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Current in vitro methods in nanoparticle risk assessment: limitations and challenges

Alexandra Kroll^a, Mike H. Pillukat^a, Daniela Hahn^a and Jürgen Schnekenburger ^{, a,}

^aDepartment of Medicine B, Westf?lische Wilhelms-University, Münster, Germany

Abstract

Nanoparticles are an emerging class of functional materials defined by size-dependent properties. Application fields range from medical imaging, new drug delivery technologies to various industrial products. Due to the expanding use of nanoparticles, the risk of human exposure rapidly increases and reliable toxicity test systems are urgently needed. Currently, nanoparticle cytotoxicity testing is based on in vitro methods established for hazard characterization of chemicals. However, evidence is accumulating that nanoparticles differ largely from these materials and may interfere with commonly used test systems. Here, we present an overview of current in vitro toxicity test methods for nanoparticle risk assessment and focus on their limitations resulting from specific nanoparticle properties. Nanoparticle features such as high adsorption capacity, hydrophobicity, surface charge, optical and magnetic properties, or catalytic activity may interfere with assay components or detection systems, which has to be considered in nanoparticle toxicity studies by characterization of specific particle properties and a careful test system validation. Future studies require well-characterized materials, the use of available reference materials and an extensive characterization of the applicability of the test methods employed. The resulting challenge for nanoparticle toxicity testing is the development of new standardized in vitro methods that cannot be affected by nanoparticle properties.

Keywords: Nanoparticles; Nanoparticle properties; In vitro toxicology; Particle assay interference; New toxicology detection systems

Article Outline

1. Introduction

2. General use and limitations of in vitro assays in toxicology

3. Current in vitro cytotoxicity assays

3.1. Cell viability

3.2. Stress response

3.3. Inflammatory response

4. Nanoparticle properties influencing in vitro toxicity assays

5. Challenges for nanoparticle in vitro test methods

Acknowledgements

References

1. Introduction

Nanoparticles are generally defined as engineered structures with at least one dimension less than 100 nm. They are manufactured as multifunctional particles in a variety comparable to that of conventional chemicals. In recent years, an increasing number of products composed of these tiny structures have entered the market. Metal oxide nanoparticles such as TiO₂, for instance, are frequently used in products which range from cosmetics to food packaging [1].

Nanoparticles also offer an extraordinary opportunity for applications in pharmacology and medicine. Carbon nanoparticles, such as fullerenes (e.g., C₆₀), or carbon nanotubes are currently tested for site-specific targeted drug delivery [2] and [3]. Nanoshells, composed of a dielectric core such as silica surrounded by a metallic shell (e.g., Au, Cu), can act as photoabsorbers and have been successfully used for photothermal ablation of tumours in mice [4]. Additionally, they have been applied in bioimaging with optical and photoacoustic tomography (for review see [5]). Even more attractive for the use in medical diagnostics are quantum dots (QDs), semiconductor nanocrystals with unique fluorescent properties. QDs of different sizes are excitable with one single light source but – depending on their size – emit light at different wavelengths, which enables an optical “bar-coding” of targets [6] and [7]. Polymer nanoparticles, like chitosans, have been used in gene therapy [8], drug delivery [9], and water treatment [10].

With the ongoing commercialization of nanotechnology products, human exposure to nanoparticles will dramatically increase and an evaluation of their potential toxicity is essential. A number of manufactured nanoparticles have recently been shown to cause adverse effects in vitro and in vivo (reviewed by [11], [12] and [13]). Their large surface area per unit mass may lead to an increased biological reactivity that enhances any intrinsic toxicity. Nanoparticles of TiO₂, for instance, have been shown to induce a much greater pulmonary–inflammatory response than larger particles of the same chemical content at equivalent mass doses [14]. Consequently, screening techniques commonly used for toxicity testing of macro-scale substances may not be appropriate for nanoparticle hazard characterization, but may have to be adapted or modified with regard to their nanospecific properties [15].

Although numerous in vitro nanotoxicity studies have already been published, most of the experiments carried out so far have used particles not well characterized regarding their composition and physicochemical properties. However, such a characterization is mandatory since nanoparticles might interact with assay components or interfere with detection systems resulting in unreliable data [16]. In this review, we shortly reflect the general use and limitations of in vitro assays in toxicology, provide an overview of currently used in vitro cytotoxicity methods, and focus on nanoparticle properties that likely interfere with these in vitro test systems. Furthermore, we discuss novel technologies enabling marker-free toxicity testing that may provide new approaches to nanoparticle risk assessment.

2. General use and limitations of in vitro assays in toxicology

In vitro toxicity assays are primarily utilized to investigate the generic cytotoxicity or genotoxicity of chemicals. While the traditional application of in vitro tests has been screening of chemicals, recent developments in molecular biology have helped to provide mechanistic information on toxicity [17]. The use of in vitro toxicity methods for the assessment of nanoparticles was a subsequent development and the applicability of a prevalent in vitro test to nanoparticles is reviewed below. Sutter et al. identified six general applications of in vitro toxicity assays, among which are the selection of the most appropriate animal model of humans and the rapid screening of series of toxicants [18]. In comparison to animal models, in vitro assays allow for a simpler, faster and more cost-efficient assessment of defined toxicity endpoints [19]. However, in vitro test systems lack the complexity of animal models or the human body [17], and the metabolic activity of standardized cell lines has often not been comprehensively characterized [20]. According to Snodin et al., in vitro systems have no value for the prediction of biodistribution and target organ toxicity for the applied chemical and its metabolites [21]. Furthermore, a dose–response relationship will probably only be reliably determined in vivo [22]. Besides these considerations, scientists have been striving to determine the correlation between the results obtained from in vitro and in vivo toxicity assessments. In the domain of particle toxicology, for example, Sayes et al. found little correlation between in vitro and in vivo pulmonary toxicity of several fine and nanoparticles [23]. On the other hand, Donaldson et al. reported that the threshold for inflammation onset was identical in vitro and in vivo when the “particle surface area burden per unit of proximal alveolar region surface area” of different low-toxicity, low-solubility particles was used as reference [24]. Contradictory in vitro results on the same test substance accumulating in the literature as well as the continuous public concern for animal welfare and safe handling of substances have driven diverse international efforts to standardize and validate in vitro tests (e.g., [25]). The OECD validation harmonization report [26], programs as the MEIC (Multicenter Evaluation of In vitro Toxicity, e.g., [27]), the German Center for the Documentation and Validation of Alternative Methods (ZEBET), and standard protocols made available via the INVITTOX database run by the European Centre for the Validation of Alternative Methods (ECVAM), for example, are important aspects of an international standardization and validation process (reviewed e.g., by [28]).

In vitro assays have general limitations for reliable risk assessment in the aspects of validation by in vivo experiments and a lack in standardized testing procedures, which are verified by reference materials or interlaboratory validation.

3. Current in vitro cytotoxicity assays

3.1. Cell viability

Cell viability is the most commonly investigated parameter in cytotoxicity testing. As cell viability is determined by various cellular processes, different endpoints are currently utilized to assess the actual state of cultured cells in vitro.

Detection of mitochondrial activity. The colorimetric MTT assay is a widely used cell viability assay based on the reduction of the yellow tetrazolium dye 3-(4,5-dimethylthiazol-2-yl) -2,5-diphenyltetrazolium bromide (MTT) to a purple water insoluble formazan in cells bearing intact mitochondria [29] and [30]. It has been improved and applied in numerous cytotoxicity studies [31], and employed to validate other methods [32] and to determine nanoparticle toxicity. Results from MTT assays have been reported, e.g., for nanoparticles composed of titanium dioxide [33], iron oxide [34], zinc oxide [23], chitosan [35], silica [36], for fullerenes [37], and for naked or chitosan-coated QD [38]. Assay protocols described in the literature differ strongly [39], which has to be considered when comparing results reported by different laboratories. The absorption spectrum of reduced MTT is pH-dependent [40], and metal ions such as Zn²⁺ interfere with the MTT reduction reaction [41]. Nanoparticles may interact with the substrate, thereby depleting free MTT and causing false negative results as described for single-walled carbon nanotubes (SWCNTs) [42], [43] and [44] (Table 1). Therefore, the suitability of MTT for in vitro toxicity assessment of SWCNTs has come into question [45]. Due to their optical properties, nanoparticles present in the reaction mixture, in or on cell culture cells may directly influence the readout by increasing the light absorption, which has already been demonstrated for sodium titanate nanoparticles [46].

Table 1. Nanoparticle interference with cytotoxicity assays

Detection of LDH release upon necrosis. The colorimetric lactate dehydrogenase (LDH) assay which is based on the oxidation of the yellow tetrazolium salt INT to a red formazan has a long tradition in the clinic to evaluate tissue or cell damage [47] and [48]. As significant amounts of LDH are released from the cytosol upon cellular necrosis [49], LDH activity is measured in the cell culture supernatant. In the cellular metabolism, LDH catalyses the balance reaction pyruvate – lactate generating NADH+H⁺ [50]. Instead of pyruvate, INT (2-(p-iodophenyl)-3-(p-nitrophenyl)-5-phenyltetrazolium chloride) is used as an alternative substrate for dehydrogenases such as LDH [51], lactyl-CoA dehydrogenase [52], and succinate dehydrogenase [53].

LDH assays using INT as substrate have been applied to assess the cytotoxic potential of various nanoparticles, for instance, silica [54], iron oxide [34], TiO₂ [55], ZnO [23], SWCNTs [56], and fullerenes [57].

LDH is significantly deactivated under low pH conditions [51], whereas a high basic pH destabilizes the substrate [58]. Furthermore, metal ions (e.g., copper) have been shown to interfere with the LDH assay [59]. Single-walled carbon nanotubes, however, do not seem to interact with the substrate INT [42].

Annexin V/propidium iodide staining for apoptotic and necrotic cells. Annexin V (VAC alpha), which is regularly used to detect apoptotic cells [60], binds strongly to phosphatidylserine in a calcium-dependent manner [61]. Phosphatidylserine is normally excluded from the extracellular side of the plasma membrane [62], but flips between the inner and the outer side upon the onset of apoptosis [63]. Fluorescently labelled Annexin V can therefore be used to detect apoptotic cells [64]. Necrotic cells will allow Annexin V to bind PS on the inner part of the plasma membrane resulting in false negative results due to cell disintegration. Hence, cells have to be co-stained with propidium iodide (3,8-diamino-5-[3-(diethylmethylammonio)propyl]-6-phenylphenanthridinium diiodide) which will exclusively stain necrotic cells [65] and [66]. Investigation of apoptosis via Annexin V staining and flow cytometry has been reported, e.g., for TiO₂ nanoparticles [67], for pure and polyhydroxylated fullerenes [68], for SWCNTs [42], and for quantum dots [69].

As apoptotic cells may easily detach from their substrate, it is essential to collect adherent and floating cells for Annexin V staining [70]. Consequently, nanoparticles in the cell culture supernatant will be present in the cell suspension despite several washing steps. Gold nanoparticles have been shown to bind propidium iodide and to be taken up by intact cell culture cells. This process leads to false positive results in the detection of necrotic cells [71] (Table 1).

Detection of intact lysosomes via neutral red uptake. Since 1894, neutral red has been used as a viability stain [72] and has since been implemented in numerous cytotoxicity, cell proliferation and adhesion assays [73] and [74]. Neutral red (3-amino-7-dimethylamino-2-methylphenazine hydrochloride) is weakly cationic, and is thought to be taken up into the cytosol by non-ionic diffusion through the cell membrane to then accumulate in the lysosomes of viable cells, whereas it is excluded from dead cells [75]. The uptake of neutral red may be detected via fluorescence or absorption measurement. So far, the neutral red uptake (NRU) in NIH3T3 mouse fibroblasts is the only validated in vitro method for toxicity testing [76] and has been incorporated into the REACH (Registration, Evaluation, Authorisation of Chemicals) directive by the European Commission for the in vitro toxicity assessment of chemicals [77]. A number of different nanoparticles such as TiO₂ [78], SWCNTs [43], multi-walled carbon nanotubes (MWCNTs) [79], and chitosan nanoparticles [35] have been tested in NRU assays. The colour and intensity of light emission of neutral red is pH-dependent [80]. It is known that neutral red has a high affinity to lipophilic structures (such as suberin and lipids [81] or phenolic substances [82]), the protonated dye interacts with negative charges [82], and covalently binds to cellular structures [81]. These properties were utilized to study the adsorption of neutral red to mercaptoethane sulfonate-protected gold nanoparticles. The optical properties of neutral red were significantly influenced, the nanoparticles exerted static and dynamic energy transfer quenching of neutral red fluorescence [83]. Furthermore, SWCNTs have been shown to interact with neutral red and deplete the dye from the cell supernatant leading to false positive results [45] (Table 1). Hence, properties that are useful for technical applications may be deleterious in cytotoxicity testing.

Detection of the apoptosis marker Caspase-3. The detection of active Caspase-3 is one of the most commonly used apoptosis assays. Apoptosis may be triggered by different elicitors activating two main signalling cascades that converge in the activation of Caspase-3 [84]. The cystein protease Caspase-3 is produced as a zymogen in the cytosol and is activated in the terminal apoptotic cascade by cleavage [85]. As soon as Caspase-3 is activated, cell death is inevitable. Activated Caspase-3 can be detected by measuring the cleavage of a Caspase-3 substrate (preferably the aminoacids DEVD) linked to a chromophore (pNA) or fluorophore (AFC, AMC) that absorbs or emits light when separated from the substrate [86]. As yet, the Caspase-3 assay has been utilized to examine apoptosis in cell culture cells upon exposure, e.g., to fullerenes [87], SWCNTs [88], silica nanoparticles [89], quantum dots [90], and to TiO₂ nanoparticles [91].