Biomaterials 35 (2014) 5549e5564

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Ultrasmall superparamagnetic iron oxide nanoparticle prelabelling of human neural precursor cells Steven S. Eamegdool a, Michael W. Weible II a, b, Binh T.T. Pham c, Brian S. Hawkett c, Stuart M. Grieve d, Tailoi Chan-ling a, * a

Department of Anatomy and Histology, Bosch Institute, Sydney Medical School, University of Sydney, NSW 2006, Australia Biomolecular and Physical Sciences, Griffith University, QLD 4111, Australia School of Chemistry, Key Centre for Polymers and Colloids, University of Sydney, NSW 2006, Australia d Sydney Translational Imaging Laboratory, Sydney Medical School, University of Sydney, NSW 2006, Australia b c

a r t i c l e i n f o

a b s t r a c t

Article history: Received 26 February 2014 Accepted 21 March 2014 Available online 13 April 2014

Stem cells prelabelled with iron oxide nanoparticles can be visualised using magnetic resonance imaging (MRI). This technique allows for noninvasive long-term monitoring of migration, integration and stem cell fate following transplantation into living animals. In order to determine biocompatibility, the present study investigated the biological impact of introducing ultrasmall superparamagnetic iron oxide nanoparticles (USPIOs) into primary human fetal neural precursor cells (hNPCs) in vitro. USPIOs with a mean diameter of 10e15 nm maghemite iron oxide core were sterically stabilised by 95% methoxypoly(ethylene glycol) (MPEG) and either 5% cationic (NH2) end-functionalised, or 5% Rhodamine B end-functionalised, polyacrylamide. The stabilising polymer diblocks were synthesised by reversible addition-fragmentation chain transfer (RAFT) polymerisation. Upon loading, cellular viability, total iron capacity, differentiation, average distance of migration and changes in intracellular calcium ion concentration were measured to determine optimal loading conditions. Taken together we demonstrate that prelabelling of hNPCs with USPIOs has no significant detrimental effect on cell biology and that USPIOs, when utilised at an optimised dosage, are an effective means of noninvasively tracking prelabelled hNPCs. Crown Copyright Ó 2014 Published by Elsevier Ltd. All rights reserved.

Keywords: MRI (magnetic resonance imaging) Biocompatibility Iron oxide nanoparticles Neural cell Stem cells Regenerative medicine

1. Introduction Neural stem cells (NSCs) in the mammalian brain can give rise to astrocytes, neurons and oligodendrocytes and functionally contribute to cognition and repair processes after nervous tissue injury. Transplanted NSCs are promising candidates for providing beneficial effects on recovery from central nervous system (CNS) trauma and neurodegenerative disorders. However, clinical applications remain elusive, positive-outcomes limited and significant roadblocks to clinical translation remain [1]. Critical shortcomings of current transplantation methodologies are low cellular survival, limited migration through host parenchyma and limited functional integration with the host neuronal circuitry [2]. Typical survival ranges recorded fall between 10 and 20% of engrafted cells [3e5],

* Corresponding author. Discipline of Anatomy and Histology, Sydney Medical School, Anderson Stuart Building F13, The University of Sydney, Sydney, NSW 2006, Australia. Tel.: þ61 2 9351 2596; fax: þ61 2 9351 6556. E-mail address: [email protected] (T. Chan-ling). http://dx.doi.org/10.1016/j.biomaterials.2014.03.061 0142-9612/Crown Copyright Ó 2014 Published by Elsevier Ltd. All rights reserved.

and of these, migration from the initial site of engraftment generally reaches a maximum distance of between 700 and 800 mm [6]. These factors mean that only a small number of cells survive transplantation, and that an even fewer number actually migrate far enough to functionally integrate into the host neuronal circuitry [7]. Traditional histopathological methods are routinely employed to visualise cell integration post-transplantation surgery, however these methods rely heavily on processing post-fixed tissue ex vivo. The ability to track endogenous precursors under pathophysiological conditions is therefore restricted and it is not possible to evaluate in vivo changes at the site of engraftment or longitudinally patterns of survival and migration in live animals [8,9]. To accurately assess the long-term efficacy of cell-replacement therapy in the nervous system, reliable methods for quantification of engraftment and visualisation technologies therefore need to be developed [10]. At present, long-term noninvasive imaging techniques for tracking donor cells include ultrasonography, radionuclide tomography, two-photon fluorescent optical imaging, and magnetic resonance [11]. MRI permits effective detection of transplanted

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stem cells following prelabelling with iron oxide nanoparticles (NP) serving as T1 and T2 contrast agents [12]. Hawrylak et al. were the first to use MRI to track iron oxide labelled fetal neural tissue after injection into the brain of live rats [13]. Since then, numerous laboratories have utilised contrast agents to track stem cell transplants prelabelled in vitro as well as endogenous adult stem cells labelled in vivo in order to assess migration and integration into the nervous system via quantitative MRI [14,15]. One of the advantages of using MRI to investigate long-term stem cell engraftment is its ability to map positional data directly onto images of detailed neuroanatomy made during the same acquisition [16]. The combination of NP contrast agents with anatomical information has the potential to be extended further through the use of connectivity measurement techniques such as resting state functional MRI (rs-fMRI), diffusion tensor imaging (DTI) and manganese-enhanced axonal tract-tracing [17]. NPs can be created from solid lipids, polymers or hybrids using metal cores of gold, gadolinium, manganese or iron [11]. Superparamagnetic iron-oxide nanoparticles (SPIOs), with their small size and coating versatility, have been shown to increase ironloading efficiency and hence confer the needed MRI signal changes required for live-cell tracking [18]. SPIO polymer coatings can be further modified and attached to functional groups, including growth factors, fluorescent dyes, and small molecules to enhance biocompatibility [14]. Depending on the desired applications, NP size, stabiliser end group charge and functionalisation, and toxicity, can be customised and finely tuned dependent on experimental needs or clinical application. Once altered, the accompanying changes in biocompatibility due to the emergent properties of the NP must then be determined through in vitro experimentation. This is because any changes made to NP composition may significantly impact cellular biological properties such as uptake, viability and/or prelabelling efficiency [19]. Furthermore, earlier available USPIOs suffered from tendency to coagulate when internalised within cells and become unstable in physiological conditions [19,20]. The coating is therefore of critical importance when determining NP physical characteristics, which in turn affect their use [21]. Dextran is the most common coating polymer for SPIOs, which has FDA approval for use in patients [22]. However, its high molecular weight, which contributes significantly to the hydrodynamic size, results in relatively lower total metal content of the final coated USPIOs and reduction in NP-labelling efficiency [23]. To overcome these limitations we have previously synthesised USPIOs, utilising RAFT diblocks to stabilise the iron oxide core, which exhibit long-term stability in water and physiological solutions [23]. Advantages of RAFT polymerisation are that the diblocks for USPIOs, with an anchoring and stabilising block, are short chains and narrow in molecular weight distribution, and therefore have a minimal addition to the overall hydrodynamic diameter of the coated particle. More importantly, with a functional group at the end of each stabilising block, these RAFT diblock copolymers can be modified by attaching targeting agents such as fluorescent molecules, antibody or receptor binding molecules (Fig. 1AeE). The stabilising method is also flexible, providing the possibility to mix and match different types of stabilising molecules, which allows the USPIOs to be fine-tuned to their physiological environment as well as to act as effective drug delivery vehicles [23]. Moreover, the anchored stabilisers remain attached to the particles and stabilise them after they are taken up within the target cells, making them ideal candidates for use in cellular labelling and MRI visualisation. The biological consequences of pre-labelling hNPCs with such USPIOs have not been thoroughly characterised previously. Given the robust nature of our sterically stabilised NPs, with functional end groups on the stabiliser, NPs synthesised with this technique have immense potential for stem cell based clinical

applications. Such NPs could be modified to optimise tracking of human neural stem cells using MRI to simultaneously monitor migration as well as changes in the surrounding parenchyma. This study set out to determine the biological impact of USPIO prelabelling on neural stem cell viability, cell cycling, proliferation, apoptosis, migration, lineage potential and intracellular calcium concentration, utilising our previously characterised population of hNPCs [24]. The uptake of USPIOs into hNPCs, at various concentrations and incubation times, was investigated quantitatively by graphite furnace atomic absorption spectrometry, and by transmission electron microscopy and super-resolution laser microscopy. Furthermore, in vitro visualisation of USPIO-labelled hNPCs was performed using MRI. Future studies will investigate the applicability of USPIO-labelled hNPCs for in vivo tracking of transplanted cells. 2. Materials and methods 2.1. Synthesis of iron oxide nanoparticles Unless otherwise indicated, all materials were obtained from SigmaeAldrich. RAFT agents 2-{[(butylsulfanyl)carbonothioyl]sulfanyl}propanoic acid and methoxypolyethylene glycol modified 2 [(butylsulfanyl)carbonothioyl] sulfanyl}propanoic acid were kindly provided by Dr Algi Serelis (DuluxGroup). 1,4-Dioxane (Fluka) was distilled under reduced pressure. Monoacryloxyethyl phosphate (MAEP) was passed through an inhibitor removal column. Acrylamide (AAm), 4,40 -azobis(4-cyanovaleric acid) (V-501, Wako), sodium hydroxide (NaOH), N-hydroxysuccinimide (NHS), 1ethyl-3-(3-dimethylamino-propyl)- carbodiimide (EDC) and 2,20 -(ethylenedioxy) bis-(ethylamine) were used as received. Maghemite iron oxide cores (g-Fe2O3) were produced, as previously described, using the Massart method [25]. RAFT diblocks for coating iron oxide cores were synthesised and prepared according to Bryce et al. [23]. Magnetic iron oxide nanoparticles were prepared by coating the iron oxide cores with the desired combination of the steric stabilisers of 95% MPEG and 5% NH2 end functionalised RAFT diblocks (Fig. 1AeE). For Rhodamine labelled particles, a desired amount of rhodamine B isothiocyanate was added to the coated USPIO in water, pH 7e8, protected from light by aluminium cover and mixed for 2 h at room temperature. The free rhodamine was washed with water and removed via centrifugation. 2.2. Characterisation of USPIOs Particles size distribution, zeta potential and transmission electron microscopy (TEM) of the iron oxide cores and coated nanoparticles have been characterised by Bryce et al. [23]. Briefly, nanoparticle morphology was visualised by TEM before and after steric stabilisation. One drop of particle dispersion in water (0.001% (w/v)) was placed on a carbon coated copper grid and left to dry at room temperature. Specimens were imaged on a Phillips CM120 Biofilter transmission electron microscope (Philips, The Netherlands) at 120 kV. Particle size distribution and zeta potential of particles in water was measured with a dynamic light scattering (DLS) instrument (Malvern, Zetasizer nano series) with a detection angle of 173 , 25  C. The iron oxide cores had diameters of 10e15 nm and zeta potential of 55 mV. The coated nanoparticles had an average hydrodynamic diameter Z-average of 60 nm (PDI ¼ 0.15) and zeta potential of 34 mV and 18 mV after Rhodamine labelling. The stability of coated NPs in physiological conditions was studied by using a dispersion of the particles in phosphate-buffered saline (PBS). Visual observation and DLS measurements (data not shown) showed no aggregation/settling of particles, and a constant hydrodynamic diameter was observed. 2.3. MR relaxation rate The T2 relaxation rate of the USPIOs was measured using a Bruker 9.4 T Biospec NMR (Bruker, Ettlingen, Germany) at iron concentrations of 1, 2, 5, 10, 20 and 50 mg/ mL in water solution at 25  C. T2 measurements were made by spin-echo sequence and T2 were calculated by a mono-exponential fit. The transverse relaxivity (R2) was then determined by linear regression of R2 versus iron concentration. The R2 relaxivity of the USPIOs was 368.2 mM1 s1 (Fig. 1F). 2.4. Cell cultures Primary cultures of hNPCs were generated from fetal cortices between the ages of 16e19 weeks gestation obtained under approval from the Human Ethics Committee of the University of Sydney with informed consent, as previously described [24]. Briefly, cortical dissections were processed by mincing tissue (twice at 90 angle for 30 s) followed by trypsinisation (0.25% Trypsin/EDTA, 37  C for 30 min), trituration with a flame polished Pasteur pipette, pelleted (centrifugation at 300  g) and washed in PBS to create a single cell suspension. Using this methodology a single cell suspension with >90% viability was produced when assayed using trypan-blue. Cells were then plated in a medium composed of Neurobasal medium enriched (eNB) with B27 supplement, GlutaMax, epidermal growth factor (EGF, 20 ng/mL),

S.S. Eamegdool et al. / Biomaterials 35 (2014) 5549e5564 and basic fibroblast growth factor (bFGF, 20 ng/mL) (Invitrogen) as previously described [24]. Viable cells were maintained as neurospheres in eNB at 37  C in 5% CO2, with 50% media changes performed biweekly. Cultures were passaged (P) with TrypLE (Invitrogen) every 2 weeks and experiments conducted on P4 e 6 cultures.

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images were captured with an EM-CCD Camera iXon DU897 mounted on an Eclipse Ti inverted Nikon N-SIM Super Resolution microscope system (Nikon, Japan).with Apo TIRF 100x 1.49 (Oil) objective. Super-resolution images were constructed using the 561 nm laser line. A minimum of 10 FOV for each sample were randomly selected for analysis.

2.5. Prelabelling of NPCs with USPIOs NPCs were seeded into glass bottom wells or glass coverslips at 7.5  104 cells/ mL sequentially coated with poly-L-ornithine (10 mg/mL), human placental laminin (5 mg/mL) and human plasma fibronectin (2.5 mg/mL) as single-cell monolayers and incubated for 24 h under standard culturing conditions as previously described [24]. The cells were then labelled with USPIOs (1e100 mg/mL) by incubation for 10 min to 48 h in eNB medium. Following incubation cells were washed with PBS to remove unattached USPIOs and cultured in eNB. 2.6. Viability proliferation and apoptosis The impact USPIOs prelabelling had on NPCs in culture was measured using the tetrazolium dye (MTT) colorimetric assay for cellular growth and viability. Briefly, labelled cells (2  105 cells/mL) were incubated with 1 mg/mL 3-(4,5dimethylthiazolyl-2)-2,5-diphenyltetrazolium bromide for 4 h at 37  C, 5% CO2. Media containing MTT was then removed and the dye was solubilised with dimethyl sulfoxide for 10 min after which the absorbance (590 nm) was read using a POLARstar Galaxy microplate reader (BMG Labtech, Germany). All data was normalised against control (0 mg/mL) conditions at each time-point. Using the absorbances, percentage viability was calculated using blank/no cell absorbances (AB), control (0 mg/mL) absorbances (Acontrol) and sample absorbances (Asample): (Asample  AB)/(Acontrol  AB)  100. 2.7. Proliferation assay Proliferation was assessed using the Click-iTÒ EdU Cell Proliferation Assay (Invitrogen) according to the manufacturers’ protocol. USPIO prelabelled cells (10 mg/mL, 2 h) were incubated for 1, 3, or 7 days in vitro (DIV), pulsed with EdU (1 mM) for 1 h and then postfixed in 4% paraformaldehyde (10 min). Cells were then permeabilised with 0.5% Triton X-100 and incubated with the Alexa Fluor 555 reaction cocktail (Invitrogen) according to the manufacturers’ protocol. Nuclei were counterstained with 40 ,6-diamidino-2-phenylindole (DAPI) (1.25 mg/mL) and imaged using a LSM 510 Meta Confocal microscope (Carl Zeiss, Germany). The mean percentage of cells positive for EdU was determined from 10 fields-of-view (FOV) for each triplicate condition and was measured for cells double positive for EdU and DAPI against total number of nuclei.

2.12. Cellular TEM NPCs (1  106 cells) prelabelled with USPIOs at 10 mg/mL for 2 h were incubated in vitro for a further 6 h and then fixed in 2% glutaraldehyde for 20 min before being washed in four changes of 0.1 M sodium cacodylate buffer (pH 7.4). Samples were then post-fixed in osmium tetroxide for 1 h before being washed with distilled water, dehydrated through a series of graded ethanol dilutions (50e100%), resin embedded, cut in 100 nm sections, and stained with lead citrate on a copper grid. A JEOL1011 transmission electron microscope (JEOL, Japna) operating at 60 keV coupled with a MegaView G2 camera system (Olympus, Germany) and iTEM software (Olympus, Germany) was used for image analysis. 2.13. hNPC differentiation The effect of USPIO prelabelling on hNPC differentiation was assessed using multimarker immunocytochemical staining. Labelled cells (10 mg/mL, 2 h) were incubated in eNB for 1, 3, or 7 days at 37  C, 5% CO2, prior to cellular fixation (Section 2.5). For detection of cytoplasmic antigens, cells were permeabilised with 0.5% Triton X-100 for 30 min, prior to incubation with primary antibodies, made up with 1% bovine serum albumin in PBS, for 16 h at 4  C. Antibodies used included those specific for; neural stem/precursor cells: nestin (mouse IgG1; 1:500; R&D Systems) and sox2 (rabbit IgG; 1:500; Cell Signalling), astrocytes: GFAP (chicken IgG; 1:2000; Abcam) and s100b (mouse IgG1; 1:1000), neurons: bIII-tubulin (chicken IgG; 1:2000; Chemicon) and double cortin (DCX, rabbit IgG; 1:500; Cell Signaling), and oligodendrocytes: GalC (mouse IgG3 hybridoma; 1:50; European Collection of Cell Cultures) and the oligodendrocyte precursors marker O4 (mouse IgM hybridoma; 1:4; ECACC) [24,26e28]. Immune complexes were detected with goat antibodies (1:200) (2 h, room temperature) to chicken-, mouse-IgG1 or IgG3, or rabbit-IgG; secondary antibodies were labelled with Alexa-Fluor 488, 555 (Invitrogen), Cy3 or Cy5 (Jackson ImmunoResearch). Nuclei were counterstained with DAPI prior to mounting onto glass slides with glycerol in PBS (1:1) and sealed with nail varnish. All images were captured using a LSM 510 Meta Confocal microscope (Carl Zeiss, Germany). The mean percentage of each cell types were determined from 10 FOV for each triplicate condition, and was measured for cells double positive for each lineage marker against total number of nuclei. 2.14. Cellular migration

2.8. Apoptosis assay Apoptosis was assessed using MuseÔ Caspase-3/7 kit and MuseÔ Cell Analyzer according to the manufacturers’ protocol. Briefly, labelled cells (1e100 mg/mL; 10 min, 2 h, or 24 h) were incubated for 1, 3, or 7 DIV at 37  C, 5% CO2, and the monolayers detached for cell retrieval using TrypLEÔ (Invitrogen). These were pelleted, washed, and resuspended in 5% bovine serum albumin (BSA) in minimal essential medium (MEM) (Invitrogen). Cell samples were incubated with MuseÔ Caspase-3/7 reagent for 30 min at 37  C, 5% CO2, and with MuseÔ Caspase 7-AAD solution for 5 min at room temperature prior to analysing using the MuseÔ Cell Analyzer. Apoptosis was induced in retrieved cell samples using heat-shock treatment, by incubating the cells for 10 min, at 56  C. The mean percentage of cells positive for Caspase-3/7 was measured against total 7-AAD positive cells for each triplicate condition. 2.9. Cell cycle analyses Cell cycle was assessed using MuseÔ Cell Cycle kit and MuseÔ Cell Analyzer according to the manufacturers’ protocol. Briefly, labelled cells (10 mg/mL) were incubated for 10 min, 2 h, or 24 h, at 37  C, 5% CO2 and retrieved using TrypLE, washed and pelleted and then resuspended in PBS prior to post-fixation in ice-cold ethanol (70%) for 24 h. Cells were then incubated with MuseÔ Cell Cycle reagent for 30 min at room temperature prior to analysing using the MuseÔ Cell Analyzer. G2/M cell cycle accumulation and G0/G1 reduction was induced in cells by incubation with 660 nM Nocodazole for 24 h prior to cell retrieval. The mean percentage of cells, for each triplicate condition, at different stages of the cell cycle was measured based on differential propidium iodide nuclei staining. 2.10. Iron quantification NPCs (2  105 cells/mL) were plated in triplicate in coated 24-well plates and labelled with USPIOs (1e100 mg/mL, 10 min to 24 h). Cells were retrieved with TrypLE washed and pelleted in PBS prior to cellular digestion with 69% ultra-pure Nitric acid (HNO3) for 24 h, at 60  C. Once the cells were fully digested, 2% ultra-pure hydrochloric acid (HCl) was added (9 parts HCl to 1 part HNO3) to the samples, and analysed using Graphite Furnace Atomic Absorption Spectrometry (GFAAS) (Agilent Technologies).

For migration analyses, hNPCs were grown as free-floating neurospheres (200e 400 mm) and plated (n ¼ 10) in triplicate in coated 24-well plates and labelled with USPIOs (10 mg/mL, 10 min to 6 days). Images of each sphere were taken over the subsequent 6 DIV using an Olympus CKX31/CKX41 Phase Contrast and Fluorescence Inverted microscope with ProgRes C5 Cooled Colour CCD Camera (Olympus, USA). Mean distance from the edge of sphere to the furthest distal cell was measured at 0, 90, 180 and 270 to the sphere, using ImageJ (NIH, USA), and was averaged between each neurosphere for each triplicate conditions. 2.15. Calcium physiology Data were collected on dynamic changes in the intracellular concentration of calcium ions [Ca2þ]I [29] through indirect measurement using the calcium indicator dye Fluo-4 AM (Molecular Probes). Cells were loaded with 1 mM dye in 0.05% (w/v) pluronic acid F-127, for 40 min at 37  C, 5% CO2. Following incubation, cells were washed and cultured an additional 30 min to ensure ester hydrolysis. Prelabelled cells, (10 mg/mL, 2 h), were seeded into wells in eNB medium for 2 h prior to adding the dye. Prior to Ca2þ imaging, cells were left in either eNB (containing Ca2þ) or calcium and magnesium free Hanks balanced salt solution (HBSS) supplemented with 2 mM 1,2-bis(o-aminophenoxy)ethane-N,N,N0 ,N0 -tetraacetic acid (BAPTA) to chelate residual Ca2þ (Invitrogen). Cells were imaged for 30 s prior to addition of 100 mM adenosine triphosphate (ATP) at which point the cells were imaged for a further 2 min. For inhibiting endoplasmic reticulum ATPase-mediated refilling of the Ca2þ store, cells were incubated with 1 mM thapsigargin (Invitrogen) for 10 min at 37  C, 5% CO2, prior to being washed with MEM and imaged in HBSS/BAPTA. Micrographs were captured on a LSM 510 Meta confocal microscope equipped with heated stage and 488 nm laser line. A minimum of 10 FOV were randomly selected from monolayers (>40 representative cells per field) and analysed. LSM Meta 4.2 (Carl Zeiss, Germany) and ImageJ software were used for image acquisition and analysis. For calcium imaging data analysis, Fluo-4 fluorescence intensity was expressed as the change in fluorescence intensity relative to baseline intensity (DF/F0), where DF is the background subtracted fluorescence intensity subtracted to F0, and F0 was the background subtracted fluorescence from each cell at rest. 2.16. In vitro MRI

2.11. USPIO intracellular location NPCs prelabelled with USPIOs conjugated with Rhodamine B (USPIO-R) at 5e 20 mg/mL for 10 min to 24 h prior to cellular fixation (Section 2.5). Epi-fluorescent

The ex vivo MR images were acquired using a vertical 9.4 T Biospec MRI (Bruker, Ettlingen, Germany). Total time for imaging was 4 h for each sample. To test the feasibility of imaging labelled cells, hNPCs neurospheres were labelled with USPIOs

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A

B

Rhodamine

C

RAFT end group Anchoring block Steric stabilizing block Functional end group, e.g., -CH3 or -COOH or -NH2

D

E

R2 Relaxation Rate (s-1)

F 200 150 100 50 0 0.0

0.1

0.2

0.3

0.4

0.5

[Fe] (mM) Fig. 1. Properties of our custom designed and manufactured USPIOs. Schematic of the iron oxide particles sterically stabilised by the RAFT diblocks (A) without and (B) with rhodamine conjugation. (C) Schematic representation of a RAFT diblock. (D) Chemical structure of 95% MPEG steric stabilising block (blue). (E) Chemical structure of 5% NH2 end polymer. (F) R2 Relaxation rates (1/T2, s1) as a function of iron concentration (mM) of USPIOs in water. The R2 relaxivity rate was 368.2 mM1 s1 (r2 ¼ 0.99, p ¼ 0.001). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

over a range of concentrations from 1 to 100 mg/mL for 24 h. Following incubation cells were washed with water to remove unbound USPIOs, then embedded in 1% agarose to set at room temperature. The agar samples were imaged using T2*weighed gradient echo imaging and T2 spin-echo techniques. High resolution 3D gradient echo sequences (3DGE) were acquired using the following parameters:

NA ¼ 1; TE (echo time) ¼ 3.5 ms; TR ¼ 60 ms; FOV, field of view: 19.4  19.4  19.4 mm; matrix: 256  256  256; resolution: 35  35  35 mm (zerofilled to 15  15  15 mm). Lower resolution images with variable TE were acquired using a TE from 3.5 to 8 ms and a resolution of 70  70  140. 3D turbo spin echo sequences (3DTSE) were acquired using the following parameters: NA ¼ 1;

S.S. Eamegdool et al. / Biomaterials 35 (2014) 5549e5564 TE ¼ 40 ms; TR ¼ 200 ms; FOV (field of view): 19.4  19.4  19.4 mm; matrix: 256  256  256; resolution: 35  35  70 mm (zero-filled to 37.5  37.5  37.5 mm). Conditions with agarose only, cells only, and USPIOs only controls were also included for imaging. 2.17. Statistical analysis Data was expressed as mean  standard error of the mean (SEM) with sample size (n). Significance was determined between the average of two groups using the independent samples t-test and for groups of three or more by two-way analysis of variance (ANOVA). When two-way ANOVA was utilised, the post-hoc analysis for pairwise comparison was the Bonferroni test. An alpha value of 5% (P < 0.05) was considered statistically significant and indicated in the figures as *(p < 0.05) or **(p < 0.01) where appropriate. Statistical analysis was performed using GraphPad Prism 5.0 (GraphPad software, USA) for Mac.

3. Results 3.1. Viability of USPIO-labelled hNPCs TEM imaging of NP-labelled hNPCs provided ultrastructural evidence for NP internalisation by hNPCs (Fig. 2A, B) and a lack of NP coalescence. The effect of NP load on hNPCs viability was colorimetrically assessed via the formation of MTT formozan

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(Fig. 2C). From 1 to 10 mg/ml, it was found that NP concentrations for all incubation time points (1e48 h) had no significant effect on cell viability, which was greater than 90%, when compared and normalised against unlabelled cells in control conditions (0 mg/mL in carrier). However our post-hoc analysis showed all concentrations higher than 20 mg/mL resulted in a significant decrease (p < 0.01) in cell viability after a 24 h incubation period: (20 mg/mL (82.63  5.83%); 50 mg/mL (73.05  2.74%); 100 mg/mL (63.47  11.96%) (Fig. 2C). The quantification of intracellular NPs using GFAAS confirmed the dose-dependent uptake of NPs by hNPCs is a function of NPs concentration and incubation time (Fig. 2D). After 10 min, we measured a ten-fold increase in intracellular iron content over control when hNPCs were incubated at 100 mg/mL NPs concentration (1.17  0.19 pg/cell; p < 0.01). Post-hoc analysis revealed that intracellular iron content was significantly higher for both 10 mg/mL and 100 mg/mL NP concentrations when incubated for 2 h (10 mg/ mL: 0.58  0.08 pg/cell (p < 0.05); 100 mg/mL: 1.68  0.14 pg/cell (p < 0.01)) and 24 h (10 mg/mL: 0.91  0.04 pg/cell (p < 0.05); 100 mg/mL: 2.9  0.6 pg/cell (p < 0.01)) (Fig. 2D). No significant difference in iron content was detected when cells were incubated

Fig. 2. Screening and optimisation of NPs labelling time and concentration. Transmission electron microscopy images of hNPCs (A) without and (B) with NP exposure showed that NPs remain dispersed as single NPs and can be localised within intra-cellular vesicles (red arrows) 6 h post-incubation at 10 mg/mL. Scale bar ¼ 2 mm. Optimisation of NP-labelling conditions on hNPCs (C) viability and D) intracellular iron content. Cell viability remained >90% (dotted line), for NP concentrations 1e10 mg/mL for all exposure times, before significantly decreasing for NP concentrations >20 mg/mL, 24 h. Significant iron uptake was achieved at NP concentrations 10 mg/mL (2 and 24 h), and 100 mg/mL (all exposure times). Significance is relative to (C) control (0 mg/mL) and (D) the respective values. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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with 1 mg/mL NPs, for all incubation time-points. By using both MTT assay and GFASS, we were able to systematically screen for the optimum NP-labelling concentration (10 mg/mL) and incubation times (2 he24 h) which had an insignificant effect on the cell viability (greater than 90%) over the long-term treatment, and showed an increasing amount of intracellular iron with time. This concentration and time point was used in the subsequent experiments. 3.2. Effect of USPIO-labelling on apoptosis, proliferation index, and cell cycle progression The rate of cellular apoptosis, proliferation, and cell cycle progression, were quantified as indicators of cell survivability following a time dependant USPIOs prelabelling treatment. Apoptosis analysis, via caspase-3/7 activity, showed that NP load did not induce cell apoptosis at concentrations between 1 and 10 mg/mL, for 10 min, 2 h, and 24 h incubation time periods (Fig. 3AeC). At 100 mg/mL concentration, 24 h incubation time, significant cellular apoptosis was measured (Live: 84.95  2.5% (p < 0.05); Early Apoptosis: 25.60  2.8% (p < 0.01); Late Apoptosis/ Dead: 5.91  0.45% (p < 0.01)) compared to unlabelled hNPCs (Fig. 3AeC). Heat-shocked induced apoptosis was used as a positive control, to confirm that hNPCs undergo caspase-3/7-dependent cellular apoptosis, and resulted in a drastic and significant

increase (p < 0.01) in early- (40.5  4.9%) and late-apoptosis/death (45.5  5.4%) (Fig. 3AeC). Following NPs incubation at 10 mg/mL, for 2 h, cellular apoptosis was analysed 1, 3, and 7 days post-incubation in order to investigate the long-term survivability of NP-labelled hNPCs, and for each of the time periods NP load did not cause a significant change in cellular apoptosis in any of the samples tested (Fig. 3D). The proliferation capacity of hNPCs following NP load was quantitatively determined with EdU incorporation, indicative of DNA synthesis, and was measured against total DAPI counterstained cells (Fig. 4AeD). Cells incubated with 10 mg/mL NPRhodamine (NP-R) for 2 h (Fig 4A), retained their ability to proliferate (Fig. 4D, left arrow). Between 1 and 7 DIV, there was no significant difference in the average numbers of NP-labelled cells undergoing proliferation compared against control (Fig. 4E). Flow cytometric cell cycle analysis was used to determine if prelabelling hNPCs with NPs had significantly altered cell cycle phases. Incubating hNPCs with 10 mg/mL NPs for 10 min, 2 h, and 24 h did not result in significant changes in the average cell cycle distribution when compared against unlabelled control populations (G0/G1: 77.3  4.2%; S: 6.1  2.2%; G2/M: 12.2  3.1%) (Fig. 5A). Importantly, this data provides good evidence that hNPC populations maintain a consistent cell cycle distribution, for all conditions, when exposed to our custom USPIOs e which was a potential concern as researchers have previously observed a

Fig. 3. Effects of NPs on cellular apoptosis. No significant difference in the percentage of (A) live, (B) early apoptotic, and (C) late apoptotic/dead cells between control and NPlabelled (1, 10 mg/mL; 10 min, 2 h, 24 h) hNPCs. NP-labelled cells labelled at 100 mg/mL, 24 h, or induced with heat-shock treatment, led to a significant (A) reduction in live cells and an increase cells undergoing (B) early- and (C) late-apoptosis. Control and NP-labelled (10 mg/mL, 2 h) hNPCs displayed similar apoptotic profiles (D) 1, 3 and 7 days postincubation. Significance is relative to control.

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Fig. 4. Proliferation of NP-labelled hNPCs. Cellular proliferation of unlabelled and NP-labelled hNPCs were (A, B, C, D) visualised by confocal microscopy and (E) quantified by EdU staining. Cells were labelled with (A) NP-R (10 mg/mL, 2 h) (red), and colocalisation between (B) EdU (green) and (C) DAPI (blue) was indicative of cellular proliferation. (D) NPlabelled hNPCs undergoing proliferation (left arrow) is displayed. Scale bar ¼ 50 mm. No significant difference in proliferation between control and NP-treated hNPCs were observed (E) 1, 3 and 7 days post-incubation. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

significant change in the rate at which cells incorporate foreign material is dependent on the phase of the cell cycle [30]. There was also no effect of NP load on hNPCs long-term cell cycling when observed at 1, 3, and 7 days post-incubation (Fig. 5B). To induce G2/M cell cycle arrest and synchronise cell cycles the microtubule inhibitor nocodazole was used. No changes in cell cycle phase patterns were observed in NP prelabelled hNPCs compared to control populations. Both were found to undergo a two-fold increase (p < 0.01) in cells entering G0/G1 phase (control: 73.0  1.5%; NPs: 74.2  2.3%; nocodazole: 47.0  0.7%) (Fig. 5C) followed by a three-fold decrease (p < 0.01) in the total numbers of cells in the G2/M phase (control: 16.7  1.3%; NPs: 13.6  1.4%; nocodazole: 38.7  0.4%) (Fig. 5C). There were no significant differences in the proportion of cells entering the S phase between control, prelabelled and nocodazole treated cells (Fig. 5C).

3.3. Intracellular localisation of NPs using super-resolution microscopy Following incubation with NP-R intracellular iron was visualised by epi-fluorescence microscopy and images reconstructed using superresolution image analysis. At 5, 10 and 20 mg/ml concentrations, 10 min incubation time, intracellular USPIOs mainly localised to the cytoplasm (Fig. 6A1, E1, I1). Prolonged incubation, for 2 h, resulted in a limited amount of USPIOs observed inside the nucleus (Fig. 6B1, F1, J1). After 12 h incubation period a significant proportion was also observed in the perinuclear region with numerous NPs found clustering around the outer nuclear membrane (Fig. 6C1, G1, K1) and at 24 h most NPs had passed through the nuclear envelope, possibly via nuclear pore complexes, and concentrated in the nucleus (Fig. 6D1, H1, L1).

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Fig. 5. Cell-cycle assessment of NP-labelled hNPCs. No significant difference in the percentage of cells at G0/G1, S, or G2/M phase between control and (A) NP-labelled hNPCs at 10 mg/mL, for 10 min, 2 h, and 24 h, and (B) NP-labelled hNPCs (10 mg/mL, 2 h) at 1, 3, and 7 days post-incubation. (C) Nocodazole treatment of hNPCs caused a significant increase in the percentage of cells in the G2/M phase (double nuclei), as well as decrease in the G0/G1 phase (single nucleus) compared to control and NP-treated hNPCs. Significance is relative to control and NP-labelled hNPCs.

3.4. Multipotency of USPIO-labelled hNPCs To examine the effects of NP load on hNPC lineage elaboration we phenotypically characterised and quantified the proportion of each cell type by morphology and multiple-marker immunocytochemistry. Due to temporal-, and spatial-specific heterogeneity, with regards to protein expression patterns [24,26e28], cell type was determined by using multiple lineage-specific markers. All experiments were conducted in the presence of growth factors (eNB) in order to specifically discern the effect of NPs on hNPCs differentiation. Under this culture system, hNPCs were previously shown to be capable of undergoing trilineage differentiation over time [24]. At 7 DIV, cells were identified as 63.7  1.3% nestinþ/ SOX2þ NPCs (Fig. 7DeG); 34.1  3.3% GFAPþ/s100bþ astrocytes (Fig. 7KeN); 19.5  1.7% bIII-tubulin/DCX neurons (Fig. 7ReU) and 0.9  0.008% GalCþ/O4þ oligodendrocyte precursors (Fig. 7YeB1) with no significant differences when compared against control. Similar results were found at 1 DIV (Fig. 7C1), and 3 DIV (Fig. 7D1). To visually confirm each lineage could be labelled with NPs (10 mg/ mL, 2 h), we also labelled the cells with NP-R under the same conditions. It was found that NPCs, astrocytes, neurons, and oligodendrocytes were equally labelled with NP-R, and that the cells exhibited long-term retention of the NPs, (Fig. 7E, L, S, Z) at 7 DIV post-NP loading. Comparisons between control and NP-prelabelled cells showed no morphological change to cells of each lineage (Fig. 7AeA1), which were consistent with our previous findings: NPCs (large nuclei with an epithelial-like cell body), astrocytes (flat, epitheliallike), neurons (compact nuclei, elongated, uni-, bi-polar), and oligodendrocytes (compact nuclei with multiple branched processes)

[24]. There was no significant difference in the population of NPCs, astrocytes, neurons, oligodendrocytes, following NP-labelling, after quantification at 1, 3, and 7 DIV post-NP labelling (Fig. 7C1eE1). These data provide evidence that NP prelabelled hNPCs retain multipotentiality and are able to continuously give rise to each of the main neural cell types following NP load (Fig. 7E1).

3.5. Influence of USPIOs on the migratory capability of hNPCs To determine the effects of NPs on hNPCs migration, hNPCs were plated as neurospheres and monolayers allowed to form. Cellular migration was quantified via serial imaging at 24 h periods, with the distance between the furthest distal cell and the edge of the neurosphere measured at four distinct locations per sphere. As we have previously shown that there was no significant difference in cellular apoptosis (Fig. 3) and proliferation rate (Fig. 4) between unlabelled and NP-labelled hNPCs, we were able to determine that any changes in migration distance could be attributed specifically to the effect of NPs on cell motility. Cells incubated with 10 mg/ml NPs for 10 min, 2 h, and 24 h showed no significant difference in the migration distance at 6 DIV (Fig. 8AeC). However, when hNPCs were incubated with NPs at non-optimal conditions (Fig. 2CeD), at 100 mg/mL for the entire 6 DIV, at 3 DIV NP-labelled hNPCs had a reduced migration when compared to optimal label and control (Control: 580.5  47.3 mm; NP-labelled: 415.4  82.2 mm) (p < 0.01) (data not shown). This effect was persistent up until post-labelling 6 DIV (Control: 1004.08  39.5 mm; NP-labelled: 506.8  39.5 mm) (p < 0.01) (Fig. 8D), indicating that prolonged exposure to NPs (100 mg/mL, 6 days) resulted in a substantial reduction in: total

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Fig. 6. Localisation of USPIOs within hNPCs. Rhodamine-tagged USPIOs were visualised in labelled hNPCs by (AeL) epi-fluorescence and using a (A1eL1) Nikon N-SIM Super Resolution microscope system. Cells were exposed to NP-R (5, 10, 20 mg/mL) for 10 min, 2 h, 12 h, 24 h, and, (A, E, I, A1, E1, I1) after entry into the cell, USPIOs were found to localise in the (B, F, J, B1, F1, J1) cytoplasm after entry into the cell, moving to the (C, G, K, C1, G1, K1) peri-nuclear rim and then concentrating in the (D, H, L, D1, H1, L1) nucleus. Scale bar ¼ 5 mm.

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Fig. 7. Representative multi-marker immunocytochemistry of post-labelled neural lineages at Day 7. Neural stem/precursor cells were identified via (A, D) Nestin and (B, F) Sox2 (C, G) colocalisation. Astrocytes were identified via (H, K) s100b and (I, M) GFAP (J, N) colocalisation. Neurons were identified via (O, R) bIII-tubulin and (P, T) DCX (Q, U) colocalisation. Oligodendrocytes were identified via (V, Y) O4 and (W, A1) GalC (X, B1) colocalisation. Cells were labelled with NP-R (10 mg/mL, 2 h) in order to visualise the presence of USPIOs in (E, L, S, Z) each lineage. Nuclei were counterstained with DAPI (blue). Scale bar ¼ 20 mm. Quantification of each lineage population indicated no significant difference between of control and NP-labelled cells at (C1) 1, (D1) 3, and (E1) 7 days post-incubation. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

migration distance and, increment in migration over 24 hrintervals, compared to unlabelled cells. 3.6. Calcium signalling in USPIO-labelled hNPCs We evaluated whether ATP-induced transients in [Ca2þ]I of hNPCs were perturbed following NP-labelling. In the absence of extracellular Ca2þ, ATP can activate metabotropic P2Y purinoreceptors (P2YR), which then induces inositol-1,4,5-trisphosphate (IP3)-mediated [Ca2þ]I and downstream signalling cascades, therefore, Ca2þ imaging of NP-labelled hNPCs was performed in HBSS (no extracellular Ca2þ) [29]. A 100 mm pulse of ATP induced a transient [Ca2þ]I response in NP-labelled (10 mg/mL, 2 h) hNPCs that significantly differed from unlabelled control cells in measurements taken in both eNB medium (contains extracellular Ca2þ), and HBSS. Addition of ATP

evoked a peak transient rise in [Ca2þ]I in NP-labelled hNPCs (1.18 at 61 s), followed by a slow decline to baseline (0.84 at 135 s), compared to control eNB (1.12 at 50 s) and control HBSS (0.80 at 59 s) conditions which had relatively faster declination to baseline (eNB: 0.32 at 135 s; HBSS: 0.05 at 135 s) (Fig. 9A). Treatment of hNPCs with thapsigargin prior to addition of ATP resulted in complete abolition of [Ca2þ]I transients, confirming that endoplasmic reticulum-mediated release of Ca2þ was largely responsible for the observed ATP-induced transients in [Ca2þ]I in the cells [29]. Additionally, the maximal ATP response of NP-labelled hNPCs (105.8  7.4%) was similar to eNB-conditions (100%), and was significantly greater (p < 0.05) than HBSS-conditions (73.1  5.7%) and thapsigargin-treated cells (16.8  1.9%) (p < 0.01) (Fig. 9B). However, NP-labelled hNPCs response to ATP recovered back down to similar levels as unlabelled hNPCs in HBSS conditions after one day post-incubation (Fig. 9CeD). This effect remained at day 3

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Fig. 8. Migration of NP-labelled hNPCs neurospheres. Cells grew over 6 days after a 24 h initial attachment period (Day 0), during which the cells were labelled with USPIOs. No significant difference in the migratory capacity was observed between control hNPCs and hNPCs labelled with NPs (10 mg/mL) for (A) 10 min, (B) 2 h, and (C) 24 h. Migration distance of hNPCs was reduced when they were labelled with NPs (100 mg/mL) for (D) 7 days. Significance is relative to control.

(Fig. 9EeF) and 7 (Fig. 9GeH) post-incubation. Taken together, we found that NP load increases ATP-evoked [Ca2þ]I levels and prolongs [Ca2þ]I transients in USPIO-labelled hNPCs.

(minIPs) (Fig 10SeX) showed that although uniform labelling was not achieved using a 5 mg/mL incubation concentration (Fig. 10S), reasonable contrast was still generated at higher TE (Fig. 10T, U).

3.7. In vitro MRI visualisation of USPIO-labelled hNPCs

4. Discussion

In order to explore the feasibility of utilising these NPs as MRI contrast agents, hNPCs neurospheres were labelled with NPs over a range of concentrations (1e100 mg/mL, 24 h), dispersed in 1% agarose, and then imaged by MRI (Fig. 10AeR). An agarose only, and a non-NP incubated hNPC neurosphere sample were included as controls (data not shown). NP-labelled neurospheres showed significant local signal loss in T2*-weighted images, corresponding to internalised NPs. Adequate uptake for full penetration of the neurospheres occurred between 5 and 10 mg/mL (Fig. 10DeF, GeI). Concentrations above this level at 20 (Fig. 10 JeL), 50 (Fig. 10MeO), and 100 mg/mL (Fig. 10PeR) resulted in more intense signal loss (Fig. 10J), consistent with the quantitative MTT and GFASS data presented above. In order to further explore the localisation and effectiveness of NP uptake we examined the effect of increasing TE on the NPlabelled hNPC neurospheres at 5 mg/mL (Fig. 10 SeU) and 20 mg/ mL (Fig. 10VeX) concentrations. There was a marked signal loss present in the 20 mg/mL sample that was especially prominent at a TE value of 10 ms (Fig. 10X) compared to a TE value of 3.5 (Fig. 10V) and 5 ms (Fig. 10W). The full FOV minimum intensity projections

Stem cell therapies have shown great potential in providing functional regeneration to damaged tissues. However, the inherent lack of knowledge of the interactions between host and transplanted cells, as well as a paucity of spatial, temporal, and behavioural information regarding the transplanted cells, have resulted in significant challenges that have inhibited clinical translation. MRI is an attractive tool for providing spatiotemporal localisation of transplanted cells, but requires cellular labelling techniques to enable specific detection of transplanted stem cells. Although NPs are widely used in research and clinical settings, their impact on cellular mechanisms and function is not fully understood, primarily due to discrepancies in NPs design and cell types [11,12,14]. We have developed sterically stabilised USPIOs which exhibit long-term stability, with narrow size distribution, and modifiable stabiliser end functionality. The ability to attach functional groups such as fluorescent-dyes and pharmacological drugs permit both cell-labelling and modulation of cellular function. In this study, the biocompatibility and imaging capability of our USPIOs on primary hNPCs, feasible candidates for neurological repair, are reported.

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Fig. 9. Rise in [Ca2þ]I of NP-labelled hNPCs neurospheres in response to ATP. Cells labelled with USPIOs (10 mg/mL), in HBSS (no extracellular Ca2þ), for (A) 2 h displayed increased [Ca2þ]I levels, similar to (B) eNB conditions (with extracellular Ca2þ), with a slower return to baseline level. After (C, D) 1, (E, F) 3, and (G, H) 7 days post-incubation, NP-labelled hNPCs (10 mg/mL, 2 h) displayed [Ca2þ]I traces, and peak response to ATP, similar to HBSS control. Significance is relative to eNB unless otherwise indicated.

We have shown that NPs have a negligible effect on cellular viability below a threshold of 10 mg/mL, and that deleterious effects were dose and time dependent above these parameters. In addition, under optimal incubation conditions, USPIOs have no effect on cellular apoptosis and proliferation as measured by Caspase-3/7

activity, EdU-labelling and cell cycle analysis. NPs were found to localise predominantly in the cytoplasm, but concentrate towards the peri-nuclear rim, and into the nucleus with increasing incubation time and concentration. NPs did not affect the trilineage potential of hNPCs in differentiating into astrocytes, neurons, or

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Fig. 10. In vitro MRI of NP-labelled hNPCs neurospheres. hNPCs incubated for 24 h with (AeC) 1 mg/mL, (DeF) 5 mg/mL, (GeI) 10 mg/mL, (JeL) 20 mg/mL, (MeO) 50 mg/mL and (PeR) 100 mg/mL NP concentrations. (A, D, G, J, M, P) Selected minimum intensity projection images (minIP) of a 20 mm thickness slice over the full field of view. (B, E, H, K, N, Q) Selected magnitude images of a smaller neurosphere (approximate size 200 mm). The FOV is 1.5 mm (C, F, I, L, O, R) Selected magnitude images of a larger neurosphere (approximately 500 mm). Dark areas indicate hypointensity, corresponding to intracellular NPs. The TE in all images is 3.5 ms. (SeX) The effect of increasing TE on contrast on NP-labelled hNPCs neurospheres at 5 mg/mL versus 20 mg/mL NP concentration. Minimum intensity projections (minIP) images over the full field of view of hNPCs neurosphere labelled with (SeU) 5 mg/mL or (VeX) 20 mg/mL, using a 5 mm slice thickness at (S, V) TE ¼ 3.5 ms, (T, W) TE ¼ 5 ms and (U, X) TE ¼ 10 ms Close-ups of a representative labelled neurosphere at (SeU) 5 mg/mL or (VeX) 20 mg/mL NP concentration with a diameter of approximately 500 mm are shown below. Scale bar ¼ 500 mm.

oligodendrocytes. Cellular migratory capacity was not severely affected by NP-labelling, unless the NPs were present with the cells for extensive periods of time (over 6 days). Finally, NP-exposed hNPCs exhibited increased intracellular calcium fluxes, however this was followed by slow recovery to baseline levels one day postexposure. We confirmed that the imaging characteristics of NPloaded neurospheres are favourable for cell-tracking, with profound signal changes demonstrated using T2*-weighted imaging. Thus, we were able to extensively characterise the effects of NPs on hNPCs biological function and cellular profile, which is important for future application of these cells in in vivo cellular tracking, MRI detection, and monitoring them in cell transplantation studies. 4.1. Impact of NPs labelling parameters on hNPCs survivability and subcellular NPs location In these studies, hNPCs were shown to remain viable and survivable under optimal NP-labelling conditions. This is consistent

with previous studies, which have found that SPIOs did not affect the viability of mesenchymal stem cells or neural stem cells [31,32]. We found that at lower NP concentrations (1e10 mg/mL) the viability of NP-labelled hNPCs remained >90% over 48 h (Fig. 2). However, at concentrations greater than 20 mg/mL, cell viability was significantly reduced. Taken together, our findings showed that hNPCs are highly responsive to the presence of foreign particulate matter in their environment at high doses. In this context, the high relaxivity of our USPIOs (T2 relaxivity: 368.2 mM1 s1, versus commercial NPs, Resovist: 61 mM1 s1; Feridex: 126.0 mM1 s1) represents an important advantage [33,34]. NPs demonstrated no effect on hNPC apoptosis in the long-term, as measured through Caspase-3/7 activity (Fig. 3). Meng et al., have previously reported that survivability of neural stem cells was compromised at higher NPs concentrations, but not lower NPs concentrations, while Geppert et al., showed that viability of rat astrocytes was substantially reduced following prolonged incubation with SPIOs [32,35]. It is known that intracellular iron overload can lead to cytotoxicity and

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functional impairment, through the generation of reactive oxygen species (ROS), formation of DNA strand breaks, and impairment to the mitochondrial function [36]. There have been numerous studies pertaining to the effects of NPs on cellular proliferation, with findings of NPs either reducing, or promoting cellular proliferation being reported [37,38]. With regards to their usage in clinical applications, and the potential for NPs to alter the tumorigenesis of the cells, it was important that we showed that hNPCs proliferation (EdU incorporation and cell cycle analysis) was not changed as a result of NP-labelling. Intracellular iron concentration depends on properties of NPs, such as charge, composition and size [12]. We found that labelling hNPCs with NPs resulted in an optimal loading capacity of 0.6 pg/ cell, and a maximal loading capacity of 3 pg/cell. Our optimal loading capacity of 0.6 pg/cell was significantly lower than that reported for one other USPIOs in mesenchymal stem cells (6 pg/ cell) [39], and Ferumoxide (Feridex) in human neural stem cells (1 pg/cell) [40]. The lower uptake by NPs in our study may be explained in part by the limitation in iron content that is associated with small core particle size (0.9 mm), which imparts higher intracellular iron content [19,41]. However, under these labelling conditions, the presence of NPs inside the cells were visible and detectable through TEM (Fig. 2), fluorescence microscopy (Fig. 6), and MRI (Fig. 10). Following internalisation of NPs into the cells, it has been shown that NPs can reside in the endosome [31], cytoplasm [35], and/or the perinuclear region [42]. NPs are gradually cleared from the cells either via lysosomic degradation or cell cycle division [36,43]. As we observed, the location of NPs in hNPCs depended on the concentration and incubation time, with NPs translocating into the perinuclear region after initially residing in the cytoplasm (Fig 6). As such, this may account for the reduction in viability observed following extensive incubation with higher NP concentrations, as NPs can cause irreversible DNA damage and perturb normal gene expression [36]. It is possible that the mode of NP entry into the cell is likely via passive diffusion, as cells of the different lineages were shown to contain NPs. Rat astrocytes were shown to take up NPs via macropinocytosis and clathrin-mediated uptake [44], suggesting that similar mechanisms could be involved in the uptake of NPs by hNPCs. 4.2. Differentiation potential of UPSIO-labelled hNPCs Neurogenesis in the mammalian CNS is thought to arise from numerous cell types, most of which originate and differentiate from neural stem cells [1]. In this study we have used multi-marker immunocytochemistry to phenotypically characterise the proportion of NPCs, astrocytes, neurons and oligodendrocytes in vitro, and have shown that NPs did not significantly favour differentiation towards either lineage, but rather that hNPCs retained their differentiation potential in the long-term, and that cells of all lineages were able to be equally labelled with NPs (Fig. 7). This is consistent with other studies that have found that mesenchymal and neural stem cells retained their multipotentiality following NP-labelling [31,32]. Despite the heterogeneous nature of our cell population in vitro, we have shown that the proportion of cells of each lineage remained constant over the long-term, and that the presence of each cell type is likely to be essential in the maintenance of cell phenotype and function. It has been postulated that astroglial cells possess an active regulatory role in neurogenesis through instructing NPCs towards the neuronal lineage [45]. Although NPCs have been widely used in cell transplantation studies [46], others groups have shown that astrocytes (spinal cord injury) [47], neurons (stroke) [48], and oligodendrocytes (spinal cord injury) [49]

are capable of providing functional recovery to the CNS. Therefore, it is worth noting that co-transplantation of NP-labelled NPCs, in addition to NP-labelled committed neural cell types, might provide the necessary conditions and regulatory cues which allows for successful stem cell therapies. 4.3. Role of NPs on functional Ca2þ stores and signalling Intracellular calcium homoeostasis is critical for cellular function, as calcium signalling is involved in many key biological aspects such as proliferation, apoptosis, metabolism, and migration [50]. Excess intracellular calcium is detrimental to cells, and has been associated with various pathological conditions such as cardiac arrhythmias [51] and amyotrophic lateral sclerosis [52], likely as a result of enzymatic cellular apoptosis (such as caspases and calpain) [53]. Currently, there are a limited number of studies which have specifically investigated the effects of iron-oxide nanoparticles on cell calcium signalling, especially in neural stem cells. Titanium-oxide nanoparticles have been shown to increase [Ca2þ]I in human keratinocytes, leading to actin reorganisation, and enhanced differentiation, likely as a result of co-localisation of nanoparticles to the Golgi apparatus and deregulation of [Ca2þ]I pools [54]. Arvizo et al., found that charged gold nanoparticles have the ability to cause mitochondrial membrane potential depolarisation, which inhibits their ability to buffer [Ca2þ]I overload, resulting in substantial increases in [Ca2þ]I upon activation [55]. We observed a significant increase in [Ca2þ]I NP-labelled hNPCs following ATP addition, and that there was a delayed reduction to baseline, compared to unlabelled hNPCs. However, cells returned to basal calcium levels after 1 day and no changes persisted thereafter. Further studies are required in order to determine the effects of elevated [Ca2þ]I on hNPCs. Previously, it has been shown that iron molecules can stimulate Ryanodine receptor-mediated endoplasmic reticulum calcium-induced calcium release, through the generation of reactive oxygen species (ROS) [56]. This in turn leads to ERK pathways phosphorylation, sustained long-term potentiation, and modulation of synaptic plasticity. ROS production has been shown to be elevated in rat astrocytes [35], and neural progenitor cells [57], after loading with iron oxide nanoparticles. Bhattacharjee et al., reported that positive-charge silicon nanoparticles can uncouple the electron transport chain, which perturbs mitochondrial membrane potential and increases intracellular ROS production [58]. Therefore, it is critical that long-term effects of NPlabelling on cell calcium signalling are determined, prior to their utilisation in cell transplantation studies. 4.4. USPIOs as potential MRI contrast agents for in vivo tracking of transplanted cells Cellular migration is essential for the development of the CNS, as it accounts for generating correct cellular compositions, spatial organisation, and brain function. Marin and Rubenstein have described two different types of migration that occur: 1) radial migration, which involves neuronal migration along glial fibres from the progenitor zone towards the brain surface for forming the cytoarchitecture, and; 2) tangential migration, where the cells migrate orthogonal to radial migration, forming cellular connectivity and spatial distribution [59]. Impaired migration is known to cause developmental abnormalities such as lissencephaly and gray matter heterotopia [60,61]. In this study, NPs were shown to have no effect on migration of hNPCs, except in conditions where prolonged NPs exposure occurred, for which a substantial reduction in hNPCs migratory capacity was observed (Fig. 8). Soenen et al., have shown that high concentrations of intracellular nanoparticles can disrupt cellular actin cytoskeleton and microtubule

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networks, thereby reducing focal adhesion points portending to reduced migratory capacity [57]. Therefore, it is essential that we have shown NP-labelled hNPCs retained their migratory capacity, as future uses in cell transplantation studies would require the cells to home into the lesion sites and provide functional integration. MRI of engrafted NP-labelled stem cells is crucial for tracking the migration of the cells towards the site of injury following transplantation, giving insight into how these cells participate in tissue regeneration. Lee et al., were able to visualise transplanted NPlabelled mesenchymal stem cells utilising MRI to demonstrate migration of transplanted cells towards the thrombotic stroke site in rats [31]. In this study, our USPIOs demonstrated a high T2 relaxivity of 368.2 mM1 s1, which was significantly greater than commercial iron oxide nanoparticles of similar core size (Resovist: 61 mM1 s1; Feridex: 126.0 mM1 s1) [33,34], indicating their superiority as MRI contrast agents. We performed in vitro MRI of NP-labelled hNPCs neurospheres embedded in agarose and found that NP-labelled neurospheres displayed excellent contrast properties, and an enhanced MRI signal-to-noise ratio, compared to unlabelled neurospheres. Optimisation of in vitro MRI provides a solid platform for the utilisation of these cells in future cell transplantation studies, as others have reported good correspondence between in vitro and in vivo detection [62,63].

5. Conclusions The biological effects of USPIO nanoparticles on the stem cell population of interest need to be determined prior to their application in cell transplantation therapies. This study provided a thorough investigation into the relationship between iron-oxide nanoparticle labelling conditions and neural stem cell function, and found that hNPCs can be efficiently loaded with NPs at optimal conditions. We also demonstrated that the presence of intracellular iron at optimal concentrations does not affect hNPCs’ viability, survivability, multipotency, and migratory capacity in the longterm. With prolonged NPs exposure, NPs translocated from the cytoplasm into the peri-nucleus region, and hNPC migration became severely limited. We also observed alterations in ATPevoked intracellular calcium transients, however this phenomenon normalised 1 day post-incubation. We were able to demonstrate that our custom designed and manufactured USPIO-labelled hNPCs can be detected with MRI in vitro, which enables these findings to be applied in future work involving experimental models of neurological damage/diseases.

Acknowledgements This work was supported by grants from the National Health and Medical Research Council (NHMRC) of Australia (No. 571100 and No. 1048082) the Baxter Charitable Foundation, the Alma Hazel Eddy Trust, the Rebecca L. Cooper Medical Research Foundation (Sydney, Australia) and the Brian M. Kirby Foundation Gift of Sight Initiative (Sydney, Australia) to TC-L. SE is a recipient of the Brian M. Kirby Foundation Gift of Sight Initiative scholarship. BTTP and BSH acknowledge the financial support from Sirtex Medical Ltd. SMG acknowledges the support of the Sydney Medical Foundation. The authors gratefully acknowledge the advice and assistance related to microscopy experiments provided by Dr. Louise Cole (Bosch Institute Advanced Microscopy Facility, University of Sydney); the iron content measurements by Atomic Absorption Spectroscopy provided by Nguyen Pham (School of Chemistry, University of Sydney). The authors declare no conflict of interest in this work.

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