Davide
Redolfi-Bristol
*acg,
Kenta
Yamamoto
c,
Elia
Marin
ahi,
Wenliang
Zhu
a,
Osam
Mazda
c,
Pietro
Riello
g and
Giuseppe
Pezzotti
*abcdefg
aCeramic Physics Laboratory, Kyoto Institute of Technology, Sakyo-ku, Matsugasaki, 606-8585, Kyoto, Japan. E-mail: davide.redolfi@unive.it; pezzotti@kit.ac.jp
bDepartment of Molecular Genetics, Institute of Biomedical Science, Kansai Medical University, 2-5-1 Shinmachi, Hiraka-ta, Osaka 573-1010, Japan
cDepartment of Immunology, Graduate School of Medical Science, Kyoto Prefectural University of Medicine, 465 Kajii-cho, Kamigyo-ku, Kyoto 602-8566, Japan
dDepartment of Dental Medicine, Graduate School of Medical Science, Kyoto Prefectural University of Medicine, 465 Kajii-cho, Kamigyo-ku, Kyoto 602-8566, Japan
eDepartment of Orthopedic Surgery, Tokyo Medical University, 6-7-1 Nishi-Shinjuku, Shinjuku-ku, 160-0023 Tokyo, Japan
fDepartment of Applied Science and Technology, Politecnico di Torino, Corso Duca degli Abruzzi 24, 10129 Torino, Italy
gDipartimento di Scienze Molecolari e Nanosistemi, Università Ca’ Foscari di Venezia, Via Torino 155, 30172 Venezia, Italy
hDepartment Polytechnic of Engineering and Architecture, University of Udine, 33100, Udine, Italy
iBiomedical Research Center, Kyoto Institute of Technology, Sakyo-ku, Matsugasaki, Kyoto 606-8585, Japan
First published on 2nd May 2024
Silver nanoparticles (AgNPs) hold great promise for several different applications, from colorimetric sensors to antimicrobial agents. Despite their widespread incorporation in consumer products, limited understanding of the detrimental effects and cellular antioxidant responses associated with AgNPs at sublethal concentrations persists, raising concerns for human and ecological well-being. To address this gap, we synthesized AgNPs of varying sizes and evaluated their cytotoxicity against human dermal fibroblasts (HDF). Our study revealed that toxicity of AgNPs is a time- and size-dependent process, even at low exposure levels. AgNPs exhibited low short-term cytotoxicity but high long-term impact, particularly for the smallest NPs tested. Raman microspectroscopy was employed for in-time investigations of intracellular molecular variations during the first 24 h of exposure to AgNPs of 35 nm. Subtle protein and lipid degradations were detected, but no discernible damage to the DNA was observed. Signals associated with antioxidant proteins, such as superoxide dismutase (SOD), catalase (CAT) and metallothioneins (MTs), increased over time, reflecting the heightened production of these defense agents. Fluorescence microscopy further confirmed the efficacy of overexpressed antioxidant proteins in mitigating ROS formation during short-term exposure to AgNPs. This work provides valuable insights into the molecular changes and remedial strategies within the cellular environment, utilizing Raman microspectroscopy as an advanced analytical technique. These findings offer a novel perspective on the cytotoxicity mechanism of AgNPs, contributing to the development of safer materials and advice on regulatory guidelines for their biomedical applications.
The toxicity of AgNPs is mainly due to the generation of reactive oxygen species (ROS), causing oxidative stress and damaging cellular components. Studies have shown that AgNPs undergo a dissolution process upon entering inside the cell, leading to the release of Ag+ ions.20–22 The exogenous metal ions induce the production of ROS, including superoxide radicals and hydrogen peroxide, within the cells.23–25 These elevated levels of ROS can lead to damage of cellular components such as lipids, proteins, and DNA if they are not properly contained by antioxidant agents.26,27 Indeed, enzymatic antioxidants, such as superoxide dismutase (SOD) and catalase (CAT), and non-enzymatic antioxidants, such as glutathione and the family of metallothioneins (MTs), play a crucial role in maintaining cellular homeostasis by counteracting exogenous metal ions and oxidative stress inside cells. However, if the toxic action is persistent and prolonged over time, it can overwhelm the initially effective defense mechanism, ultimately resulting in its failure.25,28,29
Given these reasons, it is essential to achieve a thorough comprehension of the diverse physicochemical changes resulting from exposure to silver nanoparticles. This is crucial to enhance our knowledge of cellular responses, thereby enabling better understanding of exposure limits and potential implications associated with their usage. Moreover, while the literature extensively explores the impact of AgNPs at elevated concentrations for a short period of time, an inherent lack of emphasis on investigations below toxicity thresholds underscores the need to investigate the intricate cellular defense mechanisms. Unraveling these complex interactions between AgNPs and living organisms demands sophisticated analytical techniques capable of providing detailed insights into their behavior at the molecular level.
Raman microspectroscopy stands out as a powerful analytical technique for investigating the effects of nanomaterials on cells due to its unique advantages. Unlike traditional methods, Raman spectroscopy provides label-free and non-invasive probing capabilities, allowing researchers to directly observe molecular changes within living cells.30,31 Indeed, Raman microspectroscopy has been employed for the characterization of biochemical evolution during cellular differentiation,32,33 neuronal cell networking and separation,34 metabolic changes in cancer cells35,36 and apoptosis.37,38 In the nano-toxicology context, this technique can have the advantageous ability to provide detailed information on cellular component alterations, which can be correlated with cytotoxic responses, oxidative stress, or inflammation induced by nanomaterial exposure.39–42 Furthermore, the non-destructive nature and detailed molecular information obtained through Raman spectroscopy make it a helpful tool for advancing the comprehension of nanomaterial–cell interactions in biomedical research and toxicology.31,39 In recent years, a small number of studies have been performed to assess the toxic effects induced by AgNPs through Raman spectroscopy.43–45 However, no study on the temporal evolution of the cellular stress condition has ever been carried out and, in most cases, red blood cells, which are known to be susceptible to oxidative stress due to the absence of a nucleus and other organelles, have been employed as a cellular model.
In this work, we synthesized silver nanoparticles of different sizes and tested their cytotoxicity against human dermal fibroblast (HDF) cells. We employed HDF cells as a model to better explore the consequences of dermal exposure to AgNPs, given their utilization in fabrics and wound dressings. After discovering the concentration limits for which these NPs exhibited low cytotoxicity in the short term but high cytotoxicity in the long term, an in-time investigation into intracellular molecular variation that occurs in the first 24 h of exposure was carried out using Raman microspectroscopy. Raman spectroscopy allowed the detection of slight degradations of proteins and lipids, triggered by the presence of AgNPs inside the cell. Signals deriving from the C–S bond and from the active sites of SOD and CAT proteins increased during the exposure time, caused by the increased production of antioxidant agents. Eventually, fluorescence microscopy was employed to confirm the efficacy of the overexpressed antioxidant proteins in mitigating the ROS formation during short-term exposure to AgNPs. Our work provides an elucidation about the molecular changes and remedial strategies that occur within the cellular environment using the advanced technique of Raman microspectroscopy. This novel approach allowed for real-time monitoring of molecular variations within the cellular environment, providing advanced insights into the dynamic interactions between AgNPs and cellular components. Supported by conventional end-point analyses, the focus on the temporal dynamics of cellular responses detected through Raman microspectroscopy allowed the observation of a significant increase in specific signals associated with antioxidant actions, a phenomenon rarely documented in previous studies, in particular in real-time. By uncovering these novel molecular signatures our results allow understanding, from a different perspective, the mechanism underlying the cytotoxicity of silver nanoparticles, providing valuable insights into the development of safer materials and advice on the regulatory guidelines for their use in biomedical applications.
Raman spectra of HDF cells were collected with a dedicated Raman device (RAMANtouch, Nanophoton Co., Osaka, Japan). The RAMANtouch spectroscope was operated with an excitation source of 532 nm (excitation power density = 576 mW μm−2), a 300 gr mm−1 grating and a 60× immersion objective lens (NA = 1.0). The spectral resolution was about 3 cm−1 and was recorded in the range of 400 to 1800 cm−1. The RAMANtouch spectroscope was operated in “point mode”, acquiring 6 spectra per cell (3 from the cytoplasm and 3 from the nucleus), analyzing ten cells per sample. Around 60 spectra were therefore acquired per sample. The exposure time for each measurement was 1 s and the acquisition was averaged five times. All the experiments were performed in triplicate. Reference Raman spectra of SOD and CAT were also collected with the RAMANtouch spectroscope after depositing the sample (SOD powder; CAT water solution) on the CaF2 substrate.
Raman data were processed averaging around 60 spectra per sample through RAMAN Viewer software (Nanophoton Co., Osaka, Japan), excluding those that showed too large intensity caused by the surface enhanced Raman scattering (SERS) phenomenon. On the averaged spectrum background subtraction, smoothing (Savitsky–Golay smoothing; degree 2, size 7, height 11) and baseline correction (manually selecting the points representative of the background) were performed by means of LabSpec software (version 5.5, Horiba, Japan). Subsequently, the standard normal variate (SNV) normalization was performed using SpectraGryph software (F. Menges, “Spectragryph – optical spectroscopy software”, version 1.2.16.1, 2022, https://www.effemm2.de/spectragryph/). Finally, an average of the resulting Raman spectrum for each replicate was obtained using Origin software (version 9.8.0.200, OriginLab, Massachusetts, USA). More details about the data processing are reported in Fig. S2.†
Fig. 1a shows the UV-visible spectra of AgNPs25nm seeds and their subsequent growth steps. As can be seen from the absorption bands, it is possible to observe a red-shift in the maximum absorbance as the size of the nanoparticles increases. This fact is correctly correlated with the color change of the solutions during the different growth steps. The precise maximum absorbance values are reported in Table 1.
![]() | ||
Fig. 1 (a) UV-visible spectra, (b) analytical centrifugation and (c) DLS distribution of AgNPs. SEM images of (d) AgNPs25nm, (e) AgNPs35nm and (f) AgNPs50nm (scale bar: 100 nm). |
Sample name | UV-visible (Absmax, nm) | AC – number distribution (diameter, nm) | DLS – intensity distribution (diameter, nm) | SEM – number distribution (diameter, nm) |
---|---|---|---|---|
AgNPs25nm | 408 | 24 ± 2 | 33.2 (PDI = 0.1) | 24 ± 3 |
AgNPs30nm | 421 | 30 ± 3 | 37.5 (PDI = 0.1) | 31 ± 3 |
AgNPs35nm | 429 | 34 ± 4 | 42.7 (PDI = 0.1) | 37 ± 4 |
AgNPs40nm | 434 | 40 ± 3 | 46.8 (PDI = 0.1) | 44 ± 5 |
AgNPs45nm | 438 | 45 ± 3 | 53.1 (PDI = 0.1) | 47 ± 5 |
AgNPs50nm | 442 | 49 ± 7 | 55.7 (PDI = 0.1) | 52 ± 6 |
Analytical centrifugation and DLS analysis were performed to assess the hydrodynamic diameter of AgNPs, and the results are reported in Fig. 1b and c and Table 1. AC measurements show a narrow distribution with an increase in the size of the NPs of approximately 5 nm after each growth step. These results were confirmed by DLS analysis, which also revealed a growth of around 5 nm between the steps. However, DLS results showed a broader distribution with a larger average diameter size. The slight differences in the size estimation among the techniques may be due to the different solvation effects that the particles encounter during the measurements. Fig. 1d–f shows three representative SEM images, respectively, of AgNPs25nm seeds, AgNPs35nm and AgNPs50nm obtained through different growth steps. SEM images of other growth steps are reported in Fig. S1.† Silver nanoparticles show a spherical shape with a homogeneous distribution that increases by about 5 nm between the steps (Table 1). This confirms the correct growth of the nanoparticles and that the differences in the measured size are only caused by the different characterization techniques.
After the synthesis, the AgNPs were washed and concentrated through centrifugation. Before testing the cytotoxicity, the amount of silver in the colloidal solutions was quantified through ICP-OES. This was done to properly verify that the quantity of silver tested for viability assay was the same for all the different NP types. ICP-OES results are reported in Table 2 and they highlight a large difference between the expected concentrations of the final solutions and that of the real one, except for AgNPs50nm. During the synthesis, a large quantity of silver ions did not react or, during the concentration process, a large number of nanoparticles remained in the supernatant, thus resulting in a lower concentration than expected. More studies would be necessary to discern these two possibilities.
Sample name | Expected concentration (μg mL−1) | Measured concentration (μg mL−1) |
---|---|---|
AgNPs25nm | 540 | 2.9 ± 0.1 |
AgNPs30nm | 1050 | 32.5 ± 0.2 |
AgNPs35nm | 1580 | 159.1 ± 0.3 |
AgNPs40nm | 2110 | 354.3 ± 0.3 |
AgNPs45nm | 2650 | 1093 ± 1.0 |
AgNPs50nm | 3250 | 3521 ± 2.0 |
AgNPs35nm and AgNPs50nm were selected to study the toxicity of the nanoparticles against HDF cells for 24 and 72 h at 0.2, 2 and 20 μg mL−1. These two sizes were chosen because of their distinct distribution and homogeneous diameters, while also being abundant enough to be used in toxicological tests. In addition, the concentrations were selected because they demonstrated reduced toxicity for similar NP sizes in previous studies on HDF cells.49,50 Quantification of living cells was performed by MTT assay, as reported in previous studies.49,50
The results reported in Fig. 2 indicate that nanoparticles exhibited cytotoxicity against HDF cells following prolonged exposure of 72 h, especially for AgNPs35nm; however, no discernible toxicity was observed during shorter exposures up to 24 h. After 24 h, AgNPs35nm displayed a low cytotoxicity for the concentration tested, with a slightly larger reduction in cell viability at 20 μg mL−1. An even lower cytotoxicity was observed for cells treated with AgNPs50nm, showing a maximum viability reduction of about 9% at the highest concentration tested for 24 h. In contrast, after 72 h, the toxic effect of the nanoparticles significantly increased, exhibiting a low cellular viability of 10% for AgNPs35nm and 63% for AgNPs50nm at 20 μg mL−1. As reported in the literature, it is clear that cytotoxicity is size- and time-dependent, when the same concentration in μg mL−1 is used to test them.50,51 Larger nanoparticles seem to be internalized at a slower rate when compared to smaller ones.52 Moreover, at an equivalent mass concentration (expressed in μg mL−1), their number concentration (expressed in N° of NPs mL−1) and their global surface area are lower, further diminishing their presence in the medium and within the cell. The prolonged exposure to nanoparticles can instead exert a long-lasting toxic and anti-proliferative effect. This phenomenon can be due to the fact that the AgNPs, remaining inside the cells, undergo a degradation process whose kinetics is expected to be subject to the dimension of AgNPs (smaller nanoparticles dissolve more rapidly than larger ones), ultimately leading to a size-dependent release of Ag+ ions.21,22,53 The release of Ag+ ions represents the primary cause of cytotoxicity associated with silver nanoparticles, as their presence within the cell gives rise to the generation of an excess of ROS.21,25 A longer permanence of the nanoparticles inside the cell therefore causes a continuous release of Ag+ ions and, consequently, a more prolonged cytotoxic effect over time.
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Fig. 2 Quantification of living cells after incubation with AgNPs35nm and AgNPs50nm for 24 and 72 h. *Significant differences from the negative control (P < 0.05). |
In light of the cytotoxicity results, we decided to investigate the cellular and antioxidant mechanisms that occur inside the cells during the first 24 h of exposure to AgNPs. Raman microspectroscopy was conducted over time on HDF cells treated with AgNPs35nm at a concentration of 10 μg mL−1. This size and concentration were chosen as the optimal compromise between the rapidity of cytotoxic action and the degree of toxicity between the two previously tested sizes. At different time points (0, 4, 8, 12 and 24 h) the cells were washed and fixed with formaldehyde prior to Raman microspectroscopy analysis. Although it is acknowledged that cell fixation introduces some changes to the resulting Raman spectrum, this step was performed to enable a more accurate and extensive analysis of the cells. This approach minimizes potential distortions arising from temporal gaps between measurements and from external environmental factors (e.g., temperature fluctuations and CO2 levels). Moreover, formaldehyde fixation was chosen based on previous studies indicating that it is the method that induces the least alteration to the final spectrum.54–57
The Raman spectra reported in Fig. 3a were obtained by averaging the signals from around 60 spectra per sample, followed by a final averaging per three replicates. The cytoplasmic and nuclear spectra were combined, as no significant differences relevant to our study were observed when analyzed separately. The points from which spectra were acquired were carefully chosen avoiding nanoparticle aggregates that could have given rise to exceedingly intense SERS phenomena, in order to prevent misinterpretations or spectral alterations. More details about the data processing are reported in Fig. S2.† The Raman bands are associated with diverse vibrations of specific biochemical molecules found within the cell and the assignment of these signals is presented in Table 3.
Wavenumber (cm−1) | Biological molecule | Tentative assignment of bond vibration | Ref. |
---|---|---|---|
638 | Proteins | –C–S– bond | 58 and 59 |
705–720 | Lipids | Phospholipid C–N stretch | 60 |
740–760 | Protein | Tryptophan (ring breathing) | 58, 60 and 61 |
770–785 | DNA/RNA | DNA backbone O–P–O, uracil, cytosine and thymine (ring breathing) | 58, 60 and 61 |
810–825 | RNA | RNA backbone O–P–O | 58, 60 and 61 |
920–965 | Proteins | –C–C– backbone | 58, 60 and 61 |
990–1005 | Proteins | Phenylalanine (symmetric ring breathing) | 58, 60 and 61 |
1020–1035 | Proteins | Phenylalanine (C–H in plane bending) | 58, 60 and 61 |
1075–1095 and 1115–1125 | Proteins and lipids | C–N stretching in proteins and C–C stretching in lipids | 58 and 61 |
1235–1270 | Proteins | Amide III | 36, 58 and 61 |
1280–1340 | Proteins and lipids | Amide III and CH2 and CH3 vibrations of lipids | 58 and 61 |
1430–1460 | Proteins and lipids | C–H vibration | 33, 58 and 61 |
1572 | Proteins | Superoxide dismutase and catalase | 62–67 |
1600–1670 | Proteins | Amide I | 58 and 61 |
In Fig. 3a, it is possible to observe that the Raman bands exhibit variations in intensity at different time points. Nevertheless, they maintain their characteristic shape, confirming that the formaldehyde fixing process enables a consistent and reproducible comparison between the samples. The bands at around 940, 995, 1030, 1082, 1120 and 1440 cm−1 show a small but general decrease in intensity with time, compared to the initial state (Fig. S3†). All these bands belong mainly to protein and lipid molecular structures.58,60,61 As reported in the literature, this fact indicates that in the cell a deterioration process occurs, which is caused by the presence of exogenous and toxic substances.68–70 The AgNPs that penetrate inside the cells undergo a dissolution process, releasing hazardous Ag+ ions, which impairs the regular functions of cells.21,22 The introduction of NPs and external metal ions therefore stimulates the generation of free radicals within the cellular environment, eventually contributing to the breakdown of proteins and lipids.21,22,26 However, in our case, this process seems to occur to a smaller extent and with a slower velocity. This fact could therefore be due to the action of anti-oxidation and detoxification mechanisms, that take place inside the cell to prevent the detrimental effect of AgNPs.
Over time, in Raman spectra, the appearance of a band at ∼638 cm−1 and the intensification of the one at ∼1572 cm−1 can be observed (Fig. 3). The band at ∼638 cm−1 corresponds to the vibrations of C–S bonds in peptides and proteins, originating from sulfur-containing amino acids such as methionine and cysteine.58,59 Biological molecules rich in sulfur-containing amino acids are usually involved in antioxidant and detoxification processes, and are synthesized by cells in order to reduce the harmful action of toxic compounds and ROS. The most important antioxidant molecules produced by the cells are the two enzymes superoxide dismutase (SOD) and catalase (CAT), the small molecule glutathione (GSH) and the cysteine-rich, low molecular weight family of proteins named metallothionein (MT; involved in the binding of metal ions to prevent them from catalyzing the production of ROS).25,28,71 All these molecules are rich in sulfur-containing amino acids, particularly cysteine, which are crucial for their structure and function. An increase in intensity relative to the C–S bond band can therefore be attributed to their increased synthesis due to the activation of a cellular defensive mechanism against the presence of exogenous AgNPs.22,29,72–75 The band at ∼1572 cm−1 can be instead attributed to the vibrations of bonds in the active sites of the two antioxidant enzymes SOD and CAT, which have been observed with similar laser sources. For the SOD enzyme it originates from the metal-bridging imidazole ring of histidine,65–67 while for CAT it comes from one vibration of the porphyrin skeletal structure.62–64 Therefore, the growth of this band during time additionally confirms the increased synthesis of the two antioxidant enzymes. Eventually, the temporal evolution of the two bands, fitted with an exponential curve, aligns well with the characteristic trends exhibited by biological processes.76 This reinforces the hypothesis regarding the correct assignment of signals to the ongoing biological processes.
To verify the contribution of the band at ∼1572 cm−1, we acquired the Raman spectra under 532 nm excitation of pure superoxide dismutase and catalase. The comparison between the Raman spectrum of the region at around 1572 cm−1 collected from the HDF cells exposed to AgNPs35nm for 12 h and the spectrum of the enzymes is shown in Fig. S4.† Although the enzymes are non-human-derived, for both proteins it is possible to observe a band at around 1572 cm−1, especially for the CAT enzyme, which further supports the hypothesis that this signal derives from the two antioxidants.
Being that SOD is the more abundant and ubiquitous antioxidant enzyme in cells,77,78 we quantified its total amount in cells after their exposure to AgNPs using the SOD activity assay. The SOD activity assay bases its function on the conversion of the tetrazolium salt WST-1 into a water-soluble formazan dye, upon reduction with the superoxide anion (O2−). The rate of reduction of WST-1 with the superoxide anion is linearly related to xanthine oxidase (XO) activity (the enzyme used to generate O2−), and this reduction is inhibited by the presence of SOD (Fig. 4). Therefore, a lower reduction of WST-1 into the formazan dye in the treated sample indicates the presence of a larger amount of the SOD enzyme, as SOD can catalyze the dismutation of O2− and inhibit WST-1 reduction. Fig. 4 shows the decrease with time of the conversion levels of WST-1 into the formazan dye in the samples treated with AgNPs. This observation indicates an increasing presence of SOD enzymes in the cells, which consequently prevents the formation of the superoxide anion and the conversion of WST-1. However, further investigations with immunoassay tests would be necessary to precisely quantify the overproduction of SOD and CAT enzymes.
![]() | ||
Fig. 4 Working mechanism of the SOD assay and levels of SOD expressed as the normalized absorbance intensity profile with respect to the non-treated sample (0 h). |
In addition, despite the high levels of SOD in the samples treated for 72 h, the viability of the cells was greatly reduced, as seen from the results of the MTT assay. This finding suggests that the cytotoxicity of AgNPs towards human cells may result from mechanisms other than ROS overproduction.29,79
In the literature, it has been reported that the band at 1572 cm−1 can result from vibrations of dissolved oxygen in the culture medium.48 However, during data processing, the average dissolved oxygen signal in culture media is uniformly subtracted from each sample through background subtraction. In contrast, other works assign this band to the vibrations of purine nucleobases, guanine and adenine. However, the absence of intensity variation for the band at ∼780 cm−1, related to the DNA phosphate backbone and pyrimidine bases (uracil, cytosine, and thymine),58,60,61 suggests that their vibrational contribution is only partial and that the increase in the intensity of the ∼1572 cm−1 band cannot be related to variations in the structure of the DNA/RNA. Furthermore, the fact that the ∼780 cm−1 band remains stable over time suggests that no genotoxic effect is induced by the presence of AgNPs35nm during the first 24 h of exposure. The absence of genotoxicity may also be attributed to the reduced presence of nanoparticle aggregates within the cell nuclei, which are mainly observed in the vesicles of the cytoplasmic space (Fig. 5).
Interestingly, no appearance of the band for the S–S bond at around ∼500 cm−1 occurs during the analysis. This band is generally associated with the formation of the disulfide bond between glutathione molecules, occurring during cellular antioxidative processes. This phenomenon could be caused by different factors: (i) the oxidation of glutathione is not among the predominant mechanisms for the reduction of oxidative stress; (ii) the GSH action occurs at times lower or higher than those taken into consideration.80
To confirm the action of the antioxidant molecules to regulate the formation of ROS, fluorescence microscopy was performed on HDF cells exposed to 10 μg mL−1 of AgNPs35nm for 8, 12, 24 and 72 h, using DCFH-DA as the fluorescent reporter molecule (Fig. 6). As expected, no ROS formation was observed for the cells treated with only culture media (negative control), while an increased signal resulted from those cultured in the presence of TBHP (100 μM) for 60 minutes. The cells treated with AgNPs35nm at the initial time points (8 and 12 h) showed the same conversion of DCFH-DA into the DCF fluorescent molecule as for control samples. This fact confirms that in the early stages of exposure to low concentration levels of AgNPs, the cellular antioxidant mechanism is able to control the detrimental effect of Ag+ ions and the overproduction of ROS. After 24 h, a slight increase in fluorescence intensity was observed, indicating that already at this time point the production of ROS begins to no longer be contained. Eventually, after 72 h it is possible to observe a large intensification in fluorescence, which indicates greater formation of ROS and consequently their potentially harmful effect. Nevertheless, the large standard deviation recorded for this time point suggests that several cells are not affected by the presence of AgNPs and thus do not produce ROS. As reported in the literature, ROS action could therefore not be the only mechanism that causes cytotoxicity of AgNPs towards human cells, and more studies should be carried out to confirm these hypotheses.29,79
A proposed defense mechanism of HDF cells in the presence of sub-lethal concentrations of AgNPs for short exposure times can be seen in Fig. 7 where: (i) initially, AgNPs are taken up by the cells and they enter the cellular environment; (ii) within the first 24 hours, the cells trigger their antioxidant system wherein metallothioneins, superoxide dismutase and catalase collaborate to effectively suppress the overproduction of ROS generated by the presence of AgNPs and Ag+ ions. This combined action successfully mitigates oxidative stress, contains the deterioration of proteins and lipids, and prevents damage to the DNA; (iii) as time progresses beyond 24 h, the physiological and cellular environments perpetrate the dissolution of the AgNPs, leading to an always increasing presence of Ag+ ions.22,81 Consequently, this sustained presence of Ag+ ions can induce an amplified generation of ROS. The increasingly hostile cellular environment overwhelms the initially effective antioxidant mechanism, ultimately resulting in its failure; (iv) after 72 hours of chronic exposure, the cumulative effect of heightened ROS production, coupled with the compromised defense system, is manifested in high cytotoxicity. In summary, the cytotoxic action of low concentration levels of AgNPs involves an initial defense response that succumbs to their prolonged exposure, leading to a cascade of events culminating in cytotoxicity after 72 h of chronic exposure.
HDF | Human dermal fibroblast |
AgNPs | Silver nanoparticles |
ROS | Reactive oxygen species |
SOD | Superoxide dismutase |
CAT | Catalase |
MT | Metallothionein |
GSH | Glutathione |
SERS | Surface enhanced Raman scattering |
TBHP | tert-Butyl hydroperoxide |
Footnote |
† Electronic supplementary information (ESI) available: Additional description of Raman spectra acquisition and processing steps, and magnification of specific parts of the time-dependent Raman spectra. See DOI: https://doi.org/10.1039/d4nr00462k |
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