1 Introduction

The world is currently dealing with various medical conditions such as diabetes, hypertension, cancer, stroke, etc. which seem to get more potent and advanced; such advancements in diseases also require a more scientific response. Diabetes mellitus is a group of metabolic disorders that are due to a deficiency of insulin in the body or nonreponsiveness of cells to insulin. These disorders have been classified into, three main types, namely; e type one diabetes (T1D), type two diabetes (T2D), and gestational diabetes [1]. Diabetes Mellitus is one of the most prevalent diseases in the world currently. In the year 2020, an estimated 620 million individuals globally were affected by type 2 diabetes, and in 2017, diabetes was attributed to 1 million deaths giving it the ninth position as the leading cause of death, with the rapid increase in development and urbanization. The number of individuals suffering from diabetes is expected to increase greatly [2]. Complications attributed to diabetes include kidney disease, peripheral neuropathy, retinopathy, coronary heart disease, stroke, peripheral arterial disease, and microvascular conditions while heart failure continues to produce a high mortality rate in both types of diabetes [3].

The onset of T1D is due to the loss of insulin-producing β-pancreatic cells of the islets of Langerhans found in the pancreas, and T2D is caused by the resistance to the action of the produced insulin which is said to be due to a misfunctioning insulin receptor [4]. The exact mechanism of action behind the loss of insulin production and insulin resistance is still unclear, leading to different medications being used to treat DM. T1D is treated with periodical insulin shots [5], and the first treatment of T2D is through monotherapy using metformin or through the combination of metformin and other such as treated by using a single medication such as metformin or a combination of metformin and other medications such as sulfonylureas, thiazolidinedione and DPP-4 inhibitors of DPP-4, GLP-1 and sodium-glucose cotransporter 2 [6]. The combination of various medications often leads to more complications later in life and this directs the focus to be on more natural methods including medicinal plants. Most of the approved synthetic medicines used for the treatment of diabetes, present many adverse effects including pancreatitis, genital mycosis, anorectic effect, and hepatotoxicity to mention a few [7]. Moreover, some therapeutic drugs for diabetes treatment cannot be used during pregnancy [8]. Therefore, the development of effective and non-toxic anti-diabetic drugs is necessary for the treatment of diabetes.

Recently, studies on the use of medicinal plants to combat diabetes have increased tremendously. This has been attributed to a myriad of bioactive compounds present in their extracts with potential pharmacological properties. One of the most popular medicinal plants in South Africa is the H. hemerocallidea (HH) popularly known as the African potato, and its initial discovery was in 1979 by the chairman of the company Essential Sterolin Products when he believed that the plant had anticancer properties. At that moment the plant was known to contain β-sitosterol and its glucosidase among various phytochemicals [9]. The phytosterol glycosides found in HH are hypoxoside, β-sitosterol glucoside, campesterol, stigmasterol [10] rooperol [11]. The phytosterols such as sterolins found in HH have shown antinociceptive, anti-inflammatory activity, apoptosis induction of cancer cells, cholesterol management, anti-mutation, treatment of prostate complications, antioxidant, and anti-diabetic activities [12]. Moreover, HH has been reported to induce antidiabetic properties. In a previous study, Ojewole, revealed antidiabetic properties of the rhizome ethanolic extract corm’s aqueous extract (APE) in mice and rats and the extract significantly reduced hypoglycemia in normal and diabetic rats [13]. Another study showed that HH and β-sitosterol reduced early stages of type II spontaneous diabetes in mutant mice models [14]. Some mechanism of action has been suggested, HH was shown to increase insulin release from pancreatic β-cells. With these observations, it hypothesized that nanoparticles synthesized from HH can attenuate the enzymes involved in the progression of diabetes.

In the last decades, the incorporation of medicinal plant extracts with nanoparticles has allowed for a functional programmed treatment strategy in the field of nanomedicine. Nanoparticles are an interesting nanotechnology division with vast applications such as agriculture, cell biology, chemical sensing, drug delivery, nanomedicine, textiles, antioxidants, photocatalytic organic dye, and data storage [15]. Over the years, various types of nanoparticles such as ZnO NPs, FeNPs, MnO2 NPs, AgNPs, Cu-Ni hybrid NPs, etc., have been synthesized using the green method [16,17,18,19,20]. Currently, the utilization of trace elements found inside the human body e.g., copper, zinc, magnesium, selenium, iron, etc. in the synthesis of nanoparticles are under investigation in the field of nanomedicine [21]. These nanomaterials are considered alternative candidates due to their unique plasmonic and optoelectronic properties. In particular, magnesium nanoparticles have attracted interest in diverse applications such as catalysis, adsorption, drug delivery, antimicrobial, bone implantation, and diabetes treatment [22,23,24]. As a non-metal, magnesium is an essential nutrient that is required for the optimal function of the brain and body. Major biological functions like blood clotting, heartbeat, and muscular contraction are all controlled by magnesium [25]. Previous studies have highlighted the relationship between magnesium and diabetes as patients with type 2 diabetes often have lower levels of magnesium in the blood. Magnesium regulates insulin function, especially the post-receptor actions of insulin in connection to insulin-mediated glucose absorption, as well as several enzymatic processes in glycolysis [26]. Since magnesium has been proven to be involved in processing glucose and insulin, the use of magnesium supplements was found to help mitigate glycaemic response among type 2 diabetic patients [25]. Therefore, the utilization of a magnesium source could be beneficial in preventing and reversing type 2 diabetes.

Recently, some studies have shown that magnesium oxide could result in lower absorption rates when compared to other forms of magnesium (citrate, chloride, lactate, etc.) in the body. For instance, poorly controlled diabetic patients showed enhanced glycaemic control with higher magnesium oxide dosage (1000 mg per day) after 30 days whereas patients showed improved glycaemic control with lower magnesium chloride dosage (300 mg per day) after 16 weeks in clinical trials [27, 28]. In contrast, there was no improvement in glycaemic control when patients were given magnesium aspartate (369 mg per day) for 3 months [29]. These studies demonstrated the importance of absorption rate in the effectiveness of the form of magnesium. Besides, magnesium is non-toxic, inexpensive, readily available, and has a strong affinity for oxygen, thus making the formation of magnesium oxide (a common form of magnesium in human beings) easily achievable.

The utilization of complex matrices from medicinal plant extracts in the synthesis of MgO nanoparticles has many benefits. These include improved biocompatibility, easy access to the starting material from non-petroleum sources, and frequently lower costs due to the reuse of waste material [30, 31]. Besides, some phytochemicals present in medicinal plants can persist during the synthesis process, giving the nanoparticles functionality in addition to that offered by the metal itself, thereby enhancing the biological properties It has been demonstrated that numerous secondary metabolites from plant extracts, including carbohydrates and flavonoids, can reduce M+ ion to M0 atom [32]. Consequently, the utilization of medicinal plant extract from HH in the synthesis of magnesium oxide nanoparticles could accelerate the biochemical reaction and enhance its absorption rates thereby resulting in a higher ion bioavailability and improving treatment through the synergistic effect. Therefore, this present study aims to synthesize and stabilize magnesium oxide nanoparticles from medicinal plant extract and investigate its antioxidant, cytotoxic, and antidiabetic activities.

2 Materials and methods

2.1 Chemicals and reagents

All chemicals and reagents used for the synthesis of nanoparticles were purchased from Merck (Johannesburg, South Africa) and used as received. Human Embryonic Kidney (HEK293) and Human liver (HepG2) cells. were purchased from Cellonex Separation Scientific SA (Pty) Ltd. (Johannesburg, South Africa) while Dulbecco’s modified eagle’s medium (DMEM), fetal bovine serum (FBS), and penicillin–streptomycin, were purchased from Celtic Molecular Diagnostics SA (Pty) Ltd. (Cape Town, South Africa). Dimethyl sulfoxide (DMSO), 2,2-diphenylpicrylhydrazyl (DPPH), and (2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid)) were purchased from Sigma-Aldrich® (Darmstadt, Germany).

2.2 Plant collection and extraction

Hypoxis hemerocallidea was collected from the University of South Africa, Florida Campus, Gauteng Province, South Africa. The plant materials and a voucher specimen of the plants of air-dried leaves were deposited and identified at the CERES Herbarium, University South Africa. The use of plants in the present study complies with international, national and/or institutional guidelines. To prepare the H. hemerocallidea extract, 3 g of the leaf powder was added to 100 mL of distilled water and the solution was heated at 60 oC for 30 min. The mixture was filtered, and the solution extract was stored at 4 °C until use.

2.3 Synthesis of MgO using H. hemerocallidea

For the synthesis of MgO nanoparticles, magnesium chloride salt was used as the primary source of magnesium. Briefly, 25 mL of aqueous leaf extract of H. hemerocallidea was added to a 3 g/100 mL of MgCl2·6H2O solution in a flask and the reaction was stirred at 60 °C for 1 h. The reaction pH changed from 5.58 before stirring to 4.86 after completion, indicating the formation of the nanoparticles. Following this, the obtained solution was centrifuged at 4,400 rpm for 30 min to remove unreacted precursors and extracts. The resulting pellets were washed several times with distilled water and t pellets were dried in an oven at 50 °C overnight. Afterwards, the obtained powder was calcined and dried in a ceramic crucible at 200 °C for 2 h. After allowing it to cool, the black-colored pellet obtained was grounded and used for analysis.

2.4 Materials characterization

The functional groups present in the synthesized samples and plant extract were studied using a PerkinElmer Frontier FTIR fitted with an ATR detector in the range 4000–500 cm−1. The morphology of the samples was investigated by electron microscope imaging using a transmission electron microscope (JEOL JEM 2100) running on a 200-kV voltage and equipped with an energy-dispersive X-ray (EDX) analyzer. Dynamic Light Scattering (DLS) technique on a Malvern Zetasizer (Nano-ZS, Malvern, UK) was used to assess the surface charge and stability of the nanoparticles.

2.5 Antioxidant activity

2.5.1 DPPH radical scavenging activity

The 2,2-Diphenyl-1-picrylhydrazyl (DPPH) radicals scavenging activity of NPs was performed according to [33]. Briefly, NPs at various concentrations were added to 0.1 mM DPPH methanol solution and incubated for 30 min. The absorbance was measured at 517 nm. The concentrations tested of NPs and ascorbic acid ranged from (5.0–1000 μg/mL). The antioxidant activity was evaluated using the values IC50, i.e., the concentration of the studied sample, which causes a 50% inhibition. Ascorbic acid and methanol were used as a positive control and blank, respectively.

2.5.2 ABTS radical scavenging activity

The 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) radicals scavenging activity of NPs was done following the method by [34]. The ABTS+ cation radical was produced by mixing 7 mM of ABTS powder with 2.45 mM of potassium persulfate (K2S2O8) in distilled water and kept in the dark for 16 h. The quantification of the cation mixture was adjusted with methanol to an absorbance of 0.70 (± 0.02) at 734 nm. The ABTS+ scavenging activity assay was determined as the DPPH scavenging activity assay (2.5.1). The absorbance was measured on a microplate reader (Varioskan-Flash®, Thermo Fisher Scientific, Vantaa, Finland) at a wavelength of 734 nm. Ascorbic acid was used as the positive control at the same concentrations as the NPs. The ABTS + scavenging activity percentage was calculated using the formula shown in Sect. 2.5.1.

2.6 Cell culture and cytotoxicity activity

Human embryonic kidney (HEK 293) and human liver (HepG2) cells were maintained in T75 filter screw caps (TPP, Trasadingen, Switzerland) culture flasks in Dulbecco’s Minimal Essential Medium (DMEM, Gibco) supplemented with recovery media containing 20% fetal bovine serum (FBS) and 1% penicillin/streptomycin solution. Cells were incubated at 37 °C in a humidified atmosphere and 5% CO2. When cells reached 80–90% monolayer confluency they were detached from the flasks using trypsin (Trypsin–EDTA, Sigma–Aldrich) and resuspended in complete medium in a 1:3 split ratio to perform the sub-culturing. The sub-culturing procedure was repeated depending on the confluency of cells.

The cytotoxic activity of the NPs was determined using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay following a method described [35]. The HEK293 and HepG2 cells were cultured in sterile Dulbecco’s Minimal Essential Medium (DMEM, Gibco) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin solution. 100 μL of cells (1 × 104 cells/well) were added into 96 well microplates and incubated for 24 h at 37 °C in 5% CO2. Cells were then treated with varying concentrations of NPs ranging from (5.0–100 μg/mL). Doxorubicin and untreated cells were added as the positive and negative controls. After 24 h incubation, the MTT solution (20 μL) prepared in PBS (5 mg/mL) was added to all the wells, and the plates were incubated for 4 h, followed by the addition of 100 μL of DMSO to dissolve the formazan crystals for 1 h. The plates were read at 570 nm using an ELISA plate reader. The percentage of cell viability was calculated using the equation below

$$\mathrm{Cell \; Viability\; \%}= \frac{{\text{As}}}{{\text{Ac}}}\;{ \times }100$$

where As and Ac are absorbances of samples (treated cells) and control (untreated cells).

2.7 Inhibitory effects of α-amylase and α-glucosidase

2.7.1 α-amylase inhibitory activity

The α-amylase inhibitory activity was determined using a slightly modified method as described [36]. Briefly, a reaction mixture containing 50 μL of phosphate buffer, 10 μL of α-amylase (2 U/mL), and 20 μL of NPs was added to a 96-well plate and incubated for 20 min at 37 °C. Then, 1% soluble potato starch (100 mM phosphate buffer pH 6.8) was added as a substrate and incubated at 37 °C for 30 min. The concentration of the NPs and the positive drug control acarbose ranged from 0.125 to 2 mg/mL. A negative control was done using the phosphate buffer (100 mM, pH = 6.8). After incubation, the DNS (100 μL) was added and boiled for 10 min. The absorbance was quantified using an Elisa microplate reader at 540 nm (Varioskan Flash Spectrophotometer). The results were expressed as percentage inhibition, which was calculated using the formula below.

$$\mathrm{Inhibitory\; Activity\; \%}= 1-\frac{{\text{As}}}{{\text{Ac}}}\;{\times}100$$

where As is the absorbance in the presence of test substance and Ac is the absorbance of control.

2.7.2 α-glucosidase inhibitory activity

The intestinal α-glucosidase inhibitory activity of NPs was determined using a method reported [36]. The NPs were incubated at 37 °C for 15 min in a 96-well plate containing 50 μL of phosphate buffer, 10 μL α-glucosidase (1 U/mL), and 20 μL of different NPs concentrations. Acarbose was used as a reference standard (positive control). The substrate (5 mM P-NPG) was added to 20 μL of the mixture and incubated at 37 °C for 20 min. A 0.1 M Na2CO3 (50 μL) was added to stop the reaction. The NPs and acarbose concentrations varied from 0.125 to 2 mg/mL and 100 mM of phosphate buffer (pH = 6.8) was used as negative control. The quantification of p-nitrophenol was read at 405 nm on an Elisa microplate reader (Varioskan Flash Spectrophotometer). The results were expressed as percentage inhibition, which was calculated using the following formula in Sect. 2.7.1.

3 Statistical analysis

Absorbances were measured on a microplate reader (Varioskan-Flash®, Thermo Fisher Scientific, Vantaa, Finland). GraphPad Prism software 8.2 (GraphPad Software, CA, USA) software was used for data analysis. The comparison of means was done based on Turkey’s test and differences were considered statistically significant at p < 0.05.

4 Results and discussion

4.1 Synthesis of MgO nanoparticles

For the synthesis of MgO nanoparticles, the aqueous leaf extract of H. hemerocallidea was utilized as a cap**, stabilizing, and reducing agent. In the previous study, the preliminary phytochemical screening of H. hemerocallidea plant extract revealed the presence of bioactive substances such as polyphenols, proanthocyanidins, tannins, and flavonoids, supporting its application as an environmentally safe method of synthesizing metal nanoparticles [37] The bio-reduction of Mg ion in the presence of oxygen to form MgO using the aqueous extract was monitored by a change in pH of the solution. With an initial pH of 4.86, a final pH of 5.58 was observed after the completion of the reaction, indicating the formation of MgO nanoparticles. Figure 1a shows the transmission electron microscopy (TEM) image of the synthesized MgO nanoparticles. The image revealed the formation of two-dimensional (1D) nanosheet morphology with sharp edges and no sign of agglomeration. The sheet-like nanostructure of the smooth surface MgO (average diameter of 300 nm and length of about 1 µm) would result in a higher surface area with numerous binding sites for enhanced performance. Phytochemicals from plant extracts can adsorb onto the surface of nanoparticles. The strong chelating ability of these phytochemicals plays a significant role in accelerating electron transfer between the active metal center and substrate [38]. Similarly, the corresponding selected area electron diffraction (SAED) pattern displayed distinct bright spots in concentric close circles, thus demonstrating the polycrystalline nature of the MgO nanoparticles (Fig. 1b). Energy dispersive X-ray spectroscopy was used to identify the presence of the elements in the nanoparticles. The spectrum (Fig. 1c) showed the reduction peaks of both Mg and O while the Cu and C peaks could be attributed to the carbon-coated copper grid used for the analysis. MgO NPs that were synthesized from the extract were found to have a zeta potential value of − 28.11 mV (Fig. 1d). This shows that the nanoparticles’ surfaces are negatively charged and are evenly dispersed throughout the aqueous media. Besides, using the dynamic light scattering (DLS) method, the stability of the purified nanoparticles dispersed in water was evaluated forseven days (Inset). During this time, the zeta-potential values of the MgO NPs barely increased. The strong adsorption of extract-containing bioactive compounds on the nanoparticles may be attributed to the high negative values obtained. This adsorption generates an electrostatic repelling interaction between the particles, and therefore improves their stability and prevents aggregation. The observed negative charge and good stability are consistent with previous works on utilizing plant extracts and thus hold great potential in the synthesis and stabilization of many types of nanoparticles [35, 39, 40].

Fig. 1
figure 1

a TEM image, b corresponding SAED and c EDX spectrum, and, d zeta-potentials (inset: stability) of the synthesized MgO nanosheets

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Figure 2 shows the FT-IR spectra for the synthesized samples and the aqueous plant extract of H. hemerocallidea. The aqueous extract spectrum showed distinctive peaks at wavenumbers of 1047, 1250, 1405, 1597, and 3280 cm−1. These peaks correspond to the stretching vibration bands of –C–O–C (anhydride group), –C–N (stretching amine group), –S=O (sulfate group), –COO (carboxylic group), and -OH (alcohol group), respectively. Other weak peaks at 3672, 2985, 2900, and 1508 cm−1 were also found in the extract. The presence of nearly identical functional groups can be seen in the sample, which suggests that the MgO nanoparticles were coated with biomolecules derived from the aqueous HH plant extract. Besides, an additional conspicuous peak that can be attributed to Mg–O bending vibration emerged at 600 cm−1 in the FT-IR spectra of MgO nanoparticles [41]. Numerous biomolecules, including alkaloids, flavonoids, and others in the aqueous extract that can function as reducing and cap** agents, may have produced these functional groups [15, 37]. Similar results were obtained by Muhamad et al. using Monotheca buxifolia aqueous extract for the synthesis of Zinc oxide nanoparticles, FT-IR indicated peaks at 3408, 1640, 1394, and 1038 cm−1 which were credited to the same functional groups from the extract [16]. Also, the methanol extract of H. hemerocallidea was utilized for the synthesis of gold nanoparticles and provided similar peaks observed from the powdered extract [37] These similar peaks from aqueous and methanol indicate that the extract methods result in similar phytochemicals being extracted.

Fig. 2
figure 2

FT-IR spectra of dried H. hemerocallidea plant extract and biosynthesized MgO nanosheets

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4.2 Antioxidant activity

The hallmark of the DPPH methods employed to evaluate the antioxidant potential of the MgO NPs reveals the hydrogen-donating ability or the electron-donating ability of NPs. In addition, the ABTS method is more reflective of highly hydrophilic antioxidants than the DPPH method. Table 1 shows the estimation of the DPPH and ABTS radical reduction capacity of NPs with ascorbic acid as a positive control. The results demonstrated that there were no significant differences in the reduction of both radicals and IC50 values of 57.35 ± 0.28 and 52.08 ± 0.24 µg/mL were observed for DPPH and ABTS, respectively. While the reputable antioxidant, ascorbic acid exhibited potent radical reduction with IC50 values of (DPPH = 4.11 ± 0.82 µg/mL) and (ABTS = 3.92 ± 0.25 µg/mL). Therefore, results obtained in this study show that the NPs consist of both electron-donating and hydrophilic antioxidant properties. Previous studies have shown that phytosterols, glycosides, and hypoxides to mention a few, are responsible for plants’ free radical quenching activity [42].

Table 1 Inhibitory effects (IC50) of Hypoxis hemerocallidea MgO NPs on α-glucosidase, α-amylase, α glucosidase, DPPH, ABTS, HEK293 and HepG2 cells
Full size table

4.3 Cytotoxicity activity

The application of NPs for human benefits has raised concern regarding their possibility to induce toxicity to human cells and for this reason, we were encouraged to investigate whether HH MgO NPs are toxic to human embryonic kidney (HEK293) and human liver (HepG2) cell lines. The cytotoxicity effect of the NPs in HEK293 and HepG2 cells was determined using the MTT assay. The MTT assay evaluates the mitochondrial dehydrogenase activity of viable cells where the MTT dye is reduced from a yellow to formazan crystals, which were solubilized with DMSO. Then, the color change is photometrically quantified. The NPs tested exhibited a concentration-dependent toxicity behavior in both cell lines, where an increase in the concentration of NPs led to a decrease in the percentage of cell viability. (Fig. 3). As seen in Table 1, the results estimated by inhibitory concentration (IC50) showed IC50 values of 41.63 ± 0.73 µg/mL and 32.39 ± 0.95 µg/mL, which are regarded as non-toxic. The results were compared to a standard anticancer drug, doxorubicin which was more toxic to cells (IC50 = 2.70 ± 0.32 µg/mL). Although MgO NPs are used in human application, several studies have revealed their taxological potential The H. hemerocallidea gold nanoparticles (HH-AgNPs) was toxic to malignant glioblastoma tumour cell lines U87 and U251 cells with IC50’s of 0.81 and 4.0 µg/mL, respectively [43]. Assessment of the cytotoxic nature of MgO NPs evaluated against HepG2 cell lines, demonstrated that the NPs induce significant toxicity with an IC50 values of 4.7 µg/mL [44]. According to the latter study, MgO NPs are more toxic to cancerous cells compared to non-cancerous cells. A study conducted on the toxicity of 50% methanol and petroleum ether HH extract, indicated a higher LC50 value of 95.5 \(\pm\) 13.3 µg/mL and 210.91 ± 8 µg/mL, respectively [45], these results can confirm that unmodified HH extract is relatively non-toxic to cells supporting the ethnopharmacological use of the plant.

Fig. 3
figure 3

Cytotoxicity effects of HH MgO NPs on HEK293 (a) and HepG2 (b) cell lines after 24 h treatment. The data are represented as mean ± SD

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4.4 Inhibitory effects of α-amylase and α-glucosidase

The present study reveals the inhibitory activity of H. hemerocallidea biosynthesized MgO-NPs with IC50 values of 33.03 ± 1.43 and 52.38 ± 3.06 µg/mL against α-amylase and α-glucosidase enzymes (Table 1). These results were compared with a standard reference, acarbose which had the best IC50 value of 24.54 ± 1.55 and 6.54 ± 0.27 µg/mL, respectively. These results indicate that α-amylase was the most susceptible diabetic enzyme to the NPs and acarbose since the lowest inhibitory concentrations were obtained when tested against [46] showed that different solvent fractions of H. hemerocallidea exhibit potency against α-amylase and α-glucosidase with IC50 value < 10 mg/mL. Khan et al. 2021 explored the antidiabetic by evaluating the inhibitory activity of biosynthesized Hibiscus rosa sinensis MgO-NPs against α-amylase and α-glucosidase. The MgO-NPs showed 54.32 ± 2.0% and 53.27 ± 0.84% reduction with IC50 327 ± 0.82 µg/mL and 357 ± 0.82 µg/mL on α-amylase and α-glucosidase, respectively [40].

5 Conclusions

According to the findings, MgO nanoparticles were successfully synthesized from the aqueous extracts of the H. hemerocallidea medicinal plant. Various optical, functional, and morphological analyses were used to characterize the extract-mediated MgO nanoparticles. The nanoparticles obtained were sheet-like in morphology, highly crystalline, and had an average diameter of about 300 nm. The hallmark of treating diabetes is to inhibit the enzymes α-amylase and α-glucosidase that are involved in the progression of diabetes. The synthesized HH MgO-NPs exhibited significant α-amylase and α-glucosidase inhibitory activities. On the other hand, the nanoparticles revealed antioxidant efficacy which plays a crucial role in reducing diabetes related inflammatory diseases. Furthermore, an ideal therapeutic agent must have none/less toxicity on somatic cells, and results obtained in this study confirmed that NPs showed low toxicity against normal HEK293 cells. Nevertheless, further cytotoxicity tests against different cells need to be done to confirm the safety of the HH MgO-NPs. Prospectively, we envisage conducting molecular docking studies to fully understand how these HH MgO nanoparticles may be used to treat or manage diabetes and oxidative stress-related diseases. The use of MgO in the management of diabetes is constrained by the limited in vivo research done on the efficiency of the nanoparticles, the stability of nanoparticles is dependent on environmental conditions such as temperature and pH, and variations in these conditions ultimately lead to the change in the stability and shape of nanoparticles.