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European Journal of Nanomedicine

Editor-in-Chief: Hunziker, Patrick / Mollenhauer, Jan

Managing Editor: Löffler, Beat / Salieb-Beugelaar, Georgette

Editorial Board: Alexiou, Christoph / Balogh, Lajos / Barenholz, Yechezkel / Dawson, Kenneth / Fadeel, Bengt / Husseini, Ghaleb / Krol, Silke / Lee, Dong Soo / Lehr, Claus-Michael / Mangge, Harald / Müller, Bert / Peer, Dan / Scoles, Giacinto / Serruys, Patrick / Schwartz, Simo / Szebeni, Janos

CiteScore 2016: 1.06

SCImago Journal Rank (SJR) 2016: 0.411
Source Normalized Impact per Paper (SNIP) 2016: 0.360

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Advanced in vitro systems for efficacy and toxicity testing in nanomedicine

Maria Rita Fabbrizi / Tracey Duff / Jo Oliver / Colin Wilde
Published Online: 2014-08-23 | DOI: https://doi.org/10.1515/ejnm-2014-0018


The use of nanoparticles (NPs) in nanomedicine is becoming an established therapeutic strategy for transport and delivery of bioactive molecules across biological barriers to elicit a specific cytological response in a target cell population. The utility of NP-derivatised nanomedicines and their potential therapeutic benefits, together with their biosafety, are being explored in cell models that, in order to accurately predict clinical potential, must reflect the architecture and function of the tissues from which the cells originate. These cell-based models, and their application in preclinical nanomedicine, are the subject of this review. Models are considered expressly from a commercial perspective, for their use as screening tools to identify and optimise therapeutic constructs, rather than as research tools requiring minimal throughput. Commercial utility is discussed in terms of the sourcing and assembly of cell-based model components, likelihood of data outputs highly predictive of clinical performance, and the compromise between need for advanced 3D culture architectures and the incremental difficulty in assembling those structures cost-effectively for medium-throughput analysis. The relative merits of cell lines and primary human cells are discussed, the latter with respect to their potential physiological relevance when cultured under permissive conditions conferred by a biologically-rich micro-environment. Tools for assembly of such microenvironments are reviewed, and the practitioner’s choices of commercial products enabling 3D cell culture are also critically evaluated, from the merits of cell aggregates and micro-tissues to the architectural attractions of recreating epithelial monolayers and multi-cell type reconstituted tissues. Additionally, pitfalls and practical challenges in co-assembly of cells and scaffolds/matrices are considered, with the intention of signposting routes to consistent, scalable production of 3D cell cultures capable of screening the nanomedical potential of NP-conjugated therapeutics.

Keywords: cell-based systems; commercial platforms; nanosafety


Nanomedicine can be defined as the design and development of diagnostic agents and/or therapeutics in the nanoscale dimension (with diameters ranging from 1 nm to 100 nm), with the possibility to transport and deliver a variety of biomedical molecules for the prevention, diagnosis and treatment of many diseases (1–5). Nanoparticles (NPs) have physicochemical properties that differ fundamentally from the bulk materials used for their synthesis, including their size, shape, surface area, aspect ratio, surface functionalisation, coating and charge, zeta potential, crystallinity, dissolution and redox potential. These properties can determine NP biological safety or otherwise (6); thus, a detailed understanding of the pharmacologic and toxicological properties of these nanomaterials, and a balanced and detailed evaluation of their risks and benefits to human health, are expected before their translation into clinical use (7).

The efficacy of a NP as a transporter and delivery molecule, and the potential toxicity it could engender in a biological environment, can be analysed through three different approaches: 1) By analysing the cellular response after NP treatment in-vitro, using immortalised cells or primary cell cultures (8, 10); 2) By experiments in-vivo, with the NP administered directly to experimental animals by various routes (11); 3) By ex-vivo experiments, harvesting tissue from human or animal sources followed by culturing on permissive substrata in culture wells or on coverslips (for tests based on microscopic analysis) (9, 11). Since a clear definition of “ex-vivo is not currently available, in this review we will consider these experiments as part of the in-vitro category.

The use of animals for experimental purposes has been widely criticized on both ethical and practical grounds, and many efforts have been made to limit their use in current applications of nanosafety testing. Although some research groups report that the response observed in-vitro is often far from the one observed in-vivo, advances in cell culture technology are progressively improving and enhancing in vitro models to the point where they can realistically claim to exhibit biological functions reflective of living tissue, and thereby predict the performance of test agents in the human body (12–14).

In 1951 George Otto Gey developed the first immortalised cell culture from a biopsy of human cervical cancer. Since then, researchers have found cell cultures to be a valid instrument for mimicking and analysing human biology. Enormous progress has been made in relation to extending the analytical repertoire, improving sensitivity and predictive value, and making it available to industry as an ethic reliable tool for screening candidate drugs and other therapeutics. Nowadays, protocols exist for cell isolation from tissue for the isolation and handling of progenitor cells potentially capable of tissue-specific (differentiated) function, and for the creation of long-lived cell lines from original populations (15–23). The formulation of such protocols has expanded the repertoire of the cell biologist to unprecedented – and sometimes previously unimaginable – extents.

Nano-scientific research has benefitted fundamentally from this proliferation of technical opportunity, and the literature describing NP testing on cell culture systems is substantial. So, e.g., NPs have been tested for their ability to penetrate the cell membrane (24–27), ability to induce specific cytological response (28–30), potential cytotoxicity, genotoxicity, carcinogenic potential (31–38), and to produce critical information for assessing the utility and safety of these novel materials. Moreover, NPs have been tested for their ability to carry and release specific molecules in a target cell/tissue/organ, e.g., the delivery of a chemotherapeutic agent into a target tumour cell (24, 39–43). Fundamental to the evaluation of nano-medicines, NPs have been tested extensively for their ability to target cell-biological events for diagnostic purposes (44, 45) (Figure 1). Many in-vitro systems have been used for NP testing, and there is extensive debate about their relative merits, a debate that usually revolves around whether the cell model generates a response that closely mimics the in-vivo observation. There are different schools of thought as to the most attractive approach, and yet more debate as to how, in practice, to make models with high predictive value available to the scale required by industrial end-users. The aim of this review is to compare the in-vitro systems already used for testing the efficiency and toxicity of NPs in terms of their relative advantages and disadvantages, and to review the new approaches recently developed and the commercial systems about to enter the marketplace.

NPs are used in nanomedicine for their ability to carry and release specific molecules in a target cell/tissue/organ, for their ability to target cell-biological events for diagnostic purposes.
Figure 1

NPs are used in nanomedicine for their ability to carry and release specific molecules in a target cell/tissue/organ, for their ability to target cell-biological events for diagnostic purposes.

In vitro systems: primary cells vs. immortalised cells

Across biomedical research, and in studies ranging from cancer drug screening to developmental molecular biology, the default experimental approach has been to use secondary cells, i.e., cell lines (46). They are easy to handle, readily-propagated, and relatively consistent in performance. Their disadvantage is that they acquire these features whilst losing other important characteristics, typically their tissue-specific functionality. That is, cell lines are often derived from proliferating cells in tumours, or have adapted to favour growth in culture over other cell behaviours. Thus, cell lines can often lack tissue-specific functions that influence the process under study, and may acquire a molecular phenotype quite different from cells in-vivo, especially with extended passaging (47). The result is that, whilst results can be generated relatively easily and at low cost, those results may lack predictive value, i.e., the behaviour of a process under study or a material under test may not reflect the impact of that process or material in the human body. Reports based on secondary cell cultures (cell lines) may therefore, at best, warrant verification in-vivo or in more physiological in vitro contexts, and at worst, could lead to unjustified preclinical or product-safety decisions. When this happens in the context of pharmaceutical drug discovery, the potential clinical implications can be significant, and the commercial impact of subsequent failure of candidates selected using cell line data can be expensive. The same approach to in vitro testing is likely to have similar significant human health implications (and very possibly commercially-expensive consequences) in the field of nano-medicine and nanosafety testing. In short, the same caveats of physiological relevance and predictive value are likely to apply in the field of nanotoxicology and nano-efficacy testing, with reports of consistency between in vivo and in vitro results from immortalised cell lines (48–57) being fundamentally compromised by cross-species extrapolation unjustified by model biochemical divergence.

The shortcomings of cell lines can be avoided by the use of primary cell cultures freshly-prepared from living tissue. Under the right conditions, these cells can retain tissue-specific functionality, and therefore demonstrate a repertoire of cellular functions likely to increase the predictive value of data gathered from the cells’ use as screening tools, or as models for biomedical research (58, 59). This attractive advantage comes, however, with the greater technical challenge of, firstly, making these cells and, if necessary, banking them ready for use and, on the other hand, ensuring that the “right conditions” are found to ensure retention of that advantageous functionality. There is also the significant drawback, in some applications, of limited cell-pool size, and low propagation potential, which means that extended studies may require use of several batches from different donor tissues, requiring careful use of normalising controls. Also, the incentive to passage primary cells and increase pool size must be weighed against the possibility/likelihood that passaging is likely to be inversely related to tissue-specific functionality (60, 61). On the other hand, cell batches from different donor tissues can be used to explore the biological variation of response to challenge within tissues ostensibly phenotypically similar, but genotypically different.

Many studies testing NP effects on primary cell cultures have already been published and the results obtained offer an interesting comparison with those obtained using secondary cell lines as they often show significant differences between the responses of primary and immortalised cell cultures. For example, a toxic effect has been observed in primary hepatocytes cultures from rabbit (62) and bovine tissue (63) after Se NP treatments, whilst no toxic response has been found in immortalised hepatocytes (64). Also, primary human blood cells showed a higher grade of toxicity (65) compared to the secondary cell lines (66, 67) after exposure to nanosized hydroxyapatite. These discrepancies could derive from the loss of functions that secondary cell cultures must inevitably encounter, due to genomic alterations or as a result of repeated passages in culture, which is likely to select cells with a particular culture-conditioned biochemical phenotype. However, it should be noted that, a correlation between the response obtained by using primary and immortalised cells has been observed in some situations, including studies testing NPs with a well-known grade of cytotoxic activity, i.e., Ag NPs and Zn NPs (68–74).

In vitro systems: 2D vs. 3D

Most in-vitro research into nanotoxicology is performed in 2-dimensional cell cultures i.e., on flat surfaces of cell culture vessels (Figure 2A) (75). Clearly, this is a wholly artificial environment: in a living organism, cells are located in a three-dimensional microenvironment, which is crucial for the cells’ behaviour, and controls their proliferation (if appropriate), normal metabolic activity, and death in response to physiological signals or injury (76). Importantly, the lack of a 3D environment, or even the absence of appropriate cell anchorage to a 2D surface, can cause abnormalities in 2D primary cell function that essentially eliminate the potential advantages of adopting a primary cell-culture model in the first place (77). There are now numerous examples of the use of a 3D microenvironment as the crucial element in improving in-vitro disease models for cancer research and nanotoxicology, and these increasingly offer a meaningful alternative to ethically-unattractive animal testing, as well as a physiologically-relevant advance over conventional 2D culture models (78–85). As an example, in case of cancer cells, a key characteristic is their acquisition of resistance to a variety of anticancer drugs through a variety of mechanisms. There is good evidence that unicellular mechanisms of resistance of cancer cells in 2D culture may be different to the intrinsic resistance of cells observed in 3D culture or in-vivo (86). To address the shortcomings of conventional 2D culture, there is now widespread investment in three-dimensional culture methods for a range of cell types from a variety of tissues whose organ architecture creates particular incentives to take into account the spatial organisation of the cells (87, 88).

Schematic representation of (A) two-dimensional cell culture; (B) three-dimensional cell culture; (C) spheroid.
Figure 2

Schematic representation of (A) two-dimensional cell culture; (B) three-dimensional cell culture; (C) spheroid.

3D cell culture (Figure 2B) has the potential to recapitulate the architecture, function, and cellular biochemistry of living tissues, including cell-cell communication and cell-extracellular matrix (ECM) interaction, which played a key role in cancer aetiology (89, 90) and multidrug resistance (91, 92). It is increasingly evident that this can translate into greater fidelity of response to challenge by, e.g., cytotoxic / nanotoxic agents, which better predict the behaviour of those agents in the human body. A greater enhancement in therapeutic resistance to anticancer drugs (i.e., greater increase in IC50) has been observed in cells cultured in 3D compared with a 2D environment (86, 93), probably due to the higher cell packing density and larger spheroid structure, which can cause the block of diffusion and penetration of drug (94, 95).

Strategies for building 3D culture models varies from a minimalist approach in which cells self-assemble to the most complex of pre-formed scaffolds, some of which require equally advanced cell-handling in order to achieve the desired functionality and capture the ultimate assay readout. The minimalist approach is most usefully illustrated by models based on cellular spheroids (Figure 2C); these are simple three-dimensional models that can be generated from a wide range of cell types and form due to the tendency of adherent cells to aggregate. They are typically created from single culture or co-culture techniques such as hanging drop, rotating culture, or concave plate methods (96–98). Spheroids can readily be imaged by light, fluorescence, and confocal microscopy, but also lend themselves to high-throughput outputs (99). Consequently, they have seen use not only in modelling solid tumour growth and study of metastasis but also in therapeutic-discovery studies. In nanomedicine, tumour spheroids have been used to test the ability of NPs loaded with chemotherapy drugs to cause cell death in a context that can better mimic the cancer in-vivo: Ho and colleagues found magnetite nanoparticle agglomeration stimulated by high-intensity focused ultrasound can cause cell lysis and disintegrate the whole tumour spheroid (100). Other studies analysed the different responses of monolayer cell cultures and spheroids, suggesting the latter as a better model to simulate in-vivo tumour tissue and evaluate nanoparticle penetration behaviour (13, 101, 102).

The demonstrable advantages of this minimalist 3D approach have encouraged elaborations of the models using biomaterials technology, in order to control more precisely the location of cells within 3D environment, whilst retaining the capacity for multiwell plate-enabled medium or high throughput (46). In many instances, 3D models have been developed to meet a specific experimental objective, and whilst having utility and demonstrable biological advantage, were not designed to address fundamental attributes of consistency in manufacture and use demanded of commercial products, or indeed products which can form the basis of a standardised platform for nano-medicine evaluation or nanoparticle testing. However, a number of 3D cell culture technologies are commercially available, with features that address different needs of the research community. The Perfecta3D® Hanging Drop Plates from 3D Biomatrix (Ann Arbor, MI, USA) is suitable to test embryoid bodies that should not make cellular contact with artificial surfaces or matrices (103, 104). In other products, 3D scaffolds fabricated from biodegradable materials with well-defined pores size can be used in studies focused on stem cells, tumour models and – important from a biodegradable perspective tissue engineering. In a more well-established example, Corning®, Transwell® (Corning, Tewksbury, MA, USA) permeable supports have become a standard method for culturing cells by permitting cells to uptake and secrete molecules on both its basal and apical surfaces (105, 106). Insphero GravityPLUS (Insphero, Schlieren, Switzerland) can be used for scalable microspheroid assembly applicable in toxicology and oncology applications (107, 108). Less popular, but arguably with their own attraction of pre-assembly and structural flexibility, is the process of fibre electrospinning which offers control over product architecture and can be exploited to customise not only the scaffold porosities and depths, but also selection of base polymer, and the option to functionalise polymer chemistry. Fibre-based scaffold models manufactured in multiwell plate formats, e.g., Mimetex from The Electrospinning Company (Didcot, UK) involve cell loading into consistent, pre-formed structures, and require expertise and protocols which ensure that cells remain within the scaffolds (109). The risk is that incomplete loading can create 2D and 3D cell populations within a single culture well. Nevertheless, with judicious scaffold design, cell loading can be optimised and 3D cell performance ensured: recent work using electrospun scaffolds and human breast cancer cells demonstrated the reinstatement of resistance to drug-induced apoptosis in these cells by 3D culture, reflecting their in vivo sensitivity and not the elevated apoptotic response exhibited by the same cells in 2D culture (110).

In conclusion, 3D technology has been proven, easy-access offerings have certainly multiplied, diversified and have undoubtedly enhanced both the biological advantages of 3D culture, and removed a significant barrier to entry represented by the initial investment previously needed to develop such models. However, there still appears to be some latent hesitancy in the market, such that uptake of 3D culture technology does not appear to be developing at quite the pace as might have been predicted, given the level of interest there seems to be in 3D methodology at the bench, with cost implication appearing still to be a deterrent (111).

New approaches

NP presentation

Analytical platforms testing NPs or NP-functionalised candidate therapeutics should take account of the conditions under which NPs and cells come into contact. Clearly, a static model wherein NPs presented in solution or suspension is presented once onto a 2D cell culture lacks the dynamic interactions that occur in vivo. Moreover, cell models, and nanotoxicological assays using those cell models, can be adversely affected by NP interference with the assay readout (112–119). NP presentation to more complex models may pose challenges when the NP format is incompatible with the cells’ microenvironment, e.g., the presentation of NP in aqueous solution or suspension to lung epithelial cells at an air:liquid interface (120, 121). As an alternative, particles can be nebulised over the air-exposed system using a spraying device (e.g., a Microsprayer®, Penn Century, Wyndmoor, PA, USA) (120) or an exposition chamber with an integrated aerosol generator (122). New cultivation and exposure techniques enhance the efficiency of in-vitro studies, as demonstrated by one new experimental system called CULTEX (Cultex®Laboratories GmbH, Hannover, Germany) which allows direct exposure of cells at the air/liquid interface. In this model, with human bronchial epithelial cells are cultivated on porous trans-well membranes in a device allowing intermittent medium supply by pumping it into a special modular culture unit through the trans-well membrane supporting the cells. The introduction of these new cultivation and exposure techniques offers new testing strategies for the toxicological evaluation of inhalable soluble and inert substances as well as complex mixtures (123).

Another improved in-vitro assessment method for evaluating quantum dots (QD) toxicity has been developed as a multi-compartmented microfluidic device integrated with a syringe pump, utilized in order to establish a sensitive flow exposure system capable of exposing cultured cells to variable concentrations of QD. Results with this model suggested noticeable differences in the number of detached and deformed cells, as well as the viability percentages compare to static exposure, with the flow exposure condition resembling in-vivo physiological conditions very closely. Thus, this approach can offer potential advantages for nanotoxicity research (124).

Co-culture models

3D structure is important to mimic better a real tissue environment and thereby deliver results that are predictive of behaviour in-vivo. It is also the case that cells do not function in isolation, but react to paracrine and juxtacrine signals from nearby cells. Accordingly, co-cultures of different cell types have been shown to influence fundamentally the outcome of NP testing in vitro. For example, epithelial cells, macrophages and dendritic cells continuously interact in-vivo through intercellular signalling; this cross-talk maintains homeostasis and coordinates immune responses (125). A triple cell culture in-vitro model of the human airway wall was developed to study the cellular interplay and the cellular response of epithelial cells, human blood monocyte-derived macrophages and dendritic cells to polystyrene particles (1 μm in diameter). In this model, monolayers of two different epithelial cell lines, A549 and 16HBE14o epithelia, were grown on a micro-porous membrane in a two-chamber system (126). The same group has recently modified this system by using primary alveolar type I-like cells isolated from human lung biopsies combined with two immune cells (i.e., macrophages and dendritic cells). Comparing this system with the one built with immortalised cells, the authors suggested that the triple cell co-culture model composed of primary cells offer a novel and more realistic cell co-culture system to study possible cell interactions of inhaled xenobiotics and their toxic potential on the human alveolar type I epithelial wall (127).

In vitro perfusion models

Few studies have used the protocol of the ex-vivo perfusion model, by which the mechanism of nanoparticle transport through a target organ can be studied in a situation close to that in-vivo. One disadvantage of such systems is that they allow only acute studies of a few hours: chronic treatments with low doses over a long period are not possible because of tissue degradation. The perfusion technique has been utilized to investigate whether fluorescently labelled polystyrene beads (PS NPs) with diameters of 50, 80, 240, and 500 nm can cross the human placental barrier and whether this process is size dependent: the study showed a clear size-dependent barrier capacity of a healthy human placenta for PS NPs (128, 129). A perfusion method has also been utilized in isolated perfused porcine skin flap, to quantify the local bio-distribution of two types of Ag nanoparticles, Ag-citrate and Ag-silica. This model was designed to closely reflect an in vivo situation: the perfusion flow was as close as possible the in-vivo situation of the skin compartment and the media concentrations of the two Ag nanoparticles, 0.840 mg/L and 0.480 mg/L for Ag-citrate and Ag-silica, respectively, used in this study were a similar magnitude compared with plasma concentrations noted in humans exposed to Ag-containing products (130). The study concluded that there is no difference in the distribution between the Ag-citrate and the Ag-silica particles. Apart from studies on placenta (131) and skin (132, 133), in vitro perfusion has been used to test the effects of NPs in other organs, such as lungs (134) and heart (135, 136). Despite promising results, this technique is not commonly used, probably because of the difficulty of maintaining the system and the limited lifespan of the models, and, of course, the need for a human or animal source from which to draw the organ or tissue.

Single-cell models

This review reflects a general trend towards NP evaluation in more complex in-vitro (i.e., 3D cultures, co-cultures) that mimics the in-vivo situation. In contrast a recent study proposed a new approach, using single cell mechanics derived from atomic force microscopy-based single cell compression. Its reliability and potential to measure cytotoxicity was evaluated using known systems: zinc oxide (ZnO) and silicon dioxide (SiO2) nanoparticles (NP) on human aortic endothelial cells, and indicated the reliability of single cell compression. For example, ZnO NPs caused significant changes in force versus relative deformation profiles, whereas SiO2 NPs did not. The authors reported that, even though this method does not provide specific chemical information (such as markers associated with cellular signalling cascades) the single cell based approach is intrinsically advantageous, as results appear to corroborate in-vivo investigations (137).

Commercial analytical platforms

The increasing demand for cell cultures in 3D formats has seen a resultant increase of investment in R&D focused on “smart”, i.e., functionalised scaffold materials (138), their scalable production, and their assembly for use in multi-well plate formats required by most commercial users. Some laboratories opt for scaffolds made in-house, but many groups rely on commercial scaffolds created by standardised procedures and quality-controlled manufacture. There is no accepted standard for NP evaluation (139) and, not unreasonably, pragmatic decisions are made on the basis of convenience and desired functionality (140–151). For example, Hanada and colleague used an in-vitro rat BBB model (RBT-24H, BBB Kit; PharmaCo-Cell Company Ltd., Japan) to test the ability of fluorescent silica NPs (30, 100, and 400 nm) to cross the BBB (152). The BBB model was based upon co-cultures of primary rat brain microvascular endothelial cells and pericytes separated by a macroporous 3.0 μm Millicell® membrane (Millipore, USA), which were pre-cultured with rat astrocytes to support the tight junctions of the BBB cell layers. The 30 nm silica nanoparticles were transported through the BBB model, mirroring the same result reported in an animal model (153, 154).

Another challenge in meeting industry need for high-value but user-friendly means to test NP bioactivity lies in the degree of operator input required. Clearly, such NP evaluation could be supplied as a service, with complex models being assembled and used in-house by their developers, or by specialist contract service providers. This means of analytical technology delivery would be in keeping with market intelligence, which suggests that the cell-based analysis service sector is likely to grow faster than the sector as a whole (155). To put this prediction in the context of present market scale, the cell-based analysis market, which was valued at $12.1 Billion in 2013, will reach $15.1 Billion in 2015, and is set to reach nearly $21.6 Billion by 2018, registering a 5-year CAGR of 12.4% (156). On the other hand, NP testing, especially that associated with assessment of environmental risk, sits at the other end of the spectrum of analytical options, where testing needs to occur at, or close to, the point of sample collection, or at least needs to sit comfortably in a laboratory environment where cell-based analysis is not the primary skill-set. For this, there appear to be a number of ancillary culture technologies that need to sit alongside the core multiwell culture protocols. These ancillary technologies need to enable supply of cell-based analytical tools to end-users in forms which demands the minimum of operator input, so that lack of operator expertise is de-risked. In practice, this may translate into the supply of assay components which are easily prepared and convenient to use. It may also be achieved by the supply of cells in a format most easily handled prior to dispensing into the test system, and by a protocol which demands little subsequent manipulation of cells in that system. A variety of strategies are being adopted to deliver this convenience and utility. One approach is to supply cells in pre-plated form, with the cells stabilised during transport from producer to user in chilled, gelled formats; e.g., cells may be embedded in agarose, which is removed by agarase treatment at physiological temperature. A number of manufacturer offer products based upon this approach, and whilst none is specifically designed for nano-medicine or nanotoxicology testing, extension of the technology to a widening range of cell types progressively increases the potential to deliver NP-relevant assays in this form.

A drawback of gel-based methods for pre-dispensed cell product supply in ready-to-run form is the relatively short shelf life of the product. A solution to this shelf-life limitation can be to supply cells as pre-plated frozen stock, for return the cells to culture in a ready-to-assay form. End-user convenience is counter-balanced by the challenge of reproducibly cryopreserving and thawing pre-plated cells. There are few examples of such utility in the commercial cell-based analysis products presently available, and where these exist, the cell type represented in the product tends to be a relatively robust cell line, rather than a primary human cell with its attendant advantage of physiological relevance and predictive value. Also, these products may be designed around hardware suitable for –80°C storage rather than ultra-low (≤150°C) temperature, with indefinite storage potential. Whilst this represents a daunting set of performance criteria, progress is nevertheless being made using software-managed gradient freezing, precisely-controlled ice nucleation and cold-chain supply integrity based on cryo-stable culture disposables, all offering the standardisation and platform reproducibility needed in next-generation nano-medicine and nanosafety testing.

The foregoing discussion assumes that NP testing will be performed in a laboratory environment set up for cell-based analysis. This requirement arguably creates a barrier to routine industry use of cell-based NP testing, and especially to its adoption in non-specialist locations, or as a portable system for environmental nanosafety testing in the field. This industry need for flexibility and portability can be met by use of nano-sensitive cells that inherently do not need the same technical infrastructure of mammalian cells. Accordingly, nanosafety testing has focused on so-called sentinel species that react early and sensitively to contamination of their environment. In one such species, the marine mussel Mytilus, nano-sensitivity reside principally in haemocyte cells, which possess biochemical features of the mammalian immune system in vitro and respond to NPs characterised in other systems as being nanotoxic (157). Importantly, the cells do not require a controlled culture environment, and respond rapidly, within hours, to nanoparticle challenge. Whilst significant technical issues remain in achieving scalable production of a commercially-exploitable platform, mussel haemocyte culture appears to offer the basis for a portable nanosafety assay.


The increasing need for in-vitro models to test the therapeutic potential of nano-medicines and the toxicity on NPs is leading the life science industries to develop cell-based systems which reliably predict the effect these materials have in the human body. The elaboration of cell culture models to achieve this, far beyond conventional methods of a decade ago, or less, has stimulated innovation in biomaterials, as well as grappling with the challenges of isolating and culturing physiologically-relevant cells. This activity nevertheless must take place within the context of end-user convenience, analytical throughput and financial affordability, such that systems may ultimately be constrained by what is achievable in multiwell plate formats, unless prototypic microfluidic and single-cell options mature. In any event in-vitro cell models are emerging which can realistically replace animal testing, with all the ethical advantages and operational efficiencies that will accrue.


This study was funded by the Marie Curie Initial Training Network (FP7-PEOPLE-2013-ITN) under grant agreement n° PITN-GA-2013-608373, project name Pathchooser (http://www.pathchooser.eu).

Conflict of interest statement: AvantiCell Science Ltd (ACS) is a commercial company specialising in cell-based analysis. ACS has conducted commercial business with some suppliers of cell technology critically evaluated in the article. ACS shall not benefit financially from identification of these technology providers, and has at this time no vested interest nor commercial incentive that would influence the written evaluation of the various technologies.


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About the article

Maria Rita Fabbrizi

Maria Rita Fabbrizi studied Physio-Pathological Sciences at the University of Pisa, Italy, graduating in 2009 cum laude with a work on gene polymorphisms involved in amyotrophic lateral sclerosis development. Following graduation, she attended a post-graduation training course at the Medical Genetics Unit of the Department of Translational Research and New Technologies in Medicine and Surgery, University of Pisa, where she also got a PhD in Neuroscience, defending a thesis on cytotoxic and genotoxic effects of several metal nanoparticles in different cell cultures. In February 2014 she joined Avanticell Science Ltd as experienced researched in the frame of the Marie Curie Initial Training, project Pathchooser.

Tracey Duff

Tracey Duff studied Applied Biosciences at Glasgow Caledonian University, graduating in 1998. Following graduation she joined a group within the academic haematology department of the Royal Free Hospital in London, where she worked as a medical scientific officer, focused on haematological malignancies. In 2000 Tracey took up the position of Research Assistant/Study Director within the University of Aberdeen, conducting an in vitro assay development project, focused on producing a platform that could be used to screen molecules for nephrotoxic effects. Following the end of the study Tracey moved into commercial research taking up a position as a Trainee Study Director within the Special Toxicology department of a large GLP accredited CRO. Tracey’s currently works as Project Manager for AvantiCell Science Ltd, an SME focused on cell-based assays for use in drug discovery, biomedical research and natural product testing.

Jo Oliver

Jo Oliver gained a 1st class degree in Biochemistry from University College London, before attaining a PhD in Molecular Biology from the University of Edinburgh. After leading a commercial vaccine development group for some years, she moved to head up a new CRO operating in the field of animal health. Consultancy work for the private and public life science sector followed, including running an extensive programme of grass-roots training, and acting as a ’technology translator for the venture capital community. Jo Oliver is CEO and co-founder of AvantiCell Science Ltd, an SME focussed on physiologically-relevant cell-based products and services.

Colin Wilde

Colin Wilde is Chief Scientific Officer and co-founder of AvantiCell Science Ltd. He has more than 35 years in biological scientific research, and has spent the last 10 years in commercial science, in a senior executive role. He trained in Biochemistry, graduating from the Universities of Surrey and Birmingham, before joining the UK’s Hannah Research Institute to lead research groups specialising in cell biology and cell culture technology. Presently, Colin manages AvantiCell’s contract services, product development, and its programme of research and development in the field of cell-based analysis. He is the author of ten patents or patent applications and >100 research papers in the fields of mammary biology and cell-based analysis.

Corresponding author: Maria Rita Fabbrizi, AvantiCell Science Ltd, GibbsYard Building, Auchincruive, Ayr KA6 5HW, UK, E-mail:

Received: 2014-05-14

Accepted: 2014-07-23

Published Online: 2014-08-23

Published in Print: 2014-09-01

Citation Information: European Journal of Nanomedicine, Volume 6, Issue 3, Pages 171–183, ISSN (Online) 1662-596X, ISSN (Print) 1662-5986, DOI: https://doi.org/10.1515/ejnm-2014-0018.

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