Rumen Inland NW INW1 Moscow 9/23/2022 Female Adult Cecum Inland NW INW1 Moscow 9/23/2022 Female Adult Fecal Inland NW INW2 Spokane 10/4/2022 Male Adult Rumen Inland NW INW2 Spokane 10/4/2022 Male Adult Cecum Inland NW INW2 Spokane 10/4/2022 Male Adult Fecal Inland NW INW3 Spokane 10/12/2022 Male Adult Rumen Inland NW INW3 Spokane 10/12/2022 Male Adult Cecum Inland NW INW3 Spokane 10/12/2022 Male Adult Fecal Inland NW INW4 Spokane 11/28/2022 Female Adult Rumen Inland NW INW4 Spokane 11/28/2022 Female Adult Fecal Inland NW INW5 Mt. Spokane 3/20/2023 Female Adult Rumen Inland NW INW5 Mt. Spokane 3/20/2023 Female Adult Cecum Inland NW INW5 Mt. Spokane 3/20/2023 Female Adult Fecal Inland NW INW6 Moscow/Troy 10/25/2023 Female Adult Rumen Inland NW INW6 Moscow/Troy 10/25/2023 Female Adult Cecum Inland NW INW6 Moscow/Troy 10/25/2023 Female Adult Fecal Inland NW INW7 Spokane 11/12/2024 Male Adult Rumen Inland NW INW7 Spokane 11/12/2024 Male Adult Cecum Inland NW INW7 Spokane 11/12/2024 Male Adult Fecal ALCES VOL. 60, 2024 ANDERSON AND SHIPLEY • THE GASTROINTESTINAL TRACT OF MOOSE 145 Table A2. Top 20 microbial families for moose sampled from the combined rumen, cecum, and colon in Alaska (n = 5) and the Inland NW (n = 7) eastern Washington and western Idaho), United States, from the fall and winter seasons, 2022–2024. Families are listed with their mean abundance and standard error. Asterisks denote significant differences between the two geographic regions, α = 0.10. Family Alaska Mean Abundance SE Inland NW Mean Abundance SE Enterobacteriaceae* 28.49 31.39 3.39 3.32 Oscillospiraceae* 15.49 8.57 20.95 2.03 Lachnospiraceae* 10.88 7.46 17.06 3.85 Clostridia_UCG-014 9.04 10.42 3.08 1.10 Christensenellaceae 8.31 5.73 13.12 2.66 Peptostreptococcaceae 4.86 3.19 4.86 3.19 Rikenellaceae 4.1 2.58 5.99 1.78 Prevotellaceae 3.92 2.65 6.14 1.34 Spirochaetaceae 3.42 3.31 3.42 3.31 Ruminococcaceae 2.92 2.19 4.28 2.25 Anaerovoracaceae 2.64 1.11 2.64 1.11 [Eubacterium]_coprostanoligenes_group 1.81 1.06 2.13 0.68 UCG-010 0 0 1.97 0.60 Atopobiaceae 1.44 1.21 1.44 1.21 Erysipelotrichaceae 1.44 0.84 1.44 0.84 PeH15 1.41 1.54 0 0 F082 1.38 2.01 2.28 2.72 Bacteroidaceae 1.35 1.20 2.23 0.88 Eggerthellaceae 1.33 0.55 1.33 0.55 Streptococcaceae 1.30 0.96 1.30 0.96 RF39 1.17 0.70 0 0 p-251-o5 0 0 0.96 0.24