1 Introduction

Glial tumors stand out as the most prevalent type of central nervous system tumors [44]. Within this category, glioblastoma multiforme represents the most aggressive and malignant form of glioma, posing a formidable challenge as one of the most aggressive and incurable cancers [48], with an average survival of just 12 to 15 months [39]. The limitations of surgical resection in completely eliminating malignant cells [49], coupled with the extensive side effects associated with chemotherapy [56], have spurred research into innovative approaches for selectively targeting cancer cells. Among these recent strategies, antineoplastic bionanocatalysts have emerged as a highly promising avenue [20]. Their appeal lies in several key attributes: their diminutive nanoscale size (typically < 100 nm), expansive surface area, selectivity, mesoporous structure, and catalytic activity, particularly in selectively cleaving essential molecular bonds, such as those within DNA [19].

Bionanocatalysts consist of an oxide matrix where nanostructured metallic agents are uniformly dispersed on its surface, orchestrating a harmonious synergy with the matrix [30]. These nanostructures are adept at catalyzing the selective breakdown of carbon-carbon and carbon-nitrogen bonds by lowering the activation energy required for the process [10, 52, 57]. Of significant note, these nanostructures are engineered for precision, exerting their catalytic influence specifically on the bonds found within genetic material, including DNA and RNA [30]. Moreover, the surface of bionanocatalysts is thoughtfully functionalized with organic agents, rendering these structures inherently and selectively biocompatible exclusively for cancer cells while preserving the integrity of surrounding healthy tissues [24, 26].

In this endeavor, we have synthesized a distinct variant of bionanocatalyst, rooted in a composite matrix of titanosilicate (TiSiOx), where copper-coated nanoparticles have been adroitly dispersed on its surface. Notably, these nanoparticles have demonstrated a remarkable capability to effectively diminish the viability of glioma cells. The physicochemical and structural properties of these innovative bionanocatalysts are meticulously examined through a range of characterization techniques, paving the way for an enhanced understanding of their potential in Catalytic Nanomedicine for cancer treatment.

2 Experimental procedure

2.1 Synthesis of coated bionanocatalysts

Bionanocatalysts were synthesized following a previously reported protocol [26]. Briefly, matrix precursors were added dropwise to solution of deionized water and refluxed at 70 °C until gelation of the matrix: the gel was evaporated until the obtention of a powder. Then, for coating, the appropriate amount of copper precursor (Cu(acac)2) was dissolved in deionized water and organic functionalizing agents, in which the already synthesized powder matrix was added. The amount of coating copper was stablished following the results of previous studies in which different concentrations of copper were evaluated (from 10 to 1%) [4]. Solvents were evaporated until fine power obtention, which was subjected to a manual grinding process in an Agate mortar.

2.2 Bionanocatalysts characterization

X-ray photoelectron spectroscopy (XPS) spectra was obtained using a PHI 5700 spectrometer (Physical Electronics, MN, USA) with a non-monochromatic Mg Kα radiation (300 W, 15 kV, h = 1253.6 eV) as excitation source (pass energy mode at 25.9 eV, chamber kept below 10− 7 Pa). Charge referencing was done against adventitious carbon (C 1s at 284.8 eV). PHI ACCESS ESCA-V6.0 F and Multipack 8.2b software packages were used for acquisition and data analysis. Short acquisition times (< 10 min) were used in order to avoid the photoreduction of the Cu species [45]. N2 adsorption-desorption isotherms were generated through the utilization of a Micromeritics Belsorp IIBell Japan INC instrument. The surface area was determined via the application of the Brunauer-Emmett-Teller (BET) methodology, while the pore volume and pore size distribution were assessed using the Barrett-Joyner-Halenda (BJH) approach. The size, shape, and texture of the grains were analyzed using a JEOL JSM-6010LV scanning electron microscope (SEM). The size of the particles in the samples was assessed through Transmission Electron Microscopy (TEM). Micrographs were captured using a JEOL JEM-2100 F electron microscope, operating with a 200 kV accelerated electron beam. FTIR spectra acquisition was conducted using a NICOLET FTIR spectrometer, specifically the MAGNA 560 model, which featured a resolution of 4 cm-1. A total of 100 scans were averaged for each measurement.

2.3 Cell culture

The Rat C6 glioma cell line was acquired from the European Collection of Animal Cell Cultures located in Porton Down, UK. These cells were cultured in 60-mm Petri dishes using an F-12 Ham Nutrient Mixture, which was supplemented with 10% FBS Gold serum, 2 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin. This culture was maintained in a humid environment consisting of 95% air and 5% CO2 at a temperature of 37 °C. To continue the culture, the cells were collected using a trypsin–EDTA solution twice a week and then seeded into dishes at a density of 106 cells per dish. For the purpose of experimentation, cells from the eighth and twentieth passages were utilized.

2.4 In vitro cell viability assay

C6 glioma cells, initially seeded at a density of 30,000 cells per well, were cultivated in 96-well plates under standard culture conditions. The cells were treated for 8 h with different concentrations (10, 50 and 100 µg/mL) of bionanocatalysts and coated bionanocatalysts diluted in serum free DMEM. Twenty-four hours prior to exposure, the culture medium was replaced with fresh serum-free medium. To assess cell viability and mitochondrial function, we employed the 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2 H-tetrazolium bromide (MTT) assay, which measures the reduction of MTT to MTT-formazan by cellular mitochondrial dehydrogenases. After exposure to orexin A and orexin B for 24 and 48 h, the cell cultures were rinsed with PBS before adding MTT (0.5 mg/ml). The cells were then incubated for 3 h at 37 °C. The formazan crystals were dissolved in 100% dimethyl sulfoxide (DMSO), and the absorbance, which is proportional to the number of viable cells, was assessed at 570 nm using a microplate reader (EnVision 2103, PerkinElmer). The mean inhibitory concentration (IC50) was calculated by calculating the four-parameter dose-response curve.

2.5 Immunocytochemistry and microscopy

For double labeling immunofluorescence, the cells were fixed with 2% paraformaldehyde in phosphate buffer (PBS) at room temperature (RT) for 15 min and permeated in 0.1% Triton X-100-PBS (PBS/T). After blocking for 1 h at RT (0.5% gelatin and 1.5% fetal bovine serum in PBS), cells were incubated for 1 h. at RT in a humidity chamber with antibody anti-α-tubulin monoclonal-IgG (Santacruz Biotechnology, TX, USA). The corresponding secondary antibodies to mouse-IgG were tagged with fluorescein-isothiocyanate (FITC, Jackson ImmunoResearch Labs, PA, USA), and incubated in PBS/T for 1 h at RT (1:1000 dilution). The actin cytoskeleton was stained with dye Rhodamine-Phalloidin (Invitrogen, NY, USA). In experiments with cells, in vivo labeling was done by including the fluorescent mitochondrial marker MitroTracker Deep Red (Invitrogen). Immunolabeled cells were analyzed by epifluorescence through 60x (numerical aperture (NA):1.00 W) and a 100x (NA: 1.3 Oil) Plan-Fluor Lens coupled to a Nikon Eclipse-80i Microscope (Nikon Corp, Tokyo, Japan). The images were obtained and recorded by using a Nikon digital sight-DG-Ri1 camera controlled with the Nikon NIS-Elements AR-3.0- SP7 software included in the system (Nikon Corp).

2.6 Statistical analysis

All data are presented as the mean ± standard error of the mean and 95% confidence interval. Statistical tests and biological replicates are indicated in the figure captions. All statistical analyses were performed with GraphPad Prism v8.0 software. An alpha level of p < 0.05 was set as the threshold for significance.

3 Results and discussion

3.1 Cytotoxicity through bionanocatalytic activity

The cytotoxic effect in terms of cell viability of glioma C6 cells treated with different concentrations of bionanocatalysts is shown in Fig. 1. As can be seen, the mixed oxide catalytic matrix does not generate a significant decrease in the percentage of cell viability with respect to untreated cells. In contrast, the copper-coated bionanocatalyst generated a significant shift in the number of viable cells from the concentration of 10 µg/mL. This phenomenon suggests a high intrinsic biocompatibility of the oxide matrix by not exerting considerable cell death, while the coating with copper groups considerably increases the cytotoxic activity of the bionanocatalyst. Pure copper cytotoxicity evaluation is included as a control. The cytotoxicity of copper-coated nanocomposites is widely known [15], its main mechanisms of action being DNA damage through catalytic formation of reactive oxygen species [46]. In previous work, we have demonstrated the high dispersion of copper-coating nanostructured agents in oxide matrices [11, 27, 28]. As in the present work, the coating with the cuprous derivatives gave the bionanocatalysts enhanced antineoplastic properties. In particular, these copper-coated bionanocatalysts were found to exert part of their cytotoxic activity due to the catalytic degradation of the DNA [22].

Fig. 1
figure 1

Catalytic cytotoxic effect of bionanocatalysts as a function of cell viability at different concentrations and comparison against pure copper cytotoxicity. Significant differences (*) were determined as p < 0.05

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3.2 Mechanisms of cytotoxicity

To delve deeper into these mechanisms, we examined cell morphology following the introduction of bionanocatalysts. In Fig. 2, we present an analysis of C6 glioma cell cytoskeleton interactions with copper-coated bionanocatalysts. In the brightfield images, we can discern the presence of crystals formed due to the aggregation of these nanostructures. Notably, in cells treated with the matrix the radial arrangement of the tubulin cytoskeleton (identified through anti-α-tubulin staining in the green channel) and the fibrillar actin structure (revealed via rhodamine-phalloidin staining in the red channel, white arrows) both maintain their normal and unaffected structure. In contrast, the coated bionanocatalysts lead to the disruption of the central microtubule structure, transforming it into a structure that encircles the nucleus. Consequently, this transformation results in the complete degradation of actin filaments, as indicated by the arrowheads. This outcome unequivocally demonstrates that the presence of copper coating in the matrix induces a significant alteration in the cytoskeletal structure. Previous studies suggest that such destabilization may take place through a catalytic proteolysis, in which the bionanocatalyst act as a cleavage compound for the carbon-carbon and carbon-nitrogen bonds within the cytoskeleton protein [34, 43].

Fig. 2
figure 2

Cytoskeleton integrity evaluation by double-labelling with a-tubulin and rhodamine-phalloidin showing organelle disorganization as a function of bionanocatalysts activity. The nucleus is indicated with an asterisk (*). Arrows indicate normal cytoskeleton, whereas arrowheads indicate filament degradation

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Finally, to corroborate the impact of the catalytic activity of bionanocatalysts on the metabolic activity of mitochondria in C6 cells, staining by MitoTracker Deep Red dye was performed to identify metabolically active mitochondria (Fig. 3). Under control conditions (uncoated matrix), we observed the typical morphology of the organelles. However, in cells exposed to the coated bionanocatalysts, the mitochondrial network underwent significant structural modifications. The bionanocatalysts accumulated around the nucleus, causing the mitochondria to lose their normal network-like structure. Additionally, these cells displayed an elongated morphology distinct from the majority of fibers. Recent studies have firmly established that changes in the morphology of mitochondria indicate the onset of stress, shifts in the redox state, and the initiation of apoptosis [1]. Consequently, our results clearly indicate a significant cytotoxicity and pronounced distortions in mitochondrial morphology.

Fig. 3
figure 3

Mitochondrial network integrity evaluation by MitoTracker Deep Red showing damage due to bionanocatalyst concentration in the surrounding nucleus

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3.3 Bionanocatalyst atomic composition

To further understand the cytotoxic capacity of the bionanocatalysts, a physicochemical characterization was carried out. Figure 4a shows the XPS patterns of the two bionanocatalysts evaluated. The high-resolution XPS spectra for each component are shown in Fig. 4b-f. The deconvolution of the Cu 2p region reveals several key signals. The primary peak, situated at 934.05 eV, is attributed to the Cu 2p3/2 state of Cu2+ within cupric oxide (CuO) [53]. Its accompanying Cu 2p1/2 signal is found at a 20.0 eV split, which is characteristic of copper oxides and lies in the same spectral region [5]. Additionally, though less pronounced in intensity, another component emerges at 932.57 eV (with a corresponding 20.0 eV counterpart), potentially associated with either Cu+ in Cu2O or Cu0 [18]. Unhopefully, through XPS analysis it is not possible to discern between these two states [18].

These various oxidation states of copper play a crucial role in achieving a catalytic enhancement effect [8]. This effect arises from the overlapping of the conduction bands of the oxide matrix, leading to increased catalytic activity [6]. In the case of TiO2, this phenomenon results in a shift in its photocatalytic activity from ultraviolet to visible light [16]. Furthermore, three signals, each with a complementary component at approximately 20.0 eV, are observable at 939.37, 942.26, and 944.39 eV. The first of these signals can be attributed to Cu(OH)2 [18], likely corresponding to surface copper-coating nanoparticles stabilized by hydrogen interactions with the matrix. The second signal has not been classified in the literature and may represent a unique bond formed between Cu and the oxide matrix, possibly through coordination bonds [50]. The last signal corresponds to a satellite feature associated with the Cu2+ signal [16].

Fig. 4
figure 4

(a) XPS and (b-f) high-resolution XPS spectra of coated bionanocatalysts compared to uncoated bionanocatalyst showing peaks for (b) Cu 2p, (c) Si 2p, (d) Ti 2p, (e) O 1s, and (f) C 1s

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Regarding the catalytic oxide matrix, the signal deconvolution for Si 2p (Fig. 4c) indicates the presence of different oxidation states: Si+ (suboxides SiOx), Si4+ (silicon dioxide, SiO2), and Si3+ (Si-O-Si bond) [14, 37, 38], the latter being the preponderant one in the bionanocatalyst matrix. No signals were observed for Si0 or Si2+, indicating that all Si in the bionanocatalyst matrix is oxidized and that no organic remnants are directly bound to Si, respectively [38, 55]. In the other hand, Fig. 4d shows the de-convolution of XPS spectra for Ti 2p. The peaks observed at 458.68 and 464.38 eV correspond to the Ti4+ 2p3/2 and Ti4+ 2p1/2, respectively. The shoulder peaks at 459.81 and 465.51 eV represent traces of Ti3+ 2p3/2 and Ti3+ 2p1/2. The binding energy difference (Δ) for (Ti4+ 2p3/2 - Ti4+ 2p1/2) and (Ti3+ 2p3/2 – Ti3+ 2p1/2) is 5.7 for both, which is similar to the reported by previous study [30]. The Ti3+ state is known to be presented due to oxygen deficiency and high hydrogen concentration in the network [40]. In this regard, in the O 1s region (Fig. 4e), both samples exhibit two signals: one attributed to the Si-O bonding [36], and the other attributed to oxygen vacancies [41]. Bi et al. [3] observed that TiO2 exhibits high visible light response and stable photocatalytic activity owing to the presence of rich oxygen vacancies and Ti3+, which act as traps to reduce the recombination of e-/h+ pairs, hence potentiating the catalytic activity of the oxide.

Finally, regarding the C 1s region, Fig. 4f shows four signals: the first two correspond to the C-C bond, which exhibits a sp2 hybridization state (attributable to the delocalized electrons in the acetylacetonate group), and a sp3 hybridization state (attributable to organic remnants generated during the hydrolysis process of the precursor alkoxide in the synthesis) [36]. The other additional signal corresponds to the C-O bond which is associated with the OH groups of the acetylacetonate group [12]. It is to be noted that the corroboration of the presence of all the bond signals attributed to the acetylacetonate group confirms the presence of the group. It has been found that this molecule is essential during the coating of the matrix so as to achieve high dispersion of copper. Finally, the last signal corresponds to ambient CO2 deposited on the surface of the bionanocatalyst [38].

3.4 Bionanocatalyst morphology and particle size

The morphology of bionanocatalysts is observed by SEM and TEM (Fig. 5). Coated bionanocatalysts are observed to form smooth and amorphous clusters with sizes on the order of 500 nm (see Fig. 5a). In contrast, transmission electron microscopy (TEM) analysis of the coated bionanocatalysts reveals crystalline structures (Fig. 5b) and particle sizes around 10 nm (Fig. 5c). The SAED patterns of copper-coated bionanocatalysts (Fig. 5d) in rutile phase exhibit scattered ring patterns devoid of extra diffraction spots and rings from secondary phases, indicating their strong crystalline structure.

For the uncoated matrix, SEM imaging (Fig. 5e) shows the formation of conglomerates with diameters around 200 nm. These conglomerates are spherical and have a smooth surface. TEM examination of individual particles confirms their crystallinity (Fig. 5f) and sizes to be approximately 10 nm (Fig. 5g). It is worth noting that the high dispersion and low concentration of coating copper in the coated sample seems to alter the conglomerate structure of the bionanocatalysts: the SAED patterns of oxide matrix (Fig. 5h) shows that the brightness and intensity of polymorphic ring is weak, so they are poorly crystallized and partly amorphous; however, the structure and size of the individual nanoparticle remains the same (Fig. 5c and g).

Fig. 5
figure 5

(a, e) SEM, (b, f) TEM, (c, g) high-resolution TEM micrographs, and SAED (d, h) of the compounds (a-d) coated and (e-h) uncoated bionanocatalysts

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3.5 Bionanocatalyst surface properties

Figure 6a illustrates the FTIR spectra (2000 –400 cm− 1) of coated and uncoated bionanocatalysts and the copper precursor used. In the titanosilicate spectrum, a peak is observed around 1629 cm− 1, which is attributed to the bending vibration of the O-H bond, indicative of adsorbed water molecules. Additionally, peaks at 796 and 1086 cm− 1 are attributed to the symmetric and asymmetric stretching vibrations of Si-O-Si bonds, respectively [42]. The peak at 944 cm− 1 corresponds to the Ti-O-Si asymmetric stretching vibration, confirming the formation of Ti-O-Si linkages [42]. This suggests the replacement of some silicon atoms in the silica network by titanium atoms, resulting in the titanosilicate mixed oxide. Regarding the Cu(acac)2 spectra, peaks at 1577, 1555, and 1533 cm− 1 correspond to C = C and C = O stretching vibrations, while the peak at 684 cm− 1 is attributed to Cu-O stretching vibrations between the copper-coating atom and the oxygen atoms in the acetylacetonate ions [47]. The 610 cm− 1 peak is due to ring vibrations and Cu-O stretching vibration modes, and the peaks at 451 cm− 1 and 431 cm− 1 are related to Cu-O and ring deformation modes. Upon analyzing the coated bionanocatalysts spectrum, several similarities with the uncoated and Cu(acac)2 spectra were found, confirming the presence of characteristics from both Cu(acac)2 and TiSiOx.

Fig. 6
figure 6

(a) FTIR spectra of coated and uncoated bionanocatalysts, as well as copper precursor Cu(acac)2. (b) BET isotherms of coated and uncoated bionanocatalysts

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Surface area, pore volume, and pore diameter data (Table 1) were obtained using the BET and BJH methods based on nitrogen adsorption-desorption isotherms (Fig. 6b). The uncoated sample exhibits a surface area of 320 m2/g, a pore volume of 0.423 cm3/g, and a pore diameter of 5.28 nm. Interestingly, the sample coated with copper exhibited a significant reduction in these values, especially in terms of BET surface area. This decline suggests that the copper-coating complexes modify the mesoporous structure of the matrix upon coating, as it is known that such nanoparticles fill the pores of the oxide matrix [9]. It is to be noted, however, that such a decrease in surface area does not exert a negative effect on the reactivity of the material, but, on the contrary, increases the cytotoxic activity of the nanostructures as evidenced in Fig. 1. This is due to the very nature of bionanocatalysts as catalytic ultra-nanoparticles with colloid-like behavior [21]. In normal catalysts, the surface area is directly related to the reactivity of the material. However, in ultra-nanoparticulate bionanocatalysts, their reactivity is directly linked to the metallic agents with which their surface is coated, to their overall size of 1 to 3 nm, and to the surface functionalizing agents.

Table 1 Textural properties of the bionanocatalysts
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3.6 Bionanocatalysts’ mechanisms of action in terms of structural composition

As a whole, the physicochemical characterization suggests that the bionanocatalysts are composed of a mixed matrix of titanosilicate in which copper-coating nanoparticles (mainly CuO) are homogeneously dispersed in the surface of the matrix and determine the morphological and textural properties of the bionanocatalyst clusters (without affecting the structure of the individual nanoparticles), as well as the catalytic activity of the nanostructures by overlapping the conduction bands of the matrix oxides. In turn, copper-coating nanoparticles endow the intrinsically biocompatible titanosilicate matrix with an antineoplastic capacity by allowing it to exert cytotoxic activity on the genetic material of cancer cells, destabilize the cytoskeleton and, finally, alter the oxide-reductive activity of mitochondria, leading them to the activation of programmed cell death by apoptosis.

In previous work we have demonstrated the high selectivity of this type of bionanocatalysts for exclusive cancer cells, without affecting normal cells in vitro [11]. Similarly, other studies have examined the function of the oxide matrix as a cytotoxic chemical buffering agent in healthy cells at the cellular level. For instance, Lopez et al. discovered that the nanostructured TiO2 matrix reduced the phototoxic effects of zinc phthalocyanine by up to 80% compared to the pure substance by restricting the drug’s entry into healthy cells [25]. Similarly, studies have shown that the functionalized matrix of onco-bionanocatalysts lacking transition metals does not harm cells, as evidenced by various cell viability tests [24, 26, 28]. This has also been confirmed by several clinical studies performed on chronic wounds where this type of bionanocatalysts composed of TiO2, SiO2 and superficial CuO were used as disinfectant agents in open wounds, without any observable side effects or damage to the surrounding healthy tissues for up to 6 months [29, 30]. An intriguing correlation exists between the surface coating and natural biocompatibility of the nanostructured matrix of bionanocatalysts [2]. These findings underscore the biosafety of bionanocatalysts. However, further investigation across various cell lines and tissues is necessary to fully elucidate their selective toxicity and biocompatibility.

In addition, we have proposed hypotheses related to the possible mechanisms of action of this type of nanostructures for the selective bionanocatalytic cleavage of carbon-carbon and carbon-nitrogen bonds in the genetic material of cancer cells [31]. In this regard, in a previous work, the kinetics of degradation of genetic material in C6 glioma cells guided by the formation of adducts from the first 30 min of interaction was evaluated [23]. Remarkably, after the first 60 min of incubation, the genetic material had been completely degraded, demonstrating enhanced activity with respect to cisplatin whose genotoxic potential was observed up to 180 min. However, considering the other two mechanisms of action of bionanocatalysts for the elimination of cancer cells, it is necessary to deepen our understanding of their cytotoxicity kinetics. Although in the present work a significant increase in cell death was observed after 8 h of interaction, future work should evaluate the rate of cell viability as a function of time, so that the time of action of the nanostructures can be known.

When bionanocatalysts interact with DNA, they induce the breaking of bonds throughout the genetic material at a rapid rate. This damage occurs randomly across the DNA structure, overwhelming repair mechanisms. In line with current understanding, if DNA repair mechanisms fail, cells activate programmed cell death pathways, such as apoptosis [13]. Essentially, cells in a tissue that sustain DNA damage beyond repair capacity are eliminated from the population. DNA damage often triggers death receptors outside the cell (such as Fas, CD95, Apo-1) and/or internal mitochondrial apoptotic pathways [7]. The Fas signaling pathway involves receptor activation, formation of the DIS complex (consisting of FADD, Fas-associated protein with a death domain, and procaspase-8 and − 10), and initiation of the caspase cascade, leading to DNA cleavage by CAD (caspase-activated DNAase) and degradation of intracellular proteins by caspase-3 and − 7 [54]. Conversely, the mitochondrial apoptotic pathway revolves around the regulation of cytochrome c release from mitochondria into the cytoplasm, along with proapoptotic proteins like Bax and Bak, countered by antiapoptotic proteins like Bcl-2 and Bcl-XL. Upon cytochrome c release, apoptosomes containing procaspase-9, Apaf-1, and cytochrome c assemble, initiating the caspase cascade, ultimately leading to protein and DNA breakdown [51].

In this sense, the present work deepens in the cytotoxic mechanisms of bionanocatalysts and evidences the possibility of using copper-coating nanoparticles (in contrast with previous findings using silver and platinum nanoparticles [23, 32]); for the potentiation of the matrix. The above is summarized in Fig. 7.

Fig. 7
figure 7

Cytotoxic mechanisms of action of bionanocatalysts: destabilization of cytoskeleton, mitochondria, and DNA through bond cleavage

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4 Conclusions

In the present work, the cytotoxic activity towards glioma cancer cells of a new type of bionanocatalysts based on a catalytic mixed oxide matrix, in which copper-coating nanoparticles were homogeneously dispersed on its surface for the optimization of the intrinsic catalytic properties of the matrix and the generation of cytotoxic properties, was evaluated. Copper-coated bionanocatalysts were able to significantly reduce cell viability compared to reference bionanocatalysts. Structural damage to the cytoskeleton and mitochondrial activity were observed, suggesting these pathways as part of the mechanisms of action of these nanostructures derived from their bionanocatalytic activity for bond breaking. The physicochemical studies corroborated that the functionalization with copper-coating species (mainly CuO) did not alter the individual structure of the bionanocatalysts but enhanced their cytotoxic activity. With this work, it is expected to deepen in the mechanisms of action of this potential antineoplastic treatment and to evidence the use of copper-coating nanoparticles as potentiating agents of the intrinsic properties of the bionanocatalysts.