Simple Summary: Manipulating dietary fatty acid composition is a feasible approach to improve the mutton fatty acid profiles of feedlot fattening goats. A previous study showed that flaxseed oil and grain affected the concentrations of c18:3n3, c20:5n3, c22:6n3, and n-3PUFA in the muscle tissues of Albas cashmere goats differently. The intestinal microflora may be involved in the regulation of blood lipid levels and their accumulation, as it plays an important role in the host's metabolism. The specific results showed that the effects of flaxseed oil and grain on gut microbiota diversity vary in different segments in Albas cashmere goats. And flaxseed grain is more efficient than flaxseed oil in protecting intestinal health and promoting the absorption of c18:3n3. The present study investigated the effects of flaxseed oil or flaxseed grain on the intestinal microbiota and blood fatty acid profiles of Albas cashmere goats. Sixty kid goats were allocated to three treatments and fed for 90 days with a control treatment, comprising a basal diet (CON, total-mixed ration with flaxseed meal), or experimental treatments, comprising a basal diet with added flaxseed oil (LNO) and a basal diet with added heated flaxseed grain (HLS). On day 90, two goats were randomly selected from each pen (eight goats per treatment) for euthanizing; then, five of the eight goats were randomly selected, and we collected their intestinal (duodenum, jejunum, ileum, cecum, and colon) digesta for analysis of the bacteria community. The results indicated that Firmicutes are the most predominant phylum in different segments of the intestinal digesta. Compared with the CON group, the relative abundance of duodenal Firmicutes, jejunal Saccharibacteria, and Verrucomicrobia significantly decreased in the LNO and HLS groups (p < 0.05), but there was no significant difference between the LNO and HLS groups. Compared with the CON and HLS groups, the RA of duodenal and jejunal Proteobacteria remarkably increased in the LNO group (p < 0.05), and there was no significant difference between the CON and HLS groups. Compared with the CON and LNO groups, the RA of Actinobacteria remarkably increased in the small intestine of the HLS group (p < 0.05), but there was no significant difference between the CON and LNO groups in the duodenum and ileum. The results of linear discriminant analysis (LDA) effect size (LEfSe) analysis showed that the HLS group was characterized by a higher RA of the [Eubacterium]_coprostanoligenes_group in the small intestine and the LNO group was represented by a higher RA of the Lachnospiraceae_NK3A20_group in the cecum and colon, while the CON group was represented by a higher RA of Solobacterium, Pseudoramibacter, and Acetitomaculum in the small intestine and a higher RA of norank_o__Bradymonadales, the Prevotellaceae_Ga6A1_group, and Ruminiclostridium_1 in the cecum and colon. In conclusion, the addition of flaxseed oil and grain rich in c18:3n3 to the diet could reduce the microbial diversity of the small intestinal segments and the microbial diversity and richness of the cecum and colon in Albas cashmere goats. And flaxseed grain is more efficient than flaxseed oil in protecting intestinal health and promoting the absorption of c18:3n3.
Keywords: grain; goat; different intestinal segments; bacteria; n-3 poly-unsaturated fatty acids
Low levels of saturated fatty acids (SFAs) and high levels of n-3poly-unsaturated FAs (n-3PUFAs) in the diet are beneficial to human health [[
The intestinal microbiota primarily influences the host by providing essential nutrients and non-nutrients, enhancing the host's ability to obtain nutrients through the production of gastrointestinal enzymes, the modification of intestinal histology, and the creation of a physical barrier against pathogens [[
In view of this, the effects of flaxseed oil and flaxseed supplementation on the intestinal microbiota of Albas cashmere goats should be investigated, and studies should further explore whether the shift in the blood fatty acid profiles of cashmere goats in response to dietary supplementation with flaxseed oil or grain is related to the change in intestinal microbiota composition and explain the possible mechanisms by which flaxseed oil and grain ameliorate blood fatty acid profiles.
This study was conducted at the Inner Mongolia White Cashmere Goat Breeding Farm, Wulan Town, Etuoke Banner, Ordos City, Inner Mongolia Autonomous Region, China (39°12′ N; 107°97′ E). All animal procedures were performed in accordance with the National Standard Guidelines for Ethical Review of Animal Welfare (GB/T 35892-2018) [[
Sixty 4-month-old, castrated Albas white cashmere male kid goats (average body weight 18.6 ± 0.1 kg) were selected and randomly divided into three treatments, with each treatment comprising four pens of five kid goats. The control treatment group (CON) was fed the total-mixed ration (TMR) with no supplementation. The experimental treatment group was fed the TMR with added flaxseed oil (LNO). In the second experimental treatment group, heated flaxseed grain (HLS; flaxseed contains about 36% oil and was roasted for 10 min at 120 °C) was added to the TMR, with the same flaxseed oil content as in the LNO treatment. The diet was formulated according to the nutritional requirements for meat goats [[
On day 90, two goats were randomly selected from each pen (eight goats per group) for euthanizing, and then five of the eight goats were randomly selected for microbial community analysis. Before slaughter, the goats fasted for 24 h and were prohibited from drinking water for 2 h. After slaughter, the intestine was opened and the duodenum, large intestine, and colon were separated using a suture line to avoid reflux between adjacent areas of the gastrointestinal tract. The digesta samples were collected separately and homogenized. Finally, homogenized samples from each segment of the digestive tract were frozen in liquid nitrogen and then stored at −80 °C for analysis of microbial composition.
Seventy-five digesta samples were thawed at 4 °C and kept on ice throughout the extraction process. Microbial DNA was extracted using the E.Z.N.A.
The V3-V4 regions of the bacterial 16S rRNA genes were amplified in triplicate using PCR (95 °C for 3 min of denaturation, followed by 27 cycles at 95 °C for 30 s, 55 °C for 30 s of annealing, 72 °C for 45 s of elongation, and a final extension at 72 °C for 10 min). The universal primers used in this amplification protocol were: 338F (5′-ACCHOCTACGGGAGGCAGCAG-3′) and 806R (5′-GGACTACHVGGGTWCHOTAAT-3′). The PCR reactions were conducted in a 20 μL mixture containing: 4 μL of 5 × FastPfu Buffer, 2 μL of 2.5 mM dNTPs (Deoxynucleotide Triphosphates), 0.8 μL of each primer (5 μM), 0.4 μL of FastPfu Polymerase, and 10 ng of template DNA. The PCR products were excised from a 2% agarose gel. Concerning purification, an AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, USA) was used and quantified using QuantiFluor™-ST (Promega, Madison, WI, USA) according to the instruction manual.
Purified amplicons were pooled in equimolar and paired-end sequenced (2 × 300 bp) on an Illumina MiSeq PE300 instrument (Illumina, San Diego, CA, USA) according to the standard protocols of Majorbio Bio-Pharm Technology Co. Ltd. (Shanghai, China).
Raw reads of each sample were demultiplexed and quality filtered using default parameters in Quantitative Insights into Microbial Ecology through QIIME (Quantitative Insights into Microbial Ecology, version 1.9.1) software, quality filtered using Trimmomatic, and merged using FLASH (Fast Length Adjustment of Short Reads). Low-quality reads were removed according to the following criteria: (i) The reads were truncated at any site receiving an average quality score <20 over a 50 bp sliding window and truncated reads shorter than 50 bp were discarded; (ii) reads with 2 nucleotide mismatches in primer matching or that contained ambiguous characters were removed; (iii) sequences that overlapped by at least 10 bp were assembled based on their overlap sequences. The assembled sequences were assigned to operational taxonomic units (OTUs), which are the most used microbial diversity units, were clustered with a 97% similarity cutoff using UPARSE (Highly Accurate OTU Sequences from Microbial Amplicon Reads, version 7.1,
The bacterial diversity indexes and the relative abundance of bacteria at the phylum level were analyzed in SAS (SAS Inst. Inc., Cary, NC, USA). The data were checked for normality of variance. The data obeying normal distribution were still analyzed using a one-way analysis of variance (ANOVA), and Duncan's multiple range tests were carried out. The data that disobeyed normal distribution were analyzed using the Kruskal–Wallis test. The results were presented as the mean values and standard error of the mean (SEM). Data means significance was declared at p ≤ 0.05 and tendencies were considered at 0.05 < p ≤ 0.10; there was no significance at p > 0.10. The differences in microbial community abundance at the genus level between the groups, and the effects of each differentially abundant taxon, were assessed using the non-parametric factorial Kruskal–Wallis sum-rank test and the linear discriminant analysis (LDA) effect size (LEfSe) method, which emphasized statistical significance and biological correlation. The threshold was set at an LDA level of > 2, p < 0.05. Spearman correlation was used to correlate the blood fatty acid profiles with the differential bacterial genera through CCREPE software (version 1.7.0); the data for the blood fatty acid profiles were obtained from our previous results [[
In the present study, fifteen digesta samples were collected and analyzed for three different treatments in each region of the intestine. To facilitate unified analysis, the results of the 15 samples from the three different treatments were normalized according to the minimum sequence in one sample. A total of 4,111,507 optimized sequences were obtained after quality filtering with an average of (54,820 ± 6500) optimized sequences per sample (Table 2). As shown in Figure 1, the individual-based rarefaction curves generated for each sample were sufficient to accurately describe the bacterial composition of intestinal digesta. The results showed that the sampling depth was adequate to estimate bacterial community.
As indicated in Table 3, the coverage (the coverage estimator) index of the intestinal digesta in all groups was 0.991–0.999, which indicated that the sequencing data were sufficiently representative. In the duodenum and ileum, compared with the LNO group, the Sobs (the richness estimator), Ace (the richness estimator), and Chao (the richness estimator) indexes significantly increased in the CON and HLS groups (p < 0.05), but there was no significant difference between CON and HLS groups. In the jejunum, compared with the HLS group, the Sobs, Shannon, Ace, and Chao indexes significantly increased in the CON and LNO groups (p < 0.05), but the Simpson (the diversity estimator) index remarkably decreased, and there was no significant difference between the CON and LNO groups. In the cecum and colon, compared with the CON group, the Sobs, Shannon, Ace, and Chao indexes significantly decreased in the HLS and LNO groups (p < 0.05), but the Simpson index remarkably increased, and there was no significant difference between the HLS and LNO groups. At the OTU level, the principal coordinate analysis (PCoA) plots (Figure 2) demonstrate dissimilarities between the CON group, the LNO group, and the HLS group (p ≤ 0.05).
As shown in Figure 3, a detailed overview of the bacterial composition of the intestinal digesta in each sample was illustrated at the phylum level. The Firmicutes are the most predominant phylum in different segments of intestinal digesta. As shown in Table 4, compared with the CON group, the relative abundance (RA) of duodenal Firmicutes, jejunal Saccharibacteria, and Verrucomicrobia decreased in the LNO and HLS groups (p < 0.05), but there was no significant difference between the LNO and HLS groups. Compared with the CON and HLS groups, the RA of duodenal and jejunal Proteobacteria and colonic Spirochaetae increased in the LNO group (p < 0.05); the duodenal Saccharibacteria's RA showed the opposite result (p < 0.05), and there was no significant difference between the CON and HLS groups. Compared with the CON and LNO groups, the RA of duodenal Actinobacteria and Bacteroidetes, ileal Actinobacteria, and cecal Verrucomicrobia increased in the HLS group (p < 0.05); the RA of ileal Firmicutes, Proteobacteria, Tenericutes and colonic Tenericutes showed the opposite result (p < 0.05), but there was no significant difference between the CON and LNO groups. In the jejunum, the RAs of Actinobacteria in the LNO group, CON group, and HLS group were increased (p < 0.0001). In the cecum, the Firmicutes' RA in the HLS group is remarkably higher than in the LNO group (p = 0.027), but the CON group did not differ from the LNO and HLS groups. In the colon, the RAs of Firmicutes in the LNO group, HLS group, and CON group were decreased (p = 0.009).
LEfSe (LDA > 2) was used to investigate the microbial genus that the differences between the LNO, HLS, and CON groups. As shown in Figure 4, differential expression analysis showed that the RAs of multiple genera in the duodenum, jejunum, ileum, cecum, and colon were significantly different between the LNO, HLS, and CON groups. In the duodenum (Figure 4A), 15 genera were significantly enriched in the HLS group compared to the CON and LNO groups: Enterorhabdus, the [Eubacterium]_coprostanoligenes_group, Fusobacterium, Olsenella, Brevibacillus, Atopobium, Howardella, Pseudobutyrivibrio, the Lachnospiraceae_FE2018_group, Denitrobacterium, norank_f__Rhodocyclaceae, Mycoplasma, Pseudarthrobacter, Prevotella_7, unclassified_f__Erysipelotrichaceae; three genera were remarkably enriched in the LNO compared to the CON and HLS groups: Roseburia, Succinimonas, unclassified_f__Prevotellaceae; and 15 genera were remarkably enriched in the CON compared to the LNO and HLS groups: the Lachnospiraceae_XPB1014_group, Catenisphaera, unclassified_f__Coriobacteriaceae, Anaerofustis, Marvinbryantia, Solobacterium, norank_c__Cyanobacteria, Senegalimassilia, Lachnospiraceae_UCG-002, Pseudoramibacter, Ruminiclostridium, Acetitomaculum, norank_f__Coriobacteriaceae, Staphylococcus, Tyzzerella_3.
In the jejunum, five genera were more abundant in the HLS group than in the LNO and CON groups: the [Eubacterium]_coprostanoligenes_group, Actinobacillus, Atopobium, Enterococcus, Erysipelotrichaceae_UCG-009; 12 genera were more abundant in the LNO group than in the HLS and CON groups: the Lachnospiraceae_XPB1014_group, Selenomonas_1, Ruminococcaceae_UCG-005, Ruminococcaceae_UCG-004, Phocaeicola, Butyrivibrio_2, Domibacillus, the [Eubacterium]_ruminantium_group, Lachnoclostridium_10, the Rikenellaceae_RC9_gut_group, Anaerovibrio, Psychrobacillus; and 22 genera were more abundant in the CON group than in the HLS and LNO groups: Enterorhabdus, unclassified_o__Bacteroidales, unclassified_p__Proteobacteria, unclassified_f__Coriobacteriaceae, Anaerofustis, Marvinbryantia, Solobacterium, Anaerovorax, Candidatus_Saccharimonas, Saccharofermentans, Family_XIII_UCG-002, Family_XIII_UCG-001Blautia, Senegalimassilia, Pseudoramibacter, Catenisphaera, Acetitomaculum, Terribacillus, norank_f__Coriobacteriaceae, the [Eubacterium]_cellulosolvens_group, norank_o__Mollicutes_RF9, Lachnospiraceae_UCG-002 (Figure 4B).
In the ileum, two genera were significantly enriched in the HLS group compared to the CON and LNO groups: the [Eubacterium]_coprostanoligenes_group, Rhodococcus; one genera was remarkably enriched in the LNO compared to CON and HLS groups: the Christensenellaceae_R-7_group; and five genera were remarkably enriched in the CON compared to the LNO and HLS groups: Pseudoramibacter, unclassified_f__Erysipelotrichaceae, Solobacterium, Acetitomaculum, Erysipelotrichaceae_UCG-009 (Figure 4C).
In the cecum, three genera were more abundant in the HLS group than in the LNO and CON groups: Aeriscardovia, Barnesiella, Ruminococcaceae_UCG-013; four genera were more abundant in the LNO group than in the HLS and CON groups: the Lachnospiraceae_NK3A20_group, Streptococcus, Mogibacterium, the [Eubacterium]_brachy_group; and eight genera were more abundant in the CON group than in the HLS and LNO groups: the Prevotellaceae_Ga6A1_group, Ruminiclostridium_1, Tyzzerella, Anaerosporobacter, the norank_f__Bacteroidales_BS11_gut_group, norank_o__Bradymonadales, unclassified_o__Clostridiales, Ruminococcaceae_UCG-010 (Figure 4D).
In the colon, five genera were more abundant in the HLS group than in the LNO and CON groups: Roseburia, Atopobium, Bacteroides, Anaerotruncus, the Lachnospiraceae_NK4A136_group; four genera were more abundant in the LNO group than in the HLS and CON groups: the Lachnospiraceae_NK3A20_group, Rikenellaceae_RC9_gut_group, Olsenella, Treponema_2; and nine genera were more abundant in the CON group than in the HLS and LNO groups: Hydrogenoanaerobacterium, norank_f__Christensenellaceae, the Prevotellaceae_Ga6A1_group, Ruminiclostridium_1, Family_XIII_UCG-002, Ruminococcaceae_UCG-002, norank_o__Bradymonadales, Saccharofermentans, the norank_f__Bacteroidales_RF16_group (Figure 4E).
To further identify the shared genera in different intestinal segments, as shown in Figure 5, we found one shared genus ([Eubacterium]_coprostanoligenes_group) in the digesta samples of the duodenum, jejunum, and ileum, one shared genus (Atopobium) in the digesta samples of the duodenum, jejunum, and colon, and one shared genus (Enterorhabdus) in the digesta samples of the duodenum and jejunum, which were significantly increased in the HLS group compared to the CON and LNO groups. Of these, one shared genus (Rikenellaceae_RC9_gut_group) in the digesta samples of the jejunum and colon, and one shared genus (Lachnospiraceae_NK3A20_group) in the digesta samples of the cecum and colon, were significantly increased in the LNO group compared to the CON and HLS groups. Of these, three shared genera (Solobacterium, Pseudoramibacter, Acetitomaculum) in the digesta samples of the duodenum, jejunum and ileum, eight shared genera (Lachnospiraceae_UCG-002, the Lachnospiraceae_XPB1014_group, unclassified_f__Coriobacteriaceae, norank_f__Coriobacteriaceae, Marvinbryantia, Anaerofustis, Senegalimassilia, Catenisphaera) in the digesta samples of the duodenum and jejunum, two shared genera (Saccharofermentans, Family_XIII_UCG-002) in the digesta samples of the jejunum and colon, and three shared genera (norank_o__Bradymonadales, the Prevotellaceae_Ga6A1_group, Ruminiclostridium_1) in the digesta samples of the cecum and colon were significantly increased in the CON compared to the HLS and LNO groups.
Spearman correlation analysis was conducted between the differential bacterial genera in different intestinal tracts and blood fatty acid profiles, as indicated in Table 5, Table 6, Table 7, Table 8 and Table 9. The data set of the blood fatty acid profiles was obtained from our previous research [[
In the duodenum (Table 5), the [Eubacterium]_coprostanoligenes_group was positively associated with c18:3n3 and c20:5n3, but was negatively correlated with c20:4n6, Anaerofustis, Marvinbryantia, Senegalimassilia, and Solobacterium showed the opposite effect. Acetitomaculum, Denitrobacterium, Lachnospiraceae_UCG-002, and Pseudoramibacter were negatively correlated with c18:3n3 and c20:5n3. The [Eubacterium]_coprostanoligenes_group and unclassified_f__Erysipelotrichaceae were negatively correlated with c16:0. Brevibacillus was negatively correlated with c18:0. Marvinbryantia, Pseudarthrobacter, and Pseudoramibacter were positively associated with c18:1c9, but the [Eubacterium]_coprostanoligenes_group was negatively correlated with it. Pseudarthrobacter was negatively correlated with c18:2c6. norank_f__Coriobacteriaceae and unclassified_f__Coriobacteriaceae were negatively correlated with C18:3n3. The Lachnospiraceae_XPB1014_group was positively associated with c20:4n6, but Olsenella was negatively correlated with it. Enterorhabdus was negatively correlated with c20:5n3.
In the jejunum (Table 6), the [Eubacterium]_coprostanoligenes_group was positively associated with C18:3n3, but was negatively correlated with c20:4n6; Family_XIII_UCG-001 and norank_o__Mollicutes_RF9 showed the opposite effect. Anaerofustis, Catenisphaera, Lachnospiraceae_UCG-002, Pseudoramibacter, Saccharofermentans, Solobacterium, and unclassified_f__Coriobacteriaceae were negatively correlated with c18:3n3 and c20:5n3 but were positively associated with c20:4n6. The [Eubacterium]_cellulosolvens_group was positively associated with c18:1c9 but was negatively correlated with c20:5n3. Norank_f__Coriobacteriaceae was positively associated with c16:0 but was negatively correlated with C20:5n3. Anaerofustis and Catenisphaera were positively associated with c18_1c9. Marvinbryantia and Senegalimassilia were negatively correlated with c18:3n3. Blautia was positively associated with c20:4n6, but Enterococcus was negatively correlated with it.
In ileum (Table 7), unclassified_f__Erysipelotrichaceae, Solobacterium, Erysipelotrichaceae_UCG-009, and Acetitomaculum were negatively correlated with c18:3n3 and c20:5n3, but were positively associated with c20:4n6; the [Eubacterium]_coprostanoligenes_group showed the opposite effect. Solobacterium and Erysipelotrichaceae_UCG-009 were positively associated with c16:0. The [Eubacterium]_coprostanoligenes_group was positively associated with c22:6n3, but Christensenellaceae_R-7_group was negatively correlated with it.
In the cecum (Table 8), Ruminiclostridium_1 was negatively correlated with c18:3n3 and c20:5n3. Ruminococcaceae_UCG-010 was positively associated with c18:1c9. Prevotellaceae_Ga6A1_group was positively associated with c20:4n6, but was negatively correlated with c18:3n3 and c20:5n3. Barnesiella was positively associated with c18:0. Norank_o__Bradymonadales was negatively correlated with c20:5n3. The norank_f__Bacteroidales_BS11_gut_group was positively associated with c20:4n6. The [Eubacterium]_brachy_group and Mogibacterium were positively associated with c20:5n3.
In the colon (Table 9), Roseburia, Bacteroides, and Anaerotruncus were positively associated with c22:6n3. Ruminococcaceae_UCG-002 was positively associated with c18:2c6 and c20:4n6. Ruminiclostridium_1, norank_f__Bacteroidales_RF16_group, Prevotellaceae_Ga6A1_group, Saccharofermentans, norank_f__Christensenellaceae, Family_XIII_UCG-002, Hydrogenoanaerobacterium, and norank_o__Bradymonadales were negatively correlated with c18:3n3 and c20:5n3; except for norank_f__Christensenellaceae and norank_o__Bradymonadales, the other bacteria were positively correlated with c20:4n6.
At the phylum level, 50 bacteria phyla were detected in the intestinal digesta of the Albas cashmere goats. The number of bacterial phyla detected in the duodenum, jejunum, ileum, cecum, and colon of CON, LNO, and HLS goats were 39, 30, 33; 28, 34, 33; 18, 18, 21; 18, 18, 16; 15, 14, 16, respectively. As indicated in Figure 6, the small intestinal (duodenum, jejunum, and ileum) digesta were dominated by Firmicutes (50.30–79.98%), Actinobacteria (9.47–17.11%), and Proteobacteria (5.22–22.37%). The large intestinal digesta (cecum and colon) were dominated by Firmicutes (58.27–60.26%) and Bacteroidetes (33.62–34.50%).
At the genus level, a Venn daigram was used to evaluate the common and exclusive bacterial genus in the intestinal digesta of each group. The data indicated that 708, 649, 331, 299, and 259 genera were found in the digesta of the duodenum, jejunum, ileum, cecum, and colon, respectively, and 167 genera were found to be shared among the different intestinal tracts independent of the diet (Figure 7A). The data indicated that 511, 324, 211, 249, and 215 genera were found in the digesta of the duodenum, jejunum, ileum, cecum, and colon, respectively, and 115 genera were found to be shared among the different intestinal tracts in the CON group (Figure 7B). The data indicated that 379, 542, 206, 240, and 215 genera were found in the digesta of the duodenum, jejunum, ileum, cecum, and colon, respectively, and 124 genera were found to be shared among the different intestinal tracts in the LNO group (Figure 7C). The data indicated that 454, 321, 285, 206, and 212 genera were found in the digesta of the duodenum, jejunum, ileum, cecum, and colon, respectively, and 106 genera were found to be shared among the different intestinal tracts in the HLS group (Figure 7D).
At the OTU level, PCoA was used based on the Bray–Curtis dissimilarity matrices to estimate the bacterial community structure of each group in the intestine; the figure shows that the bacterial communities of the small intestine (duodenum, jejunum, and ileum) and large intestine (cecum and colon) were very different from each other (Figure 8A–D). Small intestine samples (duodenum, jejunum, and ileum) occupied the left side of PC1, while large intestine samples (cecum and colon) occupied the right side of PC1.
This study provided a detailed picture of bacterial community dynamics along the intestinal tract segments in cashmere goats, and the relationship between the significantly differential genera and blood FA profiles. The bacterial community richness was measured using the Sobs, Chao, and Ace indexes, and the diversity was measured using the Shannon and Simpson indexes [[
The intestinal microflora is a major regulator of host metabolism. The composition and function of the gut microbiota are dynamic and are influenced by diet, such as by the quantity and profile of fatty acids. Consequently, dietary fatty acids can influence host physiology by interacting with the gut microbiota. The n-3 PUFAs are mainly absorbed in the gut, where some microorganisms can directly utilize n-3 PUFAs and produce numerous small molecules [[
Acetomaculum is mainly found in ruminants fed with a concentrate-rich diet and can use monosaccharides to produce acetate [[
At the phylum level, independent of diet (that is, in the same treatment), Firmicutes were the most abundant bacteria in the intestine of cashmere goats, and the Firmicutes, Actinobacteria, and Proteobacteria were the predominant bacterial phyla in the small intestine; the Firmicutes and Bacteroides were the predominant bacterial phyla in cecum and colon (Figure 5). The number of the major and minor phyla of bacteria indicated that the large intestine may have lower diversity but a higher abundance of bacterial phyla, and the small intestine is occupied by numerous minor phyla that may participate in a higher number of processes [[
Feedlot fattening compromised the mutton quality, which could not meet the consumers' demand for high-quality meat products. Based on the results of this study and previous research by our group, adding flaxseed grain to the diet of cashmere goats under feedlot fattening would be an efficient way to protect the intestinal health of cashmere goats and improve the quality of mutton by enrichment of c18:3n3 in muscle and fat tissue.
The addition of flaxseed oil and grain rich in c18:3n3 to the diet could reduce the microbial diversity of the small intestinal segments, and the microbial diversity and richness of the cecum and colon of cashmere goats. A reduction in the RA of the duodenal Firmicutes phylum, the jejunal Verrucomicrobia phylum, and the harmful bacteria Solobacterium, Acetitomaculum, and Pseudoramibacter in the small intestine, plus the decrease in the RA of Ruminiclostridium_1 and the Prevotellaceae_ Ga6A1_group in the cecum and colon, implied that flaxseed oil and grain could effectively protect intestinal health and promote the absorption of c18:3n3 and n-3PUFA into blood. Compared to flaxseed oil, flaxseed grain increased the RA of the probiotic Actinobacteria phylum and the [Eubacterium]_coprostanoligenes_group in the small intestine of cashmere goats, as the grain is more helpful for the absorption of c18:3n3.
Graph: Figure 1 The OTU rarefaction curves of the intestinal digesta bacterial communities. Curves were drawn using the least-sequenced sample as the upper limit for the rarefactions. Each color represents one treatment: the red curves represent kids fed the basal diet (CON), the green curves represent kids fed the basal diet supplemented with flaxseed oil (LNO), and the blue curves represent kids fed the basal diet supplemented with heated flaxseed grain (HLS). (A): duodenum; (B): jejunum; (C): ileum; (D): cecum; (E): colon.
Graph: Figure 2 Principal coordinate analysis (PCoA, using the weighted Unifrac similarity metric) of bacterial operational taxonomic units (OTUs) in the intestinal digesta of kid goats. Each symbol represents one treatment: The solid red circle represents kids fed with the basal diet (CON), the solid blue triangle represents kids fed with the basal diet supplemented with flaxseed oil (LNO), and the solid green rhombus represents kids fed with the basal diet supplemented with heated flaxseed grain (HLS). (A): duodenum; (B): jejunum; (C): ileum; (D): cecum; (E): colon.
Graph: Figure 3 The bar chart shows the phylum-level bacterial composition of intestinal digesta. The x-coordinate represents the samples; the Y-coordinate indicates the relative abundance of bacteria at the phylum level. (A): duodenum; (B): jejunum; (C): ileum; (D): cecum; (E): colon.
Graph: Figure 4 Linear discriminant analysis (LDA) identified distinct bacterial genera that were enriched in the CON, LNO, and HLS groups. Genera with LDA score > 2 and p < 0.05 were considered significant. Each color represents one treatment: the red represents kids fed the basal diet (CON), the blue represents kids fed the basal diet supplemented with flaxseed oil (LNO), and the green represents kids fed the basal diet supplemented with heated flaxseed grain (HLS). (A): duodenum; (B): jejunum; (C): ileum; (D): cecum; (E): colon.
DIAGRAM: Figure 5 Venn diagram analysis of shared distinct bacteria at genus level in the intestinal digesta of kid goat. Each color represents one site: the blue represents the duodenum, the red represents the jejunum, the green represents the ileum, the yellow represents the cecum, and the brown represents the colon.
Graph: Figure 6 Average relative abundance of the dominant phyla (phyla with average relative abundance ≥ 0.01 in at least one region) in 5 intestinal regions of cashmere goats. The X-coordinate represents the samples, the Y-coordinate indicates the relative abundance of bacteria at the phylum level. (A): All groups; (B): CON group; (C): LNO group; (D): HLS group).
DIAGRAM: Figure 7 Venn diagram analysis of generic level bacterial composition in the intestinal digesta of goat kids. (A): All groups (n = 75). (B): CON group. (C): LNO group. (D): HLS group. Each color represents one site: the red represents duodenum, the blue represents jejunum, the green represents ileum, the yellow represents cecum, and the purple represents colon. The bar chart shows the number of genera in each segment.
Graph: Figure 8 Principal coordinate analysis (PCoA, using the weighted Unifrac similarity metric) of bacterial operational taxonomic units in 5 intestinal segments of Albas cashmere goats. (A) All groups (n = 75). (B): CON group. (C): LNO group. (D): HLS group. Each symbol represents one site: the solid red circle represents duodenum, the solid blue triangle represents jejunum, the solid green rhombus represents ileum, the solid yellow square represents cecum, and the solid purple cross-shaped represents colon.
Table 1 The composition and nutrient levels of CON, LNO, and HLS groups (dry-matter basis, DM basis).
Ingredients Day 1 to 30 Day 31 to 60 Day 61 to 90 CON LNO HLS CON LNO HLS CON LNO HLS Alfalfa 25.00 25.00 25.00 15.00 15.00 15.00 12.50 12.50 12.50 Corn stalk 5.00 5.00 5.00 20.00 20.00 20.00 25.00 25.00 25.00 Oat 20.00 20.00 20.00 15.00 15.00 15.00 12.50 12.50 12.50 Corn 28.41 23.37 23.17 30.80 30.40 29.90 31.30 29.90 29.40 Soybean meal 46% 11.70 10.50 11.50 9.50 11.40 11.90 8.00 10.40 10.90 Distiller's dried grains with soluble, DDGS 3.00 7.24 7.74 4.00 0.50 0.50 4.00 0.50 0.50 Flaxseed meal 4.80 4.80 0.00 3.50 3.50 0.00 4.50 4.50 0.00 Flaxseed 0.00 0.00 5.50 0.00 0.00 5.50 0.00 0.00 7.00 Flaxseed oil 0.00 2.00 0.00 0.00 2.00 0.00 0.00 2.50 0.00 Premix (1) 0.50 0.50 0.50 0.50 0.50 0.50 0.50 0.50 0.50 Limestone 0.20 0.20 0.20 0.20 0.20 0.20 0.20 0.20 0.20 CaHPO4 0.20 0.20 0.20 0.20 0.20 0.20 0.20 0.20 0.20 NaCl 0.54 0.54 0.54 0.50 0.50 0.50 0.50 0.50 0.50 NaHCO3 0.35 0.35 0.35 0.80 0.80 0.80 0.80 0.80 0.80 MgO 0.30 0.30 0.30 0.00 0.00 0.00 0.00 0.00 0.00 Total 100.00 100.00 100.00 100.00 100.00 100.00 100.00 100.00 100.00 Nutrient levels Digestible energy, DE MJ/Kg DM (2) 12.83 13.09 13.06 12.87 13.00 12.96 12.74 13.09 13.05 Dry Matter, DM/% 88.02 88.36 88.24 89.14 89.36 89.32 87.09 87.02 87.03 Crude protein, CP g/kg DM 188.73 188.13 188.20 162.84 158.71 159.69 153.52 151.34 151.85 Ether extract, EE g/kg DM 29.12 53.97 53.97 28.99 45.84 46.14 26.85 48.99 49.92 Neutral Detergent Fiber, NDF g/kg DM 425.31 431.18 441.60 448.60 427.41 439.12 457.42 436.06 450.68 Acid Detergent Fiber, ADF g/kg DM 232.20 237.71 243.3 242.59 235.23 248.40 247.69 240.32 256.96 Calcium, Ca g/kg DM 11.25 11.11 11.00 10.48 10.89 10.78 10.26 10.67 10.56 Phosphorus, P g/kg DM 4.65 4.67 4.78 4.50 4.44 4.56 4.31 4.22 4.33
Table 2 Optimized sequencing data of intestinal digesta.
Group Optimized Sequences Average Mean_Base Mean_Length Min_Length Max_Length Duodenum CON 224,418 44,884 18,521,788 412.57 247 480 LNO 244,950 48,990 20,493,991 417.72 245 480 HLS 282,979 56,596 23,178,454 409.65 247 474 Jejunum CON 309,498 61,900 25,363,009 409.88 246 446 LNO 310,366 62,073 25,579,668 412.12 230 450 HLS 274,223 54,845 22,469,544 409.65 243 448 Ileum CON 271,857 54,371 23,336,091 429.08 291 467 LNO 281,019 56,204 24,221,281 431.32 280 481 HLS 253,906 50,781 21,967,437 432.89 283 474 Cecum CON 313,157 62,631 25,849,679 412.71 263 457 LNO 245,830 49,166 20,323,122 413.30 236 449 HLS 319,688 63,938 26,381,252 412.50 236 456 Colon CON 297,818 59,564 24,586,815 412.72 275 454 LNO 216,507 43,301 17,895,233 413.11 276 439 HLS 265,291 53,058 21,920,157 413.16 238 463
Table 3 Effects of dietary flaxseed oil or heated flaxseed grain on bacterial α-diversity index of intestinal digesta.
Items CON LNO HLS SEM Duodenum Sobs 717A 462B 626A 47.33 0.008 Shannon 3.81 2.92 3.90 0.27 0.230 Simpson 0.08 0.15 0.07 0.04 0.990 Ace 812.91A 611.61B 779.29A 43.19 0.014 Chao 865.17A 557.30B 791.07A 58.45 0.008 Coverage 0.997 0.997 0.997 0.0005 0.988 Jejunum Sobs 689A 772A 384B 55.24 0.0008 Shannon 4.07A 4.08A 3.30B 0.11 0.0003 Simpson 0.04B 0.05B 0.12A 0.01 <0.0001 Ace 860.16A 905.30A 485.12B 56.34 0.0003 Chao 839.03A 901.77A 505.29B 47.21 0.0001 Coverage 0.996 0.996 0.997 0.0005 0.312 Ileum Sobs 325A 253B 355A 13 0.038 Shannon 2.69 2.56 2.64 0.19 0.931 Simpson 0.16 0.14 0.17 0.03 0.748 Ace 418.86A 356.34B 404.77A 3.71 0.001 Chao 407.55A 354.18B 408.35A 5.75 0.019 Coverage 0.999 0.999 0.999 0.0002 0.722 Cecum Sobs 931A 803B 740B 34.52 0.006 Shannon 5.33A 4.95B 4.81B 0.07 0.001 Simpson 0.01B 0.02A 0.02A 0.002 0.014 Ace 1062.50A 908.79B 860.71B 39.92 0.01 Chao 1095.83A 919.42B 867.17B 38.27 0.003 Coverage 0.994 0.994 0.995 0.0004 0.329 Colon Sobs 847A 713B 654B 29.05 0.002 Shannon 5.28A 4.90B 4.74B 0.12 0.019 Simpson 0.013B 0.025A 0.023A 0.002 0.004 Ace 989.43A 841.48B 793.75B 34 0.004 Chao 992.68A 857.26B 800.52B 35.43 0.007 Coverage 0.991 0.992 0.992 0.0004 0.149
Table 4 Effects of dietary flaxseed oil or heated flaxseed grain on bacterial composition of intestinal digesta (phylum level).
Phylum, % CON LNO HLS SEM Duodenum p__Firmicutes 65.43A 37.73B 45.02B 3.83 0.001 p__Proteobacteria 12.00B 49.28A 4.41B 7.4 0.003 p__Actinobacteria 8.35B 4.08B 26.28A 3.99 0.005 p__Saccharibacteria 4.90A 0.55B 5.80A 0.5 0.008 p__Bacteroidetes 1.75B 1.07B 3.83A 0.56 0.046 Jejunum p__Firmicutes 64.27 70.06 60.08 7.41 0.644 p__Actinobacteria 14.42B 9.07C 29.08A 1.55 <0.0001 p__Proteobacteria 2.24B 8.63A 3.26B 0.42 0.006 p__Saccharibacteria 4.67A 0.56B 0.89B 0.37 0.008 p__Verrucomicrobia 1.40A 0.27B 0.23B 0.06 0.005 Ileum p_Firmicutes 90.80A 87.22A 75.83B 1.826 0.015 p_Actinobacteria 4.96B 8.92B 20.06A 1.449 0.001 p_Proteobacteria 0.79A 0.71A 0.37B 0.106 0.042 p_Tenericutes 1.25A 1.03A 0.62B 0.102 0.005 p_Bacteroidetes 1.48B 1.49B 2.78A 0.641 0.040 Cecum p__Firmicutes 60.74AB 56.56B 62.76A 1.41 0.027 p__Bacteroidetes 34.02 33.39 31.99 1.83 0.732 p__Spirochaetae 1.37 1.54 1.65 0.21 0.657 p__Proteobacteria 0.64 0.83 0.55 0.11 0.160 p__Verrucomicrobia 0.40B 0.30B 0.66A 0.06 0.004 Colon p__Firmicutes 53.81C 64.08A 59.54B 0.99 <0.0001 p__Bacteroidetes 30.94 33.36 36.17 1.41 0.067 p__Spirochaetae 1.37B 4.00A 1.49B 0.56 0.01 p__Verrucomicrobia 1.16 0.67 1.38 0.3 0.271 p__Tenericutes 1.38A 1.40A 0.63B 0.06 0.009
Table 5 Correlation analysis of differential duodenal bacteria and blood fatty acid composition.
Genus C16_0 C18_0 C18_1c9 C18_2c6 C18_3n3 C20_4n6 C20_5n3 C22_6n3 R (2) R R R R R R R g__[Eubacterium]_coprostanoligenes_group 0.033 −0.543 0.0002 −0.630 <0.0001 0.865 0.043 −0.548 <0.0001 0.720 g__Acetitomaculum 0.014 −0.541 <0.0001 −0.618 g__Anaerofustis 0.001 −0.698 0.010 0.627 0.004 −0.609 g__Brevibacillus 0.012 −0.560 g__Denitrobacterium 0.044 −0.504 0.012 −0.568 g__Enterorhabdus 0.000 −0.549 g__Lachnospiraceae_UCG-002 0.003 −0.673 0.0003 −0.712 g__Lachnospiraceae_XPB1014_group 0.010 0.516 g__Marvinbryantia 0.008 0.725 0.0001 −0.792 0.014 0.635 <0.0001 −0.772 g__norank_f__Coriobacteriaceae 0.046 −0.530 g__Olsenella 0.001 −0.562 g__Pseudarthrobacter 0.050 0.556 0.041 −0.526 g__Pseudoramibacter 0.021 0.761 0.001 −0.738 0.008 −0.736 g__Senegalimassilia 0.004 −0.627 0.014 0.567 0.0002 −0.640 g__Solobacterium 0.002 −0.581 0.023 0.507 0.000 −0.659 g__unclassified_f__Coriobacteriaceae 0.012 −0.534 g__unclassified_f__Erysipelotrichaceae 0.005 −0.557
Table 6 Correlation analysis of differential jejunal bacteria and blood fatty acid composition.
Genus C16_0 C18_0 C18_1c9 C18_2c6 C18_3n3 C20_4n6 C20_5n3 C22_6n3 R (2) R R R R R R R g__[Eubacterium]_cellulosolvens_group 0.006 0.566 0.04 −0.579 g__[Eubacterium]_coprostanoligenes_group <0.0001 0.718 <0.0001 −0.776 g__Anaerofustis 0.005 0.699 <0.0001 −0.751 0.023 0.556 0.0004 −0.749 g__Blautia 0.002 0.586 g__Catenisphaera 0.046 0.557 0.0002 −0.753 0.0005 0.626 <0.0001 −0.779 g__Enterococcus 0.002 −0.58 g__Family_XIII_UCG-001 0.002 −0.601 0.004 0.527 g__Lachnospiraceae_UCG- 002 0.002 −0.685 0.037 0.612 0.03 −0.534 g__Marvinbryantia 0.011 −0.52 g__norank_f__Coriobacteriaceae 0.027 0.514 0.002 −0.541 g__norank_o__Mollicutes_RF9 0.008 −0.569 0.001 0.597 g__Pseudoramibacter 0.008 −0.67 0.034 0.567 0.0001 −0.723 g__Saccharofermentans 0.0002 −0.653 0.044 0.586 0.007 −0.581 g__Senegalimassilia 0.019 −0.519 g__Solobacterium 0.001 −0.714 0.027 0.573 0.009 −0.682 g__unclassified_f__Coriobacteriaceae <0.0001 −0.732 0.021 0.682 0.002 −0.548
Table 7 Correlation analysis of differential ileal bacteria and blood fatty acid composition.
Genus C16_0 C18_0 C18_1c9 C18_2c6 C18_3n3 C20_4n6 C20_5n3 C22_6n3 R (2) R R R R R R R g__unclassified_f__Erysipelotrichaceae 0.003 −0.598 0 0.631 0.007 −0.543 g__Solobacterium 0.014 0.658 0 −0.729 0 0.718 0.003 −0.843 g__[Eubacterium]_coprostanoligenes_group 0.001 0.799 0.001 −0.863 0.022 0.62 0.032 0.585 g__Erysipelotrichaceae_UCG-009 0.001 0.724 0.001 −0.694 0.001 0.752 0.001 −0.782 g__Acetitomaculum 0.002 −0.851 0.003 0.839 0.028 −0.705 g__Christensenellaceae_R−7_group 0.032 −0.625
Table 8 Correlation analysis of differential cecal bacteria and blood fatty acid composition.
Genus C16_0 C18_0 C18_1c9 C18_2c6 C18_3n3 C20_4n6 C20_5n3 C22_6n3 R (2) R R R R R R R g__Ruminiclostridium_1 0.020 −0.540 0.000 −0.574 g__Ruminococcaceae_UCG-010 0.004 0.716 g__Prevotellaceae_Ga6A1_group 0.012 −0.625 0.001 0.582 0.003 −0.634 g__Barnesiella 0.026 0.561 g__norank_o__Bradymonadales 0.042 −0.508 g__norank_f__Bacteroidales_BS11_gut_group 0.009 0.621 g__[Eubacterium]_brachy_group 0.013 0.551 g__Mogibacterium 0.003 0.586
Table 9 Correlation analysis of differential colonic bacteria and blood fatty acid composition.
Genus C16_0 C18_0 C18_1c9 C18_2c6 C18_3n3 C20_4n6 C20_5n3 C22_6n3 R (2) R R R R R R R g__Roseburia 0.002 0.588 g__Bacteroides 0.004 0.751 g__Anaerotruncus 0.005 0.558 g__Ruminococcaceae_UCG-002 0.006 0.553 0.0001 0.658 g__Ruminiclostridium_1 0.0003 −0.642 <0.0001 0.614 0.003 −0.644 g__norank_f__Bacteroidales_RF16_group 0.006 −0.652 0.0002 0.682 0.003 −0.586 g__Prevotellaceae_Ga6A1_group 0.001 −0.763 0.0001 0.661 0.001 −0.732 g__Saccharofermentans 0.004 −0.553 0 0.514 0.006 −0.54 g__norank_f__Christensenellaceae 0.006 −0.509 0.006 −0.596 g__Family_XIII_UCG-002 0.008 −0.619 0.0005 0.636 0.0001 −0.527 g__Hydrogenoanaerobacterium <0.0001 −0.68 0 0.712 0.005 −0.551 g__norank_o__Bradymonadales 0.042 −0.52 0.004 −0.562
Conceptualization, Y.G. and S.Y.; Data curation and Formal analysis, S.L.; Funding acquisition, Y.G. and S.Y.; Investigation, Y.G., S.L. and Y.L.; Project administration, Y.G. and S.Y.; Resources, S.L. and Y.L.; Visualization, Y.G. and S.L.; Supervision, B.S. and S.Y.; Writing–original draft, Y.G. and S.L.; Writing–review and editing, X.G., Y.Z., B.S. and S.Y. All authors have read and agreed to the published version of the manuscript.
The use of the animals was approved by the Animal Ethics and Welfare Committee of Inner Mongolia Agricultural University in accordance with the Laboratory Animal Sciences and Technical Committee of the Standardization Administration of China (SAC/TC281), and performed under the national standard Guidelines for Ethical Review of Animal Welfare (GB/T 35892-2018).
Owner of the animals involved in this research was informed and understood the purpose of this experiment. He also agreed with publishing the results in the Journal Animals.
The sequencing data are available from the National Center for Biotechnology Information under the Sequence Read Archive (SRA) with the BioProject No. PRJNA809885, PRJNA810318, PRJNA810285, PRJNA810293, PRJNA810302.
The authors declare no conflict of interest.
For their help during laboratory and data analysis, the authors express deep appreciation to Qi Wen, Yongsheng Zhang, and Siqingaowa Bao from the College of Animal Science, Inner Mongolia Agricultural University, China.
The following supporting information can be downloaded at: https://
By Yongmei Guo; Shulin Liu; Yinhao Li; Xiaoyu Guo; Yanli Zhao; Binlin Shi and Sumei Yan
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