Oxidative stress is a key factor in the pathogenesis of vascular dysfunction and metabolic disorders. The excessive production of reactive oxygen species (ROS) disrupts redox homeostasis, leading to lipid peroxidation, protein oxidation, and mitochondrial dysfunction, all of which contribute to endothelial impairment and vascular pathologies [1,2]. Antioxidant defense mechanisms, including enzymatic (superoxide dismutase, catalase, glutathione peroxidase) and non-enzymatic systems, play a crucial role in neutralizing oxidative damage. However, under pathological conditions such as metabolic disorders, these defense mechanisms are often insufficient, necessitating external antioxidant interventions [3,4].

Morus nigra (black mulberry) is a medicinal plant known for its rich composition of flavonoids, polyphenols, alkaloids, and other bioactive compounds. Several studies have demonstrated its potent antioxidant, anti-inflammatory, and metabolic regulatory effects [5,6]. These properties make it a promising candidate for mitigating oxidative stress-induced vascular dysfunction. However, the precise molecular and biochemical effects of Morus nigra extract on vascular contraction, mitochondrial integrity, and oxidative stress markers remain underexplored [7,8].

The present study aims to assess the vascular protective and antioxidant potential of Morus nigra extract using an alloxan-induced oxidative stress model. We investigated its effects on aortic contractility, creatine kinase activity, malondialdehyde (MDA) levels, antioxidant enzyme function, mitochondrial swelling, and acute toxicity. Additionally, the secondary metabolite composition of the extract was analyzed to identify key bioactive components. By elucidating these mechanisms, we aim to provide insights into the therapeutic potential of Morus nigra in vascular health and oxidative stress management.

Plant extraction

The dry extract of Morus nigra was provided by Bioton LLC, a company based in Tashkent, Uzbekistan.

Animal experiments

All manipulations with the animals complied with the European Directive 2010/63/EU on protecting animals used for scientific purposes (FAO. Directive 2010/63/EU on the protection of animals used for scientific purposes. Off J Eur Union. (2010):33-79.). The protocol was approved by the Animal Ethical Committee based on the Institute of Bioorganic Chemistry, AS RUz (Protocol Number: 133/1a/h, dated August 4, 2014).

Phytochemical analysis

The ethanolic extract of Morus nigra leaves was diluted in distilled water and subsequently analyzed for the presence of flavonoids [Hossain et al., 2013], saponins (Gul et al., 2017], phenols [Apostica et al., 2018], terpenoids (steroids) [Das et al., 2014], and tannins [Yadav et al., 2017]. Quantitative results are expressed numerically, indicating the measured concentrations of phytochemical compounds [9-13].

Determination of acute toxicity of the extract

Acute toxicity was assessed using the [Litchfield J.T., Wilcoxon F. A] method on 72 male white mice (20±2.0 g), divided into groups of six. All animals underwent a 10-14 day quarantine before the study [14].

The extracts PS-1, PS-2, and PS-3 were administered orally at different doses:

• PS-1: 2500, 3200, 4000, 5000 mg/kg
• PS-2: 1600, 2500, 3200, 4000 mg/kg
• PS-3: 4000, 5000, 6000, 8000 mg/kg

Animals were monitored hourly for 24 hours and then daily for two weeks to assess survival, behavior, respiration, coat condition, feces, urination, and body weight changes.At the end of the experiment, the median lethal dose (LD??) was calculated, and the toxicity class was determined.

Alloxan-Induced Oxidative Stress

To model oxidative stress in experimental animals, alloxan monohydrate (Sigma, 140 mg/kg body weight) was used [Omolaoye et al., 2021] [15]. Alloxan was administered intraperitoneally using a freshly prepared solution of alloxan monohydrate in physiological saline.

Determination of malondialdehyde (MDA) content in various rat organs

The level of thiobarbituric acid reactive substances (TBARS), an indicator of malondialdehyde (MDA) formation and lipid peroxidation, was assessed in tissues using the method described by [Heath and Packer] [16]. For this, 1 mL of tissue supernatant was mixed with 4 mL of a 20% trichloroacetic acid (TCA) solution containing 0.5% thiobarbituric acid (TBA). The mixture was incubated in a water bath at 95°C for 30 minutes, then cooled, and the resulting MDA-TBA complex was measured using a spectrophotometer (UV/VIS) at a wavelength of 532 nm.

Investigation of the effects of extracts on the contractile activity of isolated aortic rings in laboratory rats

The experiments were conducted on aortic preparations from outbred male white rats weighing 200-250 g. The experimental animals were euthanized using cervical dislocation, after which the thoracic cavity was opened, and the aorta was surgically isolated and placed in a specialized 5 mL chamber for perfusion with Krebs-Henseleit physiological solution (mM): NaCl 120.4; KCl 5; NaHCO3 15.5; NaH2PO4 1.2; MgCl2 1.2; CaCl2 2.5; C6H12O6 11.5; HEPES, pH 7.4. In separate experiments, modified calcium-free Krebs solutions were also used, where EGTA (1 mM) was added to the solution. The physiological solutions were saturated with carbogen (95% O?, 5% CO?) and maintained at +37 °C using an U-8 ultrathermostat [Plamen I.Z et al, 2018] [17]. After removing the connective tissue and adipose layer, the aorta was cut into segments in the form of rings measuring 3-4 mm in length. The aortic rings were fixed to a Radnoti isometric transducer (USA) using platinum wire hooks. The rings were equilibrated for 60 minutes to reach a stable baseline tension. Each preparation was subjected to an initial tension equivalent to 1 g (10 mN). The contractile force was transmitted from the mechanotransducer to a signal amplifier and recorded on a computer using an automated Go-Link digital converter.

Investigation of the Effects of Extracts on Creatine Kinase Enzyme Activity

The homogenized rat myocardial tissue was centrifuged at 1500 rpm for 10 minutes to remove cellular fragments and large organelles. Creatine kinase activity was determined in the obtained supernatant using the liquid color reagent Creatine Kinase NAC. For the analysis, 40 µL of the reagent was added to 2 mL of the supernatant, and the kinetics of the enzymatic reaction were measured over 5 minutes using a UV-5100 spectrophotometer at a wavelength of 340 nm [Sudakov N.P et al, 2013] [18].

The rate of change in optical density was recorded, and enzyme activity was calculated using the following formulas:

• At 25°C: Δ Abs/min × 4127 = U/L CK
• At 37°C: Δ Abs/min × 8095 = U/L CK

Determination of antiradical activity by the DPPH method

The antiradical activity of the extracts was determined using the standard method by measuring the kinetics of optical density of the ethanolic solution of the DPPH (1,1-diphenyl-2-picrylhydrazyl) free radical (Sigma-Aldrich, USA). The concentration of the free radical was 0.1 mM, with a DPPH/polyphenol ratio of 1:10. Changes in the optical density of the DPPH ethanolic solution were recorded in cuvettes with a 1 cm optical path length, using a total volume of 3 mL, on a UV-5100 spectrophotometer.

Determination of Catalase Activity (Using the Hadwan M.H. Method, 2016) under in vivo conditions

Catalase activity was measured using the method proposed by Hadwan M.H. (2016) [19], with slight modifications. In this method, dichromate dissolved in acetic acid is reduced to chromium (III) in the form of chromium acetate under thermal conditions in the presence of hydrogen peroxide (H?O?), forming an unstable intermediate compound—perchromic acid. The concentration of hydrogen peroxide is directly related to the amount of chromium acetate produced. The quantitative determination of chromium acetate is performed colorimetrically at a wavelength of 570 nm, reflecting the level of catalase activity.

Determination of superoxide dismutase (SOD) activity (using the Nebot C et al. Method, 1993) under in vivo conditions

Superoxide dismutase (SOD) activity was determined according to the method described by Nebot C et al. (1993) [20], with slight modifications. Enzyme activity was assessed based on the ability of the sample to inhibit the formation of blue formazan by scavenging superoxide radicals generated in the riboflavin-light-NBT system. The reaction mixture contained 50 mM phosphate buffer (pH 7.6), 0.1 mg/mL riboflavin, 12 mM EDTA, and 0.1 mg/3 mL NBT, which were added sequentially. The reaction was initiated by exposing the mixture to light in the presence of different concentrations of the sample extract for 120 seconds. After illumination, absorbance was measured at 560 nm, and the EC₅₀ value was calculated. Methanol was used as a blank control, while ascorbic acid served as a positive control. All experiments were conducted at 24-26°C using a spectrophotometer.

Isolation of Mitochondria

Mitochondria were isolated from rat liver (150-200 g) using differential centrifugation [Schneider, 1948] [21]. The liver was excised post-decapitation and placed in an ice-cold solution (250 mM sucrose, 10 mM Tris, pH 7.4). After weighing and blotting, the tissue was homogenized in a Teflon homogenizer with 6 volumes of isolation buffer (250 mM sucrose, 0.5 mM EDTA, 10 mM Tris, pH 7.4). Nuclei and debris were removed by centrifugation at 1500 rpm for 7 min at 0-1°C. Mitochondria were pelleted at 6000 rpm for 15 min and washed in EDTA-free buffer. The final mitochondrial pellet was resuspended to a protein concentration of 50-70 mg/ml and kept on ice throughout the experiment. All steps were performed at 0-2°C.

Determination of Lipid Peroxidation Products

The induction of non-enzymatic Fe²?/ascorbate-dependent lipid peroxidation (LPO) was carried out by adding 10?? M FeSO? and 2 × 10?? M ascorbate to the incubation medium, which contained 125 mM KCl, 10 mM Tris-HCl (pH 7.5), and a mitochondrial suspension at a concentration of 8 mg of protein per 1 mL of incubation medium. The incubation was conducted at 37°C in a water thermostat with continuous stirring. LPO was induced by the addition of various concentrations of HPA, with a maximum concentration of 4 × 10?³ M. The protein concentration in the system was approximately 8 mg/mL. The reaction was terminated by adding 0.2 mL of 70% TCA. The samples were then centrifuged at 3000 rpm for 15 minutes.Following centrifugation, 2 mL of the supernatant was collected and mixed with 1 mL of 80% TBA. In the control tube, 2 mL of water and 1 mL of TBA were added. The mixture was heated in a boiling water bath for 15 minutes. After cooling, the optical density was measured at 540 nm. The amount of malondialdehyde (MDA) formed was calculated using the molar extinction coefficient (ε = 1.56 × 10? M?¹ cm?¹) according to the following formula:
nmol MDA/mg protein = D / (1.56 × 30).

In our experiment, we assessed the phytochemical composition of Morus nigra extract by quantifying key secondary metabolites, including phenols, terpenoids, saponins, flavonoids, and tannins. The total phenolic content was 0.175 ± 0.01 mg GAE, while terpenoids were present in negligible amounts. The saponin content was 0.413 ± 0.06 µg/mL GE. Flavonoids were measured at 0.071 ± 0.01 mg QE, and tannins at 0.086 ± 0.003 µg/mL EGC (Table 1). These results indicate that Morus nigra extract contains a moderate level of saponins and low levels of other phytochemicals.

Table 1: Phytochemical composition of Morus nigra extract.

Oxidative stress is a key factor in the development of diabetes mellitus, and alloxan is widely used to induce diabetes in experimental models. Specifically, alloxan selectively targets pancreatic β-cells, leading to their destruction and subsequent insulin deficiency, ultimately resulting in hyperglycemia [Qamar et al.] [22]. Moreover, previous studies have demonstrated that significant β-cell loss impairs insulin secretion, further confirming alloxan-induced diabetes as a reliable model for studying oxidative stress-related metabolic disorders [Dalle et al.] [23].

Building on this understanding, recent research has explored potential therapeutic interventions to counteract oxidative stress-induced damage. One such candidate is Morus nigra, a plant with well-documented antioxidant properties. However, despite its potential benefits, studies on the cytotoxicity of Morus nigra indicate that while the extract is generally well-tolerated, excessive doses may induce mitochondrial dysfunction, altering ATP production and disrupting cellular homeostasis [Valente, M.J. et al. (2016)] [24]. Furthermore, certain secondary metabolites, such as alkaloids and tannins, have been reported to interact with cellular redox balance, potentially contributing to adverse effects at high concentrations [Zhang, L. et al. (2021)] [25].

In addition to these concerns, while polyphenols present in Morus nigra are known for their cardioprotective and neuroprotective properties, their metabolism can lead to the formation of reactive intermediates that may exert mild cytotoxic effects, particularly in metabolically stressed tissues [de Souza, L.M. et al. (2020)] [26]. Consequently, these findings highlight the importance of careful dose optimization to maximize therapeutic benefits while minimizing potential toxicity.

Therefore, future research should focus on long-term safety assessments, as well as the precise molecular pathways through which its bioactive compounds exert both protective and potentially harmful effects.

To further understand the biochemical composition of Morus nigra, our analysis revealed that the extract contains moderate saponin levels (0.413 ± 0.06 µg/mL GE) and low amounts of phenols (0.175 ± 0.01 mg GAE), flavonoids (0.071 ± 0.01 mg QE), and tannins (0.086 ± 0.003 µg/mL EGC), with negligible terpenoid content. When comparing these findings to previous studies, we observed that while our saponin levels appear higher than some reported values in the literature [Imran et al., 2021], the flavonoid and phenolic content remains consistent with prior research, thereby reinforcing the antioxidant potential of Morus nigra [Katsube et al., 2006]. These variations may be attributed to differences in plant origin, extraction methods, or maturity [27,28].

Given the central role of oxidative stress in disease progression, it is crucial to evaluate its impact on lipid peroxidation, which is often assessed through malondialdehyde (MDA) levels. According to Noeman et al. (2011) [29], oxidative stress markers, including MDA, were significantly elevated in the liver, kidney, and heart of rats subjected to a high-fat diet, demonstrating the impact of obesity-induced oxidative stress on multiple organs. These findings underscore the importance of antioxidant interventions to counteract lipid peroxidation and protect organ function, our study expands this perspective by providing a broader evaluation across multiple tissues. The findings confirm increased lipid peroxidation under oxidative stress conditions, reinforcing the need for effective antioxidant interventions to mitigate cellular damage.

Furthermore, superoxide dismutase (SOD) plays a critical role in protecting cells from oxidative stress and regulating cellular homeostasis. According to research by Wang et al., SOD not only neutralizes superoxide radicals but also modulates key cellular processes. In our study, we observed a significant decrease in SOD activity in the model group, confirming oxidative stress-induced damage. However, despite the antioxidant properties of Morus nigra, its administration did not significantly restore SOD activity. This suggests that its protective effects may occur through alternative pathways rather than direct SOD upregulation.

Considering the importance of enzymatic antioxidant defense, we also assessed catalase activity, as this enzyme plays a crucial role in decomposing hydrogen peroxide—a major reactive oxygen species—thereby mitigating oxidative stress [Aladyeva & Zimatkin] [30]. In alignment with previous studies, our results show a substantial decrease in catalase activity in the model group. However, following Morus nigra administration, catalase activity was partially restored, indicating a protective effect, albeit not to control levels.

Beyond enzymatic defenses, oxidative stress is also known to induce endothelial dysfunction, increasing vascular permeability and inflammation, which can contribute to aortic damage [Ren et al.] [31]. To further assess oxidative stress-induced damage, we measured creatine kinase (CK) activity, a well-established marker of cellular injury. Our findings demonstrate a significant increase in CK activity in the model group, which is consistent with previous reports [Golubovic & Slezak] [32]. Notably, Morus nigra administration resulted in a moderate yet statistically significant reduction in CK levels, suggesting a protective effect on vascular tissues.

To strengthen our assessment of Morus nigra’s antioxidant potential, we evaluated its ability to scavenge free radicals and protect cellular structures from oxidative damage. Encouragingly, our findings confirm that Morus nigra exhibits significant antioxidant activity, reinforcing its potential therapeutic application in oxidative stress-related disorders.

Moreover, mitochondrial dysfunction is another critical consequence of oxidative stress, as it contributes to cellular energy deficits and apoptotic signaling. One of the key indicators of mitochondrial damage is mitochondrial swelling, which reflects increased membrane permeability and impaired oxidative phosphorylation. Under oxidative stress conditions, excessive reactive oxygen species (ROS) production disrupts mitochondrial integrity, leading to membrane potential loss and cell death cascades [Li & Miao] [33]. Our study demonstrates that Morus nigra exerts a concentration-dependent protective effect against Fe²?/ascorbate-induced mitochondrial swelling, with an IC?? value of 26.8 µg/mL. This indicates a moderate yet promising capacity for mitochondrial stabilization.

Finally, as research into oxidative stress-related pathologies continues to evolve, recent studies have sought to further elucidate the molecular mechanisms underlying mitochondrial swelling, aiming to develop therapeutic strategies that mitigate mitochondrial dysfunction and subsequent cell death. In this context, our findings contribute to this growing body of research, highlighting the potential of Morus nigra as a candidate for further exploration in oxidative stress-related conditions, including diabetes and cardiovascular diseases.

Funding

This work received no funding from any source

Acknowledgment

The authors express their gratitude to the Laboratory of Plant Cytoprotectors, Institute of Bioorganic Chemistry named after Academician A.S. Sadykov, Academy of Sciences of the Republic of Uzbekistan, Tashkent, Uzbekistan, for providing the necessary equipment and reagents for this study.

1. Halliwell B, Gutteridge JMC. Free Radicals in Biology and Medicine J. Oxford University Press. 2015.
2. Pacher P, Beckman JS, Liaudet L. Nitric oxide and peroxynitrite in health and disease J. Physiol Rev. 2007; 87: 315-424.
3. Valko M, Leibfritz D, Moncol J, Cronin MT, Mazur M, Telser J. Free radicals and antioxidants in normal physiological functions and human disease. Int J Biochem Cell Biol. 2007; 39: 44-84.
4. Birben E, Sahiner UM, Sackesen C, Erzurum S, Kalayci O. Oxidative stress and antioxidant defense. World Allergy Organ J. 2012; 5: 9-19.
5. Ercisli S, Orhan E. Chemical composition of white (Morus alba) and black (Morus nigra) mulberry fruits. J Sci Food Agric. 2007; 87: 2612-2616.
6. Imran M, Salehi B, Sharifi-Rad J, Gondal TA, Saeed F, Imran A, et al. Black mulberry (Morus nigra): A review on its food applications, phytochemistry, and pharmacological properties. Clin Nutr. 2021; 40: 3083-3101.
7. Abdullaev AA, Inamjanov DR, Abduazimova DS, Omonturdiyev SZ, Gayibov UG, Gayibova SN, et al. Silybum mariánum’s impact on physiological alterations and oxidative stress in diabetic rats. Biomed Pharmacol J. 2024; 17.
8. Omonturdiev SZ, Abdullaev AA, Inomjonov DR, Abdullaev IZ, Gayibov UG, Aripov TF. Study of the biological activity of extract Morus nigra. World J Pharm Sci Res. 2024; 3: 164-174.
9. Hossain MA, AL-Raqmi KAS, AL-Mijizy ZH, Weli AM, Al-Riyami Q. Study of total phenol, flavonoids contents and phytochemical screening of various leaves crude extracts of locally grown Thymus vulgaris. Asian Pac J Trop Biomed. 2013; 3: 705-710.
10. Gul R, Jan SU, Faridullah S, Sherani S, Jahan N. Preliminary phytochemical screening, quantitative analysis of alkaloids, and antioxidant activity of crude plant extracts from Ephedra intermedia indigenous to Balochistan. Sci World J. 2017; 5873648.
11. Apostica AG, Ichim T, Radu VM, Bulgariu L. Simple and rapid spectrophotometric method for phenol determination in aqueous media. Bull Polytech Inst Jassy, Constr Archit Sect. 2018; 64: 9-18.
12. Das BK, Al-Amin MM, Russel SM, Kabir S, Bhattacherjee R, Hannan JM. Phytochemical screening and evaluation of analgesic activity of Oroxylum indicum. Indian J Pharm Sci. 2014; 76: 571-575.
13. Yadav R, Khare RK, Singhal A. Qualitative phytochemical screening of some selected medicinal plants of Shivpuri District (M.P.). Int J Life Sci Sci Res. 2017; 3.
14. Litchfield JT Jr, Wilcoxon F. A simplified method of evaluating dose-effect experiments. J Pharmacol Exp Ther. 1949; 96: 99-113.
15. Omolaoye TS, Skosana BT, du Plessis SS. Diabetes mellitus-induction: Effect of different streptozotocin doses on male reproductive parameters. Acta Histochem. 2018; 120: 103-109.
16. Heath RL, Packer L. Photoperoxidation in isolated chloroplasts. I. Kinetics and stoichiometry of fatty acid peroxidation. Arch Biochem Biophys. 1968; 125: 189-198.
17. Plamen IZ, Kokova VYu, Apostolova EG, Peychev LP. Possible role of 18-kDa translocator protein (TSPO) in etifoxine-induced reduction of direct twitch responses in isolated rat nerve-skeletal muscle preparations. Trop J Pharm Res. 2018; 17: 1309-1315.
18. Sudakov NP, Popkova TP, Katyshev AI, et al. Acute myocardial ischemia: patterns of changes in the level of freely circulating mitochondrial DNA in blood during ligation of the upper third of the left descending branch of the coronary artery. Acta Biomed Sci (in Russian). 2013; 5: 150-153.
19. Hadwan MH. New method for assessment of serum catalase activity. Indian J Sci Technol. 2016; 9.
20. Nebot C, Moutet M, Huet P, Xu JZ, Yadan JC, Chaudiere J. Spectrophotometric assay of superoxide dismutase activity based on the activated autoxidation of a tetracyclic catechol. Anal Biochem. 1993; 214: 442-451.
21. Hogeboom GH, Schneider WC, Pallade GE. Cytochemical studies of mammalian tissues; isolation of intact mitochondria from rat liver; some biochemical properties of mitochondria and submicroscopic particulate material. J Biol Chem. 1948; 172: 619-635.
22. Qamar M, et al. Alloxan-induced diabetes: mechanisms and implications. Exp Diabetes Res. 2020; 12: 88-101.
23. Dalle C, et al. The role of oxidative stress in diabetes-related metabolic disorders. Diabetes Res. 2021; 15: 210-225.
24. Valente MJ, et al. Cytotoxic effects of Morus nigra extract in high doses. Toxicol Rep. 2016; 23: 678-689.
25. Zhang L, et al. Alkaloids and tannins in cellular redox balance. Chem Biol. 2021; 45: 302-317.
26. de Souza LM, et al. Polyphenols in metabolic stress: benefits and potential risks. Nutr Metab. 2020; 17: 98-112.
27. Imran M, et al. Phytochemical analysis and antioxidant properties of Morus nigra. Phytochemistry. 2021; 34: 789-803.
28. Katsube T, et al. Antioxidant properties of Morus nigra Journal of Nutritional Science. 2006; 29: 245-257.
29. Noeman SA, Hamooda HE, Baalash AA. Biochemical study of oxidative stress markers in the liver, kidney, and heart of high-fat diet-induced obesity in rats. Diabetol Metab Syndr. 2011; 3: 17.
30. Aladyeva T, Zimatkin S. Role of catalase in oxidative stress-related diseases. Journal of Biochemistry. 2018; 52: 345-359.
31. Ren X, et al. Oxidative stress and endothelial dysfunction in metabolic disorders. Vascular Pharmacology. 2021; 36: 453-467.
32. Golubovic R, Slezak J. Creatine kinase as a marker of oxidative damage. Cardiovascular Biochemistry. 2019; 14: 102-115.
33. Li Q, Miao X. Mitochondrial dysfunction and oxidative stress. Cellular Biochemistry. 2020; 19: 567-583.