Bioelectronics goes 3D: new trends in cell–chip interface engineering

F. A. Pennacchio , L. D. Garma , L. Matino and F. Santoro *
Center for Advanced Biomaterials for Healthcare, Istituto Italiano di Tecnologia, 80125, Naples, Italy. E-mail: francesca.santoro@iit.it

Received 3rd July 2018 , Accepted 2nd August 2018

First published on 6th August 2018


Bioelectronic platforms can be used for electrophysiology, monitoring and stimulating specific cellular functions. While planar electroactive materials have been extensively used, in the past decade new approaches have focused on engineering the interface with pseudo-3D micro and nanostructures and, more recently, on 3D geometries (i.e. scaffold-like). Here, we present an overview of this transition from 2D to 3D bioelectronic platforms and our recent achievements of characterizing the interface between the cells and the device.


image file: c8tb01737a-p1.tif

F. Santoro

Francesca Santoro received Bachelor's and Master's degrees in Biomedical Engineering at the ‘Federico II’ University of Naples (Italy) with specialization in biomaterials. She received a PhD in 2014 in Electrical Engineering and Information Technology in a joint partnership between RWTH Aachen and Forschungszentrum Juelich (Germany) with a scholarship by the International Helmholtz Research School in Biophysics and Soft Matter (IHRS BioSoft). In October 2014, she joined the Cui Lab – Chemistry at Stanford University (USA) and she received a research fellowship in 2016 from the Heart Rhythm Society. She joined IIT in July 2017 and is now leading the ‘Tissue Electronics’ lab at CABHC-Naples as a tenure-track principal investigator. She has been recently nominated in the Under35 Young Innovators list by the MIT Technology Review Italia for 2018.


Bioelectronic devices are conceived as electronic platforms that directly interact with biological systems, ranging from single cells to entire tissues, in order to (1) monitor cellular electrical activity and (2) modulate cellular behavior through the application of electrical fields. Here, the interplay that takes place at the interface between cells and such devices is crucial to determine the effective cell–device coupling.

The relevance of the interactions between electronics and biological components was understood early on in the study of electrogenic cells, when the first models of the cell–device interface were proposed1 (Fig. 1, panels A and C). In these studies, the electrical activity of electrogenic cells is monitored by microelectrodes that, once coupled with cell bodies, could monitor relevant cellular parameters such as the membrane potential. However, cells are in reality decoupled from the sensing electrodes by a cleft that forms at the adhesion sites between the plasma membrane and the surface of the material (Fig. 1D). This cleft has a key role in the efficiency of the capacitive coupling between the cell membrane and the electronics. Structural studies have shown that the distance between the cells and planar electrodes ranges from tens to hundreds of nanometers,2–4 which is enough to make the electrodes blind to sub-threshold potentials travelling across the cell membrane.5


image file: c8tb01737a-f1.tif
Fig. 1 (A) Schematic representation of the interface between a cell and a bioelectronic device, with a model equivalent circuit overlaid on top. Reprinted by permission from ref. 18. © 2013 Springer Nature. (B) Schematic representation of the interaction between cells and devices with different dimensionalities. (C) The current model of the cell–bioelectronic device interface. Reprinted, with permission, from ref. 50. © 2018 IEEE. (D) SEM image of a cell on top of a planar device. The bottom panel shows a magnification of the junction with the tight contact points highlighted. Republished with permission of Royal Society from ref. 2; permission conveyed through Copyright Clearance Center, Inc. © 2004 Royal Society. (E) SEM images and FIB sections of cells on top of nanopillars of different width. Reprinted with permission from ref. 20. © 2014 American Chemical Society. (F) FIB cross section of a nanopillar engulfed by a cell. Reprinted with premission from ref. 3. © 2014 American Chemical Society. (G) Cross section of a mushroom-shaped structure engulfed by a cell. Reprinted with permission from ref. 17. © 2007 IEEE. (H) Schematic representation of a 3D scaffold with integrated electrodes and seeded with cells. Reprinted by permission from ref. 22. © 2016 Springer Nature.

Given this scenario, there is a need to encourage cell–electrode coupling through the modulation of the cellular adhesion. The cell–electrode adhesion can be enhanced by engineering the material properties such as its chemical composition, mechanical tuning and dimensionality.

In fact, functionalizing metallic electrodes with biomolecules such as proteins or lipid bilayers has been shown to encourage intimate contact with adherent cells.6–9

Another approach which has gained major interest in the community is the use of polymer-based semiconductors, acting either as a functional coating or as the sole conductive material for electrodes for electrophysiology. By now, these materials have largely been shown to match the main cell-coupling requirements: a low Young's modulus, wettability and conductivity.10–12 In addition, these materials offer more micro-patterning options than the traditional metallic conductors and can be used for instance to engineer surfaces with selectively cell-adhesive or cell-repulsive areas.13

Considering the dimensionality of the devices, in the past decade another strategy has been taken by engineering the topography of planar sensing electrodes. The use of non-planar surfaces has been demonstrated to be successful to enhance cell adhesion.3 Furthermore, introducing pseudo-3D structures on the surface of the devices can have striking effects, such as recreating the concept of a patch clamp electrode at the nanoscale.14–16 Examples of these structures are presented in Fig. 1E. Alternative approaches achieved the fabrication of vertical protruding microstructures aiming to emulate the geometry of some components naturally present in tissues. One very successful example of this second application was given by Spira and coworkers, who fabricated mushroom-shaped electrodes to reproduce the shape of dendritic spines of the neuronal tissue to ‘trick’ neuronal cells and gain optimal coupling on microelectrode arrays.17 The engulfment-like process at the interface between the plasma membrane and the microelectrodes drastically reduces the cell–electrode cleft.18 An image and a cross section of these mushroom-shaped structures are shown in panels F and G in Fig. 1. Consequently, the coupling coefficient between the electrode and the cell also increases to the point where mushroom-shaped electrodes are capable of recording in-cell signals, including sub-threshold membrane potentials.19,20

Moreover, fully three-dimensional platforms have been established, taking the dimensionality of the systems one step further. These systems use scaffolds to develop and monitor cell cultures in 3D. These devices can either be made entirely of electrically conductive materials, in which case the devices are capable of monitoring collective properties of the cultures,21 or alternatively, the scaffolds can contain an embedded circuitry, so they can monitor specific parts of the culture.22Fig. 1H shows the schematic representation of one such scaffold.

Besides monitoring the electrical activity of electrogenic cells, bioelectronic devices have also been extensively used to interact with other cell types with the aim of influencing cellular behavior23 for diverse applications such as in vitro modeling of specific diseases or tissue engineering.24–26 In this framework, bioelectronic devices act as “cell–instructive” platforms, where specific functional features that mimic the cells’ physiological environment are used. Similarly, for cell-monitoring purposes, cell–device coupling is also crucial for stimulation.

One of the first pieces of evidence of the effect of electrical fields on cells’ behavior was observed in PC-12 cells. Schmidt et al. showed that neurite outgrowth of PC-12 cells grown on a polymeric planar device was enhanced upon electrical stimulation.27 Thereafter, bioelectronic platforms have been employed for influencing several cell functions including, for instance, proliferation28 and uptake capability.29 The effects of this type of platform has also been exploited for tissue engineering applications such as regeneration of nervous,30,31 cardiac,32 vascular33 or bone tissue.34 Furthermore, the interaction of cells with electrical fields can enforce differentiation pathways in multipotent cells (i.e. mesenchymal and staminal types),25,26,35 currently considered as one of the most promising cellular systems for regenerative medicine purposes.

Like in the case of devices used for cell-monitoring, bioelectronic platforms meant for stimulation have sought to close the cell–electronics interface gap by using new conductive materials and modified topographies. For instance, Smith et al. have investigated how micro-patterned composite PEG–graphene based platforms can affect cellular orientation and adhesion aiming to structurally organize newly-synthesized cardiac tissues upon electrical stimulation.36 Similarly, micropatterned polypyrrole has been used for the enhancement of axonal outgrowth in hippocampal neurons.37

Fully three-dimensional scaffolds have also been used for stimulation, employing porous materials,38 fiber-based systems30,39 or cell-embedding platforms.40 Most recently, studies have focused on the use of electroactive hydrogel-based composite materials which have found several applications,41 such as cardiac tissue control and monitoring,22,32 and triggering multipotent cell differentiation in nerve regeneration42 or bone regeneration.43

Given this overview on both monitoring and stimulation bioelectronic platforms spanning from planar to 3D systems, it is clear that the main requirement for the success of cell–chip coupling is the modulation of the interface. However, a major challenge lies in characterizing this interface at a relevant length scale. For this reason, several methods have been developed aiming to specifically characterize the plasma membrane domain which is junctional to the surface of the electroactive materials.

For instance, 2D cell cultures grown on planar chips have been investigated using fluorescence microscopy,44 surface plasmon resonance4 and electron microscopy.2 However, there are major limitations to extend these techniques to (1) micro-nanostructured and 3D material surfaces and (2) 3D complex cellular assemblies.

Electron microscopy has been shown to be the most appropriate technique to analyze cellular components and their response to the underlying devices at the nanoscale. The typical specimen embedding procedure allows for resolving plasma membrane and cell ultrastructures with nanometric resolution, however there are some major drawbacks. This procedure requires a trimming step to remove a large resin matrix which surrounds the specimen and subsequent mechanical slicing through the whole sample generating a large number of thin sections (lamellae). This procedure involves a physical separation between the cells and the material underneath which can effectively induce artefacts at the interface. The imaging of hundreds of sections is then performed individually via transmission electron microscopy (TEM) or scanning electron microscopy (SEM),45 without being selective for a specific region of interest. Although TEM can be applied to cell cultures on planar or pseudo-3D supports, when cells adhere on 3D materials (i.e. scaffolds) this technique finds major technical issues, because cell structures could collapse. Considering the above, having the possibility to expose selected regions of interest and avoiding cell–material separation is fundamental.

Although this disjunction can be avoided by treating specimens via critical point drying, several cavities form in the intracellular environment, during the transition from ‘wet’ to ‘dry’.7

Focused ion beam-assisted sectioning coupled to scanning electron microscopy imaging (SEM/FIB) has turned out to be a very suitable method for studying the cleft between cell junctional membranes and bioelectronic devices, without performing any cell–material separation.20 On the one hand, SEM provides a resolution limit of 5–10 nm46 (enough to resolve the cell membrane and organelles) and on the other, the milling capabilities of the ion beam allow the acquisition of cross sections of cells adhered on materials with different composition as well as different dimensionality at a desired location.

In order to apply SEM/FIB, the specimens undergo a simple preparation procedure: first, the sample is chemically fixated using glutaraldehyde and stained with heavy metals (ROTO procedure)3 to increase the contrast of cellular ultrastructures and then the sample is embedded in resin using an ultra-thin plasticization (UTP) procedure.46 Conventional resin infiltration generates a large polymeric matrix after polymerization in which the specimen is embedded, preventing the precise location of regions of interest. In contrast, when UTP is performed, the liquid resin penetrates mostly the intracellular domain of the cells. A schematic summary of the UTP procedure is presented in Fig. 2A.


image file: c8tb01737a-f2.tif
Fig. 2 (A) Schematic representation of the UTP procedure. SEM images of: (B) PC12 cell adhered on a planar substrate. Scale bar 10 μm. (C) U87 glioma cells adhered on a 3D planar substrate. Scale bar 50 μm. (D) HL-1 cell adhered on a groove-patterned surface. Scale bar 10 μm. (E) U87 glioma cell adhered on a 3D groove-patterned structure. Scale bar 10 μm. (F) HEK 293 cell adhered on a pillar patterned surface. Scale bar 10 μm. (G) U87 glioma cell adhered on a 3D nanopillar patterned surface. Scale bar 10 μm.

Subsequently, SEM is used to directly visualize and characterize whole cell–material interactions on different exemplary types of platforms, including planar substrates, pseudo-3D platforms and, in our current studies, 3D complex materials, such as scaffolds (Fig. 2B–G). At lower magnifications, for instance, it is possible to evaluate “macro” features such as cellular spreading (Fig. 2B and D), orientation and polarization (Fig. 2C and E–G). At higher magnifications, also other important cellular “nano-features” fundamental in understanding cell–material interactions, such as cellular filopodia organization (highlighted in Fig. 2B), can be visualized.

Furthermore, FIB milling can be employed to create an in situ cross section at a location of interest, thus allowing for direct visualization, for instance, of the plasma membrane deformation in response to diverse topographies. Furthermore, SEM/FIB supported the characterization of other cellular processes such as caveolae formation and nuclear deformation.47,48

Recently, we extended the UTP procedure also to 3D materials, characterized by a structural complexity that normally poses a serious challenge especially for what concerns the application and removal of the embedding resin.49 This last advancement gave us the possibility to (1) perform superficial imaging of whole cells adhering on 3D scaffolds (Fig. 2C, E and G), and (2) visualize the cell–material interface in complex 3D systems via FIB milling (Fig. 3D–F1); we show the overall capabilities of this method in the exemplary cross sections of cells adhering on 2D planar (Fig. 3A and A1), pseudo-3D (Fig. 3B–C1) and 3D scaffolds with flat (Fig. 3D and D1), grooved (Fig. 3E and E1) and nanopillar-like surfaces (Fig. 3F and F1).


image file: c8tb01737a-f3.tif
Fig. 3 Scanning electron micrographs showing the sections of: (A) HL1 cell adhered on a planar surface. Scale bar 5 μm. (A1) Magnification of (A). Scale bar 3 μm. (B) HL1 cell adhered on a groove-patterned surface. Scale bar 5 μm. (B1) Magnification of (B). Scale bar 1 μm. (C) HEK 293 cells adhered on a pillar patterned surface. Scale bar 5 μm. (C1) Magnification of (C). Scale bar 500 nm. (D) U87 glioma cell adhered on a planar 3D structure. Scale bar 5 μm. (D1) Magnification of (D). Scale bar 500 nm. (E) U87 glioma cell adhered on groove-patterned 3D structure. Scale bar 3 μm. (E1) Magnification of (E). Scale bar 500 nm. (F) U87 glioma cell adhered on a pillar patterned 3D structure. Scale bar 5 μm and (F1) Magnification of (F). Scale bar 500 nm.

In summary, here we reported the most recent advancements in the development and use of bioelectronic platforms for efficient monitoring and stimulation, relevant in electrophysiology and regenerative medicine applications. In this context, the chemical, physical, topographical and dimensional properties of the electroactive materials have been shown to play a crucial role in the cell–chip crosstalk and, as a consequence, in the platform efficiency. In the transition from planar to 3D platforms, the latter have gained particular attention for better recapitulating the cellular physiological environment while performing precise cell monitoring and stimulation.

Furthermore, we highlighted the unique advantages related to the application of the UTP and SEM/FIB technique in characterizing cell–chip interactions and our recent advancement in applying this method to perform electron microscopy visualization of cells on 3D materials. Thanks to the high versatility of our technique, indeed, we are currently studying systems that differ in terms of scaffold structural rearrangement (i.e. ordered vs. random), cellular density and phenotype. In this context, we are also able to distinguish and quantify the different ECM components (important elements especially in tissue engineering applications) secerned by cells that result from cell–material interactions.

In conclusion, because of its remarkable potential, this technique opens up a route to a broader comprehension of cell–material interactions and to a more rational design of the new-generation bioelectronic devices.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

The authors thank Dr Valentina Mollo for help with the procedure for the UTP embedding of cells.

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Footnote

These authors have equal contribution.

This journal is © The Royal Society of Chemistry 2018