Skip to main content
NCBI home page
ACS AuthorChoice logoLink to ACS AuthorChoice
. 2026 Jun 12;74(24):18757–18767. doi: 10.1021/acs.jafc.6c02576

Galacto-Oligosaccharides Reshape Mucin O‑Glycan Profiles to Enhance Small Intestinal Barrier Development in Early Life

Laipeng Xu †,‡, Xuan Li †, Wenlu Fan †, Weile Sun †, Chengming Meng †, Li Liu §, Josef Voglmeir §, Chunlong Mu ∥, Weiyun Zhu †,§,*
PMCID: PMC13307372  PMID: 42284521

Abstract

The mucin barrier is essential for postnatal gut homeostasis, but whether galacto-oligosaccharides (GOS) regulate mucin glycosylation remains unclear. Here, we investigated the effects of early-life GOS supplementation on mucin O-glycans and intestinal barrier development in piglets using UPLC, MALDI-TOF-MS, immunofluorescence, lectin staining, and histomorphological analyses. GOS supplementation significantly increased jejunal villus width, mucosal thickness, goblet cell numbers, and MUC2 expression during early life. Glycomic profiling revealed selective remodeling of mucin O-glycans, characterized by increased core 1/3 and α2,3-sialylated glycans and reduced core 2/4 and α2,6-sialylated structures. These alterations were accompanied by upregulation of C1GALT1, GALNTs, and ST3GALs, and downregulation of ST6GALNACs and neuraminidases. Lectin staining further confirmed enhanced α2,3-sialylation and reduced α2,6-sialylation. Collectively, these results demonstrate that dietary GOS promotes intestinal barrier development and selectively modulates the composition and sialylation of mucin O-glycans, providing novel mechanistic insight into how prebiotics support gut health in early life.

Keywords: Galacto-oligosaccharides, Mucosal barrier, MUC2, Mucin O-glycans, Piglet

graphic file with name jf6c02576_0010.jpg

1. Introduction

In modern intensive pig production systems, neonatal and suckling piglets are particularly susceptible to intestinal dysfunction due to the immaturity of their digestive and immune systems. , During this critical developmental window, the intestinal mucus layer serves as the first line of defense against luminal microbes and toxins, and its integrity largely depends on adequate mucin secretion and glycosylation. , Disruption of the mucus barrier is frequently associated with epithelial injury, microbial imbalance, and chronic inflammation, which consequently impair growth performance and increase the incidence of diarrhea and mortality. , Therefore, early life nutritional strategies that support mucus barrier formation and maturation have become essential for improving piglet health and production sustainability. ,

The intestinal mucus layer constitutes the first line of defense separating luminal microbes from the epithelium. , Its structural and functional properties largely depend on the O-linked glycans (O-glycans) decorating mucin glycoproteins secreted by goblet cells. , These O-glycans not only determine the physicochemical properties of mucus but also act as key mediators in host-microbe interactions, influencing bacterial adhesion, colonization patterns, and immune modulation. , Alterations in mucin O-glycosylation have been closely associated with intestinal barrier dysfunction and inflammatory responses in both humans and animals. − However, whether early life nutritional interventions can modulate mucin-type O-glycosylation and thereby improve mucus barrier development in piglets remains largely unexplored.

Among functional prebiotics, galacto-oligosaccharides (GOS), nondigestible oligosaccharides derived from lactose, have gained increasing attention. A number of studies have demonstrated that GOS exert beneficial impact on gut health through selectively promoting the growth and its metabolism of beneficial bacteria. , Beyond these conventional effects, emerging evidence suggests that GOS may directly interact with host glycosylation pathways. The β-galactoside residues present in certain GOS (particularly those containing lactose or β-linked galactose) may resemble terminal galactose units on mucin O-glycans, leading to the hypothesis that GOS may influence mucin synthesis or glycan remodeling. , Therefore, GOS may function as modulators of mucosal glycosylation and mucus barrier function.

This study explored the impact of early life GOS supplementation on intestinal barrier function and mucin-type O-glycan profiles using a piglet model. By integrating O-glycan profiling, glycosyltransferase expression analysis, and lectin staining, this work aimed to elucidate the mechanism by which early life GOS supplementation regulates intestinal mucosal development and mucin MUC2 O-glycosylation, thereby contributing to improved intestinal barrier function. The findings provide new mechanistic insights into prebiotic-based nutritional regulation of gut health during early life and support the development of targeted nutritional strategies for sustainable pig production.

2. Materials and Methods

2.1. Ethics Statement

All animal procedures were conducted following approval by the Ethics Committee of Nanjing Agricultural University (Nanjing, China) and in accordance with the Chinese Regulations for the Administration of Experimental Animals (Approval No. SYXK 2022-0073).

2.2. Experimental Design and Animal Management

The present study was a part of series of studies designed to investigate whether early life GOS supplementation can impact intestinal microbiome and improve intestinal development and growth performance of the pigs, data on growth performance and intestinal mucosal microbe response after GOS supplementation have been previously reported. The experiment was conducted at a commercial pig farm with the experimental design shown in Supplementary Figure S1, and detailed experiment procedures have been described previously. Briefly, ten gestating Landrace × Large White sows (parity range: 2–3) at the same physiological stage were selected, all mated with the same Duroc boar. To synchronize farrowing, all sows were intramuscularly injected with cloprostenol sodium. After farrowing, four healthy and body weight-uniform piglets (balanced for sex) were selected from each litter and randomly divided into two groups (CON and GOS), with two piglets per group per litter, resulting in 10 replicates per group and 2 piglets per replicate (total 40 piglets). GOS (Guangdong Quantum Hi-Tech Bio Co., Ltd., China; degree of polymerization (DP) > 2 and purity > 98%) administration began on postnatal day 1. Piglets in the GOS group were orally administered a GOS solution at a dose of 1 g/kg/day based on body weight, while the control (CON) group received an equal volume of saline. Piglets were fed sow milk or GOS solution daily until 21 d. At 21 d, GOS administration was ceased, all piglets were weaned, housed individually and fed a diet until 49 d, with the diet composition reported previously.

2.3. Sample Collection

Sample collection was described in our previous study. Briefly, at 21 and 49 d, one piglet per litter was randomly selected for slaughter (n = 10). The jejunum was isolated. Intestinal mucosa was gently scraped from a portion of the tissue using a sterile surgical blade, rinsed with freshly prepared diethyl pyrocarbonate (DEPC)-treated water, collected into sterile RNase/DNase-free EP tubes, and stored at −80 °C for subsequent RT-qPCR analysis. Another portion of intestinal tissue was gently rinsed with saline to remove surface debris and feces. Mucus was then carefully collected from the intestinal surface using sterile slides, transferred into sterile, protease-free EP tubes, and stored at −80 °C for glycomics analysis. In addition, intact intestinal tissues were collected and fixed in 4% paraformaldehyde for histological analysis.

2.4. Histological Analysis

2.4.1. Sialylation of Piglet Intestinal Mucosa

Sialylation was assessed by lectin staining following Xu et al. Briefly, paraffin sections were deparaffinized, hydrated, and permeabilized with 0.5% Triton X-100. Antigen retrieval was performed with sodium citrate buffer. Hydrophobic barriers were drawn, and sections were blocked with 5% BSA. Primary antibodies were SNA-FITC (1:200, Vectorlabs, USA) and MALII (1:200, Vectorlabs, USA). Secondary antibodies were FITC- or CY3-labeled streptavidin (1:200, Solarbio, Beijing). Sections were mounted with DAPI-containing antifade reagent and imaged using a ZEISS confocal microscope. Mean fluorescence intensity of α2, 3- and α2, 6-linked sialic acids was quantified using ImageJ (NIH, USA). The integrated optical density (IOD), defined as the product of the stained area and the mean fluorescence intensity, was measured in at least 3 randomly selected fields per section. Background fluorescence was subtracted prior to analysis, and the average value for each sample was calculated.

2.4.2. MUC2 Expression and Colocalization with Sialic Acids

MUC2 immunofluorescence was performed as above, substituting the primary antibody with MUC2 (1:200, Rabbit, Servicebio, Wuhan) and the secondary antibody with Alexa Fluor 594 (1:200, Goat, Abways, Beijing). For colocalization, MUC2 was combined with SNA or MALII lectin, and secondary antibody Alexa Fluor 594 was combined with FITC-labeled streptavidin (all 1:200). Imaging was performed with an LSM900 confocal microscope. Excitation/emission wavelengths: DAPI 330–380/420 nm (blue), FITC 465–495/515–555 nm (green), CY3 510–560/590 nm (red). Colocalization appears as yellow fluorescence (red + green).

2.4.3. H&E and PAS Staining of Piglet Intestinal Tissue

H&E staining: Sections were deparaffinized and hydrated, stained with hematoxylin, followed by eosin staining, dehydrated, and mounted. PAS staining: Sections were deparaffinized and hydrated, oxidized with periodic acid, rinsed with ultrapure water, stained with Schiff’s reagent in the dark, developed under running water, dehydrated, and mounted. All stained sections were observed and imaged under a light microscope. ImageJ 1.43u and ImageJ Pro Plus 6.0 were used to quantify intestinal histological parameters, including villus height, crypt depth, and goblet cell number.

2.5. Analysis of O-Glycan Structure and Abundance in Piglet Intestinal Mucus

2.5.1. Extraction of Mucins and O-Glycan Release (β-Elimination)

Mucus O-glycans were extracted and analyzed following our previous protocol. Briefly, 1 g of piglet intestinal mucus was ground in liquid nitrogen and suspended in 7.5 mL protein extraction buffer (6 M GuHCl, 0.1 M Tris-HCl, 1 mM EDTA, 0.1 mM PMSF, 5 mM NEM, pH 8.0). The mixture was shaken overnight at 4 °C, 180 rpm, and centrifuged at 20 000 × g for 1 h at 4 °C. The supernatant was reduced with 100 mM DTT at 37 °C overnight, followed by alkylation with 250 mM IAA in the dark for 4 h. After dialysis using 10 kDa MWCO cassettes (Thermo Scientific), mucins were lyophilized.

For O-glycan release, 20 mg of lyophilized mucin was incubated with 900 μL saturated ammonia at 60 °C for 16 h, with brief venting at the start to prevent tube rupture. Ammonia was removed under vacuum, and repeated cycles of dissolution in ultrapure water and drying were performed to remove residual salts. Samples were then acidified with 50 μL 1% formic acid at room temperature in the dark for 40 min, evaporated, and reconstituted in 500 μL ultrapure water for purification.

2.5.2. Purification and Fluorescent Derivatization of O-Glycans

After centrifugation at 12 000 rpm for 10 min, the supernatant was applied to a Supelclean ENVI-Carb carbon column preconditioned with 80% acetonitrile containing 0.1% TFA and ultrapure water. Columns were washed sequentially with ultrapure water, 20% acetonitrile, and 40% acetonitrile (0.1% TFA). Eluates were pooled, dried, and half of each sample was subjected to derivatization with 10 μL 2-aminobenzamide (2-AB) solution (DMSO/glacial acetic acid) at 65 °C for 4 h for fluorescent labeling.

2.5.3. UPLC and MALDI-TOF-MS Analysis of O-Glycans

After release and purification, O-glycans were fluorescently labeled with 2-AB using a commercial glycan labeling kit according to the manufacturer’s instructions. Briefly, dried O-glycan samples were dissolved in labeling reagent containing 2-AB and sodium cyanoborohydride and incubated at 65 °C for 4 h to allow reductive amination. Excess labeling reagents were removed using a cleanup cartridge supplied in the kit, and the purified labeled glycans were dried under vacuum and stored at −20 °C until analysis.

2.5.4. UPLC Analysis

UPLC analysis was performed using a Shimadzu LC-30A system (Shimadzu, Japan). Prior to injection, 10 μL of the 2-AB-labeled glycan sample was mixed with 35 μL acetonitrile and 5 μL ultrapure water, vortexed thoroughly, and centrifuged at 12 000 × g for 10 min. The supernatant was transferred to an autosampler vial and 10 μL was injected for analysis. Glycan separation was carried out on an ACQUITY UPLC Glycan BEH Amide column (130 Å, 1.7 μm, 2.1 mm × 150 mm, Waters). The column temperature was maintained at 60 °C and the flow rate was set to 0.4 mL/min. The mobile phases consisted of solvent A (100 mM ammonium formate, pH 4.4) and solvent B (acetonitrile). A linear gradient elution was applied as follows: from 75% B to 62% B over 40 min, followed by washing and re-equilibration of the column. Fluorescence detection was performed with an excitation wavelength of 330 nm and an emission wavelength of 420 nm. Chromatograms were acquired and processed using the Shimadzu LabSolutions software.

2.5.5. MALDI-TOF-MS Analysis

For mass spectrometric analysis, the remaining 2-AB-labeled glycan samples were dried under vacuum and reconstituted in ultrapure water. One microliter of the sample solution was mixed with 1 μL of 2,5-dihydroxybenzoic acid (DHB, 10 mg/mL in 50% acetonitrile containing 0.1% trifluoroacetic acid) and spotted onto a stainless-steel MALDI target plate for cocrystallization. Mass spectrometric analysis was performed using a Bruker Autoflex Speed MALDI-TOF/TOF mass spectrometer (Bruker Daltonics, USA). The instrument was calibrated externally using a standard peptide calibration mixture, achieving a mass accuracy of approximately 0.1 Da. Spectra were acquired in positive ion reflector mode with an accelerating voltage of 20 kV, laser intensity set to 7 000, and a mass range of m/z 1000–3000.

2.5.6. Data Processing and Structural Assignment

Raw mass spectra were processed using Bruker FlexAnalysis 3.3 software. Peaks with a signal-to-noise ratio greater than 4 were considered for further analysis, and the [M + H]+ ions were selected for annotation. Monoisotopic masses were extracted and compared with theoretical O-glycan compositions using GlycoWorkbench v1.1 software. Putative O-glycan structures were assigned based on accurate mass matching, known biosynthetic pathways, and previously reported mucin-type O-glycan structures.

In this study, O-glycan structures were identified by combining UPLC with MALDI-TOF-MS. First, UPLC was used to separate the sample, allowing preliminary differentiation of O-glycan components with different structures based on their chromatographic retention times. Subsequently, the fractions corresponding to each chromatographic peak were collected, and their exact molecular weights were determined by MALDI-TOF-MS. Finally, by integrating the molecular weight information with known O-glycan biosynthetic pathways and structural databases, the mass data were matched with potential glycan compositions, thereby assigning structures to the chromatographic peaks. This approach leveraged the high separation capability of UPLC and the high-sensitivity structural analysis of MALDI-TOF-MS, enabling systematic identification of O-glycans in complex samples

2.6. mRNA Preparation and Quantification by Real-Time PCR

Total RNA was extracted from piglet intestinal mucosa using TRIzol reagent (AGbio, China), and reverse transcription polymerase chain reaction (RT-PCR) was performed using Evo M-MLV Reverse Transcriptase (AGbio, China). Real-time qPCR was conducted using SYBR Green Pro Taq HS and the QuantStudio6 system (Thermo, USA). The ACTB gene was used as a housekeeping gene, and the relative expression of target genes was determined using the 2–ΔΔCt method. The primers used for qPCR of piglet intestinal mucosa tissue are listed in Supplementary Table S1.

2.7. Western Blot Analysis

To validate the transcriptional results at the protein level, three representative enzymes involved in different stages of mucin-type O-glycosylation were selected, including C1GALT1 (core 1 backbone synthesis, Abmart), GALNT12 (glycosylation initiation, Abmart), and NEU1 (terminal desialylation, Proteintech) for Western blot analysis. Proteins were separated by SDS-PAGE and transferred onto PVDF membranes. After blocking, membranes were incubated with primary antibodies against C1GALT1, GALNT12, and NEU1, followed by HRP-conjugated secondary antibodies. Protein bands were visualized using enhanced chemiluminescence. Band intensities were quantified by densitometric analysis using ImageJ (NIH, USA). Relative protein expression levels were normalized to β-actin as the internal loading control.

2.8. Correlation Analysis

Spearman correlation analysis was performed to evaluate the relationships among mucosal morphology parameters, MUC2 expression, goblet cell number, key O-glycan structures, and the expression of glycosyltransferases. Correlation analysis was performed using the OmicStudio tools at https://www.omicstudio.cn/tool.

2.9. Statistical Analysis

All experimental data were analyzed using GraphPad Prism v9.0 (GraphPad Software, USA). The two-way ANOVA test was used to compare differences between groups. Data are presented as mean ± standard error of the mean (SEM). *P < 0.05 and **P < 0.01.

3. Results

3.1. Intestinal Morphology and Mucin Expression

To evaluate whether early life GOS supplementation influences intestinal mucosal development, jejunal morphology and goblet cell differentiation were first examined at both 21 and 49 d of age. Histological analysis showed that GOS supplementation during the suckling period markedly improved the structural maturation of the jejunal mucosa. At 21 d, piglets receiving GOS exhibited significantly increased villus width, mucosal thickness, and goblet cell numbers compared with controls (P < 0.01). At 49 d, although goblet cell number remained similar to the control group, villus width and mucosal thickness were significantly higher in the GOS group (P < 0.01), indicating a sustained effect of early GOS intervention on mucosal architecture (Figure A, B). Two-way ANOVA further revealed significant time-dependent effects and diet × time interactions for villus width, mucosal thickness, and goblet cell numbers (P < 0.05), suggesting that the impact of GOS on intestinal morphology varied across developmental stages and was particularly pronounced during the suckling period. Given that goblet cells are the primary source of mucin, MUC2 expression was subsequently examined. Immunofluorescence staining demonstrated that GOS supplementation markedly enhanced MUC2 protein abundance in jejunal goblet cells at both 21 and 49 d (P < 0.01 or P < 0.05) (Figure A, B). This was further supported by RT-qPCR analysis, which showed a significant upregulation of MUC2 mRNA expression in the jejunal mucosa of GOS-treated piglets (P < 0.01) (Figure C). Notably, MUC2 expression exhibited a significant time-dependent response to GOS supplementation (P < 0.01 or P < 0.05), indicating that early life GOS intervention not only increased mucin production but also influenced its developmental trajectory. To further evaluate the effects of GOS supplementation on intestinal barrier integrity, the mRNA expression levels of representative tight junction proteins, including ZO-1, occludin, and claudin-1, were determined in the jejunal mucosa of piglets. As shown in Figure D, dietary GOS supplementation significantly increased the mRNA expression of ZO-1, occludin, and claudin-1 compared with the control group at 21 d (P < 0.05). These results indicate that GOS promotes the transcriptional expression of key tight junction components, suggesting an improvement in epithelial barrier integrity. Together, these results demonstrate that GOS supplementation during the suckling period promotes intestinal mucosal maturation and enhances the secretory capacity of goblet cells, thereby strengthening the physical basis of the mucus barrier.

1.

1

Open in a new tab

Intestinal morphology and the number of goblet cells of piglets. (A) HE and PAS staining of jejunum. (B) Villus width, mucosal layer thickness, number and density of goblet cells of jejunum. Scale bar = 300 μm. n = 10. Graphs represent mean ± SEM. The two-way ANOVA test was used to compare differences between groups. *P < 0.05, **P < 0.01.

2.

2

Open in a new tab

The expression level of intestinal mucosal MUC2 and tight junctions in piglets. (A) Immunofluorescence staining of MUC2 in jejunal mucosa of piglets. (B) The average fluorescence intensity of MUC2 in jejunal mucosa of piglets. (C) The relative expression level of MUC2 in jejunal mucosa of piglets. (D) The relative expression level of tight junction-related genes (ZO-1, Occludin and Claudin-1) in the jejunal mucosa. Scale bar = 50 μm. n = 10. Graphs represent mean ± SEM. The two-way ANOVA test was used to compare differences between groups. *P < 0.05, **P < 0.01.

3.2. Composition and Abundance of Intestinal Mucin O-Glycans

To determine whether the GOS-induced enhancement of mucin production was accompanied by alterations in O-glycosylation patterns, the composition of jejunal mucin O-glycans was analyzed using UPLC and MALDI-TOF-MS. At 21 d of age, a total of 19 distinct O-glycan structures were identified in the jejunal mucosa (Figure A and Table ), indicating a relatively high structural diversity of mucin glycosylation during the suckling period. Among these, GOS supplementation induced selective and structure-specific remodeling of O-glycan profiles. Within core 1 structures, the relative abundances of glycans with m/z 526 and 793 in the GOS group were significantly higher than those in the CON group, whereas those of glycans with m/z 607 and 688 were lower. In core 2 structures, glycan 870 and in core 3, glycans 544 and 867 in GOS group were significantly lower in abundance than in the CON group, while in core 4, glycan 930 and glycan 1153 were decreased as compared with those in CON group. At 49 d of age, a total of 17 O-glycans were identified (Figure B and Table ), suggesting a moderate simplification of glycan diversity with postweaning maturation. In this stage, GOS still exerted measurable effects, although the number of responsive glycan species was reduced compared with the suckling stage. Specifically, within core 1 structures, the relative abundances of glycans 793 and 941 in GOS group were significantly higher than in the CON group, whereas the abundance of glycan 898 became lower. In core 4, the abundance of glycan 930 was higher while that of glycan 1153 in the GOS group was lower compared with the CON group, indicating that some of the GOS-induced glycan remodeling patterns persisted beyond the suckling period. These results demonstrate that GOS supplementation induces a targeted and developmentally regulated remodeling of mucin O-glycan composition rather than a nonspecific global shift.

3.

3

Open in a new tab

The structure and relative abundance of O-glycans in the intestinal mucus of piglets. (A) UPLC spectrum in jejunal mucus O-glycans of 21-day-old piglets. (B) UPLC spectrum in jejunal mucus O-glycans of 49-day-old piglets. (C) Relative abundance of jejunal mucus fucosylated O-glycans of piglets. (D) Relative abundance of jejunal mucus sialylated O-glycans of piglets. (E) Relative abundance of jejunal mucus sialylated-fucosy O-glycans of piglets. (F) Relative abundance of jejunal mucus core sialylated O-glycans of piglets. n = 10. Graphs represent mean ± SEM. The two-way ANOVA test was used to compare differences between groups. *P < 0.05, **P < 0.01.

1. Structures and Relative Abundances of O-Glycans in Small Intestinal Mucus of 21-Day-Old Piglets.

3.2.

Open in a new tab

2. Structures and Relative Abundances of O-Glycans in Small Intestinal Mucus of 49-Day-Old Piglets.

3.2.

Open in a new tab

3.3. GOS Selectively Remodels Mucin O-Glycan Core Composition and Sialic Acid Linkage Patterns

O-glycans were classified according to fucosylation, sialylation and sialic acid linkage patterns. At both 21 and 49 d, no significant differences were observed in the total levels of fucosylation, sialylation, or fucosylated sialylation between groups (P > 0.05) (Figure C–E), indicating that GOS did not globally alter the overall extent of terminal modifications. Core 1 O-glycans were the dominant structural type at both ages, accounting for the majority of total O-glycans (Supplementary Figure S2A). However, at 21 d, GOS significantly increased the relative abundance of core 1 O-glycans and α2,3-sialylated O-glycans, while decreasing core 2, core 4, and α2,6-sialylated O-glycans (P < 0.01 or P < 0.05) (Figure F, Supplementary Figure S2A). In contrast, at 49 d, the effect of GOS was mainly reflected in sialic acid linkage patterns, with a significant increase in α2,3-sialylated O-glycans and a decrease in α2,6-sialylated O-glycans (P < 0.05), whereas the core distribution was no longer significantly affected. Two-way ANOVA revealed significant time-dependent effects for fucosylation, sialylation, core 2/4 structures, and α2,3/α2,6-sialylation (P < 0.01 or P < 0.05) (Figure C–F). In addition, long-chain O-glycans accounted for 36.9% and 33.3% of total glycans at 21 and 49 d, respectively (Supplementary Figure S2B and Supplementary Table S2), and the overall chain-length distribution patterns are shown in Supplementary Figure S2C, indicating that postnatal maturation is accompanied by moderate structural simplification. Notably, glycomics analysis showed that the maximum chain length detected in jejunal samples was six monosaccharide residues. Based on this distribution, we defined glycans containing ≤3 residues as short-chain and those with >3 residues as long-chain, in order to distinguish relatively simple core structures from more elaborated glycan forms.

3.4. Relative Expression of Glycosyltransferases

To explore the molecular basis underlying GOS-induced O-glycan remodeling, expression of key glycosyltransferases involved in O-glycan initiation, elongation, and sialylation were examined in the jejunal mucosa. Given that galactose and N-acetylgalactosamine are essential substrates for mucin O-glycosylation, enzymes responsible for these pathways were first analyzed. At 21 d, GOS significantly upregulated the expression of T-synthase (C1GALT1 and its molecular chaperone C1GALT1C1) and multiple polypeptide N-acetylgalactosaminyltransferases (GALNT1, GALNT2, GALNT6, GALNT7) (P < 0.01 or P < 0.05), indicating an enhanced capacity for O-glycan initiation and core 1 biosynthesis (Figure A). At 49 d, only GALNT2 remained significantly elevated (P < 0.01), suggesting that the regulatory effect of GOS on O-glycan biosynthesis is strongest during early life. Sialyltransferase expression exhibited a clear linkage-specific regulatory pattern. GOS markedly downregulated α2,6-sialyltransferases (ST6GALNAC1, ST6GALNAC2, ST6GALNAC4) while upregulating α2,3-sialyltransferases (ST3GAL1, ST3GAL2, ST3GAL3, ST3GAL4, ST3GAL6) (P < 0.01 or P < 0.05) (Figure B), consistent with the observed shift in sialic acid linkage distribution in O-glycan profiles. In addition, neuraminidases (NEU1, NEU2, NEU3, and NEU4), which regulate the removal and turnover of sialic acids, were significantly decreased in response to GOS supplementation (P < 0.01 or P < 0.05) (Figure C), suggesting a coordinated regulation of both sialic acid synthesis and degradation pathways. The Western blot results (Supplementary Figure S3) showed expression patterns consistent with the mRNA transcription trends presented in Figure , supporting the reliability of the gene expression profiling data.

4.

4

Open in a new tab

The relative expression levels of T-synthase, N-acetylgalactosaminyltransferase and sialyltransferase in the intestine. (A) The relative expression levels (mRNA) of T-synthase and N-acetylgalactosaminyltransferase in the jejunum. (B) The relative expression levels (mRNA) of sialyltransferase in the jejunum. n = 10. (C) The relative expression levels (mRNA) of jejunal neuraminidases (NEU1-NEU4). Graphs represent mean ± SEM. The two-way ANOVA test was used to compare differences between groups. *P < 0.05, **P < 0.01.

3.5. Levels of Sialylation and Sialylated MUC2 in the Intestinal Mucosa

To validate whether the transcriptional and structural changes in O-glycosylation were reflected at the tissue level, lectin staining was used to visualize α2,3- and α2,6-linked sialic acids in the jejunal mucosa. Consistent with the glycomic and gene expression data, GOS supplementation significantly increased α2,3-sialic acid fluorescence intensity while markedly decreasing α2,6-sialic acid signals at both 21 and 49 d (P < 0.01) (Figure A–C). Two-way ANOVA revealed significant time-dependent effects for both sialylation patterns (P < 0.01) (Figure B, C). Further, at 49 d of age, α2,6-linked sialic acid signals were nearly undetectable, and no apparent colocalization between α2,3- and α2,6-linked sialic acids was observed (Figure A), indicating a strong developmental shift toward α2,3-dominant sialylation. To further determine whether these changes occurred specifically on mucins, colocalization analysis of MUC2 with sialic acids was performed. The results showed that GOS significantly increased α2,3-sialylated MUC2 while simultaneously decreasing α2,6-sialylated MUC2 at both time points (P < 0.01) (Figure A–C). Moreover, these effects exhibited significant time-dependent patterns (P < 0.01 or P < 0.05) (Figure B, C), indicating that GOS not only alters overall sialylation but specifically reprograms the glycosylation pattern of the major secreted mucin MUC2.

5.

5

Open in a new tab

The alterations in sialylation levels and their colocalization in the intestinal mucosa of piglets. (A) Lectin staining colocalization of α2,3 sialic acid (MALII) with α2,6 sialic acid (SNA) in the jejunal mucosa of piglets. (B) The average fluorescence intensity of α2,3 sialic acid (MALII) in the jejunal mucosa of piglets. (C) The average fluorescence intensity of α2,6 sialic acid (SNA) in the jejunal mucosa of piglets. Note: The co-expression of SNA and MALII was not detected. Scale bar = 100 μm. n = 10. Graphs represent mean ± SEM. The two-way ANOVA test was used to compare differences between groups. *P < 0.05, **P < 0.01.

6.

6

Open in a new tab

The sialylation level of intestinal mucosal MUC2 in piglets. (A) Immunofluorescence colocalization of intestinal mucosal protein MUC2 with α2,3 sialic acid and α2,6 sialic acid in the jejunum of piglets. (B) The average fluorescence intensity of intestinal mucosal protein MUC2 with α2,3 sialic acid in the jejunum of piglets. (C) The average fluorescence intensity of intestinal mucosal protein MUC2 with α2,6 sialic acid in the jejunum of piglets. Scale bar = 50 μm. n = 10. Graphs represent mean ± SEM. The two-way ANOVA test was used to compare differences between groups. *P < 0.05, **P < 0.01.

3.6. Correlation Analysis among Glycosyltransferases, O-Glycan Structures, and Mucosal Barrier Parameters

Correlation analysis revealed coordinated associations among glycosyltransferase expression, O-glycan structural features, and mucosal barrier parameters. Core 1 O-glycans and α2,3-sialylation were positively correlated with MUC2 expression, goblet cell number, and mucosal thickness, whereas core 2 O-glycans and α2,6-sialylation exhibited negative correlations with these parameters. In addition, C1GALT1 and ST3GAL1 showed strong positive correlations with their corresponding glycan structures, suggesting transcriptional regulation of mucin O-glycosylation remodeling (Figure ). These results support a coordinated link between glycosyltransferase expression, glycan structural remodeling, and mucus barrier development.

7.

7

Open in a new tab

Correlation analysis among mucosal structural parameters, mucin expression, O-glycan features, and glycosyltransferase expression in the jejunal mucosa of piglets. Spearman correlation analysis was performed to evaluate the relationships among mucosal thickness, goblet cell number, MUC2 expression, key O-glycan structural features, and the expression of representative glycosyltransferases. Red indicates positive correlations and blue indicates negative correlations. Asterisks denote statistically significant correlations (* P < 0.05, ** P < 0.01).

4. Discussion

In this study, we demonstrated that supplementation with GOS during the suckling period altered the O-glycan profile of jejunal mucins in piglets, which was associated with improved mucus layer architecture and structural integrity of the intestinal barrier. Specifically, GOS supplementation reshaped the O-glycan landscape by enhancing core 1/3-type O-glycans while reducing core 2/4-type glycans. This remodeling of mucin O-glycans was accompanied by changes in the expression of key glycosyltransferases, suggesting that GOS not only modified glycosylation patterns but also influenced the enzymatic machinery that drove these changes. Furthermore, GOS supplementation enhanced α2,3-linked sialylation while suppressing α2,6-linked sialylation, a shift that aligned with the maturation of the mucus barrier and improved host defense. These molecular changes translated into functional improvements, including increased villus width, mucosal thickness, and goblet cell numbers, which collectively enhanced epithelial protection. Our findings suggest that GOS may function as a glyco-modulatory nutrient that is associated with remodeling of mucin O-glycosylation and intestinal mucus barrier properties, thereby potentially contributing to improved gut health and resilience.

To better understand the relationship between glycosyltransferase expression and the observed O-glycan structural changes, it should be noted that mucin-type O-glycosylation follows a highly ordered and enzyme-dependent biosynthetic pathway. − The initiation of O-glycosylation is catalyzed by GALNTs, which add N-acetylgalactosamine to the mucin backbone, whereas the formation of core 1 structures requires the activity of T-synthases (C1GALT1) and its molecular chaperone C1GALT1C1. , Therefore, the upregulation of GALNTs and T-synthase observed in GOS-treated piglets is consistent with the increased abundance of core 1-derived O-glycans. Moreover, the terminal sialylation pattern of O-glycans is largely determined by the relative activities of α2,3- and α2,6-sialyltransferases. , In this study, GOS markedly upregulated α2,3-sialyltransferases family members while simultaneously downregulating α2,6-sialyltransferases, which provides a direct mechanistic explanation for the shift toward increased α2,3-sialylation and reduced α2,6-sialylation observed in both the glycomic and histological analyses. In addition, neuraminidases are known to participate in the dynamic turnover of sialylated glycans, and their decreased expression may further contribute to the stabilization of sialylated structures. Together, these coordinated transcriptional changes provide a coherent biosynthetic basis for the GOS-induced remodeling of mucin O-glycan structures.

Further analysis indicated that GOS supplementation during the suckling period significantly reshaped the O-glycan landscape of jejunal mucins in piglets, particularly increasing the abundance of core 1/3-type O-glycans while reducing certain core 2/4 -type O-glycans. Such alterations suggest that GOS may influence the initiation and elongation of O-glycan chains via transcriptional regulation of key glycosyltransferases. Indeed, we observed the upregulation of GALNT1, GALNT2, GALNT6, and C1GALT1, enzymes essential for mucin-type O-glycan synthesis. These molecular changes likely underpin the observed enrichment of core 1-type structures. The regulation of glycosyltransferase gene expression by dietary oligosaccharides has been reported in other species. For example, β-galactoside-containing prebiotics have been shown to modulate GALNT and C1GALT1 expression in intestinal epithelial cells, leading to altered glycosylation of mucins and membrane glycoproteins. Such transcriptional regulation may occur through microbial fermentation products (e.g., butyrate, propionate) that act as epigenetic modulators of glycosylation pathways. For example, butyrate has been shown to stimulate the expression of core 1 β3Gal-T (core 1 synthase) and C2GnT genes in cell models. Thus, GOS appears to remodel mucin O-glycans not merely through microbiota shifts, but also by affecting host glycosylation machinery.

A striking finding in this study was that GOS supplementation enhanced α2,3-linked sialylation but suppressed α2,6-linked sialylation in both total mucin O-glycans and sialylated MUC2 proteins. This directional shift aligns with the transcriptional pattern of sialyltransferases (ST3GAL1–6 upregulated, ST6GALNAC1–4 downregulated) and reduced expression of sialidases (NEU1–4), suggesting a coordinated regulation of sialylation turnover. Functionally, α2,3-linked sialic acids are associated with increased resistance to bacterial enzymatic degradation and promote selective adhesion of commensal Lactobacillus and Bifidobacterium species, , whereas α2,6-linked forms may facilitate attachment of certain pathogens and are more prevalent in immature epithelia. The biological significance of the increased α2,3-linked sialylation observed in GOS-treated piglets deserves attention. Sialic acids are key terminal residues of mucin O-glycans and play critical roles in mucus barrier properties, host-microbe interactions, and mucosal immunity. , Compared with α2,6-linked sialylation, α2,3-linked sialic acids are more commonly enriched in intestinal mucus and have been associated with enhanced resistance to pathogen adhesion and invasion, as many enteric pathogens recognize specific sialylated glycan motifs. , In addition, α2,3-sialylated mucins contribute to mucus layer stability and barrier function. Therefore, the GOS-induced shift toward α2,3-dominant sialylation, particularly on MUC2, likely represents a maturation-associated and protective glycosylation pattern that contributes to improved mucosal defense and immune homeostasis in early life piglets.

The observed compositional shifts in O-glycans, particularly the enrichment of sialylated core 1/3-type glycans, suggest a more hydrated, less cross-linked mucus gel, which may facilitate controlled bacterial colonization in the jejunum. Such fine-tuned glycan remodeling could create a symbiotic niche that supports beneficial microbes while excluding opportunistic pathogens. GOS is known to selectively promote Lactobacillus and Bifidobacterium, which can further modulate mucin production and glycosylation via microbial metabolites and signaling molecules. , For instance, Lactobacillus rhamnosus GG can stimulate MUC2 secretion and enhance tight junction integrity through MAPK and NF-κB pathways. The synergy between GOS and commensal bacteria thus forms a positive feedback loop, strengthening mucosal defense and homeostasis.

Beyond molecular changes, GOS markedly increased villus width, mucosal thickness, and goblet cell numbers in piglets. These morphological improvements are consistent with enhanced mucin production and glycosylation, suggesting structural and functional maturation of the intestinal barrier. The upregulation of MUC2 mRNA and protein expression indicates that GOS may activate goblet cell differentiation and secretory function, potentially through the extracellular signal-regulated kinase or phosphatidylinositol 3-kinase-protein kinase B pathways previously linked to prebiotic-induced mucin synthesis. ,, Such mucosal adaptations are crucial during the weaning transition, when piglets are vulnerable to epithelial injury and microbial imbalance. Improved mucus coverage and altered O-glycan patterns together enhance epithelial resilience, reduce pathogen translocation, and stabilize gut microbial ecology. , Correlation analysis further revealed significant associations among glycosyltransferase expression, O-glycan structural remodeling, and mucosal barrier parameters, suggesting coordinated regulation of mucin glycosylation and mucus barrier development. Collectively, the results indicate that GOS directly influences intestinal mucosal development by modulating glycosyltransferase expression, resulting in a predominance of core 1/3 and α2,3-sialylated O-glycans. These alterations in O-glycan structures promote the maturation and functional enhancement of MUC2-rich mucus layers, improving epithelial protection. This perspective highlights a glyco-modulatory role of GOS that can fine-tune the properties of the intestinal mucus barrier.

In conclusion, this study reveals that GOS supplementation during the suckling period profoundly remodels intestinal mucin-type O-glycosylation in piglets, leading to a more mature and functionally protective mucus barrier. GOS selectively enhanced the expression of core 1-related glycosyltransferases (e.g., GALNT1, C1GALT1) and α2,3-sialyltransferases (ST3GALs), while downregulating α2,6-sialyltransferases (ST6GALNACs) and sialidases (NEUs), thereby promoting a structural shift toward α2,3-sialylated O-glycans. These molecular changes coincided with increased MUC2 expression, greater goblet cell density, and improved mucosal architecture, together indicating enhanced epithelial defense and microbial symbiosis. These mechanistic insights extend the current understanding of GOS beyond microbiota modulation, highlighting its potential as an early life intervention strategy to promote intestinal development and resilience in neonatal animals.

Supplementary Material

jf6c02576_si_001.pdf (562.7KB, pdf)

Acknowledgments

This research was funded by the Natural Science Foundation of China (32030104). The funders had no role in study design, data collection and interpretation, or in the decision to submit the work for publication.

Data will be made available on request.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jafc.6c02576.

  • Experimental timeline in piglets (Figure S1); structure and relative abundance of O-glycans in intestinal mucus of piglets (Figure S2); Western blot validation of representative glycosylation-related enzymes in the jejunal mucosa of piglets (Figure S3); the qRT-PCR primers (Table S1); distribution of O-glycans with different chain lengths (Table S2) (PDF)

L.X.: Conceptualization, Investigation, Methodology, Project administration, Visualization, Writingoriginal draft, Writingreview and editing; X.L.: Conceptualization, Investigation, Methodology, Project administration, Visualization; W.F.: Conceptualization, Investigation, Methodology; W.S.: Conceptualization, Investigation, Methodology; C.M.: Conceptualization, Investigation, Methodology; L.L.: Methodology; J.V.: Methodology; C.M.: Conceptualization, Methodology, Writingreview and editing; W.Z.: Conceptualization, Methodology, Funding acquisition, Supervision, Writingreview and editing.

The authors declare no competing financial interest.

References

  1. Zhang J., Chen Y., Guo X., Li X., Zhang R., Wang M., Zhu W., Yu K.. The gut microbial metabolite indole-3-aldehyde alleviates impaired intestinal development by promoting intestinal stem cell expansion in weaned piglets. J. Anim. Sci. Biotechnol. 2024;15(1):150. doi: 10.1186/s40104-024-01111-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Zhang R., Huang G., Ren Y., Wang H., Ye Y., Guo J., Wang M., Zhu W., Yu K.. Effects of Dietary Indole-3-carboxaldehyde Supplementation on Growth Performance, Intestinal Epithelial Function, and Intestinal Microbial Composition in Weaned Piglets. Front. Nutr. 2022;9:896815. doi: 10.3389/fnut.2022.896815. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Johansson M. E., Sjövall H., Hansson G. C.. The gastrointestinal mucus system in health and disease. Nat. Rev. Gastroenterol. Hepatol. 2013;10(6):352–361. doi: 10.1038/nrgastro.2013.35. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Cheng H., Li H., Li Z., Wang Y., Liu L., Wang J., Ma X., Tan B.. The role of glycosylated mucins in maintaining intestinal homeostasis and gut health. Anim. Nutr. 2025;21:439–446. doi: 10.1016/j.aninu.2025.03.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Tang X., Xiong K., Fang R., Li M.. Weaning stress and intestinal health of piglets: A review. Front. Immunol. 2022;13:1042778. doi: 10.3389/fimmu.2022.1042778. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Neurath M. F., Artis D., Becker C.. The intestinal barrier: a pivotal role in health, inflammation, and cancer. Lancet Gastroenterol. Hepatol. 2025;10(6):573–592. doi: 10.1016/S2468-1253(24)00390-X. [DOI] [PubMed] [Google Scholar]
  7. Jing Y., Mu C., Wang H., Shen J., Zoetendal E. G., Zhu W.. Amino acid utilization allows intestinal dominance of Lactobacillus amylovorus. ISME J. 2022;16(11):2491–2502. doi: 10.1038/s41396-022-01287-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Wu H., Mu C., Xu L., Yu K., Shen L., Zhu W.. Host-microbiota interaction in intestinal stem cell homeostasis. Gut Microbes. 2024;16(1):2353399. doi: 10.1080/19490976.2024.2353399. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Johansson M. E., Phillipson M., Petersson J., Velcich A., Holm L., Hansson G. C.. The inner of the two Muc2 mucin-dependent mucus layers in colon is devoid of bacteria. Proc. Natl. Acad. Sci. U. S. A. 2008;105(39):15064–15069. doi: 10.1073/pnas.0803124105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Birchenough G. M., Johansson M. E., Gustafsson J. K., Bergström J. H., Hansson G. C.. New developments in goblet cell mucus secretion and function. Mucosal Immunol. 2015;8(4):712–719. doi: 10.1038/mi.2015.32. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Jensen P. H., Kolarich D., Packer N. H.. Mucin-type O-glycosylation–putting the pieces together. FEBS J. 2010;277(1):81–94. doi: 10.1111/j.1742-4658.2009.07429.x. [DOI] [PubMed] [Google Scholar]
  12. Fekete E., Buret A. G.. The role of mucin O-glycans in microbiota dysbiosis, intestinal homeostasis, and host-pathogen interactions. Am. J. Physiol. Gastrointest. Liver Physiol. 2023;324(6):G452–G465. doi: 10.1152/ajpgi.00261.2022. [DOI] [PubMed] [Google Scholar]
  13. Luis A. S., Hansson G. C.. Intestinal mucus and their glycans: A habitat for thriving microbiota. Cell Host Microbe. 2023;31(7):1087–1100. doi: 10.1016/j.chom.2023.05.026. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Hansson G. C.. A Golgi oxygen sensor controls intestinal mucin glycosylation. EMBO J. 2023;42(2):EMBJ2022113013. doi: 10.15252/embj.2022113013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Xia B., Zhong R., Wu W., Luo C., Meng Q., Gao Q., Zhao Y., Chen L., Zhang S., Zhao X.. et al. Mucin O-glycan-microbiota axis orchestrates gut homeostasis in a diarrheal pig model. Microbiome. 2022;10(1):139. doi: 10.1186/s40168-022-01326-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Sommer F., Adam N., Johansson M. E., Xia L., Hansson G. C., Bäckhed F.. Altered mucus glycosylation in core 1 O-glycan-deficient mice affects microbiota composition and intestinal architecture. PLoS One. 2014;9(1):e85254. doi: 10.1371/journal.pone.0085254. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Yao Y., Kim G., Shafer S., Chen Z., Kubo S., Ji Y., Luo J., Yang W., Perner S. P., Kanellopoulou C.. et al. Mucus sialylation determines intestinal host-commensal homeostasis. Cell. 2022;185(7):1172–1188.e. doi: 10.1016/j.cell.2022.02.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Arioz Tunc H., Calder P. C., Cait A., Dodd G. F., Gasaly Retamal N. Y. I., Guillemet D., James D., Korzeniowski K. J., Lubkowska A., Meynier A.. et al. Impact of non-digestible carbohydrates and prebiotics on immunity, infections, inflammation and vaccine responses: a systematic review of evidence in healthy humans and a discussion of mechanistic proposals. Crit. Rev. Food Sci. Nutr. 2026;66(1):1–74. doi: 10.1080/10408398.2025.2514700. [DOI] [PubMed] [Google Scholar]
  19. Zhao J., Cao Y., Miao Y., Guo H., Wang Z., Liang X., Guan X., Sun R., Zhang X., Nie C.. et al. Structure, Enzymatic Production, Biological Activities, and Food Applications of Galacto-oligosaccharides: A Review. J. Nutr. 2025;155(12):4113–4122. doi: 10.1016/j.tjnut.2025.10.005. [DOI] [PubMed] [Google Scholar]
  20. Xu L., Li X., Han S., Mu C., Zhu W.. Galacto-oligosaccharides regulate intestinal mucosal sialylation to counteract antibiotic-induced mucin dysbiosis. Food Funct. 2024;15(24):12016–12032. doi: 10.1039/D4FO04626A. [DOI] [PubMed] [Google Scholar]
  21. Figueroa-Lozano S., Ren C., Yin H., Pham H., van Leeuwen S., Dijkhuizen L., de Vos P.. The impact of oligosaccharide content, glycosidic linkages and lactose content of galacto-oligosaccharides (GOS) on the expression of mucus-related genes in goblet cells. Food Funct. 2020;11(4):3506–3515. doi: 10.1039/D0FO00064G. [DOI] [PubMed] [Google Scholar]
  22. Fan W., Mu C., Li X., Zhang Y., Yu K., Su Y., Zhu W.. Spatiotemporal dynamics of mucosa-associated Lactobacillus along the small intestine and its response to 2’-fucosyllactose and galacto-oligosaccharides in piglets. Anim. Nutr. 2025;23:235–245. doi: 10.1016/j.aninu.2025.04.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Tian S., Wang J., Wang J., Zhu W.. Differential Effects of Early-Life and Postweaning Galacto-oligosaccharide Intervention on Colonic Bacterial Composition and Function in Weaning Piglets. Appl. Environ. Microbiol. 2022;88(1):e0131821. doi: 10.1128/AEM.01318-21. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Brockhausen I.. Pathways of O-glycan biosynthesis in cancer cells. Biochim. Biophys. Acta. 1999;1473(1):67–95. doi: 10.1016/S0304-4165(99)00170-1. [DOI] [PubMed] [Google Scholar]
  25. Corfield A. P.. Mucins: a biologically relevant glycan barrier in mucosal protection. Biochim. Biophys. Acta. 2015;1850(1):236–252. doi: 10.1016/j.bbagen.2014.05.003. [DOI] [PubMed] [Google Scholar]
  26. Schjoldager K. T., Narimatsu Y., Joshi H. J., Clausen H.. Global view of human protein glycosylation pathways and functions. Nat. Rev. Mol. Cell Biol. 2020;21(12):729–749. doi: 10.1038/s41580-020-00294-x. [DOI] [PubMed] [Google Scholar]
  27. Varki A.. Biological roles of glycans. Glycobiology. 2017;27(1):3–49. doi: 10.1093/glycob/cww086. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Miyagi T., Yamaguchi K.. Mammalian sialidases: physiological and pathological roles in cellular functions. Glycobiology. 2012;22(7):880–896. doi: 10.1093/glycob/cws057. [DOI] [PubMed] [Google Scholar]
  29. Bode L.. Human milk oligosaccharides: every baby needs a sugar mama. Glycobiology. 2012;22(9):1147–1162. doi: 10.1093/glycob/cws074. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Radhakrishnan P., Beum P. V., Tan S., Cheng P. W.. Butyrate induces sLex synthesis by stimulation of selective glycosyltransferase genes. Biochem. Biophys. Res. Commun. 2007;359(3):457–462. doi: 10.1016/j.bbrc.2007.05.165. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Schauer R.. Sialic acids as regulators of molecular and cellular interactions. Curr. Opin. Struct. Biol. 2009;19(5):507–514. doi: 10.1016/j.sbi.2009.06.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Engevik M. A., Luk B., Chang-Graham A. L., Hall A., Herrmann B., Ruan W., Endres B. T., Shi Z., Garey K. W., Hyser J. M.. et al. Bifidobacterium dentium Fortifies the Intestinal Mucus Layer via Autophagy and Calcium Signaling Pathways. mBio. 2019;10(3):e01087-19. doi: 10.1128/mBio.01087-19. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Linden S. K., Sutton P., Karlsson N. G., Korolik V., McGuckin M. A.. Mucins in the mucosal barrier to infection. Mucosal Immunol. 2008;1(3):183–197. doi: 10.1038/mi.2008.5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. McAuley J. L., Corcilius L., Tan H. X., Payne R. J., McGuckin M. A., Brown L. E.. The cell surface mucin MUC1 limits the severity of influenza A virus infection. Mucosal Immunol. 2017;10(6):1581–1593. doi: 10.1038/mi.2017.16. [DOI] [PubMed] [Google Scholar]
  35. Zhao C., Pu J.. Influence of Host Sialic Acid Receptors Structure on the Host Specificity of Influenza Viruses. Viruses. 2022;14(10):2141. doi: 10.3390/v14102141. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Topping D. L., Clifton P. M.. Short-chain fatty acids and human colonic function: roles of resistant starch and nonstarch polysaccharides. Physiol. Rev. 2001;81(3):1031–1064. doi: 10.1152/physrev.2001.81.3.1031. [DOI] [PubMed] [Google Scholar]
  37. Lee A., Mansbridge S. C., Liang L., Connerton I. F., Mellits K. H.. Galacto-Oligosaccharides Increase the Abundance of Beneficial Probiotic Bacteria and Improve Gut Architecture and Goblet Cell Expression in Poorly Performing Piglets, but Not Performance. Animals. 2023;13(2):230. doi: 10.3390/ani13020230. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Bhatia S., Prabhu P. N., Benefiel A. C., Miller M. J., Chow J., Davis S. R., Gaskins H. R.. Galacto-oligosaccharides may directly enhance intestinal barrier function through the modulation of goblet cells. Mol. Nutr. Food Res. 2015;59(3):566–573. doi: 10.1002/mnfr.201400639. [DOI] [PubMed] [Google Scholar]
  39. Zheng J., Shen H., Tan B., Yin Y., Wang J.. Dietary supplementation of Galactooligosaccharides improves growth performance and intestinal barrier function of offspring. Porcine Health Manage. 2025;11(1):37. doi: 10.1186/s40813-025-00437-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Wang Y., Li Z., Chen G., Xing Y., Wang J., Zhao Y., Kang M., Huang K., Li E., Ma X.. Dietary Galacto-Oligosaccharides Enhance Growth Performance and Modulate Gut Microbiota in Weaned Piglets: A Sustainable Alternative to Antibiotics. Animals. 2025;15(11):1508. doi: 10.3390/ani15111508. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

jf6c02576_si_001.pdf (562.7KB, pdf)

Data Availability Statement

Data will be made available on request.

Articles from Journal of Agricultural and Food Chemistry are provided here courtesy of American Chemical Society

RESOURCES