3.1 Characterization of green synthesized ZnO-Nps
3.1.1 SEM and TEM analysis
SEM measurement was carried out to analyze the morphology of ZnO-NPs. The SEM images (Fig. 2a) show that, the produced ZnO-NPs are nearly spherical in shape with agglomeration due to their water interaction and the intramolecular Van der Waals, magnetic and electrostatic interactions. The nanoscale and uniformity of ZnO-NPs are revealed by TEM images. The shape of the nanoparticles produced is of a spherical nature with an average size found to be 20 to 40 nm as shown in the (Fig. 2b). The SAED pattern in Fig. 2c indicates that the biosynthesized ZnO-NPs were polycrystalline in nature. The particle size distributions for the sample are shown in the (Fig. 2d). Low polydispersity index values indicate that the particles are homogeneous and have direct spherical forms (confirmed in the SEM analysis). Particle size distributions cover a small range of particle diameters and the smallest particles in the sample had diameters ranging from 1 to 550 nm, with an average size diameter of 122.6 nm [39].
3.1.2 FT-IR and XRD analysis
Figure 3a shows the FT-IR spectrum of ZnO-NPs. The presence of organic compounds in the sample is associated to absorption peaks in the region of 800–1600 cm− 1. The stretching vibration band of C = O of the amide group and the presence of carboxyl (COOH) and hydroxyl (OH) groups are shown by the absorption peaks at 1630.12 cm− 1 and 3437.05 cm− 1, respectively [40], while the -CH2- and C-C in the aromatic ring are responsible for the peaks at 2920.46 cm− 1 and 1411 cm− 1, respectively. Stretching vibrations of C = O can also be attributed to the absorption peaks at 1630 cm− 1. The presence of phenolic groups, alcohols, and aliphatic amines is also indicated by a peak at 1039.81cm− 1. The phenols in the capping agent bond to the surface of ZnO and cause ZnO NPs to form, while the C = O, C = O–C and C = C groups of heterocyclic compounds may act as a stabilizer [41, 42]. Then FT-IR spectrum absorbs the peak at 875.38 cm− 1 indicate the stretching vibrations of ZnO because the peaks in the range from 500–900 cm− 1 are attributed to metal Oxide bonds [43]. Using XRD analysis is useful tool to determine the phase and crystalline nature of ZnO nanoparticles. In the current study, XRD pattern of ZnO-NPs (Fig. 3b ) showed different peaks at (2ϴ) = 31.75 (1 0 0), 34.46 (0 0 2), 36.32 (1 0 1), 47.46 (1 0 2), 56.63 (1 1 0), 62.75 (1 0 3), 66.36 (2 0 0), 68.13 (1 1 2) and 69.10 (2 0 1) which agree with the JCPDS card (NO 36–1451) for a typical hexagonal phase of wurtzite form crystalline material [44, 45]. The presence of (1 0 0), (0 0 2), (1 0 1) planes in XRD designates the formation of hexagonal phase of wurtzite structure of ZnO-NPs. No impurity peaks were noted indicated the formation of pure ZnO-NPs. The prominent sharp peak at (101) represents the polycrystalline structure of ZnO-NPs. The average crystalline size was found to be 22 nm which was further confirmed by TEM analysis and SAED. These results are in accordance with previous studies on ZnO-NPs synthesized using leaf extract of Tecoma castanifolia [43] and Conyza canadensis [46].
3.2 Antimicrobial activity of ZnO-NPs
The antimicrobial effect of biosynthesized ZnO-NPs can be interpreted to different actions such as (1) generating reactive oxygen species (ROS), (2) ZnO-NPs and cell walls of microbes interaction that finally causes oxidative stress and death of cells, (3) electrostatic attraction the microbial cell and (4) entry of Zn2+ ions into the cell, [47, 48].
The ZnO-NPs nanofluid inhibit the growth of all tested pathogenic microorganisms, with antimicrobial activities by diameters ranged from 19.7 ± 0.29 to 24.6 ± 0.32 mm at concentration 1000 ug/mL as showed in Table 1 and Fig. 4, ZnO-NPs exhibit inhibition effect against Gram-negative bacteria more than Gram-positive bacteria and fungi, there is no effect for base fluid against any of the tested microorganisms, which proved that the antimicrobial activity of ZnO-NPs, different inhibition zone results for antibiotic controls were recorded which less than that obtained fromZnO-NPs.
The detect of MIC values of biomolecules against pathogenic microorganisms especially if these biomolecules are integrated into medical applications had a significant value. Effect of different concentration of ZnO-NPs (500, 250, 125, 62.5, 31.25, 15.6 and 7.8 µg/ml) against tested microbes showed that, the MIC value for Gram-positive bacteria (B. cereus and S. aureus) were 62.5 µg/ml; Gram-negative bacteria (E. coli and K. pneumonia) were 31.25 µg/ml, while MIC value for A. niger and A. flavus were 125 and 250 µg/ml respectively.
In the current study, Gram negative bacteria (E. coli) was the most sensitive microbes toward biosynthesized ZnO-NPs, and this action can be due to differences in cell wall structures between Gram negative and Gram-positive bacteria. The Gram-negative bacterial cell wall was distinguished by a thin layer of peptidoglycan and lipopolysaccharides (LPS). The positive charge of ZnO-NPs is hardly adhesive to the negative charge of LPS, and hence it is the adsorbed on the Gram-negative bacterial cell membrane that ultimately disrupts selective permeability [49]. Moreover, the inhibition effect of ZnO-NPs on fungi significantly deform conidial formation and conidiophores of Penicillium expansum and Botrytis cinerea, displaying antifungal properties [48, 50].
Table 1
Antimicrobial activity and MIC values of biosynthesized ZnO-NPs.
No. | | Mean Diameter of Inhibition Zone (mm) _ Std. Error | |
| Microorganisms | ZnO-NPs | Base fluid | Antibiotic control | MIC (ug/mL) |
1 | B. cereus | 19.7 ± 0.29 | 0.0 | 22.3 ± 0.34 | 62.5 |
2 | S. aureus | 17.9 ± 0.51 | 0.0 | 22.1 ± 0.45 | 62.5 |
3 | E. coli | 24.6 ± 0.32 | 0.0 | 17.2 ± 0.61 | 31.25 |
4 | K. pneumonia | 22.3 ± 0.25 | 0.0 | 18.7 ± 0.42 | 31.25 |
5 | A. niger | 23.3 ± 0.49 | 0.0 | 19.6 ± 0.35 | 125 |
6 | A. flavus | 21.7 ± 0.22 | 0.0 | 18.6 ± 0.57 | 250 |
3.3 Antioxidant activity of ZnO-NPs
Antioxidant activity has been found in a wide range of natural and artificial substances [51]. Antioxidant activity was assessed using DPPH radical-scavenging activity and ABTS radical-scavenging activity tests in the current study. Antioxidants are chemicals that neutralise ROS which are formed as a byproduct of biological events [52]. Several antioxidant qualities, including anti-atherosclerosis, anti-inflammatory, anti-tumor, anticancer, anti-mutagenesis, anti-carcinogenesis and anti-microbiosis, have led to their use as therapeutic agents (Fig. 5). In recent study, the antioxidant activity of ZnO-NPs was evaluated using DPPH and ABTS assays, as shown in Fig. 1. Results proved that the antioxidant activity of ZnO-NPs with IC50 = 240 µg/ml compared to 16.2 µg/ml for ascorbic acid in the case of DPPH, while IC50 of ZnO-NPs and ascorbic acid using ABTs assay were 250 and 1.6 µg/ml, respectively.
3.4 Evaluation of ZnO-NPs for seed germination
Short-term phytotoxicity of emerging contaminants, such as designed NPs, can be estimated using seed germination and root/shoot elongation experiments [53]. In this study, germination was defined as the radicle or plumule emerging from the seed coat. The germination of barley seeds exposed to ZnO-NPs was assessed after 4 days of treatment. ZnO-NPs treatment demonstrated a positive effect on barley seeds when compared to the control. The seed germination of barley was significantly increased at all ZnO-NPs doses examined. The seedlings germinated with the addition of ZnO-NPs at 2 mg/l had the highest germination rate (90%), while the control seeds had a considerably lower germination rate (63%). At 1, 4, 8, and 12 PPM of ZnO-NPs, the seed germination rate was record 83, 67, 77, and 83%, respectively (Fig. 6a). Germination of the seeds began after one day, with a proportion ranging from 60% under 1 and 2 PPM ZnO-NPs conditions to 50% under control. Zn is an important plant micronutrient that is frequently supplemented with zinc sulphate in agricultural practices to prevent Zn deficiency. Our findings resembled those of Plaksenkova et al. [54], who reported that low ZnO-NPs concentrations of 1,2, and 4 PPM significantly increased barley seed germination. In addition Upadhyaya, [55] recorded an increase in the germination of rice seeds treated with Zn NPs, which is consistent with our findings. In contrast to Xiang et al. [53], ZnO-NPs at concentrations of 1–80 mg/l had no influence on Chinese cabbage seed germination when compared to the control. Raliya et al. [56] also reported that ZnO-NPs had no influence on tomato seed germination at concentrations up to 750 mg/kg. According to Zhang et al [57], ZnO-NPs at concentrations of 10, 100, and 1,000 mg/l had no statistically significant effect on maize or cucumber germination, suggesting that germination rate is species- and concentration-dependent.
3.4 Assessment of ZnO-NPs for root and shoot length
In plants, Zn acts as a cofactor for RNA polymerases and other plant enzymes, influencing their activity. Phosphorus mobilisation enzymes such as phosphatase and phytases are stimulated by ZnO-NPs in the rhizosphere, increasing the amount of phosphorus available to plants [58, 59]. ZnO-NPs has a dual role as essential nutrients and native phosphorous mobilizers is supported by the improved physiological and biochemical responses. [60].
Both root and shoot length were influenced by ZnO-NPs as showed in (Fig. 6b). The difference in root elongation between control and all ZnO-NPs concentrations was substantial (Fig. 6b). At 2 PPM ZnO-NPs concentration, the maximum root elongation was 6 ± 1.2 cm, while the control plants' root length was 3.6 ± 1.2 cm. At a concentration of 12 PPM ZnO-NPs, the minimum root length was found to be 3.1 ± 1.3 cm. According to previous controlled studies, the toxicity of NPs during the early stages of plant growth is most likely due to the following factors: I chemical and physical properties that affect the release of ions or the aggregation of particles into more stable forms; and (ii) particle size and shape, which determine the specific surface area of NPs [61, 62]. A quick and commonly used acute phytotoxicity test method, seed germination and root elongation, has various advantages, including sensitivity, simplicity, cheap cost, and applicability for unstable substances or samples [63, 64].
The effects of suspended ZnO-NPs on young shoots are seen in (Fig. 6b). There was a positive reaction in shoots, with ZnO-NPs at all concentrations promoting significant increases in shoot length, which increased progressively as concentrations increased. At 12 PPM of ZnO-NPs concentration, the highest shoot elongation was 6.5 ± 1.2 cm, compared to 3.2 ± 0.8 cm for the control shoot. Most concentrations of ZnO-NPs (1,2,4 and 8 mg/l) in our investigation encouraged root and shoot growth in contrast to control, with the exception of 12 PPM, where root growth was similar to control. ZnO-NPs at a concentration of 10 mg/l greatly increased the root length of germinated maize, according to [58]. Our results were similar to those of Plaksenkova et al. [54], who found that low ZnO-NPs concentrations of 1, 2, and 4 PPM improved barley root and shoot lengths considerably.
Root and shoot length in tomato (Lycopersicon esculentum L.) plants treated with ZnO-NPs at doses of 2, 4, 8, or 16 mg/l were longer than in control plants, [61]. Furthermore, lower ZnO-NPs concentrations (10 and 20 mg/l) resulted in a significant increase in onion seedling shoot and root lengths, but higher concentrations (30 and 40 mg/l) resulted in a significant decrease in onion seedling shoot and root lengths [62]. In contrast to Munzuroglu and Geckil. [63], a wide range of ZnO-NPs concentrations, from low (50 mg/l) to very high (3200 mg/l), had no effect on the length of three-week-old onion plantlets grown in vitro.
On the other hand, Wang et al. [64] reported that, the ZnO-NPs suppressed garlic root growth as concentration increased. At a concentration of 50 mg/l, ZnO-NPs completely stopped root development. Low ZnO-NPs concentrations cause an increase in root and shoot length, which could be a nutritional benefit of nano zinc oxide. Zn content in seeds plays an important physiological role during seed germination and early seedling growth [65]. Zn also aids elongation and cell division by stabilising indole-3-acetic acid, the most common auxin in plants when collected in significant concentrations, however, it is hazardous due to its episodic binding to proteins and subsequent displacement of other metal ions, notably Fe [66, 67].
Finally, Variations in plant responsiveness to ZnO-NPs application may be attributed to genotype, plant part/organ (treatment of seeds, roots or leaves), experiment environment (in vitro or in vivo), NP features (size, shape and concentration), and/or exposure time [68].
3.5 Effect of ZnO-NPs on MI
In genotoxicity studies, cytogenetic analysis of root meristems with an optical microscope is a quick and effective way to determine the MI, chromosome breakages and anomalies, micronuclei [69], spindle failure, and polyploidy and aneuploidy occurrence [70, 71].The cytotoxicity and genotoxicity capacity of ZnO-NPs on barley seedlings was estimated using cytological indicators such as the MI, the number and percentage of chromosomal abnormalities. Tables 1 and 2 illustrate the cytological and chromosomal abnormalities seen in the root tip cells of barley treated with various doses of ZnO-NPs.
The control sample had a MI of 5.67 ± 0.04 and normal divisional phases, with the exception of a minor number of aberrant cells at prophase (3.53 ± 0.01). The percentage of dividing cells increased significantly until 8 mg/l ZnO-NPs was added, after which the value dropped significantly. The value of the MI (1.25 ± 0.07%) was lowered by nearly 4% at a dose concentration of 12 PPM compared to the control. This higher rise in MI at lower concentrations (1 and 2 PPM) was limited by short time exposure (3 and 6 h), but subsequent increases in nanoparticle dosage and time resulted in a significant decrease in MI until total suppression was achieved at 12 h with 4, 8, and 12 PPM concentrations. MI levels at all concentrations decreased with time as a result of long-term exposure as presented in Table 2 and Fig. 7. These results were constant with results reported by the Plaksenkova et al. [54], where the cytotoxicity generated by ZnO-NPs depend on their concentration and timeexposure. Also, there was a concentration-dependent decrease of MI in V. faba, indicating that ZnO-NPs have cytotoxic potential, according to [72].
According to our findings, exposing barley roots to ZnO-NPs produces cytotoxicity and genotoxicity. Table 2 summaries the mitotic parameters recorded before and after nanoparticle treatment. ZnO-NPs caused significant alterations in MI and a rise in chromosomal aberrations (Fig. 7) as a clear dose response impact, even when plants were exposed at low concentrations of them over a short period of time, according to the tests. CAs revealed that mitotic inhibition was linked to the gradual production of several chromosomal abnormalities.
The maximum value of MI (8.93 ± 0.06) was observed that the 3h for treatment with 1 PPM of ZnO-NPs, while the lowest (1.23 ± 0.01) was found at the 9 h for treatment with 12 PPM of ZnO-NPs. For a 12 h treatment at two doses, the drop in MI was statistically extremely significant, with a minimum value of 1.73 ± 0.06 and 1.25 ± 0.07% for (1 and 2 PPM ZnO-NPs, respectively) and complete inhibition for the other dosages. We may deduce that, the increased exposure period, ZnO-NPs concentration, and lower mitotic activity are all linked. Prophase was the most significant mitotic stage for all of the exposure doses examined, except for when 2 PPM was applied for 9 h and metaphase was the most abundant 90 ± 0.07 (Table 2 and Fig. 7).
The obtained results are in agreement with Giorgetti et al. [72], who used different ZnO-NPs treatments induce distinct increases in MI values in the V. faba root cell, with the lowest dosage (10 mg/l) causing the greatest rise. Previous research (73–75], showed that the MI values in the root tips of A. cepa and V. faba dropped when the concentration of Zn or ZnO-NPs and exposure length rose.
These suppression of MI with concentration dependent, indicating that ZnO-NPs have cytotoxic potential in barley. Kumari et al. [76] and Youssef and Elamawi. [77], described similar effects on MI, where A drop in the MI was seen as the nanoparticle concentration increased. Since ZnO-NPs appear to have a mitodepressive effect, this suggests that they may interfere with mitosis' natural development, preventing some cells from entering prophase and stopping the mitotic cycle in the interphase, therefore limiting DNA/protein synthesis [78, 79].
Table 2
Exposure to ZnO-NPs at different concentrations and for different durations of time affected the MI and phase index.
Time (h) | Conc. (PPM) | MI ± S.E | Mitotic phases % (Phase index %) |
Prophase | Metaphase | Anaphase | Telophase |
control | 5.67 ± 0.04 | 39.28 ± 0.03 | 47.67 ± 0.06 | 9.33 ± 0.04 | 3.52 ± 0.05 |
3 | 1 | 8.93 ± 0.01⁎⁎ | 44.58 ± 0.04 | 43.37 ± 0.02 | 9.63 ± 0.03 | 2.42 ± 0.01 |
2 | 8.63 ± 0.05⁎⁎ | 49.20 ± 0.08 | 44.45 ± 0.01 | 6.35 ± 0.02 | 0 |
4 | 3.25 ± 0.02⁎ | 59.09 ± 0.03⁎ | 36.36 ± 0.01⁎ | 2.28 ± 0.04⁎⁎ | 2.27 ± 0.01 |
8 | 3.47 ± 0.03⁎ | 41.91 ± 0.02 | 47.06 ± 0.04 | 8.72 ± 0.01 | 2.30 ± 0.02 |
12 | 2.79 ± 0.05⁎⁎ | 56.60 ± 0.04⁎ | 35.85 ± 0.07⁎ | 7.55 ± 0.01 | 0 |
6 | 1 | 7.69 ± 0.02⁎⁎ | 92.30 ± 0.08⁎⁎ | 7.70 ± 0.01⁎⁎ | 0 | 0 |
2 | 8.55 ± 0.03⁎⁎ | 55.00 ± 0.02⁎ | 45.00 ± 0.01 | 0 | 0 |
4 | 3.87 ± 0.05⁎ | 58.33 ± 0.06⁎ | 30.00 ± 0.03⁎ | 8.33 ± 0.02 | 3.34 ± 0.01 |
8 | 3.07 ± 0.04⁎ | 34.62 ± 0.05 | 50.00 ± 0.01 | 15.38 ± 0.03⁎⁎ | 0 |
12 | 2.60 ± 0.05⁎⁎ | 50.00 ± 0.03⁎ | 38.09 ± 0.02⁎ | 11.90 ± 0.01 | 0 |
9 | 1 | 3.23 ± 0.03⁎ | 51.51 ± 0.01⁎ | 36.37 ± 0.05⁎ | 9.09 ± 0.01 | 3.03 ± 0.01 |
2 | 2.50 ± 0.04⁎⁎ | 5.00 ± 0.02⁎⁎ | 90.00 ± 0.07⁎⁎ | 5.00 ± 0.03⁎ | 0 |
4 | 3.02 ± 0.05⁎⁎ | 48.78 ± 0.03 | 31.70 ± 0.02⁎ | 17.08 ± 0.04⁎ | 2.44 ± 0.01 |
8 | 1.87 ± 0.02⁎⁎ | 56.67 ± 0.01⁎ | 33.33 ± 0.03⁎ | 10.00 ± 0.02 | 0 |
12 | 1.23 ± 0.01⁎⁎ | 52.18 ± 0.04⁎ | 38.28 ± 0.01⁎ | 4.77 ± 0.03⁎ | 4.77 ± 0.02⁎ |
12 | 1 | 1.73 ± 0.06⁎⁎ | 52.70 ± 0.03⁎ | 44.60 ± 0.03 | 0 | 2.70 ± 0.05 |
2 | 1.25 ± 0.07⁎⁎ | 70.00 ± 0.01⁎⁎ | 30.00 ± 0.01⁎ | 0 | 0 |
4 | No divided cells (Full inhibition) |
8 |
12 |
1 S. E., Standard Error * Significant at level P < 0.05 ** Significant at level P < 0.01 |
3.6 Effect of ZnO-NPs on Chromosomal Aberration
Changes in chromosome structure caused by a break or exchange of chromosomal material are known as chromosomal aberrations. With varied quantities of ZnO-NPs suspensions, Multiple chromosomal abnormalities were found to be caused at all phases of the cell cycle, as well as changes in mitotic stages and nuclear membrane damage, according to the findings.
As the exposure length or ZnO-NPs concentration rose, the rate of chromosomal abnormality increased. For all concentrations tested, the percentage chromosomal aberration values were stated to be higher. At higher nanoparticle dosages of 12 PPM with all-time exposure and at lesser nanoparticle dosages with extended time exposure of 12 h, aberrant chromosomes were observed with their maximum percentage abnormality of 100% (Table 3). Increases in the percentage of abnormalities in root meristems indicate that the test substances are genotoxic [80]. Increased chromosomal abnormalities can be caused by a variety of reasons. The most significant is due to chemical interference during DNA repair. The clastogenicity of chemicals/nanoparticles is represented by many forms of chromosomal abnormalities.
Bridges, lagging, fragmented, disordered, micronucleous, and sticky chromosomes were found in the root meristem of barley that had been minimally exposed to various concentrations of ZnO-NPs as the most common abnormal cells (Table 3 and Fig. 8). When the total number of CAs was considered, the most common anomaly was irregular prophase, which had a high incidence in cell division stages followed by Stickiness and C-mitosis as the most common anomaly. The higher percentage from these CAs was recorded at the 2 PPM concentration as (88.18 ± 0.03 and 88.99 ± 0.04, respectively) (Table 3 and Fig. 8).
Table 3
percentage of abnormalities in all mitotic phases in barley root tips after treatment by different concentrations from ZnO-NPs.
Time (h) | Conc. (PPM) | Total abnormal cells (X ± S.E) | Abnormal mitotic phases % |
prophase | Metaphase | Anaphase | Telophase |
control | 3.53 ± 0.01 | 3.53 ± 0.01 | 0 | 0 | 0 |
3 | 1 | 72.29 ± 0.05⁎⁎ | 38.55 ± 0.07⁎⁎ | 25.30 ± 0.06⁎⁎ | 6.02 ± 0.02⁎ | 2.42 ± 0.04 |
2 | 100.0 ± 0.00⁎⁎ | 49.20 ± 0.08⁎⁎ | 44.45 ± 0.01⁎⁎ | 6.35 ± 0.02⁎ | 0 |
4 | 81.82 ± 0.05⁎⁎ | 43.18 ± 0.02⁎⁎ | 36.36 ± 0.01⁎⁎ | 2.28 ± 0.04 | 2.27 ± 0.01 |
8 | 91.39 ± 0.04⁎⁎ | 37.07 ± 0.03⁎⁎ | 44.61 ± 0.05⁎⁎ | 7.95 ± 0.03⁎ | 2.30 ± 0.02 |
12 | 100.0 ± 0.00⁎⁎ | 56.60 ± 0.04⁎⁎ | 35.85 ± 0.07⁎⁎ | 7.55 ± 0.01⁎ | 0 |
6 | 1 | 72.30 ± 0.04⁎⁎ | 64.60 ± 0.02⁎⁎ | 7.70 ± 0.01⁎⁎ | 0 | 0 |
2 | 100.00 ± 0.01⁎⁎ | 55.00 ± 0.02⁎⁎ | 45.00 ± 0.01⁎⁎ | 0 | 0 |
4 | 91.67 ± 0.03⁎⁎ | 51.67 ± 0.05⁎⁎ | 28.33 ± 0.02⁎⁎ | 8.33 ± 0.02⁎ | 3.34 ± 0.01⁎ |
8 | 100.0 ± 0.00⁎⁎ | 34.62 ± 0.05⁎⁎ | 50.00 ± 0.01⁎⁎ | 15.38 ± 0.03⁎⁎ | 0 |
12 | 100.0 ± 0.00⁎⁎ | 50.00 ± 0.03⁎⁎ | 38.09 ± 0.02⁎⁎ | 11.90 ± 0.01⁎ | 0 |
9 | 1 | 96.97 ± 0.02⁎⁎ | 51.51 ± 0.01⁎⁎ | 36.37 ± 0.05⁎⁎ | 9.09 ± 0.01⁎ | 0 |
2 | 100.00 ± 0.00⁎⁎ | 5.00 ± 0.02⁎⁎ | 90.00 ± 0.07⁎⁎ | 5.00 ± 0.03⁎ | 0 |
4 | 95.00 ± 0.01⁎⁎ | 48.78 ± 0.03⁎⁎ | 26.83 ± 0.05⁎⁎ | 16.95 ± 0.04⁎⁎ | 2.44 ± 0.01 |
8 | 96.67 ± 0.03⁎⁎ | 56.67 ± 0.01⁎⁎ | 30.00 ± 0.01⁎⁎ | 10.00 ± 0.02⁎ | 0 |
12 | 100.00 ± 0.00⁎⁎ | 52.18 ± 0.04⁎⁎ | 38.28 ± 0.01⁎⁎ | 4.77 ± 0.03⁎ | 4.77 ± 0.02⁎ |
12 | 1 | 100.00 ± 0.00⁎⁎ | 52.70 ± 0.03⁎⁎ | 44.60 ± 0.03⁎⁎ | 0 | 2.70 ± 0.01 |
2 | 100.00 ± 0.00⁎⁎ | 70.00 ± 0.01⁎⁎ | 30.00 ± 0.01⁎⁎ | 0 | 0 |
4 | No divided cells (Full inhibition) |
8 |
12 |
2 S. E., Standard Error * Significant at level P < 0.05 ** Significant at level P < 0.01 |
Sticky chromosomes were one of the most often detected abnormalities in root tips of plants treated with ZnO-NPs during metaphase, anaphase, and telophase stages of mitosis (Fig. 9). This finding contradicts the findings of [81, 82], who showed that chromosomal stickiness was the most common CA, and it backs up the significant DNA fragmentation observed in the comet assays conducted by [80, 82] using A. cepa specimens. Previously, researchers (83–85], revealed that nanoparticles have the ability to exhibit a wide range of clastogenic effects, including stickiness, aberrant metaphase, and cell wall dissociation, at various exposure dosages. Chromosomal stickiness and other chromosomal anomalies recorded in this study could be due to the ZnO-NPs binding to DNA and proteins and altering their physico-chemical properties, leading to harmful changes in their chromatin structure, nucleus condensation, or the formation of inter- and intra-chromatid crosslinks. This hypothesis is supported by (i) the strong binding affinity of ZnO-NPs for DNA [86], (ii) the formation of ZnO-NPs complexes with the ring N atom or NH site in nucleobases of DNA [87], and (iii) the interaction, as well as the development of a bioconjugate between protein and ZnO-NPs, reported by Bhunia et al. [88], who observed that, the ZnO-NPs induced structural changes/unfolding of protein. Simultaneously, many experts suggested that stickiness indicates a high toxicity of the chemical and leads to abnormal protein-protein interactions [89]. Sticky chromosomes are thought to be a kind of chromatid aberration [90]. Sticky chromosomes are defined by the loss of function of one or two types of particular nonhistone proteins that control chromatid separation and segregation, according to [91]. Stickiness, according to Darlington and McLENH [92], could be caused by chromosomal DNA degradation or depolymerization. Furthermore, the stickiness caused a disruption in the enzyme system, which may have resulted in a reduction in the rate of cell division [93]. The high values of C-metaphases (Fig. 7, 8 and Table 3) show that Zn compounds are aneugenic, which is likely due to disturbance of calmodulin, a tiny Ca2+−binding protein involved in chromosomal mobility via microtubule polymerization/depolymerization regulation [94]. In addition, very high C- mitosis frequencies could imply partial inhibition of the mitotic apparatus due to oxidative stress caused by higher ZnO-NPs concentrations.
There was evidence of structural chromosomal rearrangement and a possible clastogenic character to ZnO-NPs, since chromosomal breakage, ring chromosomes and chromatin bridges were found. During the anaphase stage, chromosomal breakage and disruption were also reported (Fig. 8). The meristematic cells that were exposed to the ZnO-NPs had bridges during anaphase and telophase. The frequency of micronuclei was seen as a marker for two doses of ZnO-NPs, 4 and 8 PPM, at all times of exposure except at 12 h, when there were no dividing cells, but there were no micronucleated cells in the distilled water as a control.
ZnO-NPs are clearly clastogenic/genotoxic and cytotoxic agents in barley root cells at certain concentrations, as shown by micronucleus. ZnO-NPs exhibited similar genotoxic effects on A. cepa and V. faba, according to the findings of [80]. In V. faba root tip cells, the micronucleus test was utilised by Manzo et al. [95] to investigate the toxicity of ZnO-NPs polluted soil. It has been suggested that micronuclei are formed by chromosomal fragments that do not integrate into either of the daughter nuclei during mitotic telophase, according to [96].
Many different metaphase abnormalities and ana-telophase chromosomal aberrations suggest Zn has genotoxic potential. Chromosome abnormalities indicate that Zn interferes with nucleic acids and has a clastogenic potential.