Elsevier

Biomaterials

Volume 34, Issue 4, January 2013, Pages 922-929
Biomaterials

Fluorescent oxygen sensitive microbead incorporation for measuring oxygen tension in cell aggregates

https://doi.org/10.1016/j.biomaterials.2012.10.019Get rights and content

Abstract

Molecular oxygen is a main regulator of various cell functions. Imaging methods designed as screening tools for fast, in situ, 3D and non-interfering measurement of oxygen tension in the cellular microenvironment would serve great purpose in identifying and monitoring this vital and pivotal signalling molecule. We describe the use of dual luminophore oxygen sensitive microbeads to measure absolute oxygen concentrations in cellular aggregates. Stable microbead integration, a prerequisite for their practical application, was ensured by a site-specific delivery method that is based on the interactions between streptavidin and biotin. The spatial stability introduced by this method allowed for long term measurements of oxygen tension without interfering with the cell aggregation process. By making multiple calibration experiments we further demonstrated the potential of these sensors to measure local oxygen tension in optically dense cellular environments.

Introduction

Oxygen is an important regulator of cell behaviour, both at a cell/tissue level and a molecular level. Based on its relatively low solubility in culture medium, oxygen gets easily depleted in static 3D culture systems [1]. This becomes most apparent in culture systems which have tight cell packing, such as cellular aggregates [2], or within a dense biomaterial carrier (for example, fibrin) [3]. Critical gradients will arise in these systems resulting from the interplay between diffusion (driven by differences in local oxygen concentration) and cellular consumption [4].

As a key component for cellular energy generation through oxidative phosphorylation, local oxygen depletion impairs cell survival when sustained over critical periods of time [5], even independent of local nutrient concentrations [6]. Most cells however are able to survive and recover from hypoxic conditions. This is accomplished by transcription factor-regulated (such as Hypoxia Inducible Factor (HIF) [7], [8]) mechanisms, that among others control energy metabolism and the cell cycle and that result in the ability to decrease oxygen utilization rate under low oxygen tension (i.e. oxygen conformity) [9], [10].

Oxygen levels also serve as a developmentally important stimulus. These oxygen regulation processes have been studied in skeletal development [11]. Endochondral ossification starts with the formation of precartilaginous condensations which originate from small cellular aggregates that expand through concerted cell activities including cell proliferation, migration, adhesion and differentiation [12]. In vitro studies have indicated that high cellularity and the avascular nature are important prerequisites for cartilage cell differentiation [13], [14], creating locally low oxygen tension and nutrient concentration, which is favourable for subsequent cartilage differentiation [15].

Investigating and understanding the mechanisms which underlie the process of chondrogenesis can be achieved in vitro using appropriately designed setups. Proper in vitro recapitulation of microenvironmental biophysical signals such as oxygen tension and cell–cell interactions is thereby mandatory [16]. An excellent model that is able to mimic condensation and chondrogenic differentiation in vitro is represented by cell aggregate cultures [17]. This culture system provides an appropriate 3D environment for synthesis, and deposition of cartilage matrix proteins, and is amenable to morphological, proteomic or transcriptional analysis [17]. Cell aggregates have been proposed as elementary building blocks for creating cell-based products using a modular design approach [18], [19], [20]. Though the influence of oxygen on cell fate is widely acknowledged, tools for non-invasively monitoring local oxygen concentrations in such dense cell environments are still limited or have major shortcomings.

Measurement of oxygen tension by the use of oxygen sensitive microfibers or micro-electrodes is complicated by the very small dimensions of cell aggregates. Furthermore such devices would strongly interfere with internal aggregate mass transport by inducing cell damage or creating ‘leaky paths’ at the probed positions. This issue is circumvented by the use of non-invasive probes [21], [22]. An example of this are perfluorocarbon (PFC) microdroplets, such as hexafluorobenzene, which rely on the paramagnetic properties of molecular oxygen to alter 19F nuclear magnetic resonance (NMR) relaxation times of the PFCs in direct proportion to the dissolved oxygen concentration [23]. These microdroplets diffuse and accumulate into interstitial spaces where they report on local oxygen tensions [24]. NMR measurements have however a rather low spatial resolution which limits their use for the small aggregate dimensions. The application of paramagnetic probes, such as trityl radicals, combined with electron paramagnetic resonance (EPR) is a very promising approach that provides reasonable resolution images (∼10–30 μm) within acceptable acquisition times (minutes) [25]. Dual luminophore oxygen-sensing beads (OSBs) also offer great possibilities for measuring oxygen tension in a fast and non-interfering manner and have been developed for measuring local oxygen tension in (poly(ethylene glycol)dimethylacrylate) hydrogel carriers [26], [27]. As the absence of controlled, site-specific delivery methods for these probes have prevented so far their practical application to cellular systems, we developed a non-invasive method for spatially controlled microbead incorporation into dense cell aggregate environments and assessed its feasibility to measure local oxygen tension.

Section snippets

Cell culture

ATDC5 cells (Riken Cell Bank, Tsukuba, Japan), a murine chondrogenic cell line [28], were grown in maintenance medium consisting of a 1:1 mixture of Dulbecco's modified Eagle's medium (DMEM) and Ham's F-12 medium (Invitrogen) supplemented with 5% foetal bovine serum (Gibco), antibiotic–antimycotic (A/A) solution (100 units/ml penicillin, 100 μg/ml streptomycin, and 0.25 μg/ml amphoterecin B; Invitrogen), 10 μg/ml human transferrin (Roche Molecular Biochemicals), and 3 × 10 m sodium selenite

Results

OSBs incorporating an oxygen sensitive luminophore (Ru(Ph2phen3)Cl2) and two reference fluorophores (Nile Blue chloride and Alexa Fluor 488) were produced and characterized (Fig. 1A). The OSBs showed in their fluorescence spectra peak values near 505 nm (Alexa Fluor 488) and 610 nm (ruthenium complex) for excitation with a 488 nm laser, and near 655 nm (Nile Blue) for excitation with a 635 nm laser (Fig. 1B), which indicates the incorporation of all fluorescent molecules by the bead. The

Discussion

Many aspects of normal cell function, ranging from basic cell respiration to integrin-mediated adhesion behaviour [1], depend on the availability of molecular oxygen. Given the important biophysical functionality, in vitro model systems would greatly benefit from in situ read-outs on local oxygen tension. These systems can for example reveal nutrient delivery problems during aggregate production in the context of tissue-engineering approaches [36], [37] or help identifying interesting

Conclusions

In this study we described the application of fluorescent oxygen sensitive microbeads (OSBs) within three dimensional cell aggregates for highly localized, quantitative measurements of oxygen tension. Successful integration of the fluorescent microbeads into cell aggregates was obtained by a binding method that relied on the interaction between biotinylated cell surfaces and streptavidin coated microbeads. We showed the non-interfering behaviour of OSB-containing aggregates through various

Acknowledgements

This work was supported by the agency for Innovation by Science and Technology (IWT-Vlaanderen, project number 090727) and the Research Foundation-Flanders (FWO-Vlaanderen, project number G.0858.12). The authors also gratefully acknowledge financial support from the Flemish Government through long-term structural funding ‘‘Methusalem’’ (CASAS Methusalem grant) and from the Hercules Foundation (HER/08/021). We thank Max Mazzone for providing the pimonidazole staining, Stefan Vinckier and

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