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. 2026 Jun 9;91(6):e71178. doi: 10.1111/1750-3841.71178

Antidiabetic Activity of Chickpea Albumin Hydrolysate in Streptozotocin‐Induced Diabetic Rats

Alicia Navarro‐Leyva 1, Gabriela López‐Angulo 1,✉, Francisco Delgado‐Vargas 1, Aimée Bastidas‐Ponce 2, Nancy Yareli Salazar‐Salas 1, Jenifer Mariana Soto‐Lozoya 1, José Ángel López‐Valenzuela 1,✉
PMCID: PMC13249347  PMID: 42263213

ABSTRACT

Chickpea (Cicer arietinum L.) is a nutritious food that contains bioactive peptides with hypoglycemic and antihyperglycemic activities. This study evaluated the antidiabetic effects of chickpea albumin hydrolysate (CAH) in rats with hyperglycemia induced by a high‐fat diet (HFD) and streptozotocin (STZ). The CAH peptide profile was analyzed by liquid chromatography‒mass spectrometry. Five groups (n = 6) were established: one healthy control (HC) and four diabetic groups: diabetic control (DC), metformin (500 mg/kg body weight, b.w.; MET), CAH (200 mg/kg b.w.; H200), and CAH (400 mg/kg b.w.; H400). Food intake, body weight, and fasting blood glucose (FBG) were assessed weekly, and an oral sucrose tolerance test (OSTT) was performed. Blood and liver were analyzed for biochemical parameters (glucose and lipid profile), renal (urea and creatinine) and hepatic function (AST, ALT, and ALP), oxidative stress markers (GSH and MDA), gluconeogenic (PEPCK and G6Pase), and pentose phosphate (G6PD) enzymes, and PI3K/AKT and AMPK signaling pathways. CAH presented peptides predicted to inhibit DPPIV and α‐glucosidase. OSTT showed that CAH (H200 and H400) reduced blood glucose levels by 18.6% and 22.8%, respectively, while metformin reduced them by 41.6%. CAH and metformin improved the lipid profile, decreased urea and creatinine levels, and attenuated elevations in hepatic enzymes. They showed antioxidant effects by increasing GSH and reducing MDA levels. CAH‐treated rats showed pancreatic tissue restoration, reduced PEPCK and G6Pase activities, and increased G6PD activity. Exploratory immunoblotting analyses revealed qualitative differences in the phosphorylation of AMPK (muscle and liver) and AKT (muscle), which may be associated with the metabolic effects of CAH. These findings suggest that chickpea bioactive peptides could be helpful in the management of diabetes.

Keywords: antidiabetic effect, chickpea hydrolysates, diabetes

1. Introduction

Diabetes mellitus (DM) is a highly prevalent chronic disease and a major health concern worldwide. In 2025, the International Diabetes Federation reported 589 million people with DM, and this number is estimated to rise to 853 million by 2050 (International Diabetes Federation 2025).

DM is characterized by persistent hyperglycemia due to metabolic alterations. Type 1 diabetes (T1D) is caused by the destruction of pancreatic β cells, while Type 2 diabetes (T2D) is mainly associated with insulin resistance and reduced insulin secretion (Oguntibeju 2019). T2D is related to metabolic and systemic complications, as evidenced by alterations in biomarkers such as lipid profile, serum urea and creatinine levels, liver enzymes (ALT, AST, and ALP), and oxidative stress markers (malondialdehyde [MDA] and glutathione [GSH]) (Schuster and Duvuuri 2002). In addition, the PI3K/AKT and AMPK signaling pathways are often dysregulated in T2D. Under normal conditions, the PI3K/AKT pathway is activated by insulin and promotes glucose uptake and storage (Feng et al. 2024), while AMPK is activated by low energy levels and promotes energy‐generating processes to reduce blood glucose (Kibur et al. 2025). In T2D, insulin resistance has been associated with inhibition of the PI3K/AKT pathway, and AMPK activation can counteract this by improving insulin sensitivity and glucose metabolism. In addition, AMPK activation mitigates oxidative stress, lipotoxicity, and inflammation, thereby ameliorating diabetes‐induced organ damage (Kakoti et al. 2024).

Currently, several drugs are used to prevent and treat T2D, but they usually have side effects. In this regard, plants could provide safe and effective bioactive compounds as accessible alternatives for treating this disease. P. Li et al. (2021) showed that a chickpea extract improved hyperglycemia, inflammatory status, and organ function in diabetic rats. Shahzad et al. (2025) also demonstrated that chickpea and barley extracts reduced hyperlipidemia, hyperglycemia, and insulin resistance in diabetic mice. Regarding bioactive peptides, it was previously shown that CAH inhibits enzymes involved in carbohydrate metabolism (Quintero‐Soto et al. 2021) and induces hypoglycemic and antihyperglycemic effects in normoglycemic mice (Navarro‐Leyva et al. 2023). Radlowski et al. (2024) administered a chickpea protein hydrolysate to an animal model of metabolic dysfunction and observed reductions in liver inflammation and liver fat accumulation. These results suggest the potential of chickpea protein hydrolysates for the treatment of T2D. Thus, the aim of this study was to analyze the antidiabetic activity (AD) of CAH in a rat model of diabetes induced by a high‐fat diet (HFD) and streptozotocin (STZ). These findings could be employed to develop chickpea‐based functional foods to prevent or treat diabetes.

2. Materials and Methods

2.1. Biological Material

The desi chickpea (Cicer arietinum L.) ICC 3761 was grown during the 2022–2023 season at the Experimental Field of the National Institute of Forestry, Agricultural, and Livestock Research (INIFAP), Culiacán, Sinaloa, México, as described by Chavez‐Ontiveros et al. (2020). Mature seeds were processed with a Retsch MM400 ball mill (Retsch GmbH, Germany) and passed through a 60‐mesh sieve.

2.2. Animals

Male Wistar rats (Bioinvert, SA de CV, México), 4 weeks old (100–120 g), were housed in acrylic boxes under controlled conditions (24 ± 1°C, 12 h light/dark cycle) with ad libitum access to water and standard rodent feed (Nutricubes, Purina). All procedures complied with the Mexican Official Standard NOM‐062‐ZOO‐1999 and were approved by the Animal Care and Use Committee (IACUC) of the Faculty of Chemical and Biological Sciences of the Universidad Autónoma de Sinaloa (Approval number: 01‐02‐CICUAL‐FCQB26).

2.3. Extraction and Quantification of the Albumin Fraction

Albumins were extracted based on Osborne fractionation as described by Navarro‐Leyva et al. (2023). Defatted chickpea flour (1 g) was resuspended in sterile deionized water (pH 8.0), stirred for 2 h at 20°C, and centrifuged (15,000 × g, 20 min; 5804R, Eppendorf, Hamburg, Germany), and the supernatant was lyophilized (25EL, VirTis Co., Gardiner, NY, USA). The lyophilized material represented the albumin fraction obtained via water‐soluble protein fractionation. The protein obtained was sequentially washed with methanol (200 mL) and acetone (200 mL) to remove nonprotein contaminants such as polyphenols. The albumins were resuspended in sterile water, and the concentration was estimated with the bicinchoninic acid (BCA) method (Brown et al. 1989).

2.4. Preparation of CAH

Albumin hydrolysates were obtained by sequential hydrolysis with pepsin and pancreatin as reported by Navarro‐Leyva et al. (2023). Approximately 1 g of albumin protein was mixed with pepsin solution (P6887, Sigma‐Aldrich, St. Louis, MO, USA) (20 mL of 0.01 M HCl, pepsin; pH 2) and stirred for 90 min at 37°C. Subsequently, the pH of the sample was adjusted to 8.0 with 1 M NaOH, pancreatin solution (P7545, Sigma‐Aldrich) (7.5 mL of phosphate buffer, pancreatin; pH 8.0) was added, and the mixture was stirred for 90 min at 37°C. The enzyme was inhibited by heating (80°C, 20 min). The mixture was centrifuged, and the supernatant was filtered (0.45 µm, PVDF). The filtrate was passed through 10 kDa cut‐off tubes and lyophilized to obtain the CAH. The degree of hydrolysis was determined using the TNBS method (Adler‐Nissen 1979).

2.5. Characterization of the CAH‐Derived Peptides

2.5.1. Peptide Profile of CAH

A 15‐µL aliquot of CAH aqueous solution (150 mg/mL) was injected into an ACCELA UPLC‐DAD system (Thermo Scientific, San Jose, CA, USA). The separation was carried out using a Luna C18 column (150 × 4.6 mm, 5 µm, Phenomenex, Torrance, CA, USA). The mobile phase consisted of water with 1% formic acid (A) and acetonitrile (B) at a flow rate of 0.2 mL/min. The gradient was as follows: 0–20 min, 95.5–96% A; 20–68 min, 96–80% A; 68–115 min, 80–52% A; 115–120 min, 52–100% A. The detection was performed at 257 and 280 nm. The peptide sequences were determined using a linear ion‐trap mass spectrometer (LTQ‐XL, Thermo Scientific, Waltham, MA, USA) with an electrospray ionization source operating in positive mode. The capillary voltage was set at 35 V, and the temperature was set at 300°C. Mass spectra (m/z 50–2000) were acquired with Xcalibur 2.1 software, and precursor ions were fragmented by collision‐induced dissociation (CID). Data were analyzed using Proteome Discoverer 1.2 with the SEQUEST algorithm and the UNIPROT C. arietinum proteome database (ID: UP000087171). Cysteine carbamoylmethylation was considered a fixed modification, and methionine oxidation was considered a variable modification. The raw files and peptide sequences identified in Proteome Discoverer were imported into Skyline (MacLean et al. 2010) to determine their relative abundances. Extracted ion chromatograms (XICs) were generated for each peptide, and the peak areas were automatically integrated. The relative abundance of each peptide was estimated based on the normalized peak areas. The potential biological activities of the identified peptides were predicted using the BIOPEP‐UWM database, considering the entire peptide sequences or specific fragments.

2.5.2. In Silico Analysis of CAH‐Derived Peptides

In silico gastrointestinal digestion of the peptides identified by UPLC–MS/MS was performed using the BIOPEP‐UWM database (Minkiewicz et al. 2008). Proteolytic cleavage was performed simultaneously using pepsin (EC 3.4.23.1), trypsin (EC 3.4.21.4), chymotrypsin (EC 3.4.21.1), and elastase (EC 3.4.21.36). The ADMET properties of the selected peptides were predicted using ADMETlab 3.0 (Xiong et al. 2021). For molecular docking analysis, the 3D structures of the peptides were generated using PeptideBuilder in MOE v.2014.09.01 (Chemical Computing Group, Montreal, Canada). Next, the ligands were prepared by assigning partial charges and adding hydrogen atoms. The MFF94 force field was applied to minimize the energy of the structures. The structures of the α‐glucosidase (PDB ID: 5NN8) and DPPIV (PDB ID: 4PNZ) proteins were obtained from the Protein Data Bank (https://www.rcsb.org/), and nonstandard molecules and water molecules were removed. Molecular docking was performed using the MOE Dock protocol. For α‐glucosidase, the analysis was directed at the active site (Asp282, Asp518, Glu521, and Asp616) and basic residues (Arg600 and His674). DPPIV was directed to the S1 active site, which contains hydrophobic residues (Tyr547, Tyr631, Val656, Trp659, Tyr662, Tyr666, and Val711), the catalytic site (Ser630, Asp708, Asn710, and His740), and the active sites S2 (Glu205, Glu206, and Arg125) and S2′ (Val207, Ser209, Arg358, and Phe357).

2.6. Establishment of a T2D‐Like Rat Model and Treatments

After a 7‐day adaptation period, 40 rats were divided into two groups: a control group (n = 8) and a group for the induction of a T2D‐like model (n = 32) based on a HFD (Abdul Kadir et al. 2015) and sucrose solution (10%) ad libitum for 2 weeks, followed by the injection of three intraperitoneal (ip) doses of STZ (35 mg/kg b.w.) at weekly intervals with overnight fasting (12 h) (M. Zhang et al. 2008). This combined approach is commonly used to generate a T2D‐like model characterized by diet‐induced insulin resistance and partial dysfunction of pancreatic β‐cells, resulting in sustained hyperglycemia (Brito et al. 2025; Gheibi et al. 2017).

STZ was resuspended in cold citrate buffer (0.1 M citric acid, pH 4.5) and immediately injected into the animals. Control rats received an equivalent intraperitoneal dose of the citrate buffer used as vehicle (pH 4.5, 1 mL/kg body weight). The rats were provided with 5% sucrose solution ad libitum for 48 h to avoid hypoglycemia. Five days after STZ injection, the rats were fasted for 5 h (7 a.m. to 12 p.m.), and blood samples were obtained from the tail vein to measure glucose levels and to confirm hyperglycemia. Rats with fasting blood glucose (FBG) levels between 180 and 300 mg/dL (Jiang et al. 2014) were considered diabetic and included in the study (24 rats), while 8 rats that did not meet this criterion were excluded.

The rats were randomly selected to form five groups (n = 6), which were assigned to the treatments based on random numbers: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400). No animals were excluded after assignment to the experimental groups. The number of animals per group was chosen based on previous studies using Wistar rats (n = 6) to evaluate the antidiabetic effect of protein hydrolysates (Ina et al. 2020; Olasehinde et al. 2023). The CAH dose of 200 mg/kg b.w. was previously shown to have an antihyperglycemic effect in mice (Navarro‐Leyva et al. 2023). A 400 mg/kg b.w. dose was included based on previous studies with legume protein hydrolysates, which have reported metabolic benefits at doses of 400–1000 mg/kg b.w. in rodent models (Liao et al. 2023; Radlowski et al. 2024). In addition, the use of metformin at 500 mg/kg b.w. has been effective in lowering blood glucose and protecting several organs in murine models of diabetes (Z. Li and Zhang 2017; Escobar‐Zuñiga et al. 2025). Furthermore, nondiabetic and diabetic rats treated with 1000 mg/kg/day metformin for one month showed normal histological findings in pancreatic, hepatic, and renal tissues (Almuttairi 2023). The treatments were given orally once daily for 30 days. Body weight and FBG levels were measured once a week.

At the end of the treatments, overnight fasted rats were anesthetized, and blood was drawn by cardiac puncture into tubes without anticoagulant. The blood was centrifuged (1,500 × g, 10 min; 5410, Eppendorf) to obtain the serum. The animals were dissected to remove the liver and pancreas. The samples were immersed in liquid nitrogen and then stored at −70°C until use. The pancreas was placed in formalin (37%) for subsequent blinded histopathological analysis.

2.7. Oral Sucrose Tolerance Test

An oral sucrose tolerance test (OSTT) was performed after three weeks of treatment. After 6 h of fasting, the rats received the treatments, and 30 min later, all groups received an intragastric sucrose load (3 g/kg b.w.). Blood samples were collected from the tail vein every 30 min for 2 h. Blood glucose concentration (mg/dL) was measured using a glucometer (Accu‐Chek Instant Kit, Roche, México), and the increase in the area under the glucose curve (mg min/dL) was calculated.

2.8. Biochemical Analyses

Biochemical analyses were performed using commercial kits according to the manufacturer's instructions: triglycerides (TGML‐0427, ELITechGroup, Puteaux, France), total cholesterol (11505, Biosystems, Barcelona, Spain), and HDL‐c (11523, BioSystems), while LDL‐c was determined using the Friedewald et al. (1972) equation:

LDL−cmg/dL=TC−HDL−TG/5

2.9. Renal Function

Commercial kits for urea (URSL‐0427, ELITechGroup) and creatinine (CRCO‐0600, ELITechGroup) were used according to the manufacturer's protocols.

2.10. Hepatic Function and Oxidative Stress Markers

2.10.1. Serum Enzymes

Commercial kits were used to evaluate the activities of serum aspartate aminotransferase (ASSL‐0430, ELITechGroup), alanine aminotransferase (ALSL‐0430, ELITechGroup), and alkaline phosphatase (PASL‐0420, Pointe Scientific Inc., Canton, MI, USA), following the manufacturers' instructions.

2.10.2. Liver MDA and Reduced GSH

MDA and GSH levels were measured from liver samples (50 mg) using commercial kits (MDA, K739‐100, Biovision, Waltham, MA, USA; GSH, CS0260‐1KT, Sigma‐Aldrich) according to the manufacturer's instructions and as described by Navarro‐Leyva et al. (2023). The results were reported as nanomoles of MDA or GSH per mg of liver tissue (nmol/mg).

2.11. Pancreatic Histopathology

The pancreatic tissue was fixed with 10% formaldehyde, dehydrated, and embedded in paraffin. Sections of 5–7 µm were obtained using a microtome (Leica Biosystems) and stained with hematoxylin and eosin (H&E). They were examined under a brightfield microscope (Axioscope 5, Carl Zeiss Microscopy, Oberkochen, Germany). The histopathologist was blinded with respect to the treatment groups.

2.12. Gluconeogenesis Regulation

2.12.1. Glucose‐6‐Phosphatase

Glucose‐6‐phosphatase (G6Pase) activity was determined as described by Algandaby et al. (2010) and Choi et al. (2017). A liver sample (100 mg) was homogenized with 900 µL of cold buffer (250 mM sucrose, 25 mM HEPES, pH 7.4, 2.5 mM EDTA, and 1 mM phenylmethylsulfonyl fluoride) using an ultrasonic homogenizer (4710 Series, Parmer Instrument Company, Chicago, IL, USA). The mixture was centrifuged (12,000 × g, 20 min at 4°C; 5410, Eppendorf), and the pellet was resuspended in 100 µL of cold homogenizer buffer. A 50‐µL aliquot of the extract was mixed with 200 µL of incubation buffer (50 mM HEPES, pH 7.2, 100 mM KCl, 2.5 mM EDTA, MgCl2, 1 mM glucose‐6‐phosphate), and the G6Pase activity was determined by monitoring the phosphate released by registering the absorbance at 660 nm (Synergy HTX, BioTek, Winooski, VT, USA). The results are reported as nmol of Pi/min/mg of protein.

2.12.2. Phosphoenolpyruvate Carboxykinase

The method described by Ramirez et al. (2014) was used. The liver sample (50 mg) was homogenized with 500 µL of cold buffer [0.25 M sucrose and 10 mM Tris‐HCl (pH 7.4)] using an ultrasonic homogenizer (4710 Series, Parmer Instrument Company). The sample was centrifuged (14,500 g × 10 min; 5410, Eppendorf), and 10 µL of the recovered supernatant was mixed with 190 µL of buffer containing 50 mM Tris‐HCl (pH 7.4), 1 mM MgCl2, 0.1 mM NADH, 2 U malate dehydrogenase, 0.5 mM phosphoenolpyruvate, and 0.2 mM guanosine 5′‐diphosphate. The absorbance was measured at 340 nm for 5 min at 37°C, 20 mM NaHCO3 was added to the mixture, and the reaction was monitored at 340 nm (Synergy HTX, BioTek) every minute for 30 min. The results were expressed as nmol of NADH/min/mg of protein.

2.13. Pentose Phosphate Pathway Regulation

2.13.1. Glucose‐6‐Phosphate Dehydrogenase

Glucose‐6‐phosphate dehydrogenase (G6PD) activity was determined according to the methods of Tian et al. (1998) and Gilglioni et al. (2018). Rat liver (50 mg) was homogenized with 500 µL of cold buffer [0.25 M sucrose and Tris‐HCl (pH 7.4)] using an ultrasonic homogenizer (4710 Series, Parmer Instrument Company). The sample was centrifuged (13,000 × g, 15 min; 5410, Eppendorf), and then 20 µL of the supernatant was mixed with 160 µL of incubation buffer (50 mM Tris‐HCl, 2 mM MgCl2, 0.2 mM NADP+). The mixture was added to 20 µL of 10 mM glucose‐6‐phosphate as a substrate. The NADPH produced was quantified by measuring the absorbance at 340 nm over 5 min (Synergy HTX, BioTek). The enzyme activity was reported as nmol of NADPH/min/mg of protein.

2.14. Analyses of PI3K/AKT and AMPK Signaling Pathways

Frozen muscle and liver samples (50 mg) were pulverized with liquid nitrogen and homogenized with 500 µL of RIPA lysis buffer [50 mM Tris‐HCl (pH 8), 150 mM NaCl, 1% NP‐40, 0.5% sodium deoxycholate, 1% SDS] supplemented with protease and phosphatase inhibitors (Halt, 100×). The samples were left on ice for 30 min and then centrifuged (12,000 × g, 15 min at 4°C) to recover the supernatant (Jiang et al. 2014). Equal amounts of protein (50 µg) from each sample were separated on a 15% polyacrylamide gel (SDS‒PAGE) and then transferred to a polyvinylidene fluoride (PVDF) membrane (Hybond‐ECL Amersham Biosciences). Nonspecific binding sites were blocked using skim milk (5%) (Svelty, Nestlé) for 1 h. The membrane was incubated for 1 h with the corresponding primary antibodies [AKT (1:1000), p‐AKT (1:1000), AS160 (1:1000), p‐AS160 (1:1000), AMPK (1:1000), p‐AMPK (1:1000), and GAPDH (1:1000)] (Cell Technology Inc., Williamsburg, VA, USA) followed by incubation for 1 h with horseradish peroxidase‐conjugated goat anti‐rabbit IgG (Sigma‐Aldrich). Detection was performed with 3,3′‐diaminobenzidine (DAB) (Sigma‐Aldrich), and images were acquired with a ChemiDoc XRS system (Bio‐Rad, Hercules, CA, USA).

2.15. Statistical Analysis

The results are reported as the mean ± standard deviation (SD). Data from variables measured repeatedly over time (glucose levels, body weight, and food intake) were analyzed using repeated‐measures ANOVA after sphericity tests and Greenhouse‒Geisser corrections. For other variables, the normality of the residuals (Shapiro‒Wilk) and homogeneity of variances (Levene's test) were verified, and one‐way ANOVA was applied, followed by Fisher's least significant difference (LSD) post hoc test. Statistical analyses were performed using Minitab v.19.1 (Minitab Inc., State College, PA, USA) and GraphPad Prism v.8.0.1 (GraphPad Software, San Diego, CA, USA).

3. Results and Discussion

3.1. Identification and In Silico Characterization of CAH‐Derived Peptides

The degree of hydrolysis of CAH was 53.8%, and 911 peptide sequences were identified using UPLC–MS/MS (Table S1). The peptides varied in length from 3 to 19 amino acids, with protonated molecular masses (MH+) ranging from 260 to 1970 Da. In silico analysis showed that the peptides have important predicted biological activities, highlighting antidiabetic activity: 97% of the identified peptides are potential DPPIV inhibitors, 16% are α‐glucosidase inhibitors, and 37% are glucose uptake stimulants. CAH was previously shown to inhibit α‐glucosidase (52.4%, 100 mg/mL) (Navarro‐Leyva et al. 2023).

Semiquantitative analysis based on peak area normalization showed that 10 peptides accounted for 47% of the total abundance, and the highest values were for EVDGVY (6.58%), NMTRc (6.58%), EDTTDT (6.5%), and ENTASc (6.58%). These 10 peptides were selected for simulated gastrointestinal digestion (Table S2). A total of 13 unique peptides were identified (5 tripeptides and 8 dipeptides).

The ADMET peptide characteristics suggested their potential as oral bioactive compounds. Some peptides have high predicted intestinal absorption (QH, QPL, EN, EV, DG, and EK). The predicted plasma protein binding (PPB) is low, which could contribute to increased bioavailability of the free active fraction. Low interaction with cytochrome P450 enzymes indicates low metabolic interference. Furthermore, T 1/2 values indicate a moderate half‐life. Similarly, no significant oral toxicity was anticipated. These data suggest the potential use of chickpea‐derived peptides as antidiabetic agents. In addition, molecular docking analysis was performed with the enzymes α‐glucosidase and DPPIV, resulting in binding energies of −4.24 to −6.13 kcal/mol and −4.70 to −7.19 kcal/mol, respectively (Table 3). PPK and QPL showed the highest affinity for α‐glucosidase and DPPIV, respectively. The binding free energies of these peptides were close to those of acarbose and omarigliptin. PPK established multiple hydrogen bonds and hydrophobic interactions with key catalytic residues of α‐glucosidase, including Asp282, Asp518, and Asp616 (Figure S1). QPL established multiple interactions with residues of DPPIV, including Tyr547, Tyr631, Tyr666, Asn710, Glu205, Glu206, and Arg125 (Figure S1).

3.2. Healthy and Diabetic Rats Treated With CAH Showed Similar Food Intake and Body Weight Patterns

The effect of the treatments on the animal's food intake (Figure 1A) and body weight (Figure 1B) was assessed during the four weeks of the experiment. The groups of diabetic rats (DC, MET, H200, and H400) had a higher initial intake than the HC group, attributable to the induced diabetes; however, the groups treated with CAH (H200 and H400) maintained a stable consumption pattern throughout the study, indicating good treatment tolerance (Figure 1A). Regarding body weight, all groups showed a progressive increase, with no significant differences among them (Figure 1B), suggesting that variations in food intake did not affect weight gain because of compensatory mechanisms in energy metabolism.

FIGURE 1.

FIGURE 1

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Effect of CAH on food intake (A) and body weight (B) in diabetic rats (DR). Values are expressed as the mean ± standard deviation (n = 6). *Significant difference from the HC (p ≤ 0.05). #Significant difference from the DC (p ≤ 0.05). Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

These results correspond with those reported in diabetic murine models treated with protein hydrolysates of whey (Gregersen et al. 2013) and potato (Asokan et al. 2019), where no relevant changes in appetite or body weight were observed. Collectively, the evidence indicates that CAH exerts beneficial metabolic effects without altering feeding behavior or compromising nutritional status, supporting its potential as a therapeutic agent in diabetes and obesity.

3.3. CAH Treatment Reduces FBG and Improves Sucrose Tolerance in Diabetic Rats

At the beginning of the treatment, the FBG levels of rats from the diabetic groups (DC, MET, H200, and H400) were above 250 mg/dL, whereas those of the HC group were close to 100 mg/dL (Figure 2A). After four weeks of intervention, the DC group maintained elevated glucose concentrations (260 ± 68 mg/dL) compared with the HC group (111 ± 5 mg/dL). In contrast, the groups treated with CAH (H200: 147 ± 18 mg/dL; H400: 147 ± 17 mg/dL) or metformin (MET: 148 ± 19 mg/dL) exhibited a significant reduction in FBG compared to the DC group, although the values were slightly higher than those of the HC group (Figure 2A). These findings indicate comparable hypoglycemic effects of CAH and metformin.

FIGURE 2.

FIGURE 2

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Effect of CAH on fasting blood glucose levels (A), glycemic response in diabetic rats (DR) after sucrose administration (3 g/kg b.w.) (B), and area under the curve (AUC) of the 0–120 min glucose increments (C). Data are expressed as the mean ± standard deviation (n = 6). *Significant difference from HC (p ≤ 0.05). #Significant difference from DC (p ≤ 0.05). Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

A similar reduction in FBG was reported for chickpea methanolic extracts in diabetic mouse models, an effect attributed to inhibition of digestive enzymes, including α‐glucosidase (Shahzad et al. 2025). Wei et al. (2019) and Liao et al. (2023) also observed hypoglycemic effects in diabetic mice treated with pea hydrolysates. The authors suggested that hydrolysates promote insulin release, improve insulin sensitivity in insulin‐responsive tissues, accelerate transmembrane glucose transport, and stimulate muscle glycogen synthesis. Furthermore, L. Li et al. (2022) reported a stabilizing effect of mung bean‐derived peptides in mice (C57BL/6) with insulin resistance.

In the OSTT, the initial glucose levels of the MET, H200, and H400 groups were significantly lower than those of the DC group but higher than those of the HC group (Figure 2B). Sucrose administration (3 g/kg b.w.) increased postprandial glucose at 30 min in all treatments, except in the metformin group; the DC group showed the highest glucose levels. After 120 min, rats treated with CAH (H200 and H400) and metformin showed glucose levels 18.6%–22.8% and 41.6% lower than those of the DC group, respectively (Figure 2C). These results indicate that CAH improves sucrose tolerance, possibly by inhibiting enzymes involved in carbohydrate metabolism (Navarro‐Leyva et al. 2023), consistent with its administration preceding the sucrose load. These effects are supported by the predicted activities of the peptides identified in the CAH by UPLC–MS (Table S1).

These results agree with those obtained in previous studies using animal models treated with plant‐derived bioactive peptides. Ina et al. (2020) evaluated rice albumin hydrolysates (133 and 67 mg/kg b.w.) in rats with induced diabetes and demonstrated that the treatment suppressed the increase in blood glucose and plasma insulin levels. L. Li et al. (2022) also showed that a mung bean hydrolysate (245 mg/kg b.w.) decreased the area under the glucose curve and reduced the postload glycemic peak in mice with insulin resistance.

Collectively, these results suggest the potential of CAH for glycemic control. The human equivalent dose (HED) was estimated by using the body surface area normalization method (Reagan‐Shaw et al. 2008). Based on this analysis, doses of 200 and 400 mg/kg b.w. in rats are approximately equal to 33 and 67 mg/kg in humans, respectively, which corresponds to an estimated intake of 2.0–4.0 g/day for a 60 kg adult. Although these doses may be difficult to achieve through a conventional diet, they could be obtained by consuming concentrated formulations or nutraceutical supplements. Human studies using hydrolyzed milk protein have shown metabolic effects with intake levels ranging from 1.4 to 2.8 g/day (Sartorius et al. 2019).

3.4. CAH Treatment Improves the Lipid Profile in Diabetic Rats

DM is frequently associated with metabolic alterations such as hypercholesterolemia and hypertriglyceridemia, increasing the risk of cardiovascular disease (Kilari et al. 2021). The effects of the treatments on lipid metabolism are shown in Figure 3. The HC group showed TG, TC, and LDL values within the normal range reported for Wistar rats (Olasehinde et al. 2023; Feyisa et al. 2019), whereas the DC group displayed significantly higher values of these parameters, reflecting an atherogenic profile induced by the HFD. Conversely, treatment with metformin and CAH (H200 and 400) improved the lipid profile by decreasing TG, TC, and LDL levels and increasing HDL levels, suggesting a hypolipidemic effect. However, the H200 and H400 treatments generated similar effects, which could be explained by the pharmacodynamic model of maximum effect. A dose‒response curve usually follows a sigmoidal model, but once the threshold effect is reached, increasing the dose does not generate proportional increases in effect due to saturation processes (Holford 2017).

FIGURE 3.

FIGURE 3

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Effect of CAH on the lipid profile of diabetic rats (DR). Values are expressed as the mean ± standard deviation (n = 6). Different letters indicate significant differences between treatments (Fisher's LSD test, p ≤ 0.05). Biochemical parameters: total cholesterol (TC), high‐density lipoprotein (HDL), low‐density lipoprotein (LDL), and triglycerides (TG). Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

Several studies have reported the effect of protein hydrolysates on lipid profiles in different animal models. Wei et al. (2019) observed that inducing diabetes in rats increased TC, TG, and LDL and reduced HDL, whereas treatment with pea oligopeptide (800, 1600, and 3200 mg/kg b.w.) attenuated these alterations. W. Li et al. (2023) demonstrated that treatment with the low‐MW fraction (< 1 kDa) of soybean hydrolysates decreased LDL levels and increased HDL in mice with STZ‐induced diabetes. Similarly, Radlowski et al. (2024) demonstrated a dose‐dependent effect of chickpea protein hydrolysates in mice fed an HFD, with the highest dose (800 mg/kg) yielding the lowest TG levels. More recently, Shahzad et al. (2025) showed that methanolic chickpea extracts normalized lipid profiles in STZ‐induced diabetic mice, restoring values comparable to those of the positive control group.

3.5. CAH Treatment Improves Renal Function in STZ‐Induced Diabetic Rats

DM is characterized by hyperglycemia, and if this condition is prolonged, it leads to chronic damage, malfunction, and failure of organs, including the kidneys. In fact, DM is the leading cause of progressive renal insufficiency requiring dialysis or transplantation (Mestry et al. 2017). Serum urea and creatinine are indicators of renal dysfunction (Anwar et al. 2024). Metformin and CAH treatments decreased the serum urea concentration compared to DC (Figure 4A). Creatinine levels in the DC group were significantly higher than those in the HC group (Figure 4B), which may be attributable to renal dysfunction. Metformin treatment significantly reduced creatinine levels compared with all groups, including HC, whereas CAH (H200 and H400) also lowered creatinine levels relative to DC, although to a lesser extent. These findings suggest that CAH has a kidney‐protective effect, consistent with prior reports. Hidayat et al. (2019) found that pea protein hydrolysates reduced serum urea and creatinine in rats with induced kidney injury, while Shahzad et al. (2025) showed that methanolic chickpea extracts decreased these biomarkers in diabetic mice. The decreases in urea and creatinine induced by metformin may be due to its effects on energy and renal metabolism. Metformin inhibits mitochondrial complex I, increasing the NADH/NAD + ratio and promoting the conversion of pyruvate to lactate, thereby reducing gluconeogenesis and urea synthesis (Hou et al. 2018; Holzhütter et al. 2025). In addition, the accumulation of metformin may promote mild lactic acidosis when kidney damage is present (Barthelmebs et al. 2003). This phenomenon may explain the decrease in creatinine below the physiological range observed in the MET group. In contrast, the CAH‐treated rats showed normal levels of renal parameters (creatinine ≈ 0.2–0.8 mg/dL; urea 20–45 mg/dL). Therefore, this effect may indicate the preservation or recovery of renal function.

FIGURE 4.

FIGURE 4

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Effect of CAH on serum levels of urea (A) and creatinine (B) in diabetic rats (DR). Data are expressed as the mean ± standard deviation (n = 6). Different letters indicate significant differences (Fisher's LSD test, p ≤ 0.05). Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

3.6. CAH Treatment Has a Hepatoprotective Effect

The liver is central to metabolic homeostasis; its impairment in T2D is associated with inflammation, oxidative stress, and hepatocellular injury, often reflected by elevated serum levels of aminotransferases (ALT and AST) and ALP (Krisnamurti et al. 2022; Teshome et al. 2019). The DC group showed significantly higher ALT, AST, and ALP levels than the HC group, indicating STZ‐induced hepatic damage (Figure 5). Treatment with metformin and CAH (H200 and H400) significantly decreased the levels of these enzymes, suggesting a hepatoprotective effect.

FIGURE 5.

FIGURE 5

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Effect of CAH on serum levels of ALT (A), AST (B), and ALP (C) in diabetic rats (DR). Data are expressed as the mean ± SD (n = 6). Different letters indicate significant differences (Fisher's LSD test, p ≤ 0.05). Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

The administration of STZ also increased reactive oxygen species (ROS) production, leading to significant alterations in hepatic antioxidant enzymes (Zafar et al. 2009; Vinothiya and Ashokkumar 2017). Oxidative stress was assessed through MDA and GSH content (Figure 6). GSH levels (Figure 6A) tended to be lower in DC, whereas treated groups exhibited higher values, suggesting that treatments improved liver antioxidant potential. As expected, the DC group showed the highest MDA levels, and the metformin and CAH treatments significantly attenuated lipid peroxidation (Figure 6B). These findings indicate that both metformin and CAH mitigate hepatic oxidative stress by restoring redox balance.

FIGURE 6.

FIGURE 6

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Effect of CAH on the liver GSH (A) and MDA (B) levels of diabetic rats (DR). Data are expressed as the mean ± SD (n = 6). Different letters indicate significant differences (Fisher's LSD test, p ≤ 0.05). Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

Comparable hepatoprotective and antioxidant effects of protein hydrolysates have been reported previously. L. Li et al. (2022) demonstrated that a mung bean hydrolysate improved the liver antioxidant status by increasing SOD and reducing MDA in mice fed an HFD. Similarly, W. Li et al. (2023) evaluated the low MW fraction (< 1 kDa) of an extruded soybean protein hydrolysate in mice with T2D. A 4‐week dietary intervention reduced MDA content, decreased ALT and AST activities, and alleviated liver damage, suggesting that soybean hydrolysate may have a beneficial effect in the treatment of diabetes. More recently, Shahzad et al. (2025) reported that methanolic extracts of chickpea decreased ALT, AST, ALP, and MDA levels while restoring GSH in diabetic mice. Collectively, these results demonstrate that CAH exerts hepatoprotective and antioxidant effects in diabetic rats, comparable to those of metformin and consistent with evidence from other plant protein hydrolysates. The nonsignificant differences observed in ALT, AST, ALP, and GSH levels between H200 and H400 doses may be due to a threshold of maximal effect.

3.7. CAH Treatment Has a Pancreatic‐Protective Effect in Diabetic Rats

The integrity of the pancreatic tissue was analyzed with histological sections stained with H&E (Figure 7). The HC group showed oval‐shaped islets of Langerhans with well‐defined boundaries and clear contours, with no apparent architectural damage (Figure 7A). In contrast, the DC group showed smaller atrophic islets with irregular shapes, poorly defined edges, and loss of cellular organization, consistent with STZ‐induced damage. Pancreatic damage was evidenced by increased interstitial space and disorganization of acinar tissue, changes consistent with mild inflammation (Figure 7B). On the other hand, the metformin‐treated rats partially recovered their pancreatic structure, with more defined boundaries than in the DC group (Figure 7C). The groups treated with CAH (H200 and H400) showed structural restoration comparable to the effect of metformin (Figure 7D,E). The pancreatic morphology of H400‐treated diabetic rats and HC rats was similar, with well‐defined boundaries, evidencing cell regeneration. These histopathological findings and biochemical analyses demonstrate the pancreas‐protective efficacy of CAH.

FIGURE 7.

FIGURE 7

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Effect of CAH on the pancreatic structure of diabetic rats (DR). AC, acinar cells; IC, islet cells. (A) healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.); (B) diabetic control (DC; DR + vehicle 1 mL/kg b.w.); (C) DR + metformin (500 mg/kg b.w.; MET); (D) DR + CAH (200 mg/kg b.w.; H200); (E) DR + CAH (400 mg/kg b.w.; H400).

Similar results have been reported for protein hydrolysates or peptides from other plant sources (Asokan et al. 2019; L. Li et al. 2022; W. Li et al. 2023; Olasehinde et al. 2023), which were able to reverse STZ cytotoxicity in pancreatic β cells.

3.8. CAH Treatment Downregulates Gluconeogenesis in Diabetic Rats

The abnormal increase in hepatic gluconeogenesis contributes to increased fasting hyperglycemia in patients with T2D (Hatting et al. 2018). The key gluconeogenic enzymes are PEPCK, FBPase, and G6Pase; therefore, their downregulation is important in managing diabetes. The activities of G6Pase (Figure 8A) and PEPCK (Figure 8B) were lower in the CAH‐treated (H200 and H400) and metformin‐treated groups than in the DC group, reaching values similar to those of the HC group. The similar enzymatic activities observed for both CAH doses suggest a saturation effect. These results demonstrate the potential of CAH to regulate gluconeogenesis. It has been proposed that metformin exerts its antidiabetic effect by reducing hepatic gluconeogenesis (Barroso et al. 2024), and this may also be the case for CAH. In this regard, Choi et al. (2017) evaluated the antidiabetic effects of protein hydrolysates (125, 250, and 500 mg/kg) from Semisulcospira libertina in mice with T2D adapted to an HFD. The treatments normalized the G6Pase and PEPCK activities. These results were considered direct evidence that the hydrolysate regulated hepatic glucose metabolism enzymes and promoted glucose homeostasis.

FIGURE 8.

FIGURE 8

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Effect of CAH on the activity of G6Pase (A), PEPCK (B), and G6PD (C) in the liver of diabetic rats (DR). Data are expressed as the mean ± SD (n = 6). Different letters indicate significant differences (Fisher's LSD test, p ≤ 0.05). Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

On the other hand, Olasehinde et al. (2023) reported that hydrolysates of Telfairia occidentalis seed protein (50, 100, and 150 mg/kg) reduced hepatic FBPase and G6Pase activities in a dose‐dependent manner in STZ‐induced diabetic rats, thereby suppressing gluconeogenesis.

3.9. CAH Treatment Upregulates the Pentose Phosphate Pathway in Diabetic Rats

T2D is associated with alterations in the pentose phosphate pathway, which produces NADPH and pentose phosphate (ribose 5‐phosphate), compounds that are important for cellular antioxidant defense and proliferation (Huang 2023). G6PD is the rate‐limiting enzyme in this pathway, and its activity was significantly lower in the livers of rats from the DC group than in those from the HC group (Figure 8C). In contrast, the groups treated with CAH (H200 and H400) and metformin showed a significant increase in G6PD activity compared to DC. These results corresponded to the GSH levels (Figure 6A): GSSG is reduced to GSH by NADPH‐dependent glutathione reductase, and the reaction catalyzed by G6PD produces NADPH (Ge et al. 2020).

Several authors have evaluated G6PD activity and its role in T2D. Ramkumar et al. (2011) and Vidhya and Udayakumar (2016) demonstrated that treatment with alcoholic and aqueous plant extracts of Gymnema montanum and Aerva lanata increased G6PD activity in diabetic mice. Regarding hydrolysates, Olasehinde et al. (2023) analyzed the potential of a T. occidentalis seed hydrolysate to regulate G6PD activity in STZ‐induced diabetic rats. The authors demonstrated that the diabetic control group exhibited a significant reduction in liver G6PD activity, whereas administration of different doses of hydrolysate significantly increased the enzyme activity. Thus, the hydrolysates may induce hepatic G6PD activation, promote cell proliferation and survival, and restore hepatic glucose oxidation. Activation of the pentose phosphate pathway plays an important role in regulating glucose levels in patients with T2D.

3.10. CAH Treatment Regulates the PI3K/AKT and AMPK Signaling Pathways

An exploratory western blot analysis showed that liver and muscle tissues from all experimental groups exhibited a protein band corresponding to AKT, but phosphorylated protein (p‐AKT) was observed only in the muscles of healthy animals and those treated with metformin and CAH (H400) (Figure 9). The lack of detection of p‐AKT in the liver may be due to the low sensitivity of the immunoblot analysis and the possible attenuation of AKT phosphorylation in the samples from overnight‐fasted rats. The AS160 protein was also observed in both tissues, but its phosphorylated form (p‐AS160) was detected only in the livers of the HC, MET, H200, and H400 groups. These apparent tissue‐specific discrepancies may suggest that liver kinases other than p‐AKT target AS160 and that something impairs AS160 phosphorylation by p‐AKT in the muscle. However, there is no evidence that supports these possibilities, and the low‐sensitivity immunoblot results cannot rule out a role for AKT in AS160 phosphorylation in both liver and muscle.

FIGURE 9.

FIGURE 9

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Effect of CAH administration on the PI3K/AKT and AMPK signaling pathways in the liver (A) and muscle (B) of rats with induced diabetes. Treatments: healthy control (HC; healthy rats + vehicle 1 mL/kg b.w.), diabetic control (DC; DR + vehicle 1 mL/kg b.w.), DR + metformin (500 mg/kg b.w.; MET), DR + CAH (200 mg/kg b.w.; H200), and DR + CAH (400 mg/kg b.w.; H400).

The AMPK protein was present in both tissues across all groups, but its phosphorylated form (p‐AMPK) was not observed in the DC group (Figure 9), consistent with the lower activity of gluconeogenic enzymes (G6Pase and PEPCK) in the HC and treated groups with respect to the DC group (Figure 8). Similar results were reported in STZ‐induced diabetic mice treated with a Ficus carica leaf (FCL) extract (Y. Zhang et al. 2019). The downregulation of AMPK in diabetic animals has been associated with high‐glucose repression via the E3 ubiquitin ligase MG53 (Jiang et al. 2021). The qualitative differences observed in the phosphorylation of AMPK (in muscle and liver) and AKT (in muscle) suggest that these signaling pathways are involved in the metabolic effects of CAH. Nevertheless, quantitative studies are required to validate these findings.

Previous studies have shown that common bean (Phaseolus vulgaris L.) peptides promote insulin secretion and glucose uptake by modulating PI3K/AKT signaling (Oseguera Toledo et al. 2016). Furthermore, pea protein hydrolysates regulate blood glucose by improving hepatic insulin sensitivity (activation of IRS, AKT, and Foxo1) in mice with diabetes induced by STZ and HFD (Liao et al. 2023).

On the other hand, AMPK is activated when the cellular energy is low, that is, when the AMP/ATP and ADP/ATP ratios increase (Steinberg and Hardie 2023). Active AMPK initiates a signaling cascade that includes phosphorylation of the target of rapamycin complex 2 (TORC2), inhibiting its nuclear translocation. These changes modulate the cAMP response element‐binding protein (CREB), a transcription factor, and the CREB‐dependent transcription of peroxisome proliferator‐activated receptor gamma coactivator 1α (PGC1α). This process reduces gluconeogenesis by decreasing the expression of PEPCK and G6Pase (W. Wang et al. 2023), an effect observed in the present study.

Peptides from different food sources promote AMPK activation in adipocytes (J. Wang et al. 2020). Wheat bran peptides (LRP and LQP) modulated hepatic AMPK activation in mice fed an HFD (Kawaguchi et al. 2017). Soy glycinin peptides (IAVPGEVA, IAVPTGVA, and LPYP) also regulate glucose metabolism and increase glucose uptake by activating the AKT and AMPK pathways in HepG2 cells (Lammi et al. 2015).

4. Conclusion

In silico analyses showed that the ten most abundant peptides in CAH inhibit DPPIV and α‐glucosidase. These results were consistent with the antidiabetic activity of CAH in rats with HFD–STZ‐induced diabetes, as evidenced by reductions in hyperglycemia, hyperlipidemia, and organ damage (kidney, liver, and pancreas). The bioactive peptides derived from chickpea could be useful for glycemic control, supporting their potential application in functional foods or nutraceuticals. Future studies quantifying changes in metabolites and components of the PI3K/AKT and AMPK pathways will provide further insights into the mechanisms underlying the antidiabetic effects of CAH.

Author Contributions

Alicia Navarro‐Leyva: investigation, validation, writing – original draft. Gabriela López‐Angulo: conceptualization, investigation, writing – review and editing, resources. Francisco Delgado‐Vargas: resources, writing – review and editing. Aimée Bastidas‐Ponce: methodology, writing – review and editing. Nancy Yareli Salazar‐Salas: methodology. Jenifer Mariana Soto‐Lozoya: methodology. José Ángel López‐Valenzuela: conceptualization, supervision, resources, writing – review and editing.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supplementary Figure 1. Molecular docking and interactions of PPK and QPL peptides with α‐glucosidase and DPPIV.

JFDS-91-0-s002.pdf (363.7KB, pdf)

Supplementary Table 1. Peptides identified by UPLC‐MS/MS in CAH.

JFDS-91-0-s001.pdf (504.7KB, pdf)

Supplementary Table 2. In silico ADMET predictions of chickpea peptides obtained after simulated gastrointestinal digestion.

JFDS-91-0-s004.pdf (131KB, pdf)

Supplementary Table 3. Molecular docking binding energies of digested peptides against α‐glucosidase and DPPIV.

JFDS-91-0-s003.pdf (85.1KB, pdf)

Acknowledgments

A.N.‐L. and J.M.S.L. acknowledge the scholarship received from SECIHTI‐México.

Contributor Information

Gabriela López‐Angulo, Email: gabylopez@uas.edu.mx.

José Ángel López‐Valenzuela, Email: jalopezvla@uas.edu.mx.

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Associated Data

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

Supplementary Materials

Supplementary Figure 1. Molecular docking and interactions of PPK and QPL peptides with α‐glucosidase and DPPIV.

JFDS-91-0-s002.pdf (363.7KB, pdf)

Supplementary Table 1. Peptides identified by UPLC‐MS/MS in CAH.

JFDS-91-0-s001.pdf (504.7KB, pdf)

Supplementary Table 2. In silico ADMET predictions of chickpea peptides obtained after simulated gastrointestinal digestion.

JFDS-91-0-s004.pdf (131KB, pdf)

Supplementary Table 3. Molecular docking binding energies of digested peptides against α‐glucosidase and DPPIV.

JFDS-91-0-s003.pdf (85.1KB, pdf)

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