Next Article in Journal
In-Field Phenotyping Using the Low-Cost and Open Access Fluorescence PhotosynQ Multispeq Sensor Together with NDVI: A Case Study with Durum Wheat
Previous Article in Journal
Research on the Prediction Model and Formation Law of Drying Cracks of Paddy Based on Multi-Physical Field Coupling
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Agriculture
Volume 15
Issue 4
10.3390/agriculture15040384
agriculture-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Effects of Fish Palm Rice and Coconut Palm Rice Oil Mixture on Intestinal Health of Weaned Piglets

by
Kaiyun Yang
1,2,3,
Wenjuan Yang
1,2,
Fei Jiang
1,2,
Bing Yu
1,2,
Zhiqing Huang
1,2,
Yuheng Luo
1,2,
Aimin Wu
1,2,
Ping Zheng
1,2,
Xiangbing Mao
1,2,
Jie Yu
1,2,
Junqiu Luo
1,2,
Hui Yan
1,2 and
Jun He
1,2,*
1
Institute of Animal Nutrition, Sichuan Agricultural University, Chengdu 611130, China
2
Key Laboratory of Animal Disease-Resistant Nutrition, Chengdu 611130, China
3
Leshan Giantstar Farming & Husbandry Co., Ltd., Leshan 614000, China
*
Author to whom correspondence should be addressed.
Agriculture 2025, 15(4), 384; https://doi.org/10.3390/agriculture15040384
Submission received: 18 December 2024 / Revised: 6 February 2025 / Accepted: 7 February 2025 / Published: 12 February 2025
(This article belongs to the Section Farm Animal Production)
Browse Figures
Versions Notes

Abstract

:
Fat not only has a high effective energy value and extra energy effects but is also an ideal energy source. As one of the most commonly used feed oils at present, soybean oil has a high cost despite its good application effect on weaned piglets. In contrast, the cost of palm oil is lower. In order to explore the effect of palm oil-dominated fat combination on intestinal health of weaned piglets, we conducted a 28-day trial, randomly dividing 72 pigs into three groups: The SO group (fed with a basal diet containing 2% soybean oil), the PRFO group (replacing soybean oil with 2% fish palm rice oil mixture), and the PRCO group (replacing soybean oil with 2% coconut palm rice oil mixture), with eight replicate pens per group (3 pigs per pen) and intestinal chyme was collected on day 29. We observed a significant increase in the concentration of propionic acid (PA) in cecal digesta of the PRFO group (895.49 mmol/L p < 0.05) compared to the SO group (626.32 mmol/L). Meanwhile, the concentration of cecal PA, butyric acid (BA), valerate and colonic valeric acid were significantly increased in the PRCO group (p < 0.05). Moreover, piglets in the PRFO group had a higher abundance of Firmicutes in the cecum and colon, but a lower abundance of Actinomyces in the cecum than those in the SO group. These results indicate that replacing soybean oil with PRFO and PRCO in weaning diet can improve intestinal microflora structure and thus improve intestinal health of piglets.

1. Introduction

With the advent of modern intensive farming, the weaning stage has acquired immense importance and has become a critical period in the life of pigs. This stage is characterized by sudden alterations in diet, environment, and social interactions [1]. Weaned piglets are often exposed to various stress-inducing factors, which can trigger a range of responses such as temporary loss of appetite, inflammation in the intestines, and imbalances in the gut microbiota [2,3]. Moreover, the transitionary environment during weaning frequently gives rise to gastrointestinal issues like Escherichia coli (E. coli)-induced diarrhea [4]. Numerous studies have demonstrated a connection between post-weaning diarrhea and intestinal infections with changes in both the weaning diet and the intestinal microbiome of piglets [5]. The gut microbiota of mammals confers numerous advantages to the host, including but not limited to carbohydrate digestion and fermentation, synthesis of vitamins, maintenance of optimal intestinal villi functionality, modulation of immune responses, and protection against pathogenic bacteria [6,7].
The intestinal mucosa consists of a solitary layer of columnar epithelial cells, underlain by the lamina propria and muscularis mucosae. These epithelial cells form a protective barrier against the potentially harmful external and internal surroundings [8,9]. The intestinal epithelial lining, which serves as the largest interface for exchange between the body and the outside world, is crucial for nutrient absorption, fluid balance, and preventing pathogen invasion due to its permeability characteristics [10]. The gut microbiota is crucial for sustaining intestinal equilibrium in humans and animals [11]. Numerous studies have shown that animals are sterile before birth, but they are quickly colonized by microorganisms from their mother and environment, in the order of aerobic and facultative anaerobic gut colonization [12]. As an animal grows and develops, the composition and number of gut microbiomes change. Weaned piglets are susceptible to microbial invasion due to incomplete intestinal development and immune function. The early establishment of microbial communities in the gastrointestinal tract of piglets is crucial for their overall well-being [13]. When a sow’s milk is no longer available, the intestinal flora of piglets changes significantly between 7 and 14 days after weaning [14]. In addition, the gut microbiome is constantly changing and vulnerable to various influences, and different feeding methods and dietary components could directly change the structure of gut microbiota [15].
Lipids, with an energy density 2.25 times higher than that of carbohydrates and proteins, have long been an excellent source of energy to address the issue of inadequate energy intake during the weaning period of piglets [16]. In animals, fat is essential for growth and metabolism, as it supplies vital fatty acids and facilitates the absorption of fat-soluble nutrients [17]. Studies have demonstrated that a high-fat diet disturbs the equilibrium between energy consumption and expenditure by altering the composition of the gut microbiota, which also exerts a lasting impact on its structure [18,19]. Studies have found that a diet rich in n-3 polyunsaturated fatty acids significantly increases the abundance of bifidobacteria and lactobacilli in the intestines of mice, which can help to sustain a balanced intestinal microbiota in the host [20]. In vitro experiments have shown that short-chain fatty acids (SCFAs) like propionate and butyrate, along with medium-chain fatty acids (MCFAs) such as octanoic acid and caproic acid, are capable of suppressing the growth of Salmonella typhimurium. Research has found that adding 15 mM of sodium octanoate can significantly reduce the abundance of Escherichia coli and Salmonella in the cecum [21]. Earlier research has demonstrated that butyric acid can reduce the amount of Salmonella in the feces of piglets infected with Salmonella typhimurium, reducing its colonization in the intestine [22].
The regulation of animal gut microbiota by lipids is influenced by their source and chemical composition [23]. It was observed that male Sprague Dawley rats consuming a diet of corn oil, palm oil, and coconut oil exhibited a markedly reduced Escherichia coli count in their cecum when compared to rats on a fat-free regimen [24]. Furthermore, another study compared the effects of coconut oil, fish oil, and lard on the number of microorganisms in the cecum of piglets, finding that compared with the lard group, coconut oil and fish oil diminished the number of Escherichia coli in the cecum content and increased the abundance of bifidobacteria and lactobacilli, while the coconut oil group was superior to the fish oil group [25]. Although these studies indicate that various lipid sources offer benefits to weaned pigs, the overall impact of combining various lipid sources is still in the initial phase of investigation.
In the present study, we hypothesized that replacing soybean oil with a fish palm rice oil mixture (PRFO) in the diet of weaned piglets will improve their intestinal health by increasing the level of beneficial SCFAs in the cecal and colonic digesta. To verify this hypothesis, we fed pigs with a corn soybean meal diet containing either 2% soybean oil or an oil mixture (fish oil and coconut oil, known as PRFO and PRCO) and then conducted tests on relevant indicators such as the concentration of volatile fatty acids (VFAs) and microbial abundance in the cecal and ileal digesta. Our study offers compelling evidence of the probiotic effects of PRFO and PRCO, providing valuable insights for making informed decisions about selecting lipid sources.

2. Materials and Methods

2.1. Experimental Design and Diets

Seventy-two healthy weaned pigs, averaging 7.80 ± 0.11 kg in weight and with uniform physical states and breeds, were randomly allocated to three distinct treatment groups. Each treatment featured eight replicates, with three piglets housed together in a single pen. The base diet consisted of corn and soybean meal, with the addition of 2% soybean oil (SO), or a mixture of 10% fish oil, 50% palm oil, and 40% rice bran oil (PRFO), or a mixture of 5% coconut oil, 80% palm oil, and 15% rice bran oil (PRCO). The formulation of the basal diet was referenced from our laboratory’s previous study (Table S1) [26], and its composition and nutritional content were formulated according to the pig feeding standards outlined by the NRC (2012). Feeding management and immunization protocols for the experimental groups followed field regulations. The experiment was conducted in 2022, starting with a 3-day pre-feeding period, followed by the main experiment that lasted for 28 days. Pigs had ad libitum access to feed and fresh water, and the room temperature was maintained between 25 and 28 °C with a relative humidity of 65 ± 5% [27]. Feed intake and body weight of the piglets were measured on days 0 and 29 of the experiment, allowing for the calculation of the average daily feed intake (ADFI), average daily gain (ADG), and feed conversion ratio (FCR) throughout the entire experimental period.

2.2. Sample Collection

On the morning of the 29th, a piglet nearest to the average weight was chosen from each cage for slaughter. After slaughter, the chyme from the cecum and colon was collected. The method for collecting cecal and colonic contents followed the study by Zhang et al. [28].

2.3. Analysis of the Concentration of Volatile Fatty Acids in Chyme

The concentrations of volatile fatty acids (VFAs) were measured using a VARIAN CP-3800, gas chromatography system, following the methods described in previous studies [29].

2.4. Determination of the Digesta pH Values

Shortly after the pigs were slaughtered, about 5 g of digesta was immediately placed into a sterile centrifuge tube that was kept on ice. Subsequently, the pH of each sample was determined with a PHS-3C pH meter (Shanghai, China).

2.5. Analysis of the Bacterial Community

The Illumina MiSeq 2000 platform was employed to conduct high-throughput sequencing of the 16S rRNA gene in the cecum and colon of piglets. Nucleic acids were extracted from 0.5 g of digesta samples using the Stool DNA kit (Omega Bio-Tech, Doraville, CA, USA). The diversity and composition of the microbial communities in the samples were analyzed using the Quantitative Insights Into Microbial Ecology (QIIME, version 2020.2.0) software package [30]. PCR amplification was used to target the V3–4 region of the 16S rRNA gene, using the following primer sequences: 338F: ACTCCTACGGGAGGCAGCAG and 806R: GGACTACHVGGGTWTCTAAT. The PCR was conducted under the following conditions: An initial denaturation at 95 °C for 5 min, followed by 35 cycles consisting of 94 °C for 30 s, 55 °C for 30 s, and 72 °C for 90 s, with a final extension at 72 °C for 10 min. Sequence amplification was performed using the Verity Thermocycler (Applied Biosystems, BIO-RAD, Foster City, CA, USA), and the raw sequencing data were stored in double-ended FASTQ format. To ensure data quality, the sliding window method was applied to filter the FASTQ sequences: With a window size of 5 bp, a step size of 1 bp, and an average base quality score ≥ Q20. Sequences were truncated from the point where the base quality dropped below Q20, and the final sequence length was required to be ≥150 bp, without any N (ambiguous bases). FLASH software (version 1.2.11) was then used to merge the quality-filtered sequences, allowing no base mismatches. Effective sequences for each sample were extracted based on index information. To remove chimeric sequences generated during PCR amplification and sequencing errors like point mutations, the QIIME platform was used for sequence filtering. To ensure accuracy, OTUs with an abundance of less than 0.001% of the total sequences were discarded. OTU annotation and clustering were performed using QIIME at a 97% similarity threshold, followed by subsequent bioinformatics analysis of the sequences [31,32]. Mothur software (version v.1.30.2) or R software (version 3.3.1) was used to generate Venn diagrams illustrating the overlap of OTUs between samples (or groups). To assess the representativeness of the sequences, all sequences were randomly subsampled to determine the number of sequences extracted and the OTUs they could represent. Dilution curves were constructed using QIIME software. Additionally, Alpha diversity analysis was performed using the summary single command in Mothur, in accordance with the abundance of each species in the OTU list for each sample [33,34]. The generation of QIIME and OTU tables was based on the study by Oberauner et al. [35]. Beta diversity analysis also uses QIIME for Unifrac analysis including unweighted Unifrac difference analysis and weighted Unifrac difference analysis. R software was used to conduct principal component analysis for classification and species abundance at the genus level and draw PCA maps. Meanwhile, genus-level classification data were clustered using cluster analysis, and a heatmap was generated. Based on the OTU abundance information, PICRUSt software was employed to predict functional genes and conduct the relevant statistical analysis [36].

2.6. Data Statistics and Analysis

The VFA data from the three experimental groups were arranged using Microsoft Excel 2019 and subjected to one-way ANOVA analysis using SPSS 24.0 statistical software [37]. The p-values indicating significance were obtained by comparing the means between treatment groups using Duncan’s multiple range test in a one-way ANOVA. All data were presented as “mean ± standard error”, with p < 0.05 considered statistically significant and 0.05 ≤ p ≤ 0.10 considered a trend.

3. Results

3.1. Effects of Different Dietary Lipid Sources on Growth Performance and Microbial Metabolites of Weaned Piglets

SCFAs, including propionate, butyrate, and valerate, are produced through the fermentation of dietary fiber by the gut microbiota. These SCFAs are crucial for maintaining gut barrier integrity, regulating immune function, and supporting overall health in piglets [38]. In the present study, we found that the cecal PA content in the PRFO group was significantly higher (p < 0.05) than that in SO group, while the content of BA and VA showed an increasing trend (0.05 < p < 0.10) (Table 1). Pigs fed with a basal diet replacing soybean oil with 2% PRCO had a significantly increased (p < 0.05) content of cecal PA, BA, and VA. In addition, the content of valeric acid in the colon of the PRCO group is also significantly higher than that in the SO group. Moreover, pigs fed with a basal diet replacing soybean oil with 2% PRFO had a significantly reduced (p < 0.05) pH of cecal digests (Table 1).

3.2. The Effect of Different Lipid Sources on the Total Bacterial Count of Cecum and Colon Chyme in Weaned Piglets

As shown in Figure 1, compared with the SO group, the abundance of total microorganisms in the cecal chyme of the PRFO and PRCO groups were significantly higher (p < 0.05) than that in the SO group. Interestingly, the same experimental results were also obtained in the colon chyme (p < 0.05).

3.3. Effects of Different Dietary Lipid Sources on Bacterial Community Structure of Weaned Piglets

The sequencing data have been stored in the National Center for Biotechnology Information (NCBI) and can be retrieved from the Short Read Archive (SRA) using the accession number PRJNA1207688. A total of 2,473,221 sequences were obtained, averaging 1,408,212 sequences per cecum sample and 1,065,009 sequences per colon sample. The average length of these sequences was 411 clipped base pairs. The sparsity and rank-abundance curves showed that sampling depth and evenness were sufficient to assess bacterial communities (Figure 2).
As depicted in Figure 3, a higher number of OTUs is observed in the colon compared to the cecum at the genus level. Specifically, the cecum sample contains 3 unique OTUs in the SO group, and 4 and 12 unique OTUs in the PRFO and PRCO groups, respectively. In total, there are 139 OTUs across the three groups. In the colon samples, the SO group, PRFO, and PRCO each have 5, 5, and 11 unique OTUs, with a sum of 147 OTUs shared by the three groups. Alpha diversity metrics were employed to assess the overall bacterial diversity in the cecum and colonic chyme of each group. Specifically, the abundance coverage estimator (ACE) was utilized to evaluate species richness, while the Simpson’s index was applied to determine species evenness [33,39]. The statistical results showed that the Shannon (p = 0.060) index of the cecal PRCO group showed a significant difference compared to the other two groups. No significant alterations were observed in the diversity of microbial communities within the colon chyme samples (Figure 3). Beta diversity analysis was performed to examine the similarity or differences in community composition across different groups by comparing species diversity in various habitats or microbial communities [40]. PCoA based on weighted UniFrac distance measurements, was employed to determine the similarity of microbial communities. Furthermore, the Anoism test method was utilized to assess whether differences existed in the changes in microbial community structure between the various samples (p < 0.05). The closer the two sample points in the figure are, the more similar the species composition of the two samples is. A tendency for separation was observed in the PRFO group of the cecal chyme samples when compared to the other two groups, with a p-value of 0.077. However, in the colon samples, no significant clustering or differentiation of chyme communities was detected (Figure 4).
The analysis indicated that, relative to the SO group, there was a successive rise in the richness of microflora in the cecum and colon of the PRFO and PRCO groups, as illustrated in Figure 5. Apart from the OTUs uniquely identified in each group, no significant difference in overall bacterial diversity was observed between the two groups. In both the cecum and colon, no significant variation in the flora was observed among the three groups. However, it was noted that the dietary lipid sources influenced the relative abundance of microbial communities at the phylum level in these regions. In this study, all eligible sequences from the cecal and colon samples were classified into 12 and 13 known phyla, respectively. Figure 2D illustrates the relative abundance distribution of the top 10 bacterial phyla in the cecum and colon across the three groups. In the SO group, Firmicutes (78.69%) and Actinomyces (17.18%) were the predominant phyla in the cecum. Nevertheless, in the PRFO and PRCO groups, an increase in the abundance of Firmicutes was noted, whereas a decrease was observed in the abundance of Actinomyces, although these alterations did not reach statistical significance (p > 0.05). In the colon samples, Firmicutes (73.32%) and Actinomyces (20.07%) were dominant in the SO group, but Firmicutes (93.12%) and Bacteroidetes (4.26%) were dominant in the PRFO group. Figure 6 illustrated that, in the cecum, Clostridium_sensu_stricto and Lactobacillus were dominant genera in the SO and PRCO groups. And in the PRFO group, Lactobacillus (21.23%), Clostridium_sensu_stricto (20.02%), and unclassified_f__Ruminococcaceae (19.60%) were the dominant classification levels. In the colon, Clostridium strictus (24.41%) and Olsenella (13.31%) were the dominant genera in the SO group, and Lactobacillus (21.63%) and Clostridium_sensu_stricto (15.73%) were the dominant genera in the PRFO group. The dominant genera in the PRCO group were Clostridium_sensu_stricto (29.88%) and Terrisporobacter (8.99%).
The sequences derived from the cecum and colon samples were classified into 12 and 13 known phyla, respectively, in addition to 50 known genera (Figure 7). Dietary supplementation with PRFO and PRCO resulted in a significant increase (p < 0.05) in the abundance of Firmicutes, at the phylum level, in both the cecum and colon, while simultaneously decreasing (p < 0.05) the abundance of Proteobacteria in these regions. At the genus level, the sequences of the cecum and colon samples were divided into 50 known genera (Figure 8). Lactobacillus and Clostridium were the two dominant genera in the cecum and colon. From Figure 7, it can be concluded that the inclusion of PRFO in the feed results in an increase in the abundance of Lactobacillus in both the cecum and colon of piglets.

3.4. Functional Profiling of the Bacterial Communities

PICRUSt (phylogenetic investigation of communities by reconstruction of unobserved states) was employed to generate the functional profile of bacterial communities, predicting the functional composition of a metagenome based on marker gene data [36]. In total 42 biological functions were identified across all samples, among which carbohydrate metabolism (26.96–27.73%), amino acid metabolism (16.71–17.22%), membrane transport (10.01–10.41%), and energy metabolism (9.82–10.08%) had the highest enrichment, and many predicted functions were related to microbial cell metabolism (Figure 8). Through COG classification of microbial community protein functions, as shown in Figure 9, a total of 23 protein functions of the microbial community were predicted. The prediction results for all three treatment groups showed that the strongest function was RNA processing and modification, followed by amino acid transport and metabolism.

4. Discussion

The gut ecosystem is significantly influenced by gut microbes, which are essential for maintaining intestinal epithelial integrity, facilitating nutrient digestion and metabolism, modulating immune responses, and preventing diseases [41,42]. Certain hard-to-digest fats, once reaching the distal intestine, serve as a nutrient source for bacteria, thus helping to regulate the balance of gut bacteria [23]. Short-chain fatty acids, which are the primary end products of colonic fermentation, can be utilized as an energy source by intestinal epithelial cells and are extensively involved in modulating cell growth, apoptosis, and a variety of inflammatory responses [43,44]. For instance, the expression of tight junction proteins like claudin-8, occludin, and claudin-1 is elevated by PA, thereby enhancing intestinal barrier function in animals. Additionally, PA stimulates the proliferation of epithelial cells [45,46]. In this study, compared with the SO group, dietary PRFO supplementation significantly increased the propionic acid concentration of cecal digesta and decreased the pH value of digesta. The concentrations of PA, BA, and VA in cecal digesta, as well as valerate in colonic digesta, were significantly elevated by PRCO. It has been demonstrated that the addition of rice bran oil and coconut oil to the diet can enhance the abundance of beneficial bacteria, which in turn indirectly boosts the production of volatile fatty acids [47,48].
In pigs, mice, and humans, Firmicutes constitute the predominant phylum of the intestinal microbiota, with many of its members being extensively implicated in energy metabolism and nutrient absorption [15]. Few studies have demonstrated that supplementing the diet with palm oil can enhance the abundance of Firmicutes in the cecum of broilers. Consistently, in this study, we found that replacing soybean oil in the piglets’ diet with 2% PRCO significantly enhanced the prevalence of Firmicutes in both the abundance of Firmicutes in both the cecum and colon of piglets. Compared to the SO group, the PRFO and PRCO groups showed an increase in Firmicutes abundance in both the cecal and colonic chyme, while a significant decrease in Actinobacteriota abundance was observed. The dominant genera in cecum and colonic digesta were Clostridium_sensu_stricto, Lactobacillus, unclassified_f__Ruminococcace and Terrisporobacter. Lactobacillus and Bifidobacterium are two key beneficial microorganisms that maintain intestinal immunity and epithelial function. TLR is an important molecule that regulates intestinal tightening, and studies have shown that Lactobacillus stimulates TLR2 to bind to ligands, thereby enhancing the function of tightening [49]. Bifidobacterium can reduce the content of Zonulin connection in serum, maintain the integrity of intestinal compact connection, reduce intestinal permeability and reduce intestinal inflammatory damage. It has been demonstrated that incorporating rice oil into the diet can enhance the prevalence of intestinal bacteria, including bifidobacterium and lactobacillus [47]. Dietary addition of coconut oil can reduce the amount of E. coli in the cecum of piglets and increase the abundance of bifidobacterium and lactobacillus [48]. In this study, compared with the SO group, dietary PRFO and PRCO can increase Bifidobacterium and Lactobacillus abundance in the cecal and colonic regions of piglets. In addition, the abundance of Eubacterium in caecum and colonic chyme in PRFO and PRCO groups also increased significantly. It has been found that Eubacterium can regulate the release of inflammatory mediators and reduce the levels of IL-2 and C-reactive protein. Numerous strains of Eubacterium are also capable of producing short-chain fatty acids, which primarily exert their anti-inflammatory effects by inhibiting the NF-κB pathway and/or histone deacetylase (HDAC) activity. This, in turn, not only leads to the downregulation of pro-inflammatory factors such as IL-6, IL-12, TNF-α and IFN-γ, but also upregulates the expression of anti-inflammatory factors like IL-10 and TGF-β [50]. In this study, despite several limitations such as the study’s brief duration, lack of environmental control, and small sample size, our results suggest that replacing soybean oil with PRFO and PRCO has the potential to enhance the intestinal health of weaned piglets by altering the composition of the gut microbiota and stimulating the production of SCFAs.
PICRUSt is a tool for functional prediction based on 16S rRNA gene sequences, which allows for the prediction of the metabolic functions of microbial communities. For instance, in a study of the gut bacteria in three equine species, PICRUSt predicted metabolism-related genes, revealing functional differences between species. These predictive results help in gaining a deeper understanding of the functional roles played by microbial communities in various hosts [51]. In the present study, we found that carbohydrate metabolism had the highest enrichment across all groups. Carbohydrate metabolism is essential for microbial fermentation in the gut, leading to the production of SCFAs which are crucial for intestinal health [52]. Firmicutes, a prominent phylum of bacteria, play an essential role in carbohydrate metabolism, particularly in the fermentation of dietary fibers into SCFAs within the gut. Studies have shown that Firmicutes are involved in the breakdown of complex carbohydrates, which are not digestible by the host, into simpler sugars and SCFAs. These SCFAs are important for maintaining gut health and providing energy to host cells, especially colonocytes. Furthermore, Firmicutes also dominate in high-carbohydrate diets and, together with Bacteroidetes, participate in the fermentation process of carbohydrates [53]. Consistently, in the present research, the high abundance of Firmicutes in PRFO and PRCO groups suggests that these bacteria may contribute to enhanced carbohydrate fermentation. Lactobacillus and Clostridium generate SCFAs and other metabolites through distinct fermentation pathways involved in energy metabolism, which not only supply energy to the gut microbiota but also exert significant effects on the host’s gut health and immune regulation. Lactobacillus ferments carbohydrates to produce lactic acid for energy. During fermentation, lactic acid bacteria convert glucose or other sugars into lactic acid while generating ATP (adenosine triphosphate), which is the primary form of cellular energy. Clostridium, an anaerobic bacterium, produces various metabolites through the fermentation of organic materials, including acetate, butyrate, and hydrogen. These metabolites play important roles in energy metabolism. For example, Clostridium uses the ethanol acetate fermentation pathway to convert ethanol and carbon dioxide into acetate and butyrate while generating hydrogen. For every 2 moles of hydrogen produced, 1 mole of acetyl CoA is generated for ATP synthesis [54]. Consistently, in this study, we found that the abundance of Lactobacillus and Clostridium in the cecal and colonic chyme of piglets in the PRFO and PRCO groups significantly increased. At the same time, PICRUSt functional prediction revealed that energy metabolism dominated. These results suggest that the enhanced energy metabolism function in piglets of the PRFO and PRCO groups may be associated with the increased abundance of Lactobacillus and Clostridium.

5. Conclusions

This experiment was conducted to examine the effects of various fat sources on the growth performance and intestinal flora of weaned piglets, as well as to analyze the differences in intestinal microbiota. The findings suggest that replacing soybean oil with PRFO and PRCO does not impact the production performance of weaned piglets. However, it appears to enhance their intestinal health by increasing the content of short-chain fatty acids, upregulating the abundance of beneficial bacteria in the microbiota, and improving the overall structure of the intestinal microbiome. Moreover, compared with SO, PRCO is the lowest, followed by PRFO. Therefore, replacing soybean oil in piglet diets with PRCO and PRFO can reduce production costs. It offers a theoretical foundation for the rational selection of lipid sources for weaned piglets, as well as for other mammals. In future studies, we recommend focusing on the application effects of PRFO and PRCO on fattening pigs or local pig breeds. Key indicators such as intestinal tight junction proteins, immunoglobulins, and inflammatory factors should be measured to reflect the impact of PRFO and PRCO on pig intestinal health.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/agriculture15040384/s1, Table S1: Experimental basal diet composition and nutrient level.

Author Contributions

J.H. conceived and designed the experiments. K.Y., W.Y. and F.J. conducted the experiment. K.Y and W.Y., write the initial manuscript. B.Y., Z.H., Y.L., A.W., P.Z., X.M., J.Y., J.L. analyzed the data. H.Y. and J.L. made the final editing and proofreading. All authors have read and agreed to the published version of the manuscript.

Funding

This study was supported by the National Key R&D Program of China (2023YFD1301200) and the Porcine Innovation Team of Sichuan Province (SCCXTD-2024-8).

Institutional Review Board Statement

All experimental protocols used in the animal experiment were approved by the Institutional Animal Care and Use Committee of Sichuan Agricultural University (No. 20181105).

Data Availability Statement

The data used to support the findings of this study are available from the corresponding author upon request.

Conflicts of Interest

Author Kaiyun Yang and Wenjuan Yang were employed by Leshan Giantstar Farming & Husbandry Co., Ltd. Author Feijiang was employed by Singao Agribusiness Development Co., Ltd.

References

  1. Gresse, R.; Chaucheyras-Durand, F.; Fleury, M.A.; van de Wiele, T.; Forano, E.; Blanquet-Diot, S. Gut microbiota dysbiosis in postweaning piglets: Understanding the keys to health. Trends Microbiol. 2017, 25, 851–873. [Google Scholar] [CrossRef]
  2. Lallès, J.P.; Sève, B.; Pié, S.; Blazy, F.; Laffitte, J.; Oswald, I.P. Weaning is associated with an upregulation of expression of inflammatory cytokines in the intestine of piglets. J. Nutr. 2004, 134, 641–647. [Google Scholar] [CrossRef]
  3. Lallès, J.-P.; Bosi, P.; Smidt, H.; Stokes, C.R. Nutritional management of gut health in pigs around weaning. Proc. Nutr. Soc. 2007, 66, 260–268. [Google Scholar] [CrossRef]
  4. Schokker, D.; Zhang, J.; Vastenhouw, S.A.; Heilig, H.G.; Smidt, H.; Rebel, J.M.J.; Smits, M.A. Long-lasting effects of early-life antibiotic treatment and routine animal handling on gut microbiota composition and immune system in pigs. PLoS ONE 2015, 10, e0116523. [Google Scholar] [CrossRef]
  5. Lallès, J.-P.; Bosi, P.; Smidt, H.; Stokes, C.R. Weaning—A challenge to gut physiologists. Livest. Sci. 2007, 108, 82–93. [Google Scholar] [CrossRef]
  6. Buffie, C.G.; Pamer, E.G. Microbiota-mediated colonization resistance against intestinal pathogens. Nat. Rev. Immunol. 2013, 13, 790–801. [Google Scholar] [CrossRef]
  7. Kamada, N.; Seo, S.-U.; Chen, G.Y.; Núñez, G. Role of the gut microbiota in immunity and inflammatory disease. Nat. Rev. Immunol. 2013, 13, 321–335. [Google Scholar] [CrossRef]
  8. Halpern, M.D.; Denning, P.W. The role of intestinal epithelial barrier function in the development of NEC. Tissue Barriers 2015, 3, e1000707. [Google Scholar] [CrossRef]
  9. Turner, J.R. Intestinal mucosal barrier function in health and disease. Nat. Rev. Immunol. 2009, 9, 799–809. [Google Scholar] [CrossRef]
  10. Neunlist, M.; van Landeghem, L.; Mahé, M.M.; Derkinderen, P.; Des Varannes, S.B.; Rolli-Derkinderen, M. The digestive neuronal–glial–epithelial unit: A new actor in gut health and disease. Nat. Rev. Gastroenterol. Hepatol. 2013, 10, 90–100. [Google Scholar] [CrossRef]
  11. Ma, X.; Wang, Q.; Li, H.; Xu, C.; Cui, N.; Zhao, X. 16S rRNA genes Illumina sequencing revealed differential cecal microbiome in specific pathogen free chickens infected with different subgroup of avian leukosis viruses. Vet. Microbiol. 2017, 207, 195–204. [Google Scholar] [CrossRef]
  12. Lauridsen, C. Effects of dietary fatty acids on gut health and function of pigs pre-and post-weaning. J. Anim. Sci. 2020, 98, skaa086. [Google Scholar] [CrossRef]
  13. Wall, R.; Ross, R.P.; Ryan, C.A.; Hussey, S.; Murphy, B.; Fitzgerald, G.F.; Stanton, C. Role of gut microbiota in early infant development. Clin. Med. Pediatr. 2009, 3, S2008. [Google Scholar] [CrossRef]
  14. Heo, J.M.; Opapeju, F.O.; Pluske, J.R.; Kim, J.C.; Hampson, D.J.; Nyachoti, C.M. Gastrointestinal health and function in weaned pigs: A review of feeding strategies to control post--weaning diarrhoea without using in--feed antimicrobial compounds. J. Anim. Physiol. Anim. Nutr. 2013, 97, 207–237. [Google Scholar] [CrossRef]
  15. Ley, R.E.; Turnbaugh, P.J.; Klein, S.; Gordon, J.I. Human gut microbes associated with obesity. Nature. 2006, 444, 1022–1023. [Google Scholar] [CrossRef]
  16. Pettigrew, J.E. Fat in swine nutrition. Swine Nutr. 1991, 133–145. [Google Scholar]
  17. Sackner-Bernstein, J.; Kanter, D.; Kaul, S. Dietary intervention for overweight and obese adults: Comparison of low-carbohydrate and low-fat diets. A meta-analysis. PLoS ONE 2015, 10, e0139817. [Google Scholar] [CrossRef]
  18. Kałużna-Czaplińska, J.; Gątarek, P.; Chartrand, M.S.; Dadar, M.; Bjørklund, G. Is there a relationship between intestinal microbiota, dietary compounds, and obesity? Trends Food Sci. Technol. 2017, 70, 105–113. [Google Scholar] [CrossRef]
  19. Liu, C.; Zhang, J.; Li, M.; Zhao, L.; Ji, C.; Ma, Q. Alterations and structural resilience of the gut microbiota under dietary fat perturbations. J. Nutr. Biochem. 2018, 61, 91–100. [Google Scholar] [CrossRef]
  20. Robertson, R.C.; Oriach, C.S.; Murphy, K.; Moloney, G.M.; Cryan, J.F.; Dinan, T.G.; Ross, R.P.; Stanton, C. Omega-3 polyunsaturated fatty acids critically regulate behaviour and gut microbiota development in adolescence and adulthood. Brain Behav. Immun. 2017, 59, 21–37. [Google Scholar] [CrossRef]
  21. Messens, W.; Goris, J.; Dierick, N.; Herman, L.; Heyndrickx, M. Inhibition of Salmonella typhimurium by medium-chain fatty acids in an in vitro simulation of the porcine cecum. Vet. Microbiol. 2010, 141, 73–80. [Google Scholar] [CrossRef] [PubMed]
  22. Boyen, F.; Haesebrouck, F.; Vanparys, A.; Volf, J.; Mahu, M.; van Immerseel, F.; Rychlik, I.; Dewulf, J.; Ducatelle, R.; Pasmans, F. Coated fatty acids alter virulence properties of Salmonella Typhimurium and decrease intestinal colonization of pigs. Vet. Microbiol. 2008, 132, 319–327. [Google Scholar] [CrossRef]
  23. Yang, F.; Zhang, S.; Tian, M.; Chen, J.; Chen, F.; Guan, W. Different sources of high fat diet induces marked changes in gut microbiota of nursery pigs. Front. Microbiol. 2020, 11, 859. [Google Scholar] [CrossRef] [PubMed]
  24. Zou Fang, Z.F.; Yu Bing, Y.B.; He Jun, H.J.; Chen DaiWen, C.D. Effects of Dietary Fat Sources on Growth and Postprandial Lipid and Glucose Metabolism in Rats. Chin. J. Anim. Nutr. 2009, 21, 769–776. [Google Scholar]
  25. Liu ZhongChen, L.Z.; Chen DaiWen, C.D.; Yu Bing, Y.B.; Lv Mei, L.M.; Mao XiangBing, M.X.; Han GuoQuan, H.G.; Chen Hong, C.H.; Mao Qian, M.Q. Effect of Different Fat Sources on Growth Performance and Lipid Metabolism in Weaned Piglets. Chin. J. Anim. Nutr. 2011, 23, 1466–1474. [Google Scholar]
  26. Yang, W.; Jiang, F.; Yu, B.; Huang, Z.; Luo, Y.; Wu, A.; Zheng, P.; Mao, X.; Yu, J.; Luo, J.; et al. Effect of different dietary lipid sources on growth performance, nutrient digestibility, and intestinal health in weaned pigs. Animals 2023, 13, 3006. [Google Scholar] [CrossRef] [PubMed]
  27. Huynh, T.T.; Aarnink, A.J.; Verstegen, M.W.; Gerrits, W.J.; Heetkamp, M.J.; Kemp, B.; Canh, T.T. Effects of increasing temperatures on physiological changes in pigs at different relative humidities. J. Anim. Sci. 2005, 83, 1385–1396. [Google Scholar] [CrossRef]
  28. Zhang, Y.; Xie, H.; Wang, L.; Hu, J.; Wang, L.; Zhang, S. Effect of weaning at 21 days of age on the content of bile acids in chyme of cecum. Animals 2022, 12, 2138. [Google Scholar] [CrossRef] [PubMed]
  29. Franklin, M.A.; Mathew, A.G.; Vickers, J.R.; Clift, R.A. Characterization of microbial populations and volatile fatty acid concentrations in the jejunum, ileum, and cecum of pigs weaned at 17 vs 24 days of age. J. Anim. Sci. 2002, 80, 2904–2910. [Google Scholar] [CrossRef]
  30. Caporaso, J.G.; Kuczynski, J.; Stombaugh, J.; Bittinger, K.; Bushman, F.D.; Costello, E.K.; Fierer, N.; Peña, A.G.; Goodrich, J.K.; Gordon, J.I. QIIME allows analysis of high-throughput community sequencing data. Nat. Methods 2010, 7, 335–336. [Google Scholar] [CrossRef]
  31. Blaxter, M.; Mann, J.; Chapman, T.; Thomas, F.; Whitton, C.; Floyd, R.; Abebe, E. Defining operational taxonomic units using DNA barcode data. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 2005, 360, 1935–1943. [Google Scholar] [CrossRef]
  32. Bokulich, N.A.; Subramanian, S.; Faith, J.J.; Gevers, D.; Gordon, J.I.; Knight, R.; Mills, D.A.; Caporaso, J.G. Quality-filtering vastly improves diversity estimates from Illumina amplicon sequencing. Nat. Methods 2013, 10, 57–59. [Google Scholar] [CrossRef]
  33. Mahaffee, W.F.; Kloepper, J.W. Temporal changes in the bacterial communities of soil, rhizosphere, and endorhiza associated with field-grown cucumber (Cucumis sativus L.). Microb. Ecol. 1997, 34, 210–223. [Google Scholar] [CrossRef]
  34. Pitta, D.W.; Parmar, N.; Patel, A.K.; Indugu, N.; Kumar, S.; Prajapathi, K.B.; Patel, A.B.; Reddy, B.; Joshi, C. Bacterial diversity dynamics associated with different diets and different primer pairs in the rumen of Kankrej cattle. PLoS ONE 2014, 9, e111710. [Google Scholar] [CrossRef] [PubMed]
  35. Oberauner, L.; Zachow, C.; Lackner, S.; Högenauer, C.; Smolle, K.-H.; Berg, G. The ignored diversity: Complex bacterial communities in intensive care units revealed by 16S pyrosequencing. Sci. Rep. 2013, 3, 1413. [Google Scholar] [CrossRef] [PubMed]
  36. Langille, M.G.I.; Zaneveld, J.; Caporaso, J.G.; McDonald, D.; Knights, D.; Reyes, J.A.; Clemente, J.C.; Burkepile, D.E.; Vega Thurber, R.L.; Knight, R. Predictive functional profiling of microbial communities using 16S rRNA marker gene sequences. Nat. Biotechnol. 2013, 31, 814–821. [Google Scholar] [CrossRef] [PubMed]
  37. Shi, Z.; Wang, T.; Kang, J.; Li, Y.; Li, Y.; Xi, L. Effects of weaning modes on the intestinal ph, activity of digestive enzymes, and intestinal morphology of piglets. Animals 2022, 12, 2200. [Google Scholar] [CrossRef]
  38. Wan, F.; Wen, X.; Zhao, H.; Tang, S.; Wang, M.; Yi, B.; Cheng, L.; Lu, Y.; Zhong, R.; Zhang, H. Chemically protected sodium butyrate supplementation improves anti-inflammatory and antioxidant capacities potentially through modulating gut microbiota and short-chain fatty acids levels in piglets. J. Funct. Foods 2024, 121, 106434. [Google Scholar] [CrossRef]
  39. Pitta, D.W.; Pinchak, W.E.; Dowd, S.E.; Osterstock, J.; Gontcharova, V.; Youn, E.; Dorton, K.; Yoon, I.; Min, B.R.; Fulford, J.D. Rumen bacterial diversity dynamics associated with changing from bermudagrass hay to grazed winter wheat diets. Microb. Ecol. 2010, 59, 511–522. [Google Scholar] [CrossRef]
  40. Evans, S.; Martiny, J.B.H.; Allison, S.D. Effects of dispersal and selection on stochastic assembly in microbial communities. ISME J. 2017, 11, 176–185. [Google Scholar] [CrossRef] [PubMed]
  41. Hooper, L.V.; Littman, D.R.; Macpherson, A.J. Interactions between the microbiota and the immune system. Science 2012, 336, 1268–1273. [Google Scholar] [CrossRef] [PubMed]
  42. Nicholson, J.K.; Holmes, E.; Wilson, I.D. Gut microorganisms, mammalian metabolism and personalized health care. Nat. Rev. Microbiol. 2005, 3, 431–438. [Google Scholar] [CrossRef]
  43. Havenaar, R. Intestinal health functions of colonic microbial metabolites: A review. Benef. Microbes 2011, 2, 103–114. [Google Scholar] [CrossRef] [PubMed]
  44. Lin, B.; Gong, J.; Wang, Q.; Cui, S.; Yu, H.; Huang, B. In-vitro assessment of the effects of dietary fibers on microbial fermentation and communities from large intestinal digesta of pigs. Food Hydrocoll. 2011, 25, 180–188. [Google Scholar] [CrossRef]
  45. Mamuad, L.L.; Kim, S.H.; Choi, Y.J.; Soriano, A.P.; Cho, K.K.; Lee, K.; Bae, G.S.; Lee, S.S. Increased propionate concentration in Lactobacillus mucosae–fermented wet brewers grains and during in vitro rumen fermentation. J. Appl. Microbiol. 2017, 123, 29–40. [Google Scholar] [CrossRef] [PubMed]
  46. Xia, Z.; Han, Y.; Wang, K.; Guo, S.; Wu, D.; Huang, X.; Li, Z.; Zhu, L. Oral administration of propionic acid during lactation enhances the colonic barrier function. Lipids Health Dis. 2017, 16, 62. [Google Scholar] [CrossRef]
  47. Hwang, Y.H.; Kim, M.K. Fatty acid composition of rice bran oil and growth-promoting effect of rice bran extract and rice bran oil on Bifidobacterium and Lactobacillus. Appli Biol Chem. 2002, 45, 121–124. [Google Scholar]
  48. Rolinec, M.; Medo, J.; Gábor, M.; Miluchová, M.; Bíro, D.; Šimko, M.; Juráček, M.; Hanušovský, O.; Schubertová, Z.; Gálik, B. The effect of coconut oil addition to feed of pigs on rectal microbial diversity and bacterial abundance. Animals 2020, 10, 1764. [Google Scholar] [CrossRef]
  49. Nighot, M.; Al-Sadi, R.; Guo, S.; Rawat, M.; Nighot, P.; Watterson, M.D.; Ma, T.Y. Lipopolysaccharide-induced increase in intestinal epithelial tight permeability is mediated by toll-like receptor 4/myeloid differentiation primary response 88 (MyD88) activation of myosin light chain kinase expression. Am. J. Pathol. 2017, 187, 2698–2710. [Google Scholar] [CrossRef] [PubMed]
  50. Mukherjee, A.; Lordan, C.; Ross, R.P.; Cotter, P.D. Gut microbes from the phylogenetically diverse genus Eubacterium and their various contributions to gut health. Gut Microbes 2020, 12, 1802866. [Google Scholar] [CrossRef]
  51. Bao, W.; Yu, J.; He, Y.; Liu, M.; Yang, X. The diversity analysis and gene function prediction of intestinal bacteria in three equine species. Front. Microbiol. 2022, 13, 973828. [Google Scholar] [CrossRef] [PubMed]
  52. Portincasa, P.; Bonfrate, L.; Vacca, M.; De Angelis, M.; Farella, I.; Lanza, E.; Khalil, M.; Wang, D.Q.-H.; Sperandio, M.; Di Ciaula, A. Gut microbiota and short chain fatty acids: Implications in glucose homeostasis. Int. J. Mol. Sci. 2022, 23, 1105. [Google Scholar] [CrossRef]
  53. Ma, M.; Ma, S.; Zeng, D.; Huang, X.; Zeng, Y.; Zhu, G.; Chen, L. Temperature-dependent microbial mechanism and accumulation of volatile fatty acids in primary sludge pretreated with peroxymonosulfate. Bioresour. Technol. 2024, 408, 131201. [Google Scholar] [CrossRef] [PubMed]
  54. Nshanian, M.; Gruber, J.J.; Geller, B.S.; Chleilat, F.; Lancaster, S.M.; White, S.M.; Alexandrova, L.; Camarillo, J.M.; Kelleher, N.L.; Zhao, Y.; et al. Short-chain fatty acid metabolites propionate and butyrate are unique epigenetic regulatory elements linking diet, metabolism and gene expression. Nat. Metab. 2025, 7, 196–211. [Google Scholar] [CrossRef]
Agriculture 15 00384 g001
Figure 1. Effects of CGA on 16S rRNA gene copy numbers by quantitative PCR analysis in the cecal (A) and colonic (B) digesta of weaned pigs. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut palm rice oil. a, b, c mean values within a row with different superscript letters were significantly different (p < 0.05).
Figure 1. Effects of CGA on 16S rRNA gene copy numbers by quantitative PCR analysis in the cecal (A) and colonic (B) digesta of weaned pigs. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut palm rice oil. a, b, c mean values within a row with different superscript letters were significantly different (p < 0.05).
Agriculture 15 00384 g001
Agriculture 15 00384 g002
Figure 2. Rarefaction curves. (A,C) Sparsity and rank-abundance curves of cecum samples; (B,D) Sparsity and rank-abundance curves of colon samples. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Figure 2. Rarefaction curves. (A,C) Sparsity and rank-abundance curves of cecum samples; (B,D) Sparsity and rank-abundance curves of colon samples. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Agriculture 15 00384 g002
Agriculture 15 00384 g003
Figure 3. The number of OTUs in the cecum and colon of two comparison methods. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Figure 3. The number of OTUs in the cecum and colon of two comparison methods. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Agriculture 15 00384 g003
Agriculture 15 00384 g004
Figure 4. Unweighted UniFrac distance was employed to conduct a PCoA of the gut microbiota in both cecum (A) and colon (B) digesta, with eight samples per group. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Figure 4. Unweighted UniFrac distance was employed to conduct a PCoA of the gut microbiota in both cecum (A) and colon (B) digesta, with eight samples per group. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Agriculture 15 00384 g004
Agriculture 15 00384 g005
Figure 5. The relative abundance of bacterial communities at the phylum level. (A) The relative abundance of species at the phylum level in cecal chyme; (B) the relative abundance of species at the phylum level in colonic chyme. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Figure 5. The relative abundance of bacterial communities at the phylum level. (A) The relative abundance of species at the phylum level in cecal chyme; (B) the relative abundance of species at the phylum level in colonic chyme. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Agriculture 15 00384 g005
Agriculture 15 00384 g006
Figure 6. The relative abundance of bacterial communities at the genus level. (A) The relative abundance of species at the genus level in cecal chyme; (B) the relative abundance of species at the genus level in colonic chyme. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Figure 6. The relative abundance of bacterial communities at the genus level. (A) The relative abundance of species at the genus level in cecal chyme; (B) the relative abundance of species at the genus level in colonic chyme. SO, soybean oil; PRFO, fish palm rice oil; PRCO, coconut-palm-rice oil.
Agriculture 15 00384 g006
Agriculture 15 00384 g007
Figure 7. Effects of different fat combinations on the structural composition of caecum and colon of phylum in weaned piglets. (A) The relative abundance of species at the phylum level in cecal chyme. (B) the relative abundance of species at the phylum level in colonic chyme. SO, soybean oil; PRFO, fish palm rice oil, PRCO, coconut palm rice oil.
Figure 7. Effects of different fat combinations on the structural composition of caecum and colon of phylum in weaned piglets. (A) The relative abundance of species at the phylum level in cecal chyme. (B) the relative abundance of species at the phylum level in colonic chyme. SO, soybean oil; PRFO, fish palm rice oil, PRCO, coconut palm rice oil.
Agriculture 15 00384 g007
Agriculture 15 00384 g008
Figure 8. Functional annotation of the bacterial communities. (A) Functional annotation of the bacterial communities in cecum; (B) Functional annotation of the bacterial communities in colon. SO, soybean oil; PRFO, fish palm rice oil, PRCO, coconut palm rice oil.
Figure 8. Functional annotation of the bacterial communities. (A) Functional annotation of the bacterial communities in cecum; (B) Functional annotation of the bacterial communities in colon. SO, soybean oil; PRFO, fish palm rice oil, PRCO, coconut palm rice oil.
Agriculture 15 00384 g008
Agriculture 15 00384 g009
Figure 9. Functional prediction at the COG level of cecum and colon microbes. SO, soybean oil; PRFO, fish palm rice oil, PRCO, coconut palm rice oil.
Figure 9. Functional prediction at the COG level of cecum and colon microbes. SO, soybean oil; PRFO, fish palm rice oil, PRCO, coconut palm rice oil.
Agriculture 15 00384 g009
Table 1. Effect of different lipid sources on performance and the content of volatile fatty acids in cecal and colonic chyme of weaned piglets.
Table 1. Effect of different lipid sources on performance and the content of volatile fatty acids in cecal and colonic chyme of weaned piglets.
ITEMTreatmentsSEMp-Value 4
SOPRFOPRCO
ADG, g/d362.96378.27380.199.470.736
ADFI, g/d611.43604.52633.1511.920.736
FCR1.691.601.660.020.436
Cecum(mmol/L)
AA1500.941926.481815.3986.040.108
PA626.32 b895.49 a970.01 a54.660.023
IBA35.7528.2739.184.780.671
BA507.17 b697.27 ab877.14 a58.080.028
IVA51.4160.4674.856.480.347
VA40.31 b82.19 ab153.16 a17.210.021
pH value 6.35 a6.27 b6.28 ab0.0150.042
Colon(mmol/L)
AA1452.851247.521168.2463.730.153
PA800.75902.13842.9883.320.866
IBA50.8445.6640.894.390.561
BA650.45652.58586.3438.770.775
IVA93.9499.3885.887.690.775
VA42.23 b58.59 b141.99 a16.100.012
pH value 6.516.506.510.0070.834
AA, acetic acid; PA, propionic acid; IBA, isobutyric acid; BA, butyric acid; IVA, isovaleric acid; VA, valeric acid. 1 Mean and total SEM are listed in separate columns (n = 8). 2 a, b mean values within a row with different superscript letters were significantly different (p < 0.05). 3 SO pigs were fed with a basal diet; PRFO pigs were fed with 2% mixed oil 1; PRCO pigs were fed with 2% mixed oil 2. 4 The p-values indicating significance were obtained by comparing the means between treatment groups using Duncan’s multiple range test in a one-way ANOVA.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Yang, K.; Yang, W.; Jiang, F.; Yu, B.; Huang, Z.; Luo, Y.; Wu, A.; Zheng, P.; Mao, X.; Yu, J.; et al. Effects of Fish Palm Rice and Coconut Palm Rice Oil Mixture on Intestinal Health of Weaned Piglets. Agriculture 2025, 15, 384. https://doi.org/10.3390/agriculture15040384

AMA Style

Yang K, Yang W, Jiang F, Yu B, Huang Z, Luo Y, Wu A, Zheng P, Mao X, Yu J, et al. Effects of Fish Palm Rice and Coconut Palm Rice Oil Mixture on Intestinal Health of Weaned Piglets. Agriculture. 2025; 15(4):384. https://doi.org/10.3390/agriculture15040384

Chicago/Turabian Style

Yang, Kaiyun, Wenjuan Yang, Fei Jiang, Bing Yu, Zhiqing Huang, Yuheng Luo, Aimin Wu, Ping Zheng, Xiangbing Mao, Jie Yu, and et al. 2025. "Effects of Fish Palm Rice and Coconut Palm Rice Oil Mixture on Intestinal Health of Weaned Piglets" Agriculture 15, no. 4: 384. https://doi.org/10.3390/agriculture15040384

APA Style

Yang, K., Yang, W., Jiang, F., Yu, B., Huang, Z., Luo, Y., Wu, A., Zheng, P., Mao, X., Yu, J., Luo, J., Yan, H., & He, J. (2025). Effects of Fish Palm Rice and Coconut Palm Rice Oil Mixture on Intestinal Health of Weaned Piglets. Agriculture, 15(4), 384. https://doi.org/10.3390/agriculture15040384

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop