HWE is the most widely used method for CPP extraction (32, 33). It is simple to operate, inexpensive, and highly efficient and is suitable for the extraction of most plant polysaccharides (15). Optimization of extraction conditions is necessary for improving the extraction efficiency and quality of polysaccharides (34). For instance, the ideal rise in extraction temperature enhances the solubility of polysaccharides; however, excessive temperatures trigger degradation. Similarly, extended extraction time guarantees the total release of polysaccharides, while excessive exposure will decline bioactivity. Adjusting the solid-to-liquid ratio improves the efficiency of solubility and reduces solvent wastage, while repeated cycle extraction enhances yield (35). Experiments revealed that, under optimized conditions of 80 °C, a solid-to-liquid ratio of 1:16, an extraction time of 80 minutes, and a particle size of 180 μm, the maximum yield of CPPs reached 15.56% (36). Shi et al. optimized RSM extraction conditions and found optimum parameters of extraction time 2.3 h, extraction temperature 89 °C, and solid-to-liquid ratio 1:24, under which 15.66% CPPs existed (37). Similarly, Fu et al. employed RSM to optimize fructan extraction from C. pilosula and reported a yield of 20.6% obtained with an extraction temperature of 100 °C, a solvent-to-material ratio of 40 mL/g, and an extraction time of 2.5 hours in two cycles (38). Li et al. further optimized extraction conditions and the authors concluded that an extraction temperature of 85 °C, an extraction time of 1.5 hours, and a solid-to-liquid ratio of 1:12 provided a final yield of 22.57% after two extraction cycles (39). Another work reported achieving a 25.7% final yield at 90 °C, using an extraction time of 45 minutes and a solid-to-liquid ratio of 1:20 (40). In conclusion, the optimization of extraction parameters, such as temperature, time, solid-to-liquid ratio, and extraction cycles, useing OED and RSM is a primary focus in maximizing of both the efficiency and quality of CPPs extraction.

The UAE utilizes the cavitation process and the mechanical force of ultrasound waves to rupture the cell walls of plant cells, improve the penetration of the solvent, and permit the release of polysaccharides (41). Compared to conventional extraction technologies, UAE offers numerous advantages including higher extraction rates, lower energy utilization, and improved yield, making it easier to operate and more environmentally friendly (42). In addition, UAE lowers the degradation of heat-sensitive polysaccharides and preserves their biological activity, making it an extremely efficient and sustainable extraction technique (43). Cao et al. investigated the effects of various factors, such as the solid-to-liquid ratio, ultrasonic power, extraction time, and temperature, on the yield of CPPs in single-factor experiments. They further optimized the extraction conditions and found the best parameters to be:solid-to-liquid ratio = 1:30 (g/mL), power of ultrasonication = 280 W, time of extraction = 75 minutes, and temperature of extraction = 74 °C. Under the above conditions, the extraction yield was 18.42% (16). Similarly, Wang et al. also performed single-factor tests to investigate the influence of the temperature, the solid-to-liquid ratio, extraction time, and ultrasonic power on extraction efficiency. They then optimized the extraction process using a uniform design method, and found the optimum conditions to be a 70 °C temperature, a 25:1 solid-to-liquid ratio, 35 minutes of extraction time, and 135 W of ultrasonic power. These conditions produced an average extraction yield of 31.26% (44). Furthermore, Wang et al. demonstrated that UAE with an ultrasonic power of 180 W, a frequency of 60 kHz, and a treatment time of 30 minutes increased the content of CPPs by 14.7%, while also effectively preserving polysaccharide bioactivity and other functional ingredients (45). In conclusion, UAE is an extremely effective, environmentally friendly, and green extraction technique that holds a vast potential for the extraction of CPPs. Further research and optimization of UAE parameters can contribute to enhancing the functional use and industrial applications of CPPs.

EAE utilizes specific enzymes for the degradation of plant cell walls or other structural components, thereby facilitating the efficient release of target molecules under mild conditions (46). This method has several advantages, including high extraction efficiency, high selectivity, and the ability to retain heat-sensitive bioactive compounds,making it widely applicable for polysaccharide extraction (47, 48). Fan et al. optimized the polysaccharide extraction process from yeast-fermented fresh C. pilosula through single-factor experiments combined with RSM. The results indicated that the optimal extraction condition was 63 °C of temperature, 6 hours of extraction time, and 4% of enzyme concentration. Under these conditions, extraction yield was as high as 17.46%, significantly higher than the 12.69% achieved by conventional HWE (49). Gao et al. also optimized the extraction of CPPs by single-factor experiments combined with RSM. The optimal conditions were found to be an enzyme concentration of 0.2%, an enzymatic hydrolysis time of 1.5 hours, a hydrolysis temperature of 50 °C, and a pH of 4.2. Under the optimal conditions, the extraction yield was 25.23% (18). In general, compared to traditional extraction methods, EAE is energy-efficient and environmentally friendly, lowering impurities and reducing extraction time. Besides, the simplicity of enzyme selection based on raw material characteristics will enable better extraction efficiency and quality of polysaccharides. Continued advances in enzyme selection and process optimization will also contribute to the broader application of CPPs in various applications.

Apart from the already described extraction methods, scientists arecontinually seeking simpler and faster ways to obtain CPPs. For instance, MAE induces fast oscillation of polar molecules inside the material using microwave electromagnetic fields, therefore producing both thermal and non-thermal effects. The rapid heating of the solvent, material causes cell wall rupture and thereby increasing extraction efficiency (50, 51). Yu et al. examined CPPs extraction by OED about solid-to-liquid ratio, microwave radiation power, and extraction time. Water was found to be the ideal solvent; a temperature of 70 °C, a solid-to-liquid ratio of 1:40, an extraction period of 20 minutes, and a microwave power of 500 W, therefore producing an extraction rate of 14.8% (17). Utilizing the lowered polarity of water at high temperatures and pressures, subcritical water extraction (SWE) extracts bioactive compounds. Particularly appropriate for heat-sensitive materials, this approach is also environmentally benign and reasonably affordable (52, 53). By employing a temperature of 150 °C, an extraction duration of 45 minutes, and a liquid-to-material ratio of 12 mL/g, Zhang et al. refined the SWE process for CPPs, thereby establishing optimal conditions and an extraction rate of 19.51% (54). A liquid-liquid extraction system that generates immiscible phases between ethanol and ammonium sulfate, therefore allowing the separation of target chemicals, is the water-ethanol/ammonium sulfate aqueous two-phase system (55, 56). Using this approach, Lu et al. extracted CPPs by optimizing the conditions to a concentration of 17% ammonium sulfate, 30% ethanol, an extraction temperature of 40 °C, and a pH of 6. Under these circumstances, the extractive yield was 31.57% (57). In conclusion, these advanced extraction methods offer higher efficiency and selectivity compared to traditional techniques, enabling the extraction of bioactive compounds under milder conditions. These techniques show great promise for both commercial and medicinal uses, as they enhance the bioactivity of polysaccharides and increase extraction efficiency.

The molecular weight of polysaccharides plays a crucial role in determining their biological activities and is therefore considered an important parameter reflecting their physicochemical properties and functional potential (65). Determining molecular weight facilitates a deeper understanding of the relationship between polysaccharide structure and function and is commonly estimated using statistical averaging methods (60). Traditional techniques, such as viscometry, light scattering, and osmometry, have been widely used, however, they tend to be complex and susceptible to experimental errors  (66, 67). Currently, more accurate and reliable methods are preferred, including high-performance liquid chromatography (HPLC), gel permeation chromatography (GPC), high-performance gel permeation chromatography, and high-performance size-exclusion chromatography (14, 38, 68, 69). Due to variations in the types of CPPs, as well as differences in extraction, purification processes, and analytical methods, the reported average molecular weights of CPPs vary across studies (12, 70). For instance, in hot-water extraction methods, temperature and extraction duration significantly affect the resulting molecular weight. Increasing the temperature and extending the extraction time within an appropriate range can enhance the yield of high-molecular-weight polysaccharides. However, excessive heat may cause polysaccharide degradation, resulting in a decrease in molecular weight (71). Zou et al., for example, extracted pectic polysaccharides from C. pilosula roots at 50 °C and 100 °C, yielding 50WCP-II-I (71.6 kDa) and 100WCP-II-I (53.2 kDa), respectively, highlighting the influence of temperature on molecular size. Moreover, the pore size of ultrafiltration membranes can also affect the molecular weight distribution of polysaccharides (72). Li et al. separated CPPs into three fractions using membranes with different molecular weight cut-offs, yielding CPPS-I (< 60 kDa), CPPS-II (60–100 kDa), and CPPS-III (> 100 kDa), demonstrating the impact of membrane filtration on polysaccharide size profiling (14).

Monosaccharide composition serves as a fundamental basis for elucidating the structural features, physicochemical properties, and structure-activity relationships of polysaccharides (73). Variations in purification techniques or raw material sources can lead to differences in monosaccharide ratios, glycosidic bond types, and linkage sequences (28). To analyze the monosaccharide profile of CPPs, commonly employed techniques include gas chromatography (GC), HPLC, NMR, and IR (74, 75). Su et al. separated a homogeneous polysaccharide, CPP-1, from the roots of C. pilosula, which consisted of mannose (Man), glucose (Glc), galacturonic (Gal), and arabinose (Ara) with a molar ratio of 5.86:51.69:34.34:8.08 (70). Similarly, Bai et al. isolated two polysaccharides, CPP1a and CPP1c; CPP1a consisted of rhamnose (Rha), Ara, galacturonic acid (GalA), Gal, and Glc in a molar ratio of 1:12:1:10:3. However, CPP1c had a higher proportion of GalA, which may be accountable for its more active biological activity compared to CPP1a (11). Zhang et al.identified eight monosaccharides present in CPPs, namely Man, Rib, Rha, glucuronic acid, GalA, Glc, Gal, and Ara,with Glc being the most prevalent at 49.87%. The molar ratio of the monosaccharides was found to be 0.53:0.60:0.51:3.50:9.55:21.73:4.78:2.38 (74).

Due to the complex branching and linkage patterns of different CPPs, structural characterization is typically challenging (8). Structural analysis of polysaccharides primarily involves the identification of monosaccharide linkage sequences and glycosidic bond conformations. Common analytical techniques for CPPs structural elucidation include FT-IR, NMR, and GC-MS (9, 11, 70). A full structural characterization of polysaccharides typically requires the integration of multiple analytical techniques. FT-IR spectroscopy provides valuable information regarding functional groups and bond types within the polysaccharide molecule (76). The broad band of absorption at 3300–3450 cm⁻¹ corresponds to the stretching vibration of the free hydroxyl (-OH) group, indicative of the high hydroxyl content in polysaccharides, which is directly related to their solubility in water (77). Bands of absorption 2800–2950 cm⁻¹ are attributed to C-H stretching vibrations (methyl or methylene groups), which correspond to the carbon skeleton of the sugar ring (76). Common signals in 1000–1200 cm⁻¹ correspond to the pyranose ring and glycosidic linkage stretching vibrations (77, 78), while 1025–1152 cm⁻¹ bands of absorption also confirm the presence of pyranose rings (38, 70). In addition, bands at 852 cm⁻¹ and 893 cm⁻¹ of absorption are characteristic of the presence of α- and β-glycosidic linkages, respectively (14, 76). Bands at 1600–1750 cm⁻¹ show vibrational characteristics of carboxyl (C=O) and ester (COOCH₃) groups, which are usually present in pectic polysaccharides (12, 79, 80). Detailed determination of glycosidic bond configuration and linkage structures is achieved with ¹H NMR spectroscopy (81). The chemical shift of the anomeric carbon (C1) can determine the configuration of the sugar ring, and α-anomers would typically be in the range of 5.0–5.5 ppm (14), whereas β-anomers appear in 4.5–5.0 ppm (76). ¹H NMR also identifies signals for branched structures or distinct functional groups such as methylated high-galacturonic regions (11). GC-MS also aids in polysaccharide characterization, providing quantitative information on monosaccharide composition. The interplay of these analysis techniques gives a clearer vision of the structural complexity of polysaccharides (82, 83). Zou et al. isolated two pectic polysaccharides, 50WCP-II-I and 100WCP-II-I, from the roots of C. pilosula. The backbone of 50WCP-II-I primarily consists of a long homogalacturonan (HG) region composed of GalA units, some of which are methyl-esterified. Embedded within the HG backbone are short rhamnogalacturonan-I (RG-I) regions, in which Rha residues at the O-4 position are substituted with arabinogalactan type I (AG-I) and arabinogalactan type II (AG-II) side chains ( Figure 3A ). In contrast, 100WCP-II-I contains two major branched regions: one in which GalA residues at the O-2 position are substituted with AG-I side chains and another where Rha residues at the O-4 position are substituted with AG-II side chains in the RG-I domain. The backbone also contains methyl-esterified HG region, and Rha residues are mainly 1,2-linked ( Figure 3B ) (72). Yang et al. Found another pectic polysaccharide, CPP1b, from C. pilosula, whose backbone is constituted by 1,4-linked α-D-GalA and methyl-esterified α-D-GalA, alternately interspersed with minor amounts of 1,2-linked β-L-rhamnose, 1,2,6-linked α-D-Gal and terminal α-L-Ara. Methylation analysis and NMR spectroscopy results demonstrated that side chains contain Ara and branched Gal residues ( Figure 3C ). In summary, the overall structure can be considered as a pectin containing RG-I-type rhamnogalacturonan regions consisting of a linear backbone with sparse branching. Ultrastructurally, CPP1b were amorphous, ovoid granules with sheet-like surface morphology (83). CPPS3, a water-soluble polysaccharide extracted from the roots of C. pilosula, was isolated. Its branched backbone is constituted by (1→3)-linked β-N-acetylgalactosamine, (1→3)-linked α-rhamnose and (1→2,3)-linked β-Gal. The degree of branching of this water-soluble polysaccharide is approximately 26%, meaning that one side chain exists for every four backbone residues. Its side chains consist of (1→5)-linked α-arabinofuran, (1→4)-linked β-Gal and (1→2)-linked α-rhamnose–Ara disaccharide chains. Its side chains were found to contain β-glycosidic linkages by infrared spectroscopy ( Figure 3D ) (84). Sun et al. Found a water-soluble polysaccharide from C. pilosula roots. Its backbone is constituted by (1→3)-linked β-D-pyranogalactose, (1→2,3)-linked β-D-pyranogalactose and (1→3)-linked α-D-pyranorhamnose. The O-2 position of Gal residues is substituted, in a 1:1 ratio, with side chains of a disaccharide, which consists of (1→5)-linked α-L-arabinofuranose units ( Figure 3E ) (85).

There are two types of immune organs: central immune organs (such as the bone marrow and thymus) and peripheral immune organs (including the spleen and lymph nodes). The Immune organ index is the ratio of the weight of the immune organ and the body weight (BW). It can reflect the development and function of the immune organ (an important indicator of immune function) (99). In addition, the immune organs can also secrete cytokines, such as IL-2, IL-6, IFN-γ, and TNF-α. These cytokines can induce the activation, proliferations, and differentiation of immune cells and are involved in both inflammatory and anti-infection processes. It is very essential to ensure structure and function of the immune organ to maintain the immune homeostasis, prevent pathogen invasion,and induce the correct response to vaccines (100).

CPPs have strong protective and regulatory effects in the CP-induced immunosuppressive mouse model. In the present experiment, the mice were injected intraperitoneally with CP at 60 mg/kg·BW·d for three consecutive days and then administered different doses of CPPs (50, 100, 200 mg/kg/day) orally for 7 days. The spleen index of all groups was significantly increased compared to the model group, and the spleen index of the 100 mg/kg/day group was even higher than that of the standard control group. The thymus index of the 200 mg/kg/day group was significantly increased and approached the normal group. In addition, the liver index of the 100 and 200 mg/kg/day groups was significantly increased compared with the model group (101). These findings indicate that CPPs effectively alleviate CP-induced damage to immune organs, particularly improving spleen atrophy, and also exert protective and enhancing effects on thymus and liver-related immune functions. CPPs may improve immunity by increasing immune organ indices and relieving immune organ suppression, providing experimental evidence for their potential as natural immunomodulators.

Immune factors are important molecules that mediate the regulation of the immune system, such as cytokines, antibodies, and complement. Cytokines include interleukins, interferons and tumor necrosis factors (102). They can mediate the proliferation, differentiation and activation of immune cells. They are involved in both innate and adaptive immune responses. Antibodies are secreted by B cells to mediate the neutralization of target cells, enhance phagocytosis of target cells, and induce antibody-dependent cellular cytotoxicity. Complements can directly cause damage to the pathogen’s membrane and cause inflammation. They can also enhance the effects of the antibody (103).

CPPs possess multidimensional pharmacological effects on regulating immune factors. On one hand, CPPs significantly regulate the secretion of multiple cytokines with bidirectional modulation. For example, low-molecular-weight (<30 kDa) β-(2→1)-fructan-rich fractions have been shown to bind TLR4 and activate MyD88-dependent NF-κB signaling pathways, resulting in increased production of pro-inflammatory cytokines such as TNF-α and IL-6, which are beneficial for applications in vaccine adjuvants and tumor immunotherapy (81). Conversely, pectic polysaccharides enriched in rhamnogalacturonan-I regions with high galacturonic acid content (>60%) exhibit complement C3 inhibitory activity (CH₅₀=2.06 mg/mL) and upregulate anti-inflammatory cytokine IL-10, suggesting potential utility in autoimmune and chronic inflammatory diseases such as COPD (104). When CPPs were applied to lipopolysaccharide-stimulated THP-1 inflammatory models, CPPs suppressed the expression of pro-inflammatory cytokines TNF-α and IL-1β while enhancing the activity of immune function in non-stimulated THP-1 cells, indicating that CPPs have the potential to balance the expression of immune factors (105). When applied to the chronic obstructive pulmonary disease model, CPPs downregulated the expression of pro-inflammatory cytokines (TNF-α, IL-6, IL-1β) and upregulated the expression of the anti-inflammatory cytokine IL-10 by suppressing the NF-κB signaling pathway, thereby reducing the expression of pro-inflammatory cytokines and alleviating airway inflammation (106). When applied to CP-induced immunosuppressed mice, CPPs significantly increased the contents of IL-2, IL-6 and TNF-α in serum and enhanced the immune response (107). When applied to poultry models, CPPs increased the titers of Newcastle disease virus antibodies, indicating that CPPs can enhance the humoral immune response and improve the vaccine effectiveness (108). When applied to piglets, CPPs significantly upregulated the protein and mRNA expression of SIgA, IgG and IgM in the small intestinal mucosa, indicating that CPPs can participate in local intestinal immunity (109).

Emerging evidence suggests that gut microbiota modulation may represent a primary mechanism underlying the systemic immunoregulatory effects of CPPs. Specifically, CPPs have been shown to enrich butyrate-producing bacterial taxa, leading to increased production of short-chain fatty acids (SCFAs) such as butyrate. Butyrate acts as a ligand for G-protein-coupled receptor 43 (GPR43), a receptor expressed on colonic epithelial cells and immune cells. Engagement of GPR43 can drive the differentiation and expansion of regulatory T cells (Tregs), thereby contributing to mucosal and systemic immune tolerance (9). From a translational perspective, it remains to be elucidated whether gut microbiota modulation constitutes the dominant axis of CPPs immunoregulation in vivo or acts in parallel with direct immune cell binding. Comparative studies evaluating germ-free versus conventional animal models could clarify the relative contributions of microbiota-dependent and microbiota-independent mechanisms.

Complement is an essential component of innate immunity (110). It consists of some plasma proteins and can be activated by the classic, alternative, and lectin pathways. After activation, complement can directly lyse pathogens and help phagocytes eliminate antigens by releasing active fragments, such as C3a and C5a and attracting immune cells to initiate an inflammatory response. Moreover, it synergizes with antibodies to amplify adaptive immunity and plays irreplaceable roles in infection control, immune surveillance, and maintaining immune balance (111).

Research has shown that CPPs possess definitive regulatory effects on the complement system, with anti-complement activity confirmed in several studies. For example, Lu CPPs significantly inhibit the activation of both classical and alternative complement path. The half-maximal inhibitory concentration (CH₅₀) for the classical pathway was 2.061 ± 0.127 mg/mL, and for the alternative pathway (AP₅₀) it was 6.725 ± 0.895 mg/mL, with effects showing explicit dose dependency. Mechanistic studies revealed that Lu CPPs significantly suppressed complement C3 expression in TNF-α-stimulated human hepatoma HepG2 cells. With increasing LCPP concentrations, C3 mRNA levels declined in a dose-dependent manner, suggesting inhibition of inflammation-mediated complement activation (104). Additionally, pectic polysaccharides 50WCP-II-I and 100WCP-II-I, extracted from C. pilosula roots, demonstrated complement-fixing activity. Enzymatic hydrolysis showed that activity was primarily mediated by branched regions, with high molecular weight hydrolysates (e.g., 50WCP-II-Ia and 100WCP-II-Ia) exhibiting greater activity than the native polysaccharides (72).

In vivo studies have also shown that CPPs positively regulate immune parameters. For instance, CPPs significantly increased serum levels of immunoglobulins (IgA, IgM, IgG) and complement components C3 and C4 in weaned piglets, thereby enhancing overall immune function (112). These findings provide pharmacological evidence for the use of CPPs in treating diseases associated with complement overactivation, including autoimmune disorders.