According to our previous work with minor modifications (18), CE was prepared as follows: 8 g of NS were weighed, dispersed in 100 mL phosphate buffer pH 4.6 (0.1 M), placed in a water bath at 100°C, and heated for 20 min.

The boiled starch solution was adjusted to 58°C, and pullulanase was added. The temperature was maintained for 12 h to achieve debranching, then mid-temperature α-amylase (1 U/g) was added for enzymolysis for 5 min. The supernatant was transferred to a boiling water bath at 100°C to inactivate enzymes, centrifuged at 4,000 rpm for 5 min, and placed at 4°C for 24 h. Finally, the recrystallized starch was washed three times at 4,000 rpm for 3 min, dried overnight at 40°C, ground, and filtered with a 200-mesh sieve for further analyses.

The simulated digestive process includes the oral, gastric, and intestinal phases. The preparation of simulated oral, gastric, and intestinal fluids, including the preparation of electrolyte stock solution and the addition of digestive enzymes at different stages, was performed as described (19). The experimental digestion procedure was carried out following the method of Chang et al. (20) with some adaptations. When used, the electrolyte stock solutions were diluted to 400 mL with Milli-Q-purifed water.

Oral phase: 5 mL of CE suspension (10%, w/v) was mixed with 3.5 mL simulated salivary fluid (SSF, pH = 7.0), α-amylase solution (0.5 mL, 1,500 U/mL) prepared with SSF, 25 μL of 0.3 M CaCl₂, and 975 μL of Milli-Q-purified water. The solution was placed in a 37°C water bath and shaken for 2 min to obtain salivary digestive CE solution. We added 0.2 mL 1 M HCl to inactivate α-amylase and terminated digestion. The suspension was centrifuged at 5,000 × g for 15 min and washed in water three times. The precipitate was collected and dried overnight at 40°C to obtain a dried CE sample after saliva digestion (SSF-CE).

Gastric phase: The salivary digestive CE solution (a total volume of ~10 mL) was added with 7.5 mL simulated gastric fluid (SGF, pH = 3.0), 1.6 mL of 25,000 U/mL porcine pepsin solution prepared with SGF, 5 μL of 0.3 M CaCl₂, 0.2 mL of 1.0 M HCl to reach pH 3.0, and 0.695 mL of Milli-Q-purified water. The mixture was placed in a 37°C water bath and shaken for 2 h to obtain gastric digestive RS solution. We added 0.2 mL 1 M NaOH to inactivate porcine pepsin. The suspension was centrifuged at 5,000 × g for 15 min and washed in water three times. The precipitate was collected and dried overnight at 40°C to obtain a dried gastric digestive CE sample (SGF-CE).

Intestinal phase: The gastric digestive CE solution (a total volume of ~20 mL) was mixed with 11 mL of simulated intestinal fluid (SIF, pH = 7.0), 5.0 mL of 800 U/mL pancreatin solution prepared with SIF, 2.5 mL of 8.0 mM cholate solution, 0.1 mL amyloglucosidase, 40 μL of 0.3 M CaCl₂, 0.15 mL of 1.0 M NaOH to reach pH 7.0, and 1.31 mL of water. The time of contact with the enzyme was 2 h in a 37°C water bath, then 0.2 mL 1.0 M NaOH was added to inactivate pancreatin. The suspension was centrifuged at 5,000 × g for 15 min and washed in water three times. We collected the precipitate and dried it at 40°C overnight to obtain an intestinal RS sample (SIF-CE).

Initially, the RS content was measured by employing the Megazyme RS assay kit in accordance with the AOAC official method 2002.02 (21, 22). Subsequently, the mass loss at different stages of digestion was determined by weighing dried samples before digestion and the different digestive processes using an analytical balance. The following equation was used to express the digestion rate.

Results were expressed as mean ± standard deviation for three replicates. Statistical differences were determined using GraphPad Prism 9.0 (Inc., California, United States). Differences between groups were analyzed using a one-way analysis of variance, with differences considered statistically significant at p < 0.05.

The XRD pattern of RS includes the crystalline and amorphous regions (30). The crystalline region is composed of stacked straight-chain starch double helixes, while the amorphous region is composed of disordered starch. As shown in Figure 1A, the CE samples at various digestive stages showed a typical B-crystalline morphology. Diffraction peaks appeared at 5.6°, 15°, 17°, 22°, and 24° (2θ) (31).

The level of crystallinity showed an upward trend with the progression of digestion (Table 2). The RC of CE was determined to be 38.92%. Following in vitro digestion with different digestive enzymes, the RC was 41.70% for SSF-CE, 46.01% for SGF-CE, and 49.34% for SIF-CE, separately. The depolymerization of starch structure usually occurs in the amorphous regions. The structural changes in CE samples manifest as the action of digestive enzymes and various ions on the amorphous regions, leading to a significant increase in crystallinity. Nevertheless, there was no substantial effect on the crystal type.

Figure 1B showed the FTIR spectra between wave number 4000–400 cm⁻¹ of CE and its corresponding digestive samples. All samples exhibited a broad absorption bands at 3400 cm⁻¹ which can be attributed to O-H stretching vibration of starch (32). The absorption peaks at 2928 cm⁻¹ and 1643 cm⁻¹ are ascribed to C-H stretching (33) and O-H bending vibration (34), respectively. The infrared spectra showed that no peaks disappeared or new peaks were generated after the various stages of digestion or enzymatic digestion, suggesting that no new covalent bonds or functional groups were generated after the in vitro digestion of CE.

The infrared peak range of 1200–800 cm⁻¹ (Figure 1B, red dotted coil) was deconvoluted to analyze the short-range order of starch. The absorbance of starch at 1047 cm⁻¹ and 1022 cm⁻¹ reflects the ordered crystalline structure as well as the amorphous structure of starch (35). The absorbance ratio of 1047/1022 (R_1047/1022) can be used to indicate the degree of ordered structure (DO) values of starch crystals. The absorbance ratio of 995/1022 cm⁻¹ (R_995/1022) for starch reflects the degree of double helix (DD) values between starch molecules (36). The DO and DD of CE and its digestive samples are shown in Table 2. These calculations revealed that DO and the DD of CE did not differ significantly between the groups after the action of different digestive enzymes; there was an upward trend, coinciding with the XRD pattern.

As shown in Figure 2, a-3, CE exhibited a smooth surface and a compact structure, which is likely due to the retrogradation of amylose chains. Cracks began to appear on the surface of CE particles at the oral phase, indicating the initial mechanical breakdown of the particles during chewing (37). Despite these cracks, the surface remained relatively smooth, suggesting limited enzymatic activity at this stage (Figure 2, b-3). During the gastric phase, the surface of the CE granules developed honeycomb pores (Figure 2, c-3), possibly as a result of the breakdown of the amorphous portions of the resistant starch in the acidic environment. The final digestion product, SIF-CE, exhibited a rough surface with large cracks and a homogeneous honeycomb structure. This morphology indicated extensive enzymatic degradation. However, the 500 × images (Figure 2, a-2; b-2; c-2; d-2) showed that the size of the CE particles did not change significantly at different stages of digestion.

SIF-CE was subjected to enteric bacterial fermentation experiments in healthy people using a BHI medium at a 1% addition rate. Initially, the pH of all three groups was approximately 7.2 (Figure 3A). However, within the first 4 h, there was a sharp decline in pH observed in the control, SIF-CE, and FOS groups. This finding can be attributed to the rapid consumption of nutrients within the medium by fecal microbiota, resulting in their extensive proliferation and subsequent acid production.

During the anaerobic fermentation process, a noticeable deviation in pH levels emerged between the SIF-CE and control groups after 8 h. Subsequently, the pH levels in both the control and SIF-CE groups reached 6.31 and 5.95 over time, and the SIF-CE group had a significantly lower pH than the control group. The pH of the FOS group continued decreasing until it stabilized at around 4.33 at 24 h. These findings suggested that the consumption of CE and FOS led to a considerable reduction of pH after fermentation by enteric bacteria.

The SCFAs levels in the supernatants after microbial fermentation of feces in a healthy population were measured. Figure 3 showed, six SCFAs were detected in all three groups after 24 h of fermentation, including acetic, propionic, isobutyric, butyric, isovaleric, and valeric acids. Notably, the introduction of SIF-CE for fermentation led to a significant increase in the levels of propionic and valeric acids, as depicted in Figures 3C,G. Conversely, the FOS group exhibited significantly lower SCFAs levels in the FOS group compared to the control group.

SCFAs levels in the fermentation supernatants supplemented with SIF-CE were significantly higher than in the FOS group. SIF-CE was more potent than FOS in promoting SCFAs production.

The degradation of carbohydrates by intestinal microorganisms produces organic acids, and lactic acid is a primary organic acid. Therefore, we examined the lactic acid content in the supernatants after 24 h of fermentation. Figure 3H reveals that the lactic acid content in the FOS group was significantly higher than the control group by approximately 60-fold. Fecal microorganisms use FOS to produce large amounts of lactic acid. There was no significant change in lactate content in the SIF-CE group compared with the control group. These findings suggest that the pH reduction of the fermentation system caused by SIF-CE and FOS is achieved by different mechanisms, the former by promoting the content of SCFAs and the latter by increasing the lactic acid content to achieve low pH.

Gut microbiota diversity analysis used 97% sequence similarity genes as the operational taxonomic unit delineation threshold. Alpha diversity indexes measure intra-group colony diversity from various perspectives. The alpha diversity index, Chao1 index, and Ace index are commonly used to characterize the species richness of a colony. The Shannon and Simpson indexes combine the richness and evenness of the community. Combining all alpha diversity indexes (Figures 4A–D), there was no significant difference in the effect of SIF-CE addition on the internal diversity of the fecal microbial community and no significant effect on species richness compared to the control group. In contrast, FOS had significantly lower Ace, Shannon’s index (p < 0.01), and Simpson’s index (p < 0.05), suggesting that FOS significantly reduced species richness and diversity.

The β-diversity index focuses on the differences between samples, using primary coordinates analysis to assess the β-diversity of fecal microorganisms after 24 h fermentation. Figure 4E shows that the groups are separated. The first (PC1) and second principal components (PC2) explained 64.64 and 23.79% of the total variables, respectively. Combined with the Anisom analysis, the differences in intestinal microbiota between the groups were statistically significant (p < 0.01).

To investigate the differences in community structure among different fermentation groups, the changes in species abundance at the phylum level are shown in Figure 5A. The three groups on the phylum level are mainly composed of Firmicutes, Proteobacteria, Bacteroidota, and Actinobacteriota. The Kruskal-Wallis test showed significant differences in abundance between the three groups found for Proteobacteria and Actinobacteriota, (Figure 5, A1). Compared with the control group, the Proteobacteria content in the SIF and FOS groups decreased significantly after 24 h of fermentation (Figure 5, A2). SIF-CE showed the same ability as the FOS group to upregulate Actinobacteriota significantly, and the amount of Actinobacteriota was significantly higher in the SIF-CE group than in the FOS group (Figure 5, A3).

At the genus level, the Kruskal-Wallis test showed that the relative abundance of 15 genus was statistically significant (Figure 5, B1). In the FOS group, the abundance of Anaerostipes, Holdemania, and Coprobacillus was significantly decreased to a proportion of approximately 0% (p < 0.01). These genus showed the same trend of being downregulated in the SIF-CE group and were also significant (p < 0.05).

The well-known probiotics such as Bifidobacterium and Lactobacillus, which can produce lactic acid (38) were studied. As shown in Figures 5, B2, B5, in the control and SIF-CE groups, the abundance of Lactobacillus was minimal and barely observable on the bar graph; however, its abundance increased significantly and significantly in the FOS group. The abundance of Bifidobacterium was 0.108% in the control group and 4.775% in the SIF-CE group, approximately 44 times more in the SIF-CE group than in the control group. FOS also showed an upregulation of Bifidobacterium.

In addition to these results, we found that the proportion of Enterococcus (Figure 5, B6) abundance in the FOS group was large and significant relative to the control group, and Dorea (Figure 5, B4) was significantly upregulated in the SIF-CE group. Dorea is essential in non-alcoholic steatohepatitis and is one of the primary regulators of microbiota-dependent protective phenotypes in mitochondrial microbiota interactions (39). Enterococcus is usually capable of producing lactic acid (40). The SIF-CE and FOS groups showed the same ability to downregulate Escherichia-Shigella (Figure 5, B3).