**Preface to "The Two Faces of Nanomaterials: Toxicity and Bioactivity"**

Since the early 2000s, a growing number of nanomaterials (NMs) have received attention due to the advances in nanomedicine, including the use of various nanoparticles for therapeutic and diagnostic purposes. NMs have different properties compared with larger materials, and these properties can be used in a wide spectrum of biomedical areas, such as theragnosis, drug delivery, imaging, sensing, and tissue engineering. In this context, the safety (or toxicity) profile of NMs and their impact on health must be evaluated to attain their biocompatibility and desired activity for their development. However, certain controversies remain. Despite certain inconsistencies in several detailed experimental results from numerous reports, some in vitro and in vivo studies clearly showed no particular risks posed by NMs, whereas others have indicated that NMs might become health hazards.

In this book, two review papers and five original research works focus on a better understanding the correlation of the biological effects of NMs with their intrinsic physicochemical and thermomechanical properties. This book provides novel scientific findings on the bioactivity of NMs and some perspectives on potential risks to their future development in biomedical science and engineering as well as potential applications to various clinical fields.

> **Dong-Wook Han, Timur Sh. Atabaev** *Special Issue Editors*

### *Review* **Toxicity of Zero- and One-Dimensional Carbon Nanomaterials**

**Iruthayapandi Selestin Raja 1,**†**, Su-Jin Song 2,**†**, Moon Sung Kang 2, Yu Bin Lee 2, Bongju Kim 3, Suck Won Hong 2, Seung Jo Jeong 4, Jae-Chang Lee 5,\* and Dong-Wook Han 2,\***

Received: 5 August 2019; Accepted: 23 August 2019; Published: 28 August 2019

**Abstract:** The zero (0-D) and one-dimensional (1-D) carbon nanomaterials have gained attention among researchers because they exhibit a larger surface area to volume ratio, and a smaller size. Furthermore, carbon is ubiquitously present in all living organisms. However, toxicity is a major concern while utilizing carbon nanomaterials for biomedical applications such as drug delivery, biosensing, and tissue regeneration. In the present review, we have summarized some of the recent findings of cellular and animal level toxicity studies of 0-D (carbon quantum dot, graphene quantum dot, nanodiamond, and carbon black) and 1-D (single-walled and multi-walled carbon nanotubes) carbon nanomaterials. The in vitro toxicity of carbon nanomaterials was exemplified in normal and cancer cell lines including fibroblasts, osteoblasts, macrophages, epithelial and endothelial cells of different sources. Similarly, the in vivo studies were illustrated in several animal species such as rats, mice, zebrafish, planktons and, guinea pigs, at various concentrations, route of administrations and exposure of nanoparticles. In addition, we have described the unique properties and commercial usage, as well as the similarities and differences among the nanoparticles. The aim of the current review is not only to signify the importance of studying the toxicity of 0-D and 1-D carbon nanomaterials, but also to emphasize the perspectives, future challenges and possible directions in the field.

**Keywords:** carbon nanomaterials; unique properties; biomedical applications; in vitro toxicity; in vivo toxicity

#### **1. Introduction**

Nanotechnology has been a rapidly developing field, producing many nanomaterials with alterations in different physical and physicochemical properties such as size, shape, crystalline nature, and interaction with biological systems [1–3]. These materials have found adaptability in biomedical applications such as nanomedicines, cosmetics, bioelectronics, biosensors, and biochips [4]. However, the fact that possible health risks are associated with the increasing development of nanotechnology cannot be set aside. Nanoparticles may be either organic or inorganic based on the composition of elements. Mostly, inorganic nanomaterials are based on transition metals such as silver, iron, gold, zinc, copper, etc. whereas carbon nanomaterials are mainly composed of the carbon element, which constitutes various spatial arrangements in different nanoscales from zero (0-D) to three dimensions (3-D) [1,5–7]. In the present review, we will discuss the toxicity of 0-D carbon nanostructures (carbon

black, nanodiamond, carbon nanodots and fullerene) and 1-D nanomaterials (single and multi-walled carbon nanotubes) from the research that has been conducted over the past two decades. The structure of carbon nanomaterials is shown in Figure 1.

Carbon dots are carbon-based nanomaterials with unique properties such as chemical inertness, optical stability, and wavelength-dependent photoluminescence [8]. Carbon quantum dots (CQDs) are typically quasi-spherical nanoparticles with a diameter less than 10 nm and composed of carbon, oxygen, hydrogen, nitrogen, and other elements. Because of their hydrophilic nature and cell permeation, CQDs have replaced traditional metal-based quantum dots in many applications, including photovoltaics, photocatalysis, and drug targeting [9]. The oxidized CQDs may contain 5–50% oxygen depending on synthetic procedures. Carbon quantum dots typically present two optical absorption bands in the UV-vis spectrum, which are attributed to π–π\* and n–π\* transitions in C=C and C=O bonds, respectively [10]. When the carbon nanodots are represented as a π-conjugated single sheet, with a size of 2–10 nm, they are called graphene quantum dots [11]. It has been reported that graphene quantum dots (GQDs) exhibit magnetic, electronic, and optical properties [12].

Nanodiamonds (NDs) are carbon-based crystalline nanoparticles inheriting diamond structure at the nanoscale with excellent properties such as optical transparency, hardness and chemical inertness [13]. The sp3 tetrahedral structure of the nanodiamond presents Raman signal at 1332 cm−1 and is capable of fluorescing due to point defects. However, the non-fluorescing nanodiamond displays a strong coherent anti-Stokes Raman scattering effect [14]. The quantitative analysis of cellular uptake of NDs is promising for the applications of bio labeling and bio imaging. The oxidized form of the nanodiamond has been reported to damage DNA in embryonic stem cells [15].

Carbon black nanoparticles (CBNPs) are the zero-dimensional carbon-based nanomaterials, which are produced in large quantities in different ways, such as partial combustion and thermal decomposition of hydrocarbons either in liquid or gaseous state [16]. The poor water-soluble carbon black poses a threat to health when exposed to the lungs through inhalation. The core portion of the insoluble particle yields reactive oxygen species (ROS), which render toxicity to the experimental animals [17]. Recently, the International Agency for Research on Cancer (IARC) listed carbon black nanoparticles as carcinogenic to human beings [16]. In toxicological studies, carbon black nanoparticles (CBNPs), with diameters less than 100 nm, have been reference material for diesel exhaust particles [18]. The aciniform aggregates of carbon black are basically fine powder in the size range of 100–1000 nm in a closed reaction chamber and form larger agglomerates due to van der Waals forces in the final step of the manufacturing process [19]. The term 'carbon black' should not be confused with such words as black carbon and soot, which are the carbonaceous materials emitted from incomplete combustion of fuels, such as waste oil, diesel, gasoline, wood, paper, plastic and rubber [20]. It is important to note that carbon black nanoparticles have certain physicochemical properties in common with another insoluble carbonaceous material, including graphene [16]. CBNPs have been widely used as conductive fillers due to their low aspect ratio, being economically inexpensive, and having good conductivity [21,22].

Among the carbon-based nanomaterials, fullerene (C60) is a generic term for a cluster composed of 60 carbon atoms that appears as a soccer-ball structure. The C60 contains 30 carbon atoms to readily interact with free radicals, and therefore is known as a free radical sponge [23,24]. The versatile applications of C60 include use in superconducting devices, energy device materials and catalysts [25]. The water-soluble polyhydroxylated fullerene, known as fullerenol (C60(OH)n), has been explored for its potential as being an anticancer, anti-HIV and skin rejuvenating cosmetic [25,26]. Fullerenol was reported to protect experimental animals from hepatotoxicity and doxorubicin-induced cardiotoxicity [26,27]. In nature, fullerene is available as its analogues including C70, C80, and C94, because of its tendency to aggregate and form a crystal-like structure with a diameter of 100 nm [23]. The research studies revealed that skin contact and nasal inhalation are the most likely routes of exposure to fullerenes for the workers in industries [25].

**Figure 1.** The structure of zero- and one-dimensional carbon nanomaterials have been shown. Carbon quantum dot (CQD) and graphene quantum dot (GQD), reproduced with permission from [11], Copyright Royal Society of Chemistry, 2010; nanodiamond (ND) and fullerene (C60), reproduced with permission from [7], Copyright American Chemical Society, 2013; carbon black nanoparticle (CBNP), reproduced with permission from [28], Copyright Elsevier, 2014; single-walled carbon nanotube (SWCNT) and multi-walled carbon nanotube (MWCNT), reproduced with permission from [29], Copyright Elsevier, 2017.

The unique property of CNT is its high aspect ratio, which promotes its superior properties to the encapsulating matrix polymers and has advantages over traditional reinforcements [30]. The most widely used techniques for the synthesis of carbon nanotubes (CNTs) are laser furnace, chemical vapor deposition, and arc discharge [31]. Their biomedical applications include biosensors, orthopedic prostheses, anticancer therapy, and tissue engineering [32]. The literature reports reveal that maternal exposure of CNTs might develop developmental toxicity such as teratogenicity [33]. The threat of nanotoxicity of CNTs is an increasing trend, as the global production of CNTs reaches several thousand tons per year [32]. Based on morphology, the carbon nanotube is generally classified into the two viz. single-walled and multi-walled carbon nanotubes. When one or several graphene sheets are rolled up to a cylindrical form concentrically, they yield single-walled carbon nanotubes (SWCNTs) and multi-walled carbon nanotubes (MWCNTs), respectively. Meanwhile, MWCNTs differ from SWCNTs in some physicochemical properties, such as the number of layers, the surface area and width [34,35]. The preparation of both CNTs also varies with different experimental conditions. For example, in the electric arc discharge method, SWCNTs are synthesized in the form of soot when a graphite rod comprising a metal catalyst acts as an anode and pure graphite as a cathode. Meanwhile, the production of MWCNTs is achieved strictly in the presence of inert gas such as helium. In the laser vaporization method, generation of SWCNTs mainly depends on the type of metal catalyst and the furnace temperature, whereas the yield of MWCNTs requires a pure graphite target and an optimum temperature of 1200 ◦C [36]. The nanotubes strongly interact with each other by van der Waals forces and hence exhibit hydrophobicity, which limits their biomedical applications. Hypochlorite, myeloperoxidase, and eosinophils peroxidase have been reported to degrade nanotubes within phagosomes and in the inflammation sites [37]. Researchers have adopted different approaches to modify pristine CNTs to impart hydrophilic behavior. The π-conjugated skeleton of CNT was covalently modified through different chemical reactions such as sidewall halogenation, hydrogenation, plasma activation, cycloaddition, radical, nucleophilic and electrophilic additions. The non-covalent modification occurs by physical attachment of various functional molecules and the endohedral filling takes place at the inner empty cavity of CNT [38].

SWCNTs have been used in a wide range of commercial applications such as earthquake-resistant buildings, dent-resistant car bodies, stain-resistant textiles and transistors [39]. The diameter of SWCNTs is approximately 1–2 nm and their toxicity is more substantial in comparison to MWCNTs (10–20 nm) and other carbonaceous nanomaterials such as carbon black and fullerene [40]. Despite being an attractive structural material with a high aspect ratio of length to width, carbon nanotubes threaten living organisms with potentially hazardous effects [41]. As far as the drug administration of SWCNTs is concerned, the inhalation route of exposure has more serious effects than the aspiration route in terms of oxidative stress, inflammatory responses, fibrosis and collagen deposition [42]. It has been reported that the agglomerates of SWCNTs caused granulomas in the proximal alveoli, and dispersed SWCNTs instigated interstitial fibrosis in the distal alveoli [43]. Similar to asbestos, MWCNTs have been reported to possess pathogenicity, owing to their larger durability and needle-like shape [32]. They found a wide variety of industrial applications in rechargeable batteries, water filters and sporting goods [44]. It was informed that non-branched MWCNTs had a higher potential to cause mesothelioma than the tangled MWCNTs [45].

#### **2. In Vitro Cellular Toxicity of Zero- and One-Dimensional Carbon Nanomaterials**

The in vitro toxicity effects of carbon nanomaterials (0-D and 1-D) have been listed in Table 1. The cytotoxic effect of the polyethylenimine (PEI) coated CQDs based nanohybrid, with a diameter of 6.5 ± 2 nm, was investigated at various concentrations (200, 400, 600 and 800 μg/mL) on kidney epithelial cells derived from the African green monkey. The MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay revealed that the nanohybrid killed 39% of cells at concentration 600 μg/mL, despite there being no sign of significant toxicity at lower concentrations [46]. The pristine fluorescent carbon quantum dots (~7 nm) were evaluated for its cytotoxicity assessing total ROS, glutathione, and lactate dehydrogenase activity on human bronchial epithelial cells (16 HBE). The data revealed that CQDs preferentially located on the surface of cells and that its exposure induced oxidative stress and decreased cell viability [47]. A comprehensive study was presented to describe the critical role of functionalized nanoparticles in cytotoxicity using mouse embryonic fibroblasts (NIH-3T3). The CQDs synthesized from candle soot were negatively charged. The pristine CQDs were then functionalized with PEG (polyethylene glycol) and PEI to impart neutral and positive charges on the surface of nanoparticles, respectively. The results of in vitro cellular toxicity measurements revealed that the neutral charged CQDs did not induce any abnormalities in the cell cycle, cellular trafficking and cell morphology up to the concentrations of 300 μg/mL. Meanwhile, the negatively charged pristine CQDs arrested the cell cycle at the G2/M phase, enhanced cell proliferation, and caused oxidative stress. Being the most cytotoxic, the positively charged CQDs triggered a significant alteration in the cell cycle at the G0/G1 phase, at a concentration of 100 μg/mL [48].

GQDs have also shown different cellular uptake in MC3T3 osteoblast cell lines derived from mouse calvaria and exhibited low cytotoxicity due to their small size and high oxygen content [49]. The adverse effects of hydroxyl-modified GQDs (OH-GQDs) were studied on human lung carcinoma cell lines H1299 and A549. The OH-GQDs with hydrodynamic diameter of 10.3 ± 1.9 nm, at a concentration 50 μg/mL, decreased cell viability and intracellular ROS generation at a significant level. The cell signaling pathway analysis exposed that hydroxylated GQDs induced G0/G1 arrest, cell senescence, and inhibition of Rb phosphorylation in both types of cells [50].  The nanoparticles significantly permeated into both cytoplasm and nucleus of the cells following caveolae-mediated endocytosis, but they did not affect cellular morphology. In addition, the nanoparticles exhibited lower cytotoxicity to MGC-803 cells when compared to MCF-7 cells [51].

Genotoxicity of NDs was analyzed on mouse embryonic stem cells and the results revealed that NDs of 4–5 nm expressed an elevated level of DNA repair proteins such as p53 and MOGG-1. Further, oxidized NDs were described to have more influence on triggering DNA damage than the pristine NDs. However, it was demonstrated that NDs, either in oxidized form or pristine, were not severe in toxicity when compared to MWCNTs [52]. Intracellular ROS, mitochondrial activity, apoptosis, colony formation, and cellular uptake were studied to provide elucidative information

about the toxicity of NDs in two different cell lines HaCaT and A549. At concentration of 1.0 mg/mL, inhibition of colony formation and small degree apoptosis were observed in cells. However, it was found that NDs did not have any significant influence on cell viability and ROS production [53]. Treated with RAW 264.7 murine macrophages, the cytotoxicity of NDs were examined in various sizes (6–500 nm) and concentrations (0–200 μg/mL). Cell proliferation and metabolic activity were found reduced in a concentration dependent manner. Flow cytometry analysis revealed that the nanoparticles caused necrosis, leading to significant cytotoxicity, irrespective of particle size [54]. In vitro toxicity measurements were carried out in human blood cells and the reports exposed that NDs could change the kinetics of active oxygen production, cause erythrocyte hemolysis and destruct white cells [55].

The in vitro genotoxic and mutagenic potential of NDs were investigated in human lymphocytes and the nanoparticles were reported to inhibit cell proliferation-inducing apoptotic cell death above 50 μg/mL. The cellular oxidative stress generated by the nanoparticles was found to be dose-dependent. Significant changes in chromatin stability followed by DNA oxidative damage were established, even at a concentration of 1 μg/mL. NDs had the potential to stimulate micronuclei augmenting centromeric signals at 10 μg/mL [56]. The viability of human umbilical vein endothelial cells (HUVEC-ST) was investigated following the treatment of NDs, which was synthesized by the detonation method. The results of the MTT assay revealed that NDs showed a concentration-dependent cytotoxicity and ROS production in cells [57]. In a study, the cytotoxicity effect of nanodiamond particles was explored by correlating different surface functional groups on the nanoparticles, such as –OH, –COOH and –NH2. It was shown that NDs were cytotoxic to HEK293 cells when the concentration was above 50 μg/mL. The cationic nanodiamond had the potential to permeate negatively charged cell membrane and hence exhibited cytotoxicity. In addition, carboxylated nanodiamond (ND–COOH) was reported to possess embryotoxicity as well as teratogenicity [58].

The in vitro toxicity effect of CBNPs (260 ± 13.7 nm) was evaluated on A549 human alveolar basal epithelial cells and suggested that ultrafine particles induced a greater oxidative stress with prolonged inhibitory effects than fine particles [59]. Printex 90, a commercial name of carbon black nanoparticles with a diameter of 14 nm, exhibited an oxidative damage response in HepG2 cells at 25 mg/L, which was revealed from formamidopyrimidine DNA glycosylase (Fpg)-modified comet assay [60]. In another comet (Fpg) assay, it was discovered that an increased level of oxidized purines was observed when the nanoparticles were investigated in the FE1-MML Muta Mouse lung epithelial cell line. The mutant frequency was noticed in carbon black exposed cells following eight repeated 72 h incubations with a cumulative dose of 6 mg nanoparticles [61]. The western blot analysis exposed that ultrafine carbon black nanoparticles, at 30.7 μg/cm2, stimulated proliferation of human primary bronchial epithelial cells through oxidative stress and epidermal growth factor-mediated signaling pathway [62]. The cytotoxic and genotoxic effects of CBNPs were investigated on the mouse macrophage cell line RAW 264.7. The particle size and specific surface area was 14 nm and 300 m2/g, respectively. The data confirmed acentric chromosome fragments at all concentrations and there was a slight increase in micronuclei frequencies at 3 and 10 mg/L [63]. It was reported that CBNPs (100 μg/mL) could induce DNA single-strand breaks and induce AP-1 and NFκB DNA binding in A549 lung epithelial cell line after 3 h of exposure [64]. The toxicity measurements of CBNPs in THP-1 derived monocytes and macrophages exemplified that the nanoparticles supported endothelial activation and lipid accumulation in THP-1 derived macrophages. In addition, the nanoparticles influenced increased cytotoxicity, LDH levels and intracellular ROS production in a dose-dependent manner [65].

It was discovered that C60 fullerene of approximately 0.7 nm was less toxic than carbon black and diesel exhaust particles when FE1-MutaMouse lung epithelial cells were exposed to nanoparticles. The results of the comet assay revealed that C60 significantly increased the quantity of formamidopyrimidine-glycosylase sites (22%) and oxidized purines (5%), though the nanoparticles did not involve breaking DNA strands [66]. Genotoxic effects of C60 sized 0.7 nm were investigated by micronuclei test in the human lung cancer cell line (A549) at a concentration range of 0.02–200 μg/mL and increased micronuclei frequencies were observed in nanoparticles treated cells in a dose-dependent

manner [67]. The genotoxic studies of colloidal C60 in human lymphocytes had shown genotoxicity at 2.2 μg/L, whereas the ethanolic solution of C60 had exhibited the same at 0.42 μg/L [68]. The polyhydroxylated C60 fullerenol presented a dose-dependent decrease in micronuclei frequency and chromosome aberration when the nanoparticles were treated with Chinese hamster ovary cells (CHO K1). However, the study did not show any genotoxic effects in the concentrations of 11–221 μm [27]. The cytotoxicity of hydroxylated fullerene was analyzed in vascular endothelial cells at different concentrations, 1–100 μg/mL, and a dose-dependent decrease in cell viability was perceived. Furthermore, it was reported that fullerenes affected cell growth and cell attachment with the potential to cause cardiovascular disease after a long period of exposure (10 days) [69].

The toxicity effect of SWCNTs was explored on human embryonic kidney cells (HEK293T) and reported that the nanoparticle exposure resulted in a decrease in cell adhesion, inhibition in cell proliferation and induction in apoptosis, depending on the dosage and time. In addition, a nodular structure was formed due to the nanoparticle aggregation and overlap of cells [70]. The agglomeration of CNTs had a larger impact on triggering cellular toxicity in human MSTO-211H cells. It was found that the agglomerated CNTs were more toxic compared to monodispersed CNTs [71]. The geometric structure of the nanoparticles played a pivotal role in determining cytotoxicity. A comparative study was provided in describing cytotoxicity of SWCNTs, MWCNTs, and C60 fullerenes on guinea pig alveolar macrophages. The order of displaying toxicity was as follows, SWCNTs > MWCNTs > C60 fullerenes [72]. The intracellular distribution of functionalized SWCNTs was studied in murine 3T3 and human 3T6 fibroblast cells. The length of the nanotube varied from 300 to 1000 nm and the outer diameter was 1 nm. The analyses revealed that SWCNTs resided either in the cytoplasm or nucleus after crossing the cell membrane, and exhibited toxicity when the concentration of nanoparticles reached above 10 μM [73]. It was confirmed that exposure of SWCNTs induced cutaneous and pulmonary toxicities in human bronchial epithelial cells (BEAS-2B) and human keratinocyte cells (HaCaT). The microarray analysis revealed that the nanoparticles triggered alteration of genes followed by transcriptional responses. Cellular morphology, integrity and ultrastructure were affected as the nanoparticles depleted antioxidants in the cells [74,75]. Functionalization of the nanoparticles had taken advantage of reducing the toxic level of nanoparticles. The derivatized SWCNTs were reported to have fewer toxic effects than pristine SWCNTs from in vitro cytotoxicity measurements in human dermal fibroblasts [76]. The introduction of SWCNTs into normal and malignant human mesothelial cells produced ROS causing cell death, DNA damage and H2AX phosphorylation [77]. It was reported that SWCNTs, with a primary particle size of 0.4–1.2 nm and specific surface area of 26 m2/g, had the potential to induce DNA damage in lung V79 fibroblasts [78].

The cytotoxic and genotoxic effects of single and multi-walled CNTs were studied on the mouse macrophage cell line RAW 264.7, and it was demonstrated that the exposure of nanoparticles stimulated ROS release, chromosomal aberrations, necrosis, and apoptosis, but they did not cause any inflammatory responses. In addition, MWCNTs were reported to penetrate the cell membrane and reside in the nuclear envelope [63]. Electron microscopic studies indicated that highly purified MWCNTs expressed higher cytotoxic effects by damaging the plasma membrane of mouse macrophages (J774.1). It was found that the cytotoxicity of MWCNTs was significantly larger than crocidolite, a fibrous form of sodium iron silicate [79]. The higher concentrated MWCNTs caused a decrease in cellular viability and an increase in inflammation on prolonged exposure to human epidermal keratinocytes (HEK) cells. The nanoparticles had the potential to penetrate the cell membrane and change the expression level of various proteins. The nanoparticles were reported to be abundantly present within cytoplasmic vacuoles of the cells after cell permeation [80]. The toxicity of MWCNTs of approximately 30 nm was evaluated in human skin fibroblasts (HSF42) and the results revealed that the nanoparticles disrupted intracellular signaling pathways, causing an increase in apoptosis and necrosis, and activated the genes associated with cellular cycle regulation, metabolism, cellular transport, and stress response [81]. Interestingly, oxidized MWCNTs were described to exhibit more toxicity than pristine MWCNTs. Both were reported to induce apoptosis in T lymphocytes depending on the time period and dose [82].

**Table 1.** The in Vitro Toxicity Effects of 0-D and 1-D Carbon Nanomaterials.

**Table 1.** *Cont*.

Abbreviations: PS, particle size; HD, hydrodynamic diameter; SSA, specific surface area; L, length; W, width; n/a, not available.

#### **3. In Vivo Toxicity of Zero-and One-Dimensional Carbon Nanomaterials**

In some studies, the researchers performed in vivo animal studies of carbon nanomaterials after the careful evaluation of their in vitro toxicity measurements, and some of studies are listed in Table 2.

**Table 2.** *Cont*.

**Table 2.** *Cont*.

Abbreviations: PS, particle size; IS, interlayer spacing; HD, hydrodynamic diameter; GMD, geometric mean diameter; SSA, specific surface area; L, length; W, width; n/a, not available.

The toxicity of carbon quantum dots was investigated in different species such as zebrafish, zooplankton, and phytoplankton. The primary particle size was less than 10 nm, with interlayer spacing of 0.32 nm. It was found that zooplankton was more sensitive to CQDs than zebrafish and phytoplankton species and suffered oxidative stress, water acidification, insufficiency of nutrients and no photosynthesis in a time and dose-dependent manner [83]. When the nanoparticles were administered intravenously to ICR male and female mice with a single dose, it was observed that male mice are more sensitive than female mice, and that the nanoparticles treated male mice suffered severe acute inflammatory responses [84]. The intraperitoneal injection of CQDs (8 ± 2 nm) into male ICR mice affected cell membrane, immune system and liver clearance rate [8]. While investigating the in vivo toxicity of CQDs (2–6 nm) in embryos/larvae of male and female rare minnow, concentration-dependent embryos yolk agglutination, decreased spontaneous movements, and increased heart rate were observed [9]. The toxicity studies of GQDs in AB strains of wild-type zebrafish embryos/larvae revealed that the nanoparticle had the potential to decrease heart rate, causing disrupted embryonic development in a concentration dependent manner. However, the treatment of nanoparticles did not have significant toxicity at lower doses [85,86]. The toxicity of functional GQDs in an animal model was studied to understand the influence of functional groups attached

on the surface of nanoparticles. The polyethylene glycol modified GQDs (PEG-GQDs) exhibited no significant toxicity when the nanoparticles were instilled intraperitoneally into female BALB/c mice [87]. Likewise, carboxylated GQDs (COOH-GQDs) triggered no obvious damage to SD rats after 21 days of intravenous post-administration [88].

The microinjection of NDs (0.5 mg/mL) to wild type young Caenorhabditis elegans had shown no detectable toxicity in brood size and longevity of animals. The hydrodynamic diameter of the nanoparticles in solution was approximately 120 nm [89]. When NDs of approximately 4 nm were intratracheally injected into male ICR mice at a concentration of 1.0 mg/kg, the nanoparticles produced lung burden during the whole exposure time, but there was no event of lipid peroxidation in lung tissue [90]. A dose-dependent toxicity was observed in the lung tissue of male Kun Ming mice after the NDs were intratracheally administered at different concentrations 0.8, 4.0 and 20 mg/kg [13]. While investigating possible toxicity of bovine serum albumin functionalized nanodiamond (ND-BSA, ~100 nm) in AB strain zebrafish embryos at a concentration range of 1–5 mg/mL and 4–96 h post-fertilization (hpf), it was found that the control and NDs treated groups had no significant differences in embryonic development at concentration of 1 mg/mL. However, a higher concentration of NDs affected the pharyngula stage of embryos and caused fin curve in larvae during the hatching stage [14].

There were many reports that demonstrated the toxicity of carbon nanomaterials in animal models, which included pulmonary inflammation, DNA breaks, oxidative stress and elevated expression of mRNAs [17,91–106]. The intratracheally administered CBNPs (67 μg/animal) to female pregnant mice did not trigger significant germline mutation when compared to the control [107]. When the rats were exposed to 7.1 and 52.8 mg/m3 of CBNPs for 13 weeks, a significant dose-dependent increase in hypoxanthine-guanine phosphoribosyltransferase (hprt) mutation frequency was observed in rat alveolar epithelial cells. The nanoparticles impaired lung clearance, causing lung burden, and changed the expression of bronchoalveolar lavage fluid (BALF) markers of inflammation and lung injury [108]. Various immunohistochemical measurements were established to quantify DNA damage markers such as poly (ADP-ribose), 8-hydroxyguanosine, and 8-oxoguanine DNA glycosylase after intratracheally instilling CBNPs into rats for 3 months. The analyses revealed that the nanoparticles had significantly increased the expression of DNA damage markers, though the genotoxicity was less pronounced [109]. Genotoxic effects, acute phase and inflammatory responses were examined while exposing C57BL/6JBomTac mice to CBNPs. Even at low exposure doses of nanoparticles (0.67, 2, 6 μg), an increase in DNA strand breaks occurred in bronchoalveolar lavage (BAL) cells. It was reported that DNA damage was triggered by primary genotoxicity without inflammatory responses [110]. The pulmonary toxicity of carbon black nanoparticles was studied in C57BL/6 female mice administering a single dose of 0.162 mg. An increase in expression of miRNAs such as miR-135b, miR-21, and miR-146b, which are associated with pulmonary inflammation, was observed [111]. The polycyclic aromatic hydrocarbon modified CBNPs (PAH-CBNPs) were demonstrated to express the noticeable amount of keratinocyte chemoattractant and IL-6 mRNA, when compared to uncoated CBNPs and air control when male Wistar rats were subjected to nasal inhalation exposure for 2 weeks at a concentration of 6 mg/m3. The primary particle size and specific surface area of functionalized CBNPs was 14.2 ± 0.1 and 115 ± 3 m2/g, respectively [94].

The toxicity of fullerene of 96 nm was studied after subjecting male Wistar rats to whole-body inhalation for 4 weeks. The experiment was carried out for 6h/day with the exposure of 0.12 mg/m3. No significant changes were reported in the gene expression of CINC-1, CINC-2αβ, and CINC-3 in lung tissue [95]. In another similar study, the upregulation of genes associated with inflammation, oxidative stress and apoptosis was noted after one month of nanoparticle exposure. The geometric mean diameter of fullerene nanoparticles was 96 nm and specific surface area of them was 0.92 m2/g [96]. The intratracheal instillation of C60 to gpt delta transgenic mice at a single dose of 0.2 mg/mouse induced mutant frequencies with 2–3-fold increase in comparison to the control. When administered at multiple doses (4 times), the nanoparticles brought about transversion of A:T to T:A in treated animals [67]. The

intratracheally instilled C60 (46.7 ± 18.6 nm) increased the expression of pro-inflammatory cytokines including tumor necrosis factor-α (TNF-α), interleukins (IL-1 and IL-6) and T-cell distribution in ICR male mice [97]. It was demonstrated that single oral intragastric administration of fullerene to female Fisher 344 rats generated oxidative damage along with the expression of mRNA 8-oxoguanine DNA glycosylase (8-oxodG) in the lung at high dose [98]. No acute oral toxicity was reported for the C60 treated Sprague-Dawley male and female rats for 2 weeks [26]. The intratracheally administered fullerenol (C60(OH)n) showed increased neutrophil influx in the lungs causing inflammation in BALB/c female mice after 24 h of post-administration of 200 μg/mouse [99].

The DNA damage was examined in rats following intragastric instillation of SWCNT at a concentration of 0.64 mg/kg body weight. SWCNTs were demonstrated to elevate the levels of 8-oxodG in liver and lung tissues of rats. The length and width of the nanoparticles was less than 1 μm and 0.9–1.7 nm, respectively [98]. The aortic mitochondrial alteration was studied using oxidative stress assays in SWCNTs exposed C57BL/6 mice. The intra-pharyngeal instilled SWCNTs (40 μg/mouse) activated heme oxygenase 1, which is indicative of oxidative stress. The nanoparticles exhibited increased mitochondrial DNA damage accompanied by the changes in aortic mitochondrial protein carbonyl and glutathione levels [112].

The general toxicity effects of MWCNTs were inflammation, granuloma and fibrosis when in vivo toxicity measurements were performed in experimental animals [103,104,106]. The induction of mesothelioma in p53+/− mouse was studied by the intraperitoneal application of multi-wall carbon nanotube. It was found that intraperitoneally administered, micro-sized MWCNTs (10–20 μm) stimulated mesothelioma such as the positive control, crocidolite [113]. The immune and inflammatory responses of MWCNTs were tested following intraperitoneal administration of a single dose of 2 mg/kg body weight to female ICR mice. After 1 week of post-exposure, the expression of leukocyte adhesion molecules and cluster of differentiation on granulocytes were found increased. The number of monocytes, leukocytes, and granulocytes were also present in peripheral blood significantly. MWCNTs were reported to exhibit sustained immune responses with the overexpressed ovalbumin specific IgG1 and IgM. The original morphology of the liver had also suffered changes to a rounded shape along with the appearance of MWCNTs on internal organs [114].

#### **4. Conclusions and Perspectives**

In this review, we have discussed the toxicity effects of 0-D and 1-D carbon nanomaterials in different cell lines and animal models. It was demonstrated that differential toxicity of carbon nanomaterials was inherited from various factors such as size, dispersion, cell permeability, and functionalization. Though the researchers studied the toxicity of carbon nanomaterials in both in vitro and in vivo intensively, there are still some issues to be addressed. (1) Many researchers showed experimental results with the aim of comparing the toxicity of two or more carbon-based nanoparticles for the same cell line and animal model. A comparative study is required for different cell line sources and animal species for the same kind of nanoparticle. (2) There are many studies that emphasize the role of the encapsulating agents on the nanoparticles in altering the overall functionality. The differential toxicity depending on the charge on the surface of nanoparticles has also been demonstrated. However, a systematic study is needed to corroborate the toxicity results with the surface charge of the nanoparticles (either positive or negative) with subtle differences. (3) The toxicity studies of the same kind of carbon nanoparticle prepared from different techniques should also be examined. The following suggestions are put forth for future research in this field: (1) A comprehensive study on the toxicity of carbon nanomaterials using different physicochemical and biological parameters to exemplify toxicity limitation and prove the effectiveness of the materials. (2) A systematic study to ensure that the carbon nanoparticles exhibit toxicity towards cancerous cells but not normal cells at the established concentration range. Undoubtedly, the knowledge of the toxicity of carbon nanomaterials will help the researchers with interdisciplinary backgrounds to deliver more successful biocompatible materials to society in the future.

**Author Contributions:** I.-S.R. and D.-W.H. developed the idea and structure of the review article. I.S.R. and S.J.S. wrote the paper using the materials supplied by M.S.K., Y.B.L., B.K., S.W.H., and S.J.J., J.-C.L. revised and improved the manuscript. D.W.H. supervised the manuscript. All the authors have given approval to the final version of the manuscript.

**Funding:** This research was supported by the Bio & Medical Technology Development Program of the National Research Foundation (NRF) funded by the Korean government (MEST, No. 2015M3A9E2028643), Ministry of Trade, Industry and Energy (MOTIE, Korea, No. N0002310 and Technology Innovation Program No. 20000397), and Korea Research Institute of Chemical Technology (KRICT, Daejeon and Ulsan, Korea) and Ulsan City (SI1941-20, BS.K19-251).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Effect of Surface Coating of Gold Nanoparticles on Cytotoxicity and Cell Cycle Progression**

#### **Qian Li, Chun Huang, Liwei Liu, Rui Hu \* and Junle Qu**

Key Laboratory of Optoelectronic Devices and Systems of Ministry of Education and Guangdong Province, College of Optoelectronic Engineering, Shenzhen University, Shenzhen 518060, China<EMAIL_ADDRESS>(Q.L<EMAIL_ADDRESS>(C.H<EMAIL_ADDRESS>(L.L<EMAIL_ADDRESS>(J.Q.)

**\*** Correspondence<EMAIL_ADDRESS>Tel.: +86-0755-2673-3319; Fax: +86-0755-2653-6237 Received: 26 November 2018; Accepted: 12 December 2018; Published: 17 December 2018

**Abstract:** Gold nanoparticles (GNPs) are usually wrapped with biocompatible polymers in biomedical field, however, the effect of biocompatible polymers of gold nanoparticles on cellular responses are still not fully understood. In this study, GNPs with/without polymer wrapping were used as model probes for the investigation of cytotoxicity and cell cycle progression. Our results show that the bovine serum albumin (BSA) coated GNPs (BSA-GNPs) had been transported into lysosomes after endocytosis. The lysosomal accumulation had then led to increased binding between kinesin 5 and microtubules, enhanced microtubule stabilization, and eventually induced G2/M arrest through the regulation of cadherin 1. In contrast, the bare GNPs experienced lysosomal escape, resulting in microtubule damage and G0/G1 arrest through the regulation of proliferating cell nuclear antigen. Overall, our findings showed that both naked and BSA wrapped gold nanoparticles had cytotoxicity, however, they affected cell proliferation via different pathways. This will greatly help us to regulate cell responses for different biomedical applications.

**Keywords:** cell cycle; nanoparticle location; surface biocompatibility; microtubule; proteomics

#### **1. Introduction**

As the engineering of nanoparticles has been extensively developed over the past decades, various nanoparticles with unique physical and chemical properties have been designed for potential medical applications [1,2]. Improving our understanding of the interactions between nanoparticles and biological systems, especially at the cellular level, is crucial for their risk control and for evaluating their potential applications as drug delivery vehicles or therapeutic agents [3].The study of interactions between nanoparticles and biological systems, with an emphasis on elucidating the relationship between the physicochemical properties of nanoparticles and biological responses, is essential [4,5]. Such studies are important prerequisites for designing and engineering nanoparticles with intentionally enhanced or suppressed cellular responses and toxicity. However, the mechanisms mediating cellular responses to nanoparticles remain unclear, particularly about the effects of nanoparticles on cell cycle arrest at different phases.

As the cell cycle is closely related to cell proliferation and cytotoxicity, elucidating the mechanisms of different nanostructures on the regulation of cell cycle will be of great importance [6].Nanoparticles have been shown to cause cell cycle arrest, including G2/M and G0/G1 arrest. The type and extent of cell cycle arrest varies depending on the composition, size, size distribution, surface modification, and subsequent surface derivatization of nanoparticles [7–9]. G0/G1 arrest can be caused by DNA damage and microtubule damage, while nanoparticles in combination with oxidative stress and/or lysosome rupture could lead to G0/G1 arrest. However, the mechanisms and the factors behind the G2/M cell cycle arrest caused by nanoparticles are still unclear. Recently, Mahmoudi et al. speculated that the effects of nanoparticles on the cell cycle may depend on the intracellular location of the

nanoparticles [6]. Additionally, Choudhury et al. reported that gold nanoparticles (GNPs) with lysosomal escape ability localized to the tubulin/microtubule system and caused cell cycle arrest at G0/G1 phase through induction of microtubule damage [10]. However, whether the intracellular localization of nanoparticles is linked with G2/M cell cycle arrest is still unknown.

GNPs have been recognized as promising nanoprobes in biomedical applications for clinical translation. Although they were once believed to be biocompatible, they are now known to cause cell cycle arrest and show unexpected toxicity to mammalian cells. In addition, the results of various studies have differed due to the use of GNPs with different physicochemical properties [10–12]. Thus, further studies are needed to evaluate the mechanisms through which GNPs cause cell cycle arrest. In this study, two types of GNPs with cetyltrimethylammonium bromide (CTAB)/bovine serum albumin (BSA) coatings were evaluated using raw 264.7 macrophage cells. Although there are earlier reports about BSA coated gold nanoparticles, their mechanism for modulating the cell cycle is still unknown. The correlations between biocompatibility, subcellular localization and cell cycle arrest of GNPs were investigated. We believe that our findings will improve the current understanding of the mechanisms behind which nanoparticles induce cell cycle arrest at different phases.

#### **2. Materials and Methods**

#### *2.1. Materials*

Hydrogen tetrachloroaurate (HAuCl4·3H2O), silver nitrate (AgNO3), CTAB, sodium salicylate, ascorbic acid, sodium citrate, poly(4-styrenesulfonic acid-co-maleic acid) sodium salt (PSSMA), and BSA were obtained from Sigma (Saint Louis, MO, USA). Rat monoclonal anti-α-tubulin antibodies conjugated with Alexa Fluor 647, rat monoclonal anti-α-tubulin antibodies and horseradish peroxidase (HRP)-labeled goat anti-rat IgG were purchased from Abcam (Cambridge, UK). Dulbecco's modified Eagle's medium (DMEM), fetal bovine serum (FBS), and phosphate-buffered saline (PBS) were purchased from Gibco (New York, NY, USA). The annexin V-fluorescein isothiocyanate (FITC) apoptosis detection kit, and propidium iodide (PI) cell cycle assay kit, phalloidin-FITC cytopainter, and 4 ,6-diamidino-2-phenylindole (DAPI) were purchased from Beyotime (Shanghai, China). Lyso Tracker Green DND-26 was obtained from Invitrogen (Carlsbad, CA, USA).

#### *2.2. Preparation and Characterization of GNPs*

GNPs were prepared as previously described [13]. Briefly, 5 mL of 0.5 mM HAuCl4 was mixed with 5 mL of 0.2 M CTAB solution, and 1 mL of 6 mM NaBH4 was then added. The solution was stirred for 2 min and incubated for 30 min to prepare the seed solution. To prepare colorless growth solution, 9.0 g CTAB plus 0.8 g sodium salicylate were dissolved in 250 mL warm water. Next, 6 mL of 4 mM AgNO3 solution and 1 mL of 0.064 M ascorbic acid were added. Finally, 0.8 mL seed solution was injected into the growth solution, stirred for 30 s and left undisturbed at 30 ◦C for 12 h for GNP growth. To obtain CTAB-GNPs, the precipitates after centrifugation were re-dispersed in 10 mL distilled water. According to previous reports [14,15],the CTAB-GNPs were further coated with the polyelectrolyte PSSMA and BSA under stirring to obtain BSA-GNPs.

The morphology of the nanoparticles was observed by TEM at an accelerating voltage of 100 kV (JEOL Co.,Tokyo, Japan). The size stability of nanoparticles was determined with a Brooke Haven Nanosizer at 37 ◦C with GNPs in DMEM medium (pH 7.4) at the concentration of 15 pM. Ultraviolet-visible (UV-vis) measurements were carried out using a Shimadzu UV 2700 spectrophotometer (Kyoto, Japan).

#### *2.3. Flow Cytometry Analysis of the Cell Cycle and Apoptosis*

RAW264.7 cells were obtained from the Shanghai Institutes for Biological Sciences (Shanghai, China) and routinely cultured in DMEM supplemented with 10% FBS at 37 ◦C in a humidified atmosphere with 5% CO2 in air. Progression of cells through the cell cycle was examined by flow cytometry in RAW264.7 cells treated with GNPs. RAW264.7 cells were plated at 4 × 104 cells/cm2 in

six-well plates and grown for 18 h. After incubation of cells with GNPs (0–30 pM) for 2 h, the medium was replaced with DMEM, and the cells were incubated for an additional 0–14 h. The interaction time between cells and GNPs in this study refers to incubation with GNPs in DMEM for 2 h plus incubation with GNP-free DMEM for 0–14 h. The cells were fixed in 70% ethanol, and DNA was stained with PI in the presence of 40 mg/mL DNase-free RNase A for 30 min at 37 ◦C in the dark. Cell cycle analysis was performed using flow cytometry according to the manufacturer's instructions. Early apoptotic cells were quantified by annexin V-FITC, whereas late apoptotic and necrotic cells were identified by PI staining. After treatment with GNPs, cells were trypsinized, harvested, washed with PBS, incubated with annexin V-FITC/PI for 15 min at room temperature in the dark, and analyzed on a flow cytometer (FACSCalibur BD, CA, USA).

#### *2.4. Confocal Microscopy Analysis*

Intracellular localization of GNPs was visualized using a Leica SP8 fluorescence confocal microscope (Wetzlar, Germany). RAW264.7 cells were treated with GNPs as described in the previous section and then co-incubated with Lyso Tracker Green DND-26 at 37 ◦C for 0.5 h. After washing with PBS, live cell imaging of green fluorescence with Lyso Tracker Green and diffusion reflection of GNPs irradiated at 630 nm were measured using a Leica SP8 (Wetzlar, Germany).

The morphology of the cytoskeleton was determined by immuno-staining RAW264.7 cells with a microtubule/microfilament fluorescent probe. Briefly, cells were fixed with 4% formaldehyde for 10 min, permeabilized with 0.1% Triton X-100 for 5 min, blocked with 1% BSA for 1 h, and then incubated with anti-tubulin antibodies conjugated with Alexa Fluor 647 and/or phalloidin-FITC according to the manufacturer's instructions for staining microfilament and microtubules. The cells were co-incubated with DAPI for nuclear staining. Polychromatic images of cells were measured using a Leica SP8 for green fluorescence indicating microfilament, red fluorescence indicating microtubules, and blue fluorescence indicating nuclei. Diffusion reflection of GNPs was defined as yellow, green, or red to avoid confusion with fluorescence.

#### *2.5. Western Blot Analysis*

Changes in the ratios of free tubulin within the polymerized microtubules were measured by western blotting. After treatment with GNPs, cells were lysed with 0.2 mL lysis buffer (20 mM Tris-HCl (pH 6.8), 0.5% NP-40, 1 mM MgCl2, 2 mM EGTA, 1 mM orthovanadate, and 20 mg/mL aprotinin, leupeptin, and pepstatin). Centrifugation yielded soluble tubulin dimers in the supernatant and polymerized microtubules in the pellet. Pellets were solubilized with sodium dodecyl sulfate (SDS) lysis buffer (Beyotime; Shanghai, China). Cell lysates containing equal amounts of protein were separated by SDS-polyacrylamide gel electrophoresis, transferred, probed with specific antibodies against α-tubulin, and detected on X-ray films using the chemiluminescence technique.

#### *2.6. qRT-PCR*

RAW264.7 cells were incubated with GNPs and then harvested for examination of gene expression by qRT-PCR. Briefly, total RNA was extracted with TRIzol reagent (CW0580S; CWBIO, Beijing, China). cDNA was then synthesized using a SuperScriptFirst-Strand Synthesis kit (CW2569M; CWBIO). The specific primers used in this study were as follows: *p53*, (forward) 5 -GCTCCTCCCCAGCATCTTA-3 and (reverse) 5 -GGGCAGTTCAGGGCAAA-3 ; kinesin 5A, (forward) 5 -GGCGGAGACTAACAACGAA-3 and (reverse) 5 -CTTGGAAAATGGGGATGAA-3 ; glyceraldehyde 3-phosphate dehydrogenase, (forward) 5 -AAGAAGGTGGTGAAGCAGG-3 and (reverse) 5 -GAAGGTGGAAGAGTGGGAGT-3 . The relative expression of mRNA was calculated by the 2−−ΔΔct method.

#### *2.7. Proteomics Analysis*

Cells were incubated with GNPs (30 pM) for 6 h, and cellular protein was extracted, digested, and desalted. The resulting peptide mixture was labeled with an iTRAQ Reagent-8 plex Multiplex Kit (AB Sciex U.K. Limited, Sheffield, U.K.) according to the manufacturer's instructions. Next, the labeled samples were fractionated using high-performance liquid chromatography (Thermo DINOEX Ultimate 3000 BioRS, Waltham, USA) using a Durashell C18 column (5 μm, 100 Å, 4.6 × 250 mm, Tianjin, China). Liquid chromatography electrospray ionization tandem mass spectrometry (MS/MS) analysis was performed on an AB SCIEX nanoLC-MS/MS (Triple TOF 5600 plus) system. Briefly, samples were chromategraphed on a C18 column (3 μm, 75 μm × 150 mm) with a 90-min gradient elution. Buffer A (0.1% formic acid and 5% acetonitrile) and buffer B (0.1% formic acid and 95% acetonitrile) served as the mobile phase. MS1 spectra were collected in the range of m/z 350–1500 for 250 ms, and the 30 most intense precursor ions were selected for fragmentation. MS2 spectra were collected in the range of m/z 100–1500 for 50 ms. Precursor ions were excluded from reselection for 15 s. The original MS/MS file data were submitted to ProteinPilot Software v4.5 (AB Sciex Pte Ltd. Waltham, MA, USA; https://sciex.com/products/software/proteinpilot-software) for data analysis. For protein identification, Paragon algorithm2, which was integrated into ProteinPilot, was employed against Uniprot Mus musculus 20171124.fasta (84434 items, updated in November 2017) for database searching. The parameters were set as follows: Instrument, TripleTOF 5600; iTRAQ quantification; cysteine modified with iodoacetamide. The biological modifications included ID focus and trypsin digestion, and the quantitate, bias correction, and background correction was used for protein quantification and normalization. Only proteins with at least one unique peptide and an unused value of more than 1.3 were considered for further analysis.

#### **3. Results**

#### *3.1. Proertiesof GNPs withBSA/CTABCapping Agents*

As the surfactants have poor biocompatibility, several shells such as carbon shells and biopolymer shells have been used to reduce toxicity of surfactants [16]. We chose BSA as a model molecule for its biocompatibility. Zeta potential of CTAB-GNPs and BSA-GNPs are 28.4 ± 2.6 mV and −20.5 ± 2.1 mV respectively, indicating positive CTAB and negative BSA on the surface of GNPs. The physicochemical properties of BSA-GNPs and CTAB-GNPs are shown in Figure 1. Transmission electron microscopy (TEM) showed that both types of nanoparticles were rod shaped with similar particle sizes in distilled water (Figure 1A,B). Although the hydrodynamic diameter deduced from the Stokes-Einstein equation was not accurate when regarding nano-rods as nano-spheres, the diffusion coefficient determined by dynamic light scattering (DLS) was still accurate. The particle size peak can be a signature for determining the nano-rod aggregation formation [17]. DLS analysis showed that the particle size of CTAB-GNPs increased from 45 to 79 nm as the incubation time increased. In contrast, no significant changes in particle size were observed for BSA-GNPs (Figure 1C). These results indicated coating with BSA enhanced the stability of GNPs, which can be attributed to good dispersity of BSA in high salt solution.The extinction spectra of the nanoparticles showed a peak at around 630 nm, which corresponded to the longitudinal surface plasmon resonance of the rod-shaped GNPs (Figure 1D).

**Figure 1.** Characterization of GNPs. (**A**) TEM image of CTAB-GNPs in distilled water. (**B**) TEM image of BSA-GNPs in distilled water. (**C**) Hydrodynamic size of GNPs in DMEM. (**D**) UV-vis spectra of GNPs in distilled water.

#### *3.2. Effects of GNPs on the Cell Cycle and Apoptosis*

Murine macrophages RAW264.7 were used owing to their strong nanoparticle phagocytosis and short cell cycle period. Apoptosis assays revealed that incubation of RAW264.7 cells with 15 pM of BSA-GNPs yielded 38.82% ± 4.30% early apoptotic cells and 33.98% ± 4.37% late apoptotic cells, whereas incubation of cells with 15 pM of CTAB-GNPs yielded 59.72% ± 1.52% early apoptotic cells and 10.23% ± 1.57% late apoptotic cells (Figure 2A,C). The apoptosis rate increased as the concentration of GNPs increased. Thus, for subsequent cell cycle analyses, we chose a dosage of 15 pM. Compared with the control group (7.71% ± 1.64% in G2/M phase), 18.54% ± 1.40% of cells were in the G2/M phase after treatment with 15 pM BSA-GNPs for 2 h. This indicated that BSA-GNPs induced cell cycle arrest at G2/M phase. Notably, for the cells treated with 15 pM CTAB-GNPs for 2 h, 62.88% ± 3.01% of cells were found to be in the G0/G1 phase, compared with 48.56% ± 1.57% in the control group. BSA-GNPs and CTAB-GNPs also induced G2/M and G0/G1 arrest after 16 h of treatment (Figure 2B,D), respectively.

**Figure 2.** Apoptosis and cell cycle distributions of RAW264.7 cells before and after GNP treatment, with untreated cells used as control. (**A**) Flow cytometry images of GNPs inducing cell apoptosis after incubation for 16 h. (**B**) Flow cytometry images of cell cycle arrest in RAW264.7 cells treated with 15 pM BSA-GNPs and CTAB-GNPs for 2 or 16 h. (**C**) Barchart showing intensity of cell apoptosis. (**D**) Barchart showing distributions of the cell cycle.

#### *3.3. Intracellular Localization of GNPs*

Lysosomes, sliding on microtubules, play important roles in the intracellular transportation of nanoparticles [18]. As microtubules greatly affect the cell cycle, interactions of GNPs with lysosomes/microtubules were investigated. Figure 3 shows fluorescent images of GNP-treated cells in which the cell nucleus, lysosomes, microtubules, and GNPs were labeled in different channels. As shown in Figure 3A, the green fluorescence from Lyso Tracker Green DND-26 disappeared in most of the regions of the cells treated with CTAB-GNPs, indicating the disruption of lysosomes by CTAB-GNPs. This could be attributed to the surfactant CTAB, which facilitates lysosome escapees reported in previous reports [19]. However, different results were found in BSA-GNP-treated cells. The colocalization of green fluorescence from lysosomes and scattering reflection from BSA-GNPs (in red) showed that BSA-GNPs were accumulated in lysosomes. In addition, the accumulated green fluorescence of the Lyso Tracker Green DND-26 indicated an accumulation of lysosomes.

Figure 3B,C show the cytoskeleton morphology upon GNP treatment, as determined by laser confocal microscopy. Compared with the PBS treated cells, CTAB-GNPs caused shrinkage of microtubules, microfilaments, and nuclei after 2 h of nanoparticle treatment [20]. Additionally, CTAB-GNPs were aggregated into small circulars with diameters between 0.6–0.9 μm, and they were matched with the red fluorescence from α-tubulin after 16 h of treatment. These suggest that CTAB-GNPs induced tubulin aggregation. In contrast, BSA-GNPs treated cells showed increased microtubule and nuclear organization in the mitosis phase [21]. No overlap between the BSA-GNPs and microtubule-tubulin system was observed after 16 h of nanoparticle treatment.

**Figure 3.** Effect of GNPs on subcellular organelles, with untreated cells used as control. (**A**) Confocal microscopy images of cell lysosome after incubation with GNPs for 16 h, showing colocalization of BSA-GNPs (red) with lysosomes (green). (**B**) Fluorescence microscopy images of cell cytoskeleton after incubation with GNP for 2 h, showing shrinkage of microtubules (red) and microfilaments (green) in CTAB-GNPs and increased microtubules (red) and nuclear (blue) organization in the mitosis phase in BSA-GNPs. (**C**) Fluorescence microscopy images of cell cytoskeleton after incubation with GNP for 16 h, showing colocalization of GNPs (green) with microtubules (red). (**D**) Western blot analysis of free tubulin and polymerized microtubule in cells treated with GNPs. (**E**) Relative mRNA levels of kinesin 5A and P53 in cells treated with GNPs.

#### *3.4. Effects of Nanoparticles Ondepolymerization/Polymerization of Microtubules*

To investigate the potential effects of nanoparticles on the depolymerization/polymerization of microtubules, western blotting was performed after separating the free tubulin from polymerized microtubules. In these cells, treatment with BSA-GNPs increased polymerized microtubules compared with the untreated control cells (Figure 3D), suggesting microtubule stabilization and inhibition of microtubule depolymerization. However, cells treated with CTAB-GNPs showed increased free tubulin, which may be due to the inhibition of microtubule polymerization and assembly of tubulin into small aggregates [10,22].

#### *3.5. Protein Identification and Quantification by Quantitative Real-Time Reverse Transcription Polymerase Chain Reaction (qRT-PCR)*

Kinesin 5A is a microtubule motor protein associated with lysosomes and acts as a microtubule polymerase by promoting tubulin polymerization and inhibition of tubulin depolymerization [23,24]. Compared with the control group, the mRNA level of kinesin 5A increased 1.26- and 1.91-fold in CTAB-GNP and BSA-GNP treated cells, respectively (Figure 3E). The increase in kinesin 5A could be attributed to the accumulation of lysosomes on microtubule during GNP transport. However, kinesin 5A levels in CTAB-GNP treated cells were much lower than those in BSA-GNP-treated cells, potentially because of the subsequent lysosome rupture induced by CTAB-GNPs. Overall, the significant increase of kinesin 5A (*p* < 0.01) suggested lysosome accumulation on microtubules and microtubule stabilization in BSA-GNP treated cells. As shown in Figure 3E, p53 mRNA levels were decreased by 8.92% in BSA-GNP-treated cells, yet increased by 1.21fold (*p* < 0.05) in CTAB-GNP treated cells. The increase of P53 can be contributed to microtubule disruption [25].

#### *3.6. Protein Identification and Quantification by Isobaric Tags for Relative and Absolute Quantitation (iTRAQ)*

To further explore the cell cycle arrest mechanism induced by GNPs, we used iTRAQ proteomics to identify and quantify protein changes in RAW264.7 cells before and after GNP treatment. In this study, 3341 and 3348distinct proteins were identified using iTRAQ-based proteomic technology in BSA-GNP and CTAB-GNP treatment, respectively (Figure S1a, Supporting Information). To improve our understanding of the roles of these proteins, differentially accumulated protein analysis was based on the fold-change >1.5 or <0.667 (*p* < 0.05). For the cells treated with BSA-GNPs, 159 proteins were found to be differentially expressed compared with the control, including 65 up-regulated and 94 down-regulated proteins. Moreover, 102 proteins were found to be differentially expressed in CTAB-GNP treated cells, including 55 up-regulated and 47 down-regulated proteins. As shown in the Venn diagram, 36 differentially expressed proteins were common in both GNP-treated groups (Figure S1b, Supporting Information). Gene ontology (GO) classification of these differentially expressed proteins were divided into three classes (biological processes, cellular components, and molecular functions). Cells treated with BSA-GNPs or CTAB-GNPs have shown differences in all the three classes (Figure S2, Supporting Information).

#### *3.7. Effects of Nanoparticles on Cell Cycle-Related Protein Expression*

Kyoto Encyclopedia of Genes Genomes (KEGG) annotation analysis of all differentially expressed proteins was used to explore the underlying pathways and processes, and the top 10 altered pathways are shown in Figure S3 (Supporting Information). Down-regulation of cell cycle-related proteins was observed following BSA-GNP treatment. Down-regulation of actin cytoskeleton-related proteins, which are closely related to the cell cycle, were observed following CTAB-GNP treatment. We adopted KEGG annotation analysis to explore the underlying pathways of the cell cycle (Figure 4). Our results showed that three of these unique proteins (cadherin 1 (Cdh1), minichromosome maintenance complex component 5 (MCM5), 14-3-3 protein) were related to the cell cycle in BSA-GNP-treated cells. Of these proteins, the expression of Cdh1 increased 2.22 fold in response to mispositioned

spindles. Cdh1 is an antagonist of the spindle assembly checkpoint and its over-expression could lead to the silencing of mitotic cyclin-dependent kinase 1 (CDK1) activity and consequently the cell cycle arrest at G2/M phase. MCM5, which was up-regulated in the transition from the G0 to G1/S phase of the cell cycle [26], was decreased 0.59 fold. Therefore, the reduction of MCM5 is implicated in low numbers of cells in the G0/G1 and S phases. The 14-3-3 protein zeta/delta, 14-3-3 protein gamma, and 14-3-3 protein tau were down-regulated 0.36–0.57 fold. The 14-3-3 protein directly binds to kinesin heterodimers and acts as a phospho-Ser/Thr-binding factor [27]. Phosphorylation of kinesin 5A inhibits its binding to microtubules [28]. Thus, we conclude that the down-regulation of 14-3-3 has weakened the phosphorylation of kinesin 5A and thus promoted the binding of kinesin 5Ato spindle microtubules. As a result, the microtubule was stabilized by BSA-GNPs. In CTAB-GNPtreated cells, proliferating cell nuclear antigen (PCNA) protein was up-regulated by 1.52 fold as compared with that in the control group. PCNA, as an accessory factor for DNA polymerases, is up-regulated rapidly in the G1phase through early S phase and is then down-regulated in late S and G2/M phases. Increased levels of PCNA can cause cell cycle arrest in G0/G1 through the inactivation of CDK4/6. Moreover, increased levels of p53 and PCNA can contribute to microtubule damage [25,29].

**Figure 4.** KEGG pathway analysis of the cell cycle in GNP-treated cells. PCNA was up-regulated in CTAB-GNP treated cells. Cdh1 was up-regulated, whereas 14-3-3 protein and MCM5 were down-regulated in BSA-GNP-treated cells. Single line frames refer to BSA-GNPs, and double line frames refer to CTAB-GNPs. Red frames indicate up-regulated proteins, and green frames indicate down-regulated proteins.

#### **4. Discussion**

In this study, we found that GNPs causing cell cycle arrest was dependent on biocompatibility of GNP surfaces. Coating of GNPs with biocompatible BSA induced G2/M arrest through microtubule stabilization, while residual toxic CTAB on the surface of GNPs typically caused cell cycle arrest in G0/G1 phase microtubule disruption. Kim et al. have shown that the cell cycle affects the intracellular transport of nanoparticles [30]. Nanoparticles internalized by cells are not exported from cells but are split during G2/M. Indeed, we found that the intracellular transport/location of nanoparticles had an effect on cell cycle progression (Figure 5). The accumulation of BSA-GNPs in lysosomes increased the level of kinesin 5A and caused subsequent stabilization of microtubules (including the promotion of tubulin polymerization and inhibition of tubulin depolymerization) [23,24], blockage of chromosome segregation, and induction of cell cycle arrest in G2/M viaCdh1 elevation [31]. In contrast, CTAB on the surface of GNPs caused lysosome/endosome rupture and subsequent microtubule damage through tubulin aggregation and the inhibition of tubulin polymerization. These changes induced G0/G1 arrest through the regulation of p53 and PCNA. Overall, biocompatibility properties of GNPs plays an important role in cell cycle progression. Biocompatible coated GNPs could inhibit lysosome rupture caused by residual surfactant and switched G0/G1 arrest to G2/M arrest. Similar results are expected when using other biocompatible molecule coated GNPs, including polyethylene glycol and fluoro-6-deoxy-d-glucose, in accordance with previous reports [32,33].

**Figure 5.** Mechanism through which GNPs causes cell cycle arrest was dependent on the biocompatible property of GNP surface. Coating of GNPs with biocompatible molecules, such as BSA, inhibited lysosome rupture and switched G0/G1arrest to G2/M arrest. The accumulation of BSA-GNPs in lysosomes increased the level of kinesin 5A and caused subsequent stabilization of microtubules (including promotion of tubulin polymerization and inhibition of tubulin depolymerization), blockage of chromosome segregation, and induction of cell cycle arrest in G2/M via Cdh1 elevation. In contrast, toxic CTAB on the surface of GNPs caused lysosome rupture and ssubsequent microtubule damage through tubulin aggregation. These changes induced G0/G1 arrest through regulation of p53 and PCNA.

Microtubules are the major components of cytoskeletal systems that are responsible for regulation of the cell cycle. Many commonly used drugs, including paclitaxel (a microtubule-stabilizing agent), nocodazole (a microtubule-destabilizing agent), and vinblastine (a microtubule-destabilizing agent) induce G2/M cell cycle arrest through regulation of microtubules. Choudhury et al. reported that bare GNPs induce G0/G1 arrest by causing microtubule damage [10]. In this study, we demonstrated that BSA-coated GNPs stabilized microtubules and caused G2/M arrest by inducing interactions between lysosomes and microtubules. Nanoparticles are taken up and transported within subcellular structures that are surrounded by one or two layers of membranes, including endosomes, lysosomes, mitochondria, and multivesicular bodies [34]. The motility of these subcellular structures is based on microtubules. Therefore, the transport of nanoparticles can affect dynamic changes in microtubules. Microtubule-interfering drugs affect the cell cycle distribution by impairing the mitotic checkpoint and regulating the activity of cyclins and CDKs. Both stabilization and destabilization of microtubules could impair the mitotic checkpoint and cause G2/M arrest. For example, microtubule-stabilizing drug paclitaxel regulates the mitotic checkpoint proteins Bub1, CDK1 and CDK2 [35,36]. Microtubule-destabilizing drug nocodazole caused mitotic slippage through precocious activation of Cdh1 and inhibition of CDK1 [37,38]. BSA-GNPs in our study regulated CDK1 through up-regulation of Cdh1, so BSA-GNPs stabilizing microtubules may also lead to a potential cancer therapy.

#### **5. Conclusions**

GNPs causing cell cycle arrest was highly dependent on the surface biocompatibility of GNPs. Residual toxic CTAB on the naked GNPs typically caused cell cycle arrest in G0/G1 phase, whereas the coating of GNPs with BSA resulted in the inhibition of lysosome rupture ability, microtubule stabilization, and a switch to G2/M arrest. This will greatly help us to regulate the cell cycle progression through modulating surface coating and biocompatibility of nanoparticles and direct us to set the guidelines for the formulation of nanoparticles in different biomedical applications.More importantly, BSA-GNPs caused G2/M arrest through microtubule stabilization similarly to the mechanisms of many well-known anticancer drugs, and the recognition of this mechanism could be applied as a new therapeutic target of nanoparticles in tumor therapy.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2079-4991/8/12/1063/ s1. Additional images related to proteomics analysis of identified and differentially expressed proteins in GNPs treated raw 264.7 cells from iTRAQ proteomics, Gene ontology (GO) classification of differential expressed proteins in GNPs treated cells, Top 10 changed pathways based on proteome analysis in GNPs treated cells.Figure S1: Identified and differentially expressed proteins in GNPs treated raw 264.7 cells from iTRAQ proteomics. Figure S2: Gene ontology (GO) classification of differential expressed proteins. Figure S3: Top 10 changed pathways based on proteome analysis.

**Author Contributions:** Author Contributions: Q.L. and C.H. conducted the experiments and wrote the paper. L.L. and J.Q. funded the paper. R.H. initiated this study and guided the experiments.

**Funding:** This research was funded by [National Basic Research Program of China] 2015CB352005; [National Natural Science Foundation of China] 61525503/61620106016/81727804/61722508; [Natural Science Foundation of Guangdong Province Innovation Team] 2014A030312008; and [Science and Technology Innovation Commission of Shenzhen] KQJSCX20170327151457055, JCYJ20170817094609727, JCYJ20170412105003520, JCYJ20150930104948169, JCYJ20160328144746940, GJHZ20160226202139185 and [China Postdoctoral Science Foundation] 2017M622756.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

© 2018 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Dose- and Time-Dependent Cytotoxicity of Layered Black Phosphorus in Fibroblastic Cells**

#### **Su-Jin Song 1,†, Yong Cheol Shin 2,†, Hyun Uk Lee 3, Bongju Kim 4, Dong-Wook Han 1,\* and Dohyung Lim 5,\***

Received: 10 May 2018; Accepted: 4 June 2018; Published: 6 June 2018

**Abstract:** Black phosphorus (BP) is a monolayer/multilayer two-dimensional (2D) nanomaterial, which has recently emerged as one of the most attractive 2D nanomaterials due to its fascinating physicochemical and optoelectronical properties. Layered BP may have promising applications in biomedical fields, such as drug delivery, photodynamic/photothermal therapy and bioimaging, although its intrinsic toxicity has not been fully elucidated yet. In the present study, the cytotoxicological effects of layered BP on both cell metabolic activity and membrane integrity were investigated. Layered BPs were prepared using a modified ultrasonication-assisted solution method, and their physicochemical properties were characterized. The dose- and time-dependent cytotoxicity of layered BP was assessed against L-929 fibroblasts. Our findings indicate that the cytotoxicity of BPs is proportionally dependent on their concentration and exposure time, which is affected by the oxidative stress-mediated enzyme activity reduction and membrane disruption. On the other hand, layered BPs did not exhibit significant cytotoxicity at concentrations lower than 4 μg/mL. Therefore, it is suggested that layered BPs can be effectively utilized as therapeutic delivery carriers and imaging agents.

**Keywords:** black phosphorus; 2D nanomaterial; cytotoxicity; biomedical application

#### **1. Introduction**

Over the last decade, tremendous research has been conducted to understand and explore the various types of two-dimensional (2D) nanomaterials. This research has found that 2D nanomaterials have a promising potential in a variety of applications, such as optoelectronics, photonics, energy storage and conversion, and biomedicine [1,2]. Among monolayer/multilayer 2D nanomaterials, layered black phosphorus (BP) has recently emerged as an attractive novel one due to its distinct structure, with phosphorenes stacked in several layers via van der Waals forces, and has been acknowledged as one of the most stable allotropes of the phosphorus family [3–8]. Some studies have already shown the potential of BP in biomedical applications, such as drug delivery, photodynamic/photothermal therapy and bioimaging [1,9–13]. However, several controversial results regarding the toxicity of BP have been reported, which means that an in-depth understanding of the cytotoxicity and underlying mechanism of BP is of utmost importance.

A series of studies reported that layered BP has little to no toxic effects, which means that it can be employed as a biomedical material [1,6,13]. It has been found that, while the BPs can induce cell apoptosis and necrosis owing to the transient intracellular reactive oxygen species (ROS)-mediated oxidative stress, the induced oxidative stress can be gradually restored to normal levels with no long-term inflammatory reaction or obvious damage to an in vivo mouse model [14]. Moreover, BP nanosheets can be used as drug delivery vehicles because they have pH- or photo-responsive drug release characteristics as well as a high drug loading efficiency [1,9]. Additionally, BP has been found to possess both outstanding near-infrared photothermal performance and photodynamic activity, which allows it to be utilized for photothermal and photodynamic therapy [9–12,15]. However, although these studies on the biomedical potential of BP could provide valuable guidelines for the essential understanding of the biological effects of BP, the issue of the potential toxicity of BP remains unresolved. In particular, the toxicity of layered BPs is highly varied depending on their concentration, size, shape, surface chemistry, and exposure time, which is similar to the other 2D nanomaterials, such as graphene and its derivatives [14,16,17]. Therefore, prior to the use of layered BP in biomedical applications, it is urgently necessary to investigate its toxicological effects. Hence, in the present study, we assessed the cytotoxicity of layered BP on fibroblastic cells according to its concentration and exposure time, using cytotoxicity assays with different end-points, including the cell metabolic activity, membrane integrity and intracellular ROS production. Our findings revealed that layered BPs showed dose- and time-dependent cytotoxicity, which are caused by oxidative stress-mediated enzyme activity reduction and membrane disruption, but they did not exhibit significant cytotoxicity at a low concentration. These dose- and time-dependent cytotoxicity profiles of layered BPs can be quite informative and useful for their development as biocompatible therapeutic delivery carriers and imaging agents.

#### **2. Materials and Methods**

#### *2.1. Preparation and Characterization of Layered BP*

Layered BP was prepared by exfoliation of bulk BP crystals using a modified ultrasonication-assisted solution method, as described elsewhere [13]. Fourier transform infrared (FT-IR) spectroscopy was used to characterize the layered BP. The FT-IR spectrum of layered BP was collected using an FT-IR spectroscope (Nicolet Co., Madison, WI, USA) with a resolution of 4.0 cm−1 and 16-times scanning in the wavelength range of 750–4000 cm−1. The surface topography of layered BP was analyzed by atomic force microscopy (AFM; NX10, Park Systems Co., Suwon, Korea) in air at room temperature. Imaging was carried out in non-contact mode with a Multi 75 silicon scanning probe at a resonant frequency of ~300 kHz. The average hydrodynamic size of layered BPs was determined using a Zetasizer (Nano ZS, Malvern Instruments, Worcestershire, UK).

#### *2.2. In Vitro Assays for Cytotoxicity Evaluation of Layered BP*

L-929 fibroblastic cells were routinely cultured in Dulbecco's modified Eagle's Medium (DMEM, Welgene, Daegu, Korea) supplemented with 10% fetal bovine serum (Welgene) and 1% antibiotic-antimycotic solution (Sigma-Aldrich Co., Saint Louis, MO, USA) at 37 ◦C in a humidified atmosphere containing 5% CO2. The cell viability of L-929 cells, treated with layered BP for 24 h, 48 h and 72 h, was assessed by a cell counting kit-8 (CCK-8) assay (Dojindo, Kumamoto, Japan) according to the manufacturer's instructions. Briefly, L-929 fibroblasts were seeded at a density of 1 × 104 cells/mL on a 96-well plate and incubated for 24 h. Subsequently, the cells were treated with various concentrations of layered BP suspended in culture medium (0 to 125 μg/mL) and then incubated with a CCK-8 solution for the last 2 h of the culture period (24 h, 48 h and 72 h) at 37 ◦C in the dark. The absorbance was measured at 450 nm using an enzyme-linked immunosorbent assay (ELISA) reader (SpectraMax® 340, Molecular Device Co., Sunnyvale, CA, USA). The cell viability was determined to be the percentage ratio of the absorbance values in the cells (incubated with layered BP) to those in untreated control groups (0 μg/mL).

The cell membrane integrity was investigated by monitoring the release of lactate dehydrogenase (LDH) using an LDH assay kit (Takara Bio Inc., Shiga, Japan). After 24 h of incubation with various concentrations of layered BP, the supernatant from each cell culture was transferred to a new 96-well plate. Next, the LDH assay solution was added to each well and then incubated for 30 min at room temperature in the dark. The absorbance was measured at 490 nm using an ELISA reader.

The intracellular ROS production was detected using an ROS assay kit (OxiSelect™; Cell Biolabs, Inc., San Diego, CA, USA). Typically, L-929 cells were plated in a 96-well plate (1 × 104 cells/mL) and incubated for 24 h. The cells were treated with increasing concentrations of layered BP for 24 h. Each cell culture was washed with Dulbecco's phosphate-buffered saline (DPBS, Gibco, Rockville, MD, USA) and then incubated with 2 ,7 -dichloroflurorescein diacetate (DCFH-DA), a cell-permeable fluorogenic probe, for 30 min at 37 ◦C in the dark. The cells were then imaged using an inverted fluorescence microscope (IX81, Olympus, Melville, NY, USA); the fluorescence intensity was determined by a fluorescence plate reader (VICTOR3 Multilabel Counter, PerkinElmer, Inc., Waltham, MA, USA) with excitation and emission wavelengths of 480 nm and 530 nm, respectively. The fluorescence intensity was expressed as the fold-increase over the values of the untreated control groups.

For morphological observations, the time-lapse images of L-929 cells treated with 10 μg/mL of layered BP were acquired every 1 h for 12 h of incubation. The percentage of live cells was estimated by calculating the ratio of the number of attached cells, defined as cells with a spindle-like morphology (i.e., aspect ratio larger than 1) or specialized subcellular structures, such as lamellipodia, filopodia, stress fibers, and membrane protrusions, to the total number of cells [18–22].

#### *2.3. Statistical Analysis*

All variables were tested in three independent cultures for each experiment, which were repeated twice (*n* = 6). All presented data were expressed as average ± standard deviation. Statistical comparisons were carried out by a one-way analysis of variance (SAS Institute Inc., Cary, NC, USA), followed by a Bonferroni test for multiple comparisons. A value of *p* < 0.05 was considered statistically significant.

#### **3. Results and Discussion**

#### *3.1. Characteristics of Layered BP*

The physicochemical properties of layered BP were characterized by FT-IR spectroscopy and AFM (Figure 1). The FT-IR spectrum of layered BP showed the characteristic peaks of BP crystals (Figure 1a). A noticeable peak was observed near 1000 cm−1, attributed to the stretching vibrations of P–O [23]. The peaks found near 1140 and 1620 cm−1 represented the P=O stretching modes of layered BP [23,24]. On the other hand, broad absorption bands were observed, ranging from 2400 cm−1 to 3500 cm−1, which could be attributed to the CO2 stretching and OH stretching vibrations due to exposure of the layered BP to ambient atmosphere. The surface topographic image of layered BP is presented in Figure 1b. Most layered BP were found to have a 2D layer structure, and the average height was about 6.87 ± 0.58 nm (Figure 1b,c). Considering the thickness of the BP monolayer (0.53 nm), the layered BP was composed of several BP monolayers [25]. Moreover, the hydrodynamic size of 2D nanomaterials is of great importance in biomedical applications, because it has a marked effect on the interactions between 2D nanomaterials and cells [17,26–29]. The hydrodynamic size of the BPs used in the present study was found to be 960 ± 303 nm (Figure 1d).

**Figure 1.** Characterizations of layered BP. (**a**) FT-IR spectrum of layered BP; (**b**) AFM image and (**c**) the height profile of layered BP along the black line marked in (**b**); (**d**) Hydrodynamic size distribution histogram of layered BP.

#### *3.2. Dose-Dependent Cytotoxicity of Layered BP*

To investigate the cytotoxic effects of layered BP on L-929 fibroblasts according to its concentration, cells were treated with increasing concentrations of layered BP (0 to 125 μg/mL) for 24 h, and the morphology of the cells was observed (Figure 2a). There were no significant differences in the number and morphology of L-929 fibroblasts at concentrations of up to 4 μg/mL of layered BP. On the other hand, the cells with aggregated BPs exhibited an abnormal morphology and a significant decrease in cell number at concentrations higher than 8 μg/mL, clearly indicating that layered BPs exhibit dose-dependent cytotoxicity. From the CCK-8 assay, based on the cell metabolic activity (Figure 2b), it was found that the cell viability of L-929 fibroblasts decreased as BP concentration increased. At relatively low concentrations (~4 μg/mL), over 82% of fibroblasts were viable, whereas the cell viability of the control at 62 μg/mL decreased to approximately 37%. These findings are inconsistent with previous reports, which found that BP derivatives, including BP nanosheets and nanodots, were nontoxic to several types of cells even when BP concentration was as high as 1000 μg/mL [1,6,9,13]. These conflicting results may be due to size effects. It was demonstrated that layered BPs show a size-dependent cytotoxicity; larger BPs (with lateral size of ~880 nm) were more cytotoxic than smaller ones (with lateral size of ~210 nm) [17]. As shown in Figure 1d, the average lateral size (~960 ± 303 nm) of layered BPs used in this study was relatively larger than that used in other investigations, which can result in greater toxic effects on cells.

**Figure 2.** (**a**) Representative optical microscopy images of L-929 fibroblasts cultured with layered BP (0, 0.5, 2, 4, 8, 16, 31 and 62 μg/mL); (**b**) Cell viability and (**c**) LDH release profile of L-929 fibroblasts after 24 h of incubation with various concentrations of layered BP; (**d**) Correlation coefficient plot between metabolic activity and LDH release for cells cultured with layered BP at concentrations ranging from 0 to 62 μg/mL.

#### *3.3. Membrane Disruption and ROS Production Induced by Layered BP*

On the other hand, interesting results were found concerning the cytotoxicity of layered BPs. The cytotoxic effects of layered BPs can be ascribed to membrane disruption [17,30]. Therefore, we investigated the cytotoxicity of layered BPs using LDH assays based on the cell membrane integrity (Figure 2c). The extracellular release of LDH has been extensively used for investigating cell membrane integrity, because LDH, a stable cytoplasmic enzyme, can only be released into extracellular fluids upon plasma membrane disruption [31]. As shown in Figure 2c, a significant LDH release was detected at high concentrations of layered BP (≥16 μg/mL). The LDH release increased to approximately 140% of the control at 16 μg/mL of BP, indicating that high concentrations of layered BPs induced a significant membrane disruption. A slight decrease in LDH release, observed at 125 μg/mL, can be due to the decrease in the total cell number. For CCK-8 and LDH assay results, the calculated value of the corresponding correlation coefficient was −0.91 (Figure 2d), implying that the effects of layered BP on cell metabolic activity and membrane integrity were shown to have a high negative correlation.

At the same time, the dose-dependent cytotoxicity of layered BPs can also be due to oxidative stress. To further investigate the cytotoxicity of layered BPs, the effects of BPs on intracellular ROS generation were evaluated using an ROS-sensitive fluorogenic probe DCFH-DA. The DCFH-DA, a cell-permeable fluorophore, can be readily diffused into cells and subsequently deacetylated by cellular esterases to non-fluorescent DCFH (2 ,7 -dichlorodihydrofluorescin). The internalized DCFH is quickly oxidized to highly fluorescent DCF by intracellular ROS. Hence, the intracellular fluorescence of DCF reflects the oxidative stress attributed to the intracellular ROS production. As shown in Figure 3a, the minimal fluorescence was detected at low concentrations of layered BP (≤4 μg/mL), while obvious green fluorescence was detected in L-929 cells after incubation with concentrations of layered BP higher than 8 μg/mL. In addition, the fluorescence intensity was significantly (*p* < 0.05) enhanced with increasing concentrations of layered BPs (Figure 3b). It has been documented that the cytotoxicity of BP nanomaterials causes oxidative stress, such as the reduction of enzyme activity, lipid peroxidation and DNA breaks, caused by intracellular ROS production [14]. Thus, even though the size of the layered BP used in the present study was different from that used in previous studies, the cytotoxicity of layered BPs is proportionally dependent on their concentration, which can be attributed to the reduction of metabolic activity owing to oxidative stress. From our in vitro cytotoxicity assay results with different end-points (the cell metabolic activity, membrane integrity and intracellular ROS production), it was revealed that the dose-dependent cytotoxicity of layered BPs was due to both membrane disruption and oxidative stress-mediated metabolic activity reduction.

**Figure 3.** (**a**) Representative fluorescence microscopy images of oxidized DCF fluorescence in L-929 fibroblasts treated with various concentrations of layered BP (0, 4, 8, 16, 31 and 62 μg/mL) for 24 h; and (**b**) quantification of oxidized DCF fluorescence intensity. The scale bars are 100 μm.

#### *3.4. Time-Dependent Cytotoxicity of Layered BP*

To further evaluate the toxic effects of layered BPs on cells, we observed the morphological changes of L-929 fibroblasts and estimated the number of live cells. The time-lapse images of cells, treated with 10 μg/mL of layered BP for an initial 12 h at an interval of 1 h, are shown in Figure 4a. The number of live cells was estimated by quantifying the ratio of the number of attached cells to the total number of cells (Figure 4b). Because adherent cells, including fibroblastic cells, have to be attached to appropriate substrates in order to survive, the cells, which did not show typical fibroblastic morphology, were considered to be dead [18–22]. It was observed that the number of cells with apoptotic morphology (marked in red) increased throughout incubation with layered BPs for the initial 12 h (Figure 4a). In particular, the live cells decreased significantly (*p* < 0.05) after 6 h of incubation with layered BPs (Figure 4b). The morphological changes were clearly observed by comparing optical microscopy images, taken every hour for 12 h (Figure 4c). These results implied that the cytotoxicity of layered BPs is also dependent on their exposure time.

**Figure 4.** (**a**) Time-lapse images of L-929 cells treated with 10 μg/mL of layered BP for an initial 12 h at an interval of 1 h; (**b**) Quantification of the percentage of live cells for 12 h; (**c**) Optical microscopy images of L-929 fibroblasts treated with 10 μg/mL of layered BP for 0 and 12 h. The scale bars are 100 μm.

The cell viability of L-929 fibroblasts, incubated with layered BP for 48 and 72 h, was evaluated to further examine the time-dependent cytotoxicity of layered BP, as shown in Figure 5a,b, respectively. The cytotoxic effects of layered BPs after 48 and 72 h are also dose-dependent, which is similar to the results after 24 h. Additionally, the cell viability after 48 and 72 h decreased more than it did after 24 h, as the incubation time with layered BPs had increased, and the decrease in cell viability after 72 h was more significant than after 48 h. At a concentration of 16 μg/mL, the cell viability after 48 and 72 h decreased to approximately 60% and 45% of the control, respectively. These results indicated that the cytotoxic effects of layered BPs were also dependent on exposure time. Consequently, it was revealed that the layered BP exhibited dose- and time-dependent cytotoxicity, as a result of membrane disruption and oxidative stress-mediated metabolic activity reduction caused by the accumulation of intracellular ROS as well as the interactions between layered BPs and cells. However, it is worth noting that the layered BPs were not significantly cytotoxic at concentrations lower than 4 μg/mL, suggesting that layered BPs in the range of only a few μg/mL can be effectively used in biomedical applications, such as therapeutic delivery carriers and imaging agents. Furthermore, to improve biocompatibility and biological activity, BPs can be conjugated or modified with various functional compounds, such as biocompatible polymers, nanoparticles and drugs [1,10,12,15]. It has been revealed that the encapsulation of BPs with poly(lactic-*co*-glycolic acid), a biodegradable polymer, allows not only the enhancement of biocompatibility, but also the degradation of nontoxic phosphate and phosphonate [12]. These results indicated that, although the cytotoxicity of BPs is closely dependent on their concentration and exposure time, the BPs with the desirable modification can be compatibly employed in biomedical applications, even at concentrations higher than 8 μg/mL. In summary, it is suggested that BP has a promising potential as a biomedical material.

**Figure 5.** Cell viability profiles of L-929 fibroblasts after (**a**) 48 and (**b**) 72 h of incubation with various concentrations of layered BP.

#### **4. Conclusions**

This study aimed to investigate the dose- and time-dependent cytotoxicity of layered BPs against L-929 fibroblasts. It was revealed that the cytotoxicity of layered BPs was proportionally dependent on their concentration and exposure time. These cytotoxic effects of layered BPs are found to be due to both oxidative stress-mediated enzyme activity reduction and membrane disruption. On the other hand, the cytotoxicity of layered BPs is not significant at concentrations lower than 4 μg/mL. Taken together, this work suggests that layered BPs can be effectively used in biomedical applications, such as therapeutic delivery carriers and imaging agents, although further comprehensive studies are undoubtedly necessary to fundamentally explore and understand the more detailed mechanisms behind the toxic effects of BPs.

**Author Contributions:** S.-J.S. and Y.C.S. designed the experiments, performed the in vitro assays and drafted the manuscript. H.U.L. carried out the preparation and characterization of layered BPs. B.K. performed the statistical analysis and helped to interpret the data. D.-W.H. and D.L. conceived of the study, participated in its design and coordination, and helped to draft the manuscript. All authors read and approved the final manuscript.

**Funding:** This work was supported by a 2-year research grant from Pusan National University.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

© 2018 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Review*