**THE CHRONIC CHALLENGE - NEW VISTAS ON LONG-TERM MULTISITE CONTACTS TO THE CENTRAL NERVOUS SYSTEM**

# **Topic Editors Ulrich G. Hofmann and Jürgen Krüger**

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**ISSN** 1664-8714 **ISBN** 978-2-88919-508-4 **DOI** 10.3389/978-2-88919-508-4

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# **THE CHRONIC CHALLENGE - NEW VISTAS ON LONG-TERM MULTISITE CONTACTS TO THE CENTRAL NERVOUS SYSTEM**

#### Topic Editors:

**Ulrich G. Hofmann,** Albert Ludwigs University of Freiburg and Cluster of Excellence "BrainLinks-BrainTools", Germany

**Jürgen Krüger,** AG Hirnforschung, Germany

Sagittal slice of a rat brain with a flexible polyimide multisite probe in situ, implanted for 12 weeks. Immunostained glial fibrillary acidic protein (GFAP) labeled with Alexa Fluro 488. Picture by Nadja Martini and Yijing Xie, University of Freiburg, Freiburg.

Have you ever heard of a Hype-Cycle? It is a description that was put forward by an IT consultancy firm to describe certain phenomena that happen within the life cycle of new technology products. As Fenn and Raskino stated in their book (Fenn and Raskino 2008), a novel technology - a "Technology Trigger" - gives rise to a steep increase in interest, leading to the "Peak of Inflated Expectations". Following an accumulation of more detailed knowledge on the technology and its short-comings, the stake holders may need to traverse a "Trough of Disillusionment", which is followed by a shallower "Slope of Enlightenment", before finally reaching the "Plateau of Productivity".

In spite of the limitations and criticisms levied on this over-simplified description of a technology's life-cycle, it is nonetheless able to describe well the situation we are all experiencing within the brain-machine-interfacing community. Our technology trigger was the development of batch-processed multisite neuronal interfaces based on silicon during the 1980s and 1990s (Sangler and Wise 1990, Campbell, Jones et al. 1991, Wise and Najafi 1991, Rousche and Normann 1992, Nordhausen, Maynard et al. 1996). This gave rise to a seemingly exponential growth of knowledge within the neurosciences, leading to the expectation of thought-controlled devices and prostheses for handicapped people in the very near future (Chapin, Moxon et al. 1999, Wessberg, Stambaugh et al. 2000, Chapin and Moxon 2001, Serruya, Hatsopoulos et al. 2002). Unfortunately, whereas significant steps towards artificial robotic limbs could have been implemented during the last decade (Johannes, Bigelow et al. 2011, Oung, Pohl et al. 2012, Belter, Segil et al. 2013), direct invasive intracortical interfacing was not quite able to keep up with these expectations. Insofar, we are currently facing the challenging, but tedious walk through the Trough of Disillusionment.

Undoubtedly, more than two decades of intense research on brain-machine-interfaces (BMI's) have produced a tremendous wealth of information towards the ultimate goal: a clinically useful cortical prosthesis. Unfortunately even today - after huge fiscal efforts - the goal seems almost to be as far away as it was when it was originally put forward. At the very least, we have to state that one of the main challenges towards a clinical useful BMI has not been sufficiently answered yet, regarding the long term – or even truly chronic – stability of the neural cortical interface, as well as the signals it has to provide over a significant fraction of a human's lifespan. Even the recently demonstrated advances in BMI's in both humans and non-human primates have to deal with a severe decay of spiking activity that occurs over weeks and months (Chestek, Gilja et al. 2011, Hochberg, Bacher et al. 2012, Collinger, Kryger et al. 2014, Nuyujukian, Kao et al. 2014, Stavisky, Kao et al. 2014, Wodlinger, Downey et al. 2014) and resolve to simplified features to keep a brain-derived communication channel open (Christie, Tat et al. 2014).

#### **References:**

# Table of Contents

*102 Smaller, Softer, Lower-Impedance Electrodes for Human Neuroprosthesis: A Pragmatic Approach*

and Toshikazu Nishida

Elisa Castagnola, Alberto Ansaldo, Emma Maggiolini, Tamara Ius, Miran Skrap, Davide Ricci and Luciano Fadiga

#### *119 A Simple Implantation Method for Flexible, Multisite Microelectrodes into Rat Brains*

Anja Richter, Yijing Xie, Anett Schumacher, Susanne Löffler, Robert D. Kirch, Jaafar Al-Hasani, Daniel H. Rapoport, Charli Kruse, Andreas Moser, Volker Tronnier, Sandra Danner and Ulrich G. Hofmann

Ahmed Eleryan, Mukta Vaidya, Joshua Southerland, Islam S. Badreldin, Karthikeyan Balasubramanian, Andrew H. Fagg, Nicholas Hatsopoulos and Karim Oweiss

## The chronic challenge—new vistas on long-term multisite contacts to the central nervous system

Ulrich G. Hofmann1, 2 \* and Jürgen Krüger 3

1 Section for Neuroelectronic Systems, Clinic for Neurosurgery, Albert-Ludwigs-University Freiburg, Freiburg, Germany, 2 Cluster of Excellence "BrainLinks-BrainTools" EXC 1086, Freiburg, Germany, 3 AG Hirnforschung, Universität Freiburg, Freiburg, Germany

Keywords: multisite neuronal recording, gliosis, indwelling implants, compliance match hypothesis, flexible microprobes, hydrogel coating, polymeric microelectrodes

This Special Research Topic at hand is a collection of contributions from eminent research groups shedding light on several aspects of the still unresolved problem of a truly chronic cortical interface to enable long term brain-machine interfacing to human patients.

The hypothesis article of Fernandez et al. (2014) adds to the three generally agreed-on features for biocompatibility (bio-safety, bio-stability, and bio-functionality) with a fourth one that mirrors the demand for "bio-tolerability." Sommakia et al. (2014a) study aims to reduce the almostimmediate adsorption of non-cellular tissue components upon insertion by dip-coating polyethylene glycol (PEG) as a "stealth" cover. It points toward a beneficial alteration of adsorption on the probe, but cautions PEG's immediate use for long term implants in the brain. In fact, in a second contribution based on a mixed-brain culture (Sommakia et al., 2014b), they show evidence for a complex response of glia cells on micro-wires dip-coated with PEG and/or lipopolysaccharides (LPS), but not of neurons, which is somewhat contradictory to pure in vivo findings.

In the same context of passive probe coatings, De Faveri et al. (2014) moved toward a more "natural" method by coating glass-insulated micro-wires with fibrin hydrogel, as a biological cushion between brain and probe.

Edited and reviewed by: Laura Ballerini, University of Trieste, Italy

#### \*Correspondence:

Ulrich G. Hofmann, ulrich.hofmann@coregen. uni-freiburg.de

Received: 18 February 2015 Accepted: 27 February 2015 Published: 18 March 2015

#### Citation:

Hofmann UG and Krüger J (2015) The chronic challenge—new vistas on long-term multisite contacts to the central nervous system. Front. Neuroeng. 8:3. doi: 10.3389/fneng.2015.00003

Beyond the modulatory effects of passive surface coatings, two articles review organic coatings for micro-contacts in the nervous system. The contribution by Asplund et al. (2014) concisely reviews electrodes based on conductive polymers, not only for improving site-tissue coupling, but also for electrically eluting anti-inflammatory drugs using various stimulation patterns. They go to great depth on how to apply this elution process to a living being, since the active elution technique inherently requires compliance with demanding bio-compatibility issues.

The review of Aregueta-Robles et al. (2014) addresses the topic of organic and nanoscopic coatings with a wider perspective, and thus provides an excellent overview regarding a huge variety of the various reported approaches including the "living electrode" of Ochiai et al. (1980) and their adaptation by Richter et al. (2010, 2011).

Extending the time frame of all above mentioned studies, Prasad et al. (2014) investigate whether the brain's foreign body response is the sole cause for poor electrode yield using Pt/Ir micro wire arrays. They state that leading aspects include the suboptimal construction of the micro wires, as well as the severing of the blood brain barrier upon insertion. In order to achieve a deeper insight into the suboptimal micro array construction, another study of the same group (Sankar et al., 2014) analyses long-term impedance spectra using FEM simulation, and concludes that the initial increase in electrode-tissue impedance in vivo can be attributed to cell attachment and gliosis on the micro wires. Furthermore they show that the long-term decrease is probably caused by de-lamination and cracks in the wire's insulation layer.

Even though the contribution of Castagnola et al. (2014) discusses the use of nano materials for improving the signal quality from brain micro-recordings as well, its main theme is the interpretation of the so called "compliance match hypothesis" (Stieglitz and Meyer, 1999) by making cortical interfaces softer reducing the permanent mismatch between rigid micro probes and the brain's softness. Obviously, comparing elastic moduli of brain and probe materials reveals a discrepancy of several orders of magnitude, and so the positive evidence of De Faveri et al. (2014) may be based on a compliance adaptation between both due to the fibrin layer. However, Castagnola et al. (2014) as well as Richter et al. (2013), Sohal et al. (2014) and Xie et al. (2014) all use substrate materials for their multisite arrays, which are still far away from brain's bulk modulus. More importantly, they are flexible ("soft" when compared to brain tissue) due to their geometrical setup. We hypothesize that this flexibility might be the reason for Krüger's 7 year recording record in the non-human primate using ultra-thin metal wires (Krüger et al., 2010). Castagnola et al. (2014) approaches a similar goal by electro-depositing microscopic spheres at the working end of very thin wires—intended for subarachnoidal recordings. Sohal et al. (2014) in contrast, intracortically implants a geometrically "wavy" structure and shows evidence for good recordings from a limited number of animals for up to 2 years. The histological findings from their and Richter's thin multisite implants depict a reduced gliosis, and thus corroborates the need for a matching

### References

compliance and intracortical softness. Richter's probes are made from polyimide (Rubehn et al., 2010), and they explain in detail how to implant them with help of a simple removable support.

This removable support in turn provides additional value, as Xie et al. (2014) state. Their use of label-free, endoscopic, in vivo Optical Coherence Tomography by the very same fiber supporting implantation demonstrates the first optical online monitoring of deep indwelling processes. This method may provide new vistas on the traditional problem of foreign body response, and may therefore shorten the time we have to spend in the Trough of Disillusionment.

Whatever effort is required, and whichever ideas will finally prove optimal for reaching the Plateau of Productivity for brainmachine interfaces, we are thankful that the need for biotolerability and reliability of brain implants over a substantial period of time, has already sunk into the community. Eleryan et al.'s (2014) single neuronal signal tracking method demonstrates this spirit of pro-active optimism by providing a highly valuable algorithm for following the progress of an individual neuron's participation in information processing.

This Special Research Topic's collection won't be the final word with respect to chronic implants, but for now it sets a roadsign toward truly stable neuro-cortical interfaces that lie on the horizon.

#### Acknowledgments

We are indebted to Dr. Richard C. Pinnell for making our manuscript comprehensible.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Hofmann and Krüger. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

### Acute human brain responses to intracortical microelectrode arrays: challenges and future prospects

**Eduardo Fernández 1,2\*, Bradley Greger 3 , Paul A. House4 , Ignacio Aranda5 , Carlos Botella6 , Julio Albisua7 , Cristina Soto-Sánchez 1,2 , Arantxa Alfaro1,2 and Richard A. Normann8**

1 Bioengineering Institute, Miguel Hernández University of Elche, Elche, Spain

2 CIBER-BBN, Zaragoza, Spain

3 School of Biological and Health Systems Engineering, Arizona State University, Tempe, AZ, USA

4 Department of Neurosurgery, University of Utah, Salt Lake City, UT, USA

5 Department of Pathology, Hospital General Universitario, Alicante, Spain

6 Department of Neurosurgery, Hospital La Fe, Valencia, Spain

7 Department of Neurosurgery, Fundación Jimenez Díaz and Hospital Rey Juan Carlos, Madrid, Spain

8 Department of Bioengineering, University of Utah, Salt Lake City, UT, USA

#### **Edited by:**

Ulrich G. Hofmann, University of Freiburg, Germany

#### **Reviewed by:**

Winnie Jensen, Aalborg University, Denmark Tobias Holzhammer, ATLAS Neuroengineering, Belgium

#### **\*Correspondence:**

Eduardo Fernández, Bioengineering Institute, Miguel Hernández University of Elche, Avda. Universidad s/n, Elche 03202, Spain e-mail<EMAIL_ADDRESS>
The emerging field of neuroprosthetics is focused on the development of new therapeutic interventions that will be able to restore some lost neural function by selective electrical stimulation or by harnessing activity recorded from populations of neurons. As more and more patients benefit from these approaches, the interest in neural interfaces has grown significantly and a new generation of penetrating microelectrode arrays are providing unprecedented access to the neurons of the central nervous system (CNS). These microelectrodes have active tip dimensions that are similar in size to neurons and because they penetrate the nervous system, they provide selective access to these cells (within a few microns). However, the very long-term viability of chronically implanted microelectrodes and the capability of recording the same spiking activity over long time periods still remain to be established and confirmed in human studies. Here we review the main responses to acute implantation of microelectrode arrays, and emphasize that it will become essential to control the neural tissue damage induced by these intracortical microelectrodes in order to achieve the high clinical potentials accompanying this technology.

**Keywords: neural prosthesis, intracortical microelectrode, in vivo recording, biocompatibility, neurosurgery**

#### **INTRODUCTION**

Since 1990, the field of neuroprosthetics has grown at an ever increasing rate. Although current research is also pursuing noninvasive techniques to acquire signals from the human brain, important scientific and clinical information has been gained from *in vivo* experiments with microelectrode arrays able to record or stimulate the nervous system with the aim of improving or replacing motor or sensory abilities that have been lost due to disease or injury. For example, auditory brainstem implants are being used to restore auditory function (Merkus et al., 2014); deep brain stimulators have been implanted successfully in patients for control of motor disorders, tremor, dystonia and chronic pain (Beitz, 2014; Nardone et al., 2014) and micro-array type devices have been implanted in artificial vision systems (Grill et al., 2009; Hatsopoulos and Donoghue, 2009; Normann et al., 2009; Fernandez and Hofmann, 2011). Moreover advances in prosthetic limbs and brain-machine interfaces are now providing hope of increased mobility and independence for amputees and paralyzed patients (Raspopovic et al., 2014) and intracortical micro-electrodes are being used to study epileptiform discharges in cases of intractable epilepsy (Weiss et al., 2013a,b). As more and more patients have benefited from this approach, the interest in neural interfaces has grown significantly and a new generation of penetrating microelectrode arrays are providing unprecedented access to the neurons of the central nervous system (CNS).

These studies have shown the utility of microelectrode recordings to resolve neural activity at the single neuron level and to interpret brain derived commands for fine-grained control of movements and translation of these signals into command signals that are able to control external devices (Hochberg et al., 2006; Andersen et al., 2010; Rao and Donoghue, 2014). However most penetrating microelectrode arrays currently have maximum *in vivo* lifetimes from several months to a few years (Suner et al., 2005; Prasad et al., 2012; Barrese et al., 2013). Therefore much work still needs to be done before these penetrating microelectrodes can be used for many clinical purposes. In this work we focused on the acute effects of implantation of penetrating microelectrode arrays in the human brain, emphasizing the relevance of surgical techniques and biocompatibility approaches to control the neural tissue damage induced by these intracortical probes in order to achieve the therapeutic benefits envisioned by these neural interfaces.

#### **CURRENT MULTIELECTRODE-ARRAY TECHNOLOGIES**

Technology partially achieved during the development of cardiac pacemakers has been successfully used in many other applications of implanted neural prostheses. During these years great efforts have been made to develop penetrating multi-electrode arrays with dimensions similar to the cortical neurons they target for recording or stimulation and that are able to maintain a stable signal in the CNS. The two main dominant approaches are multiple insulated metal microwires (Nicolelis and Lebedev, 2009; Freire et al., 2011; Carmena, 2013) and penetrating microelectrode arrays fabricated with micro-electro-mechanical-system technologies. These devices use various substrate materials that can either be flexible and based on polymers (Rousche et al., 2001; Chen et al., 2009; Kozai and Kipke, 2009; Hassler et al., 2011) or rigid such as the Utah Electrode Array (UEA; Normann, 2007), the Michigan array (Seymour and Kipke, 2007) or the Neuro-Probes arrays (Neves, 2007; Calixto et al., 2013). However today, only a few of these devices are commercially available and do not yet exist as commercial, wireless, implantable, many-channel devices that can provide reliable recording and stimulation for many years.

An ideal neural interface would consist of an array containing many microelectrodes, which can "listen" and "talk" to stillfunctioning parts of the brain enabling bi-directional communication with ensembles of neurons. Each microelectrode would either record the electrical activity of a small population of neurons surrounding it or, when electrical current is passed through the electrode, activate small population of neurons. This approach has been difficult to realize because of the acute and chronic inflammatory reactions that can induce significant changes at the brain-electrode interface. Thus all neural probes need a stable electronic/neural interface that enables selective recording and/or activation of specific groups of neurons without deterioration of the electrodes or surrounding neural tissue. Consequently three areas have to be considered: the "*biosafety*", the "*biofunctionality*" and the "*biostability*" of the devices. Biosafety means that the microelectrodes do not harm the brain tissue in any significant way, biofunctionality is related to the ability of the microelectrodes to perform their intended function, and biostability means that the whole microelectrode array must not be susceptible to attack of biological fluids, proteases, macrophages or any metabolic byproducts (Marin and Fernandez, 2010). In addition successful devices should manifest "biotolerability" or the ability of the multielectrode-array to reside in the CNS for long periods of time. All these considerations impose extreme demands on stability and function of neural implants and place unique constraints on the architecture, materials, and surgical techniques used in the application of intracortical microelectrodes.

#### **SURGICAL PROCEDURES FOR IMPLANTATION OF INTRACORTICAL MICROELECTRODES ARRAYS**

Microelectrode arrays aimed to simultaneously record or stimulate neuronal populations have been successfully used in many animal models including non-human primates (Normann, 2007). However the only intracortical microelectrode array that has been FDA approved for long-term human studies is the UEA (Nordhausen et al., 1996). This microelectrode array consists of 100 small diameter silicon microneedles, built on a square grid with 400 µm spacing, that were designed to be inserted into the cerebral cortex to a depth of 1.5 mm, the level of thalamic input to the cerebral cortex (**Figure 1A**).

The UEA has been used in BrainGate clinical trials (Hochberg et al., 2006; Homer et al., 2013; Perge et al., 2013) and in research with epilepsy patients (Normann et al., 2009; Truccolo et al., 2011; Weiss et al., 2013a,b), and we have performed a number of preliminary experiments designed to establish the safety of the implantation procedures (House et al., 2006). These experiments were performed in persons suffering from epilepsy or brain tumors that had to undergo a surgical resection of a brain region. Briefly after exposure of the implantation site using standard neurosurgical equipment and procedures, the pia arachnoid was cut to allow access to the surface of the cerebral cortex. Even though the individual electrodes of the UEA are very sharp (with radius curvatures typically on the order of a few microns) we found that trying to push 100 electrodes into cortical tissues only depressed the surface of the tissue and resulted in only partial insertion of the electrode array. A pneumatically actuated precision instrument that allows the complete and safe insertion of the array in under 200 ms was developed to circumvent this difficulty (Rousche and Normann, 1992). Interestingly we found that it was necessary to fine adjust the parameters of the pneumatic inserter depending of the age of the patients, and that it was easier to get a complete insertion in young than in aged patients using low insertion pressures (around 15– 20 psi).

#### **HUMAN ACUTE REACTIVE RESPONSES AROUND IMPLANTED MICROELECTRODES**

Implantation of any neural probe is always a traumatic procedure that implies some local damage of neurons, vasculature and other cells (**Figures 1C,D**). Thus, when a neural probe is inserted into the brain, some neurons and glial cells are killed or injured during insertion, blood vessels are disrupted, and the bloodbrain barrier is compromised (**Figures 2A,B**). As a result, with most of our microelectrode array insertions in human cortex, we typically observed interstitial microhemorrhages emanating from the electrode tracks that extended in one or more directions. These microhemorrhages were limited to within a few millimeters of the microelectrode tracks although these were more evident around the edges of the arrays. This damage seemed to result from a combination of the numerous blood vessels encountered in the path of the penetrating electrodes plus some mechanical damage of small capillaries, especially at the borders of the array. Furthermore although the insertion of the UEA did not result in clinically relevant hemorrhages, some times we found petechial hemorrhages located below the tips of the electrodes (**Figures 2C,D**).

These microhemorrhages usually stopped spontaneously or after gentle irrigation with normal saline and we do not have any case where we had to remove the array due to the bleeding.

However, due to this small bleeding various serum plasma proteins and blood cells stick on the surface of the electrodes and trigger complement system activation, platelet activation, clot formation and a large network of changes including the release of cytokines, invasion of blood-borne macrophages and edema. **Figure 2E** shows an example of the tip of an electrode after explantation. Many red blood cells are seen in close contact with electrode materials (see **Figure 2F**). Thus, microhemorrhaging is an important issue that should be taken into account to reduce the adverse nature of the reactions and maintain an ideal environment for the microelectrodes.

Array implantation also causes an early activation and migration of microglial cells towards the microelectrodes, which could reflect highly specific interactions mediated by selectins, integrins, cytokines and carbohydrate-binding receptors (Kaur et al., 2010; Linnartz et al., 2012). These microglial cells are very sensitive to pathological conditions, even if they respond only to variations in local extracellular ionic concentrations (Kreutzberg, 1996; Freire et al., 2011) and we have found that there is a rapid activation, in a matter of minutes, of these cells in human brain. **Figure 1D** shows an example in an experiment in which the UEA was kept in place for <10 min.

Usually these scavenger cells form a network of immune alert resident macrophages with a capacity for immune surveillance and control. Therefore the initial activation and migration of microglial cells are likely beneficial and include production of neurotrophic substances and cell adhesion molecules, which support injured neurons and appear necessary for restorative events to take place (Eddleston and Mucke, 1993). However, this largely beneficial initial phase can result in a more adverse longterm response that is dependent on the extent of the injury. Consequently more effort is needed to control these responses and ensure the function of the implant without eliciting either structural, cellular or metabolic changes that compromise the microelectrode array performance and/or that result in tissue degeneration around the implanted microelectrodes.

#### **ELECTROPHYSIOLOGICAL RECORDINGS**

As has been shown from experiments in rodents and non-human primates (Oliveira and Dimitrov, 2008), we found that the quality of the surgical implantation procedure plays a major role for tissue preservation and for the outcome of recordings. Our experimental results have demonstrated that high-quality microelectrode recordings from cerebral cortical neurons can be consistently obtained in both acute (intraoperative) and short term chronic (in an epilepsy monitoring unit) settings. An example of action potentials recorded with a UEA in the temporal cortex of a patient with medically intractable epilepsy is shown in **Figure 1B**. Single units are easily identifiable with a quality similar to that observed in rat, cat or non-human primate recordings. Furthermore,

temporal cortex. **(B)** Once the array has been removed there are some evident microhemorrhages. **(C)** Horizontal section showing blood in the

> neural tissue (Parker et al., 2011). This problem can be mitigated with sophisticated action potential identification and daily recalibration.

in close contact with electrode materials. **(F)** Detail of the red blood cells on the surface of the microelectrodes. Calibration bars A, B, C and D= 2 mm.

#### **CHALLENGES AND FUTURE WORK**

Intracortical microelectrode arrays have several advantages over traditional macroelectrode brain electrophysiological techniques such as their ability to selectively access individual or small groups of cortical neurons and the possibility to deliver relevant spatiotemporal patterns of stimulation. This highlights their potential to restore some lost neural function through selective electrical stimulation or by recording activity from selected populations of neurons. Furthermore the procedures for the implantation of penetrating microelectrodes are straightforward and well within the reach of most well-trained neurosurgeons. Altogether these results suggest that intracortical microelectrodes could form the basis of new neuroprosthetic devices for treating many disorders of the nervous system. For example they are presently being used to obtain volitional command signals from primary motor cortex of people with high cervical spinal injuries or to provide a limited but useful sense of vision in profoundly blind. However the presence of acute inflammatory reactions that can lead to chronic inflammatory reactions affecting both the neural tissue and the surface of the microelectrodes must be better understood and new approaches pursued to mitigate these reactions. Controlling and reducing the neural tissue damage induced by these intracortical microelectrodes should aid in

most of the presently available intracortical microelectrode arrays (including the UEA) allow recording of local field potentials that are very reliable and that can be used to detect changes in recording quality over time. Also, recording quality is affected by other surgical procedures: most drugs used for the induction and maintenance of general anesthesia decrease action potential firing rates and the information processing capacity in the neocortex (Hanrahan et al., 2013). In addition local fluctuation in K+ and Ca+ due to acute initial trauma can induce the silencing of neurons in the proximity of microelectrodes. Consequently recording sessions are usually initiated between 3–10 days after implantation.

The ability of intracortical microelectrodes arrays such as the UEA to record single units from human cortical neurons demonstrates that the implantation can be done without major complications, and emphasizes its potential role for studying encoding and processing of sensory and motor information by large neuronal ensembles. However significant variations in spike waveforms across time have been reported (Parker et al., 2011). These variations could reflect the initial acute response that can lead to chronic inflammatory reactions. Such reactions can negatively impact neurons and microelectrodes via induction of glial proliferation that produces a slow progressive decline in spike amplitude and in the number of viable channels. Furthermore recordings from individual neurons made with penetrating microelectrodes are often lost but then recover, possibly because of micro-motion of the device relative to the keeping these devices biologically and electrically viable for many years.

Factors affecting brain tissue reactions to intracortical microelectrodes include the mechanical trauma during insertion, implantation method, biological acute responses and the physical properties of the microelectrodes. Because the brain is so richly vascularized, we have observed that there is inevitable bleeding associated with electrode array insertion and this suggests that bleeding is an important issue that should be taken into account to reduce the neural tissue damage induced by the microelectrodes. Moreover the reliability of recording the same spiking activity over long time periods still remains an unresolved problem that must be solved in longer term human studies before these penetrating microelectrodes can be used for some clinical purposes.

In this framework, important criteria for clinical success are tissue and microelectrode preservation and the maintenance of a stable signal in the CNS for the longest time possible. Remarkable progress has been reported in the use of these penetrating microelectrode arrays, but the electrode-tissue interface remains one of the major obstacles. Intracortical microelectrodes need a stable electronic/neural interface that facilitates selective recording and/or activation of specific groups of neurons without damage to the electrodes or surrounding neural tissues. Consequently it is essential to better understand the signals that lead to neuroglial activation in human brain and to create a targeted intervention strategy to prevent or at least to control this response.

Progress in this area relies on scientists being able to integrate and utilize methodologies from disparate disciplines, such as biomedical engineering, biomaterials, neuroscience, neurology, neurosurgery, information and communication technologies, molecular biology, etc. We are optimistic that with an emphasis on collaboration and a concerted push for additional clinical trials, these technologies will form the basis of devices and therapies that will substantially reduce the burden of lost neurological functions. However progress will be incremental and researchers must avoid creating false expectations that could damage the credibility of these new technologies.

#### **ACKNOWLEDGMENT**

This work has been supported in part by grants MAT2012- 39290-C02-01 from the Spanish Government, by the National Organization of the Spanish Blind (ONCE).

#### **REFERENCES**

**Conflict of Interest Statement**: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 29 March 2014; accepted: 19 June 2014; published online: 21 July 2014*. *Citation: Fernández E, Greger B, House PA, Aranda I, Botella C, Albisua J, Soto-Sánchez C, Alfaro A and Normann RA (2014) Acute human brain responses to intracortical microelectrode arrays: challenges and future prospects. Front. Neuroeng. 7:24. doi: 10.3389/fneng.2014.00024*

*This article was submitted to the journal Frontiers in Neuroengineering*.

*Copyright © 2014 Fernández, Greger, House, Aranda, Botella, Albisua, Soto-Sánchez, Alfaro and Normann. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms*.

## Resistive and reactive changes to the impedance of intracortical microelectrodes can be mitigated with polyethylene glycol under acute in vitro and in vivo settings

#### **Salah Sommakia1,2 , Janak Gaire3 , Jenna L. Rickus 1,2,4 and Kevin J. Otto1,3\***

1 Weldon School of Biomedical Engineering, Purdue University, West Lafayette, IN, USA

2 Physiological Sensing Facility at the Bindley Bioscience Center and Birck Nanotechnology Center, Purdue University, West Lafayette, IN, USA

3 Department of Biological Sciences, Purdue University, West Lafayette, IN, USA

4 Department of Agricultural and Biological Engineering, Purdue University, West Lafayette, IN, USA

#### **Edited by:**

Ulrich G. Hofmann, Albert-Ludwigs-University Freiburg, Germany

#### **Reviewed by:**

Xuefeng F. Wei, The College of New Jersey, USA Abhishek Prasad, University of Miami, USA

#### **\*Correspondence:**

Kevin J. Otto, Weldon School of Biomedical Engineering, Purdue University, 206 S. Martin Jischke Drive, West Lafayette, IN 47907, USA

e-mail<EMAIL_ADDRESS>
The reactive response of brain tissue to implantable intracortical microelectrodes is thought to negatively affect their recordable signal quality and impedance, resulting in unreliable longitudinal performance. The relationship between the progression of the reactive tissue into a glial scar and the decline in device performance is unclear. We show that exposure to a model protein solution in vitro and acute implantation result in both resistive and capacitive changes to electrode impedance, rather than purely resistive changes. We also show that applying 4000 MW polyethylene glycol (PEG) prevents impedance increases in vitro, and reduces the percent change in impedance in vivo following implantation. Our results highlight the importance of considering the contributions of non-cellular components to the decline in neural microelectrode performance, and present a proof of concept for using a simple dip-coated PEG film to modulate changes in microelectrode impedance.

**Keywords: intracortical microelectrodes, foreign-body reaction, impedance spectroscopy, polyethylene glycols, dip coating**

#### **INTRODUCTION**

Failure of intracortical microelectrodes typically manifests as increased impedance and decreased signal to noise ratio, and is thought to be associated with the formation of a dense glial scar and loss of neuronal density. *In vivo* impedance monitoring is a common tool to assess the functionality of implanted intracortical microelectrodes, and has been used to infer the progression of the reactive tissue response to implanted intracortical microelectrodes (Williams et al., 1999, 2007; Vetter et al., 2004). Recent research shows that changes in electrical properties monitored by impedance spectroscopy do not always perfectly correlate with cellular responses (Prasad et al., 2012; Prasad and Sanchez, 2012), implicating additional biotic and abiotic factors. *In vitro* testing in 3D gel constructs reveal that different glial cells adhered to the surface of a microelectrode have different impedance profiles (Frampton et al., 2010). One factor that has not been well investigated is the adsorption of proteins and other biomolecules. While adsorbed proteins have been implicated in the biological response (Leung et al., 2008), their effects on the electrical impedance of intracortical microelectrodes have not been previously described with impedance spectroscopy. Prevalent electrical circuit models of the tissue electrode interface assume that adsorbed proteins result in purely resistive impedance changes (Johnson et al., 2005; Otto et al., 2006; Williams et al., 2007), but there is not sufficient empirical verification of this assumption. To the best of the authors' knowledge, there are no reports in the literature on the effects of adsorbed proteins or non-cellular components on the impedance of intracortical microelectrodes.

Another aspect to the problem of biomolecule adsorption is the question of preventing detrimental changes to the electrical characteristics of intracortical microelectrodes using simple and cost effective approaches. For implantable devices in other biological systems, protein-resistant or anti-fouling treatments are commonplace (Salacinski et al., 2001; Bluestein et al., 2010; Li and Henry, 2011). One of the most common materials used to enhance the biocompatibility of biomedical implants is polyethylene glycol (PEG). Due to its hydrophilic nature, PEG prevents the adsorption of proteins by reducing access to the more hydrophobic surface onto which proteins prefer to bind (Michel et al., 2005). Typically, PEG is chemically grafted onto a substrate and reliably reduces protein adsorption (Sharma et al., 2004a,b; Muthusubramaniam et al., 2011). In the context of intracortical microelectrodes, PEG has traditionally been used as a scaffold for thick drug eluting hydrogels (Winter et al., 2007; Rao et al., 2012), the size scale of which might exacerbate neuronal displacement. Thinner conformal microgel coatings which incorporate PEG as a cross linker have been investigated, but do not significantly improve the chronic tissue response (Gutowski et al., 2014). Free-floating PEG injected intravenously has been reported to improve cellular and behavioral recovery following traumatic brain injury (Koob et al., 2005, 2008; Koob and Borgens, 2006). Because of the complexity of the reactive tissue response of the brain to implanted microelectrodes, and the fact that the cause of the majority of chronic microelectrode failures are unknown (Barrese et al., 2013), the long-term effects of a simply applied anti-fouling coating on the tissue response cannot be confidently predicted. In the short-term, however, it is possible that such a simply applied anti-fouling coating might improve the electrical properties of acutely implanted neural microelectrodes, which, in turn, might improve the accuracy of impedance monitoring as a predictive tool for the progression of the tissue response. The authors are not aware of any reports in the literature that examine the effects of a simple dip-coated PEG film on the electrical properties of neural microelectrodes under acute *in vitro* or*in vivo* settings.

The primary objective of this paper is to quantify the acute effects on microelectrode impedance of adsorbed proteins*in vitro*, and non-cellular molecular components *in vivo*. A secondary objective is to present a proof of concept on the use of an aqueous dip-coated PEG film in preventing impedance changes to neural microelectrodes during acute timescales. For this proof of concept experiment, a molecular weight of 4000 Da was chosen according to literature reports demonstrating optimal antifouling properties at this molecular weight (Su et al., 2009). We first present an analysis of changes in intracortical microelectrode impedance following immersion in a model protein solution mimicking *in vivo* brain protein concentration. Total impedance, resistance, and reactance are analyzed at different frequency values to quantify the contribution of adsorbed proteins to the impedance changes affecting electrode performance. We show that a dip-coated film of a relatively high molecular weight PEG prevents changes in impedance upon immersion in protein solution. We then demonstrate in an acute *in vivo* experiment that increases in microelectrode impedance after insertion into the cortex can be reduced by applying the same PEG treatment to the electrode shank and implantation site.

#### **MATERIALS AND METHODS**

#### **IN VITRO STUDY WITH MODEL PROTEIN SOLUTION**

Electrochemical measurements of 16-channel single shank Michigan electrode arrays (CNCT, Ann Arbor, MI) were made using an Autolab potentiostat PG-STAT12 with a built-in frequency response analyzer (EcoChemie, Utrecht, The Netherlands). For this study, a three-electrode cell configuration was used with the microelectrode site functioning as the working electrode, a large-area Pt wire functioning as the counter electrode, and an Accumet, gel-filled, KCl saturated calomel electrode (Thermo Fischer Scientific, Fair Lawn, NJ) functioning as the reference electrode. This three-electrode setup was chosen for this study because of its ability to isolate the electrode/solution interface impedance component.

For each electrode array, cyclic voltammetry (CV) was performed by sweeping the applied voltage from −0.6 to +0.8 V at a scanning rate of 1 V/s to determine sites with broken or poor connections, designated as those sites exhibiting a maximum current below 1 nA. These sites were discarded from the analysis, thus yielding a total of 30 functional sites on three different electrode arrays.

Electrochemical impedance spectroscopy (EIS), using the PGSTAT12, was used to measure the impedance of the electrode sites with the application of 15 sequentially applied sinusoidal waves at logarithmically spaced frequencies ranging from 46 Hz to 10 kHz, with an amplitude of 25 mVRMS. For each electrode, impedance spectroscopy was performed in PBS following each of three different treatments: (a) no treatment; (b) immersion in a 10% solution of bovine serum albumin (BSA) (Sigma-Aldrich, St. Louis, MO) in PBS, the concentration of which was chosen to mimic protein concentration in rat cerebral cortex (Banay-Schwartz et al., 1992); and (c) immersion in a 20% solution of 4000 MW PEG (Alfa-Aesar, Ward Hill, MA), air-drying for 1 min, then immersion in BSA. Immersion and subsequent removal from the described solutions was done at a controlled velocity using a micro-manipulator. An additional volume of 100 µl of PEG was further applied directly onto the microelectrode shank as it was immersed in BSA to mimic a topical application. Electrodes were rinsed with deionized water and anodically cleaned between the different treatments using 10-s long DC pulses, as described previously (Sommakia et al., 2009). Bode and Nyquist plots were generated for all treatment groups. Comparisons of the resistance, reactance and total impedance were done at 50 Hz, 100 Hz, 1 kHz, and 10 kHz.

#### **ACUTE IN VIVO STUDY**

The laboratory animal protocol for this work was approved through the Purdue Animal Care and Use Committee (West Lafayette, IN, USA), and conforms to the guidelines of the US National Institutes of Health. Three Sprague Dawley rats (Harlan Laboratories, Indianapolis, IN) were used for this study. Each rat was anesthetized with 2% isofluorane, then transferred to a stereotactic frame and maintained with 0.5–1% isofluorane delivered through a nose cone. The head was shaved and swabbed with alternating washes of betadine and alcohol, and an eye lubricating ointment applied. Lidocaine was injected subcutaneously at multiple positions in the head, and then a midline incision approximately 2 cm long was made along the cranium with a scalpel. The underlying pericranium was removed to expose the skull. A single burr hole was made with a dental drill towards the back of the head, slightly anterior to the ears, and a stainless steel bone screw attached to a segment of platinum wire was threaded into it to serve as a counter electrode. Bilateral craniotomies about 2.5 mm in diameter were made with a dental drill, approximately 2.5 mm anterior to Bregma and 2 mm lateral to the midline. For each craniotomy, a slit was made in the dura using surgical microscissors. One craniotomy serving as a control was wetted with 0.9% sterile saline before electrode insertion, while the other craniotomy was wetted with 20% w/v solution of 4000 MW PEG in MilliQ water prior to electrode insertion.

16-channel single-shank Michigan probes were also used for this study. Because of the difficulty of achieving a three-electrode setup in a surgical setting, a two-electrode setup was chosen instead. Prior to insertion into the cortex, baseline impedance was established *in vitro* by performing the same EIS procedure described above, but in a two-electrode setup. The electrode was then manually inserted into the exposed cortex of the control craniotomy using a magnet-stabilized micromanipulator, and EIS was measured 5 min after insertion. The electrode was then removed, rinsed with MilliQ water, and cleaned by applying a DC bias of 1.5 V for 10 s in PBS, as described previously (Sommakia et al., 2009). After verifying the return of the 1 kHz impedance to baseline, the electrode was dip-coated at a controlled velocity using a micromanipulator in a 20% solution of 4000 MW PEG in MilliQ water for 1 min and allowed to dry for 2 min. The PEG-coated electrode was then inserted using the micromanipulator into the other craniotomy wetted with PEG, and EIS was performed again.

#### **STATISTICAL ANALYSIS**

Statistical analysis was performed using the SAS 9.3 statistical package (SAS Institute, Cary, NC). A general linear model (GLM) procedure was used to perform a one way ANOVA with block to remove variations between the different electrodes by treating electrodes as a statistical block. Tukey *post-hoc* tests were used to identify statistically significant differences between the groups at a significance level of α = 0.05. Plots were generated using MATLAB (The MathWorks Inc., Natick, MA).

#### **RESULTS**

#### **ANALYSIS OF IMPEDANCE CHANGES WITH MODEL PROTEIN SOLUTION AND PEG IN VITRO**

An examination of Bode plots reveals that the gain for electrodes immersed into is higher than for uncoated controls for all frequencies, indicating an overall increase in impedance. Electrodes treated with PEG prior to BSA immersion show more congruence with the gain of uncoated electrodes (**Figure 1A**). The phase for non-PEG treated electrodes immersed in BSA exhibits lower angles at the lower end of the frequency spectrum, and the difference is most pronounced in the middle of the spectrum between 200–500 Hz (**Figure 1B**). In **Figure 2**, the Nyquist plot for non-PEG treated electrodes immersed in BSA reveal a shift up and the right compared to the controls, indicating both increased resistance and reactance, with the most pronounced divergence occurring toward the middle of the plot. In contrast, the plot for electrodes pretreated with PEG shows more congruence with uncoated controls in the middle of the plot and slight divergence at the lower end of the plot, corresponding to the higher frequencies. To better understand the details of these changes and their potential implication for mitigation attempts, we examined the percent changes in resistance, reactance, and total impedance at various points across the frequency spectrum.

**Figure 3A** shows percent changes in the real component of the impedance, i.e., resistance, relative to the control at four frequency values across the spectrum. Electrodes immersed in BSA without PEG pretreatment exhibited statistically significant increases in the resistance compared to the uncoated control at examined frequencies. The highest resistance increase relative to control was in the middle of the frequency spectrum, specifically at 1 kHz, with a 30.7% increase in resistance. At 50 Hz, the increase in resistance for the BSA coated electrodes without PEG pretreatment was 14.5%, at 100 Hz, the resistance increase was 23.9%, and 10 kHz, the resistance increase was 17%. The electrodes pretreated with PEG prior to BSA immersion, on the other hand, did not exhibit any significant differences in resistance compared to the uncoated controls at all examined frequencies.

**Figure 3B** shows percent changes in the imaginary component of the impedance, i.e., reactance, relative to the control at four frequency values. For electrodes not pretreated with PEG prior to immersion in BSA, no significant increase in reactance was observed at 50 Hz or 100 Hz, while increases of 12% and 15% were observed at 1 kHz and 10 kHz, respectively. In contrast, electrodes pretreated with PEG prior to immersion in BSA exhibited modest decreases in the reactance relative to control at all frequencies (−6.3% at 50 Hz, −6.3% at 100 Hz, −4.2 at 1 kHz, and −4.8% at 10 kHz). **Figure 3C** shows the percent changes in total impedance relative to the control at four frequency values. For electrodes not pretreated with PEG prior to BSA immersion, no significant difference in total impedance was observed at 50 Hz, but progressive increases in the impedance were observed at the higher frequencies (4.5% at 100 Hz, 13.5% at 1 kHz, 15.3% at 10 kHz). For electrodes pretreated with PEG prior to BSA immersion, modest decreases in total impedance were observed at the lower frequencies (−5.8% at 50 Hz, −5.6% at 100 Hz, −3.9 at 1 kHz), while no significant difference in the total impedance compared to the control was observed 10 kHz.

#### **ANALYSIS OF IMPEDANCE CHANGES IN VIVO WITH AND WITHOUT PEG**

**Figure 4A** shows the percent increase of the real component of the impedance, i.e., resistance, between the *in vitro* baseline and the *in vivo* measurement at four frequency values across the spectrum. In both cases of no treatment and PEG treatment, the resistance exhibited a significant increase when measured *in vivo* compared to the *in vitro* baseline. Insertion into the cortex without PEG treatment, however, resulted in a larger increase from the *in vitro* baseline at all frequencies compared to insertion with PEG treatment. The percent change in resistance was also frequency-dependent. For the no treatment condition, the percent increase from baseline was as follows: 71.8 ± 2.99% at 50 Hz, 89.8 ± 3.77% at 100 Hz, 209.5 ± 9.1% at 1 kHz, and 290.5 ± 13.7% at 10 kHz; while for the PEG treatment condition, the percent increase from baseline was: 44.6 ± 3% at 50 Hz, 58 ± 3.9% at 100 Hz, 149.5 ± 9.3% at 1 kHz, and 223.3 ± 14% at 10 kHz. The percent increase in resistance from baseline for the no treatment condition was significantly different from the percent increase in resistance from baseline for the PEG treatment at all frequencies.

**Figure 4B** shows the percent increase of the imaginary component of the impedance, i.e., reactance, between the *in vitro* baseline and the *in vivo* measurement at four frequency values across the spectrum. In both cases of PEG treatment or no treatment, the reactance increased significantly when measured *in vivo* compared to the *in vitro* baseline. Insertion into the cortex without PEG treatment, however, resulted in a larger increase in reactance from baseline compared to insertion with PEG treatment. The percent change in reactance was also frequencydependent. For the no treatment condition, the percent increase from baseline was as follows: 45.4 ± 1.6% at 50 Hz, 48.1 ± 2%

**FIGURE 1 | Bode plots. (A)** For electrodes immersed in BSA with no PEG coating, the total impedance magnitude plot is higher than the control, while electrodes coated with PEG prior to BSA immersion show an impedance magnitude plot indistinguishable from control. **(B)** Electrodes immersed in

at 100 Hz, 66.4 ± 2.3% at 1 kHz, and 146.8 ± 6.2% at 10 kHz; while for the PEG treatment condition, the percent increase from baseline was: 30.8 ± 1.6% at 50 Hz, 31.4 ± 2% at 100 Hz, 41 ± 2.4% at 1 kHz, and 97.1 ± 6.4% at 10 kHz. The percent increase in reactance from baseline for the no treatment condition was significantly different from the percent increase in resistance from baseline for the PEG treatment at all frequencies. The amount of percent change in reactance is lower than the amount of percent change in resistance at each respective frequency. **Figure 4C** shows the percent increase in total impedance between the *in vitro* baseline and *in vivo* measurement at four frequency values across the spectrum. In both cases of PEG treatment or no treatment, the total impedance increased significantly when measured *in vivo* compared to the *in vitro* baseline. Insertion into the cortex without PEG treatment, however, resulted in a larger increase from

frequencies compared to the control, while the phase angle for electrodes coated with PEG prior to immersion in BSA exhibit smaller phase angles at all frequencies.

baseline compared to insertion with PEG treatment. The percent change in total impedance was also frequency-dependent. For the no treatment condition, the percent increase in total impedance from baseline was as follows: 47.4 ± 1.7% at 50 Hz, 52.2 ± 2.1% at 100 Hz, 88.62 ± 2.9% at 1 kHz, and 214.5 ± 9.8% at 10 kHz; while for the PEG treatment condition, the percent increase in total impedance from baseline was: 33.6 ± 1.6% at 50 Hz, 33.5 ± 2.1% at 100 Hz, 57.1 ± 3% at 1 kHz, and 162 ± 10% at 10 kHz. The percent increase in total impedance from baseline for the no treatment condition was significantly different from the percent increase in total impedance from baseline for the PEG treatment at all frequencies.

#### **DISCUSSION**

#### **RATIONALE**

The prevalent narrative in the literature suggests that the *in vivo* reactive tissue response is an aggregate of amplified biological processes that begin with device insertion and accompanying trauma. The indwelling implant acts as a sink for various proinflammatory proteins, as well as a substrate for cell attachment (Leung et al., 2008). To the best of the authors' knowledge, the impedance changes resulting from protein adhesion onto neural microelectrodes have not been previously quantified, nor have the effects of anti-fouling treatments on impedance of neural microelectrodes. Most current efforts towards mitigating the reactive tissue response focus on targeting and quantifying the cellular component of the reactive tissue response. Recent findings, however, indicate that strong cellular responses to implanted microelectrodes do not necessarily correspond to similar changes in electrode impedance (Prasad and Sanchez, 2012). We posit that extracellular components of the reactive tissue response might be more instrumental in altering the electrical properties of implanted electrodes, and thus offer an attractive alternative for quantification and mitigation. The primary objective of this paper was to identify non-cellular components as potential modulators of impedance changes in neural microelectrodes, and to quantify the effects of a specific type of non-cellular component *in vitro*

(proteins). The secondary objective was to demonstrate a proof of concept at acute time scales for using PEG applied via a simple dip-coating process to prevent impedance changes caused by noncellular components both *in vitro* and *in vivo*.

#### **EXPLANATION OF RESULTS**

For the *in vitro* study with the model protein solution, a three-electrode setup was chosen for its ability to isolate the electrode/solution interface impedance. Changes in impedance measured using this three-electrode setup indicate changes only at the electrode/solution interface (Bard and Faulkner, 2001). Because of the combination of low currents, short timescales, micro-scale working electrodes, and a large area counter, it was assumed that very minimal changes could have occurred at the recording surface. Using this measurement paradigm, our first finding was that significant increases in resistance at all examined frequencies occur immediately upon exposure to a model protein solution with a concentration mimicking *in vivo* concentrations. Furthermore, significant increases in the reactance are observed at higher frequencies, including the physiologically relevant 1 kHz. The aggregate impedance effect is that of a significant increase at frequencies higher than 50 Hz. These observed *in vitro* changes in impedance might not exactly match *in vivo* changes, given the difference in protein composition in the brain, and the presence of additional biomolecules with widely varying degrees of hydrophobicity, such as lipids and polysaccharides (O'Brien and Sampson, 1965; Pease, 1966; Margolis and Margolis, 1974; Norton et al., 1975). This finding of changes in both resistive and capacitive components following protein adsorption challenges current assumptions inherent in prevalent electrical circuit models of the device tissue interface. Such models typically assume that protein adsorption causes purely resistive changes in impedance (Johnson et al., 2005; Otto et al., 2006; Williams et al., 2007). We demonstrate that the exposure of microelectrodes to a protein solution

results in both resistive and capacitive changes in impedance, rather than purely resistive changes. These findings suggest a need for the reexamination of assumptions upon which prevalent electrical circuit models for the tissue electrode interface are built. More accurate or representative equivalent circuit models might improve the utility of impedance monitoring to accurately deduce the progression of the reactive tissue response.

Our study further demonstrates that treating electrodes with a PEG film via dip coating prior to immersion in a protein solution negates these increases in impedance at physiologically relevant frequencies. That electrodes pretreated with PEG exhibit slight decreases in reactance supports our working hypothesis that the PEG film forms a hydrated layer close to the electrode surface that prevents proteins from accessing the surface of the electrode, while simultaneously avoiding detrimental effects on charge transfer at the electrode-electrolyte layer.

For the acute *in vivo* study, a two-electrode setup was used due to the difficulty of implementing a three-electrode setup within a surgical setting. The use of a two-electrode setup means that the impedance of the tissue/electrode interface and the impedance of the tissue are lumped into a single impedance source. The expected contribution of the tissue impedance component is confirmed by the considerable increase observed in both the resistance and capacitance following the insertion of the electrode into the rat brain. For this study, we wanted to examine the contribution of non-cellular components to the change in impedance, and therefore chose a short time scale during which no tissue remodeling occurs. Since cellular responses in the brain following electrode implantation are not observed until several hours post implantation (Kozai et al., 2012), our impedance measurement needed to be conducted within that time frame. We chose an arbitrary time point of 5 min post-insertion to minimize the exposure of the animals to unnecessary anesthesia, but it is possible for impedance changes to be time sensitive.

The application of an aqueous solution of a higher molecular weight PEG by dip-coating onto the microelectrode and directly into the craniotomy resulted in a reduction in the magnitude of the increase from the *in vitro* baseline impedance. Because the tissue impedance component is presumed not to change within this short time frame, we can assume that the difference in impedance with PEG treatment compared to the no treatment condition is due primarily to the modulation of the electrode/tissue interface impedance component. In the control craniotomy, we posit that a considerable portion of the immediate impedance increase comes from hydrophobic biomolecules that come into contact with the electrode surface and hinder charge transfer. The application of free-floating high molecular weight PEG appears to confer some degree of protection from the effects of aforementioned biomolecules, and results in lower impedance. With this simple method of applying PEG via dip-coating, there are concerns about the uniformity of coatings between electrodes. While we did not directly quantify the morphology of our dip-coated PEG films, we did attempt to control the deposition process to minimize variability. Since the dip-coating velocity is a major contributing factor to the uniformity of dip-coated films (Scriven, 1988), we controlled our dip-coating velocity by using a micromanipulator. Because we did observe statistically significant changes in impedance in response to our dip coatings, we did not make further attempts to identify and reduce individual contributors to inter-electrode variations.

Much more research needs to be conducted into the use of PEG to understand its potential long-term effects on the functional longevity of implantable intracortical microelectrodes. While prior research in the field has found that conformal microgel PEG coatings that do prevent cellular adhesion *in vitro* do not result in significant improvement to the cellular composition of the chronic electrode tissue interface (Gutowski et al., 2014), the effects of such PEG-containing coatings on the electrical properties of functional electrodes have not been tested. It is possible that significant protective effects using grafted coatings have not been observed due to the use of lower molecular weights. For our proof of concept demonstration for PEG, we used a molecular weight of 4000 Da, which has been shown to be optimal for reducing biofouling in comparison with lower molecular weights (Su et al., 2009). With biopolymers, there are concerns about degradability which might limit the long-term potential for such coatings to modulate the reactive tissue response. PEG, however, is generally not considered easily degradable in physiological conditions, and degradable PEG hydrogels generally need to be engineered with special cleavage sites to facilitate biodegradability (Drury and Mooney, 2003). On the other hand, the effects of long-term presence of large polymeric molecules in the brain should be considered before employing such molecules as neurointegrative coatings.

These complex considerations surrounding such antifouling coatings present an attractive research area. While chemically immobilized coatings are a commonplace choice for neurointegrative coatings, there might be additional merits to weakly attached PEG films. For example, intravenous treatment with higher molecular weight PEG has been shown to improve cellular and behavioral recovery in traumatic brain injury (Koob et al., 2005, 2008; Koob and Borgens, 2006), due to its fusogen properties which allow it to induce membrane sealing of damaged cells and tissue (Shi, 2013). It is possible that these properties of higher molecular weight PEG can be similarly effective in disrupting deposition of proteins and other molecules at the electrode vicinity, in addition to mitigating chronic blood-brain barrier damage, a major factor in chronic device failure (Saxena et al., 2013). Before dip-coated PEG films can be prescribed as a long-term solution for preventing the reactive tissue response and increase the functional longevity of implantable neural microelectrodes, more extensive parametric studies need to be conducted. These studies should consider the molecular weight of the PEG used, in addition to the morphological characteristics of the applied films, such as roughness and thickness, and related properties such as degradation kinetics. The delivery method of PEG films is also an open question. It might be necessary to combine multiple delivery methods; for example, concurrently immobilizing PEG onto the electrode surface and applying free-floating PEG. One potential approach under investigation is the use of thin-film silica solgel coatings, which have the potential to incorporate released and immobilized molecules, and do not adversely affect electrical properties (Pierce et al., 2009). Such future experiments with PEG should combine impedance monitoring with histological analysis of the reactive tissue response on a chronic time scale. Our findings provide a first step by highlighting the potential contribution of non-cellular molecular components to impedance changes, and offer a proof of concept for dip-coated PEG films as a potential component within a holistic neurointegrative strategy.

#### **ACKNOWLEDGMENTS**

Funding for this research was provided by the Purdue Research Foundation, the Indiana Spinal Cord and Brain Injury Research Grant Program (Fund # 00015115), and the Defense Advanced Research Projects Agency (DARPA) Microsystems Technology Office (MTO), under the auspices of Dr. Jack W. Judy (jack.judy@ darpa.mil) and Dr. Doug Weber<EMAIL_ADDRESS>as part of the Reliable Neural Technology Program, through the Space and Naval Warfare Systems Command (SPAWAR) Systems Center (SSC) Pacific grant No. N66001-11-1-4013. Thanks to members of the Neuroprostheses Research Laboratory for feedback on the manuscript.

#### **REFERENCES**

**Conflict of Interest Statement**: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 30 April 2014; accepted: 11 July 2014; published online: 04 August 2014*. *Citation: Sommakia S, Gaire J, Rickus JL and Otto KJ (2014) Resistive and reactive changes to the impedance of intracortical microelectrodes can be mitigated with polyethylene glycol under acute in vitro and in vivo settings. Front. Neuroeng. 7:33. doi: 10.3389/fneng.2014.00033*

*This article was submitted to the journal Frontiers in Neuroengineering*.

*Copyright © 2014 Sommakia, Gaire, Rickus and Otto. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms*.

### Glial cells, but not neurons, exhibit a controllable response to a localized inflammatory microenvironment in vitro

#### **Salah Sommakia1,2 , Jenna L. Rickus 1,2,3 and Kevin J. Otto1,4,5\***

1 Weldon School of Biomedical Engineering, Purdue University, West Lafayette, IN, USA

2 Physiological Sensing Facility at the Bindley Bioscience Center and Birck Nanotechnology Center, Purdue University, West Lafayette, IN, USA

3 Department of Agricultural and Biological Engineering, Purdue University, West Lafayette, IN, USA

4 Department of Biological Sciences, Purdue University, West Lafayette, IN, USA

5 J. Crayton Pruitt Family Department of Biomedical Engineering, University of Florida, Gainesville, FL, USA

#### **Edited by:**

Ulrich G. Hofmann, Albert-Ludwigs-University Freiburg, Germany

#### **Reviewed by:**

Hugues Berry, INRIA, France Ian D. Dryg, University of Washington, USA

#### **\*Correspondence:**

Kevin J. Otto, J. Crayton Pruitt Family Department of Biomedical Engineering, University of Florida, 1064 Center Drive, NEB 363, Gainesville, FL 32611, USA e-mail<EMAIL_ADDRESS>
The ability to design long-lasting intracortical implants hinges on understanding the factors leading to the loss of neuronal density and the formation of the glial scar. In this study, we modify a common in vitro mixed cortical culture model using lipopolysaccharide (LPS) to examine the responses of microglia, astrocytes, and neurons to microwire segments. We also use dip-coated polyethylene glycol (PEG), which we have previously shown can modulate impedance changes to neural microelectrodes, to control the cellular responses. We find that microglia, as expected, exhibit an elevated response to LPS-coated microwire for distances of up to 150 µm, and that this elevated response can be mitigated by co-depositing PEG with LPS. Astrocytes exhibit a more complex, distance-dependent response, whereas neurons do not appear to be affected by the type or magnitude of glial response within this in vitro model. The discrepancy between our in vitro responses and typically observed in vivo responses suggest the importance of using a systems approach to understand the responses of the various brain cell types in a chronic in vivo setting, as well as the necessity of studying the roles of cell types not native to the brain. Our results further indicate that the loss of neuronal density observed in vivo is not a necessary consequence of elevated glial activation.

**Keywords: intracortical microelectrodes, foreign body response, primary cortical cultures**

#### **INTRODUCTION**

Implantable intracortical microelectrodes hold great potential as neural prostheses for the treatment of a wide range of traumatic and degenerative injuries to the central nervous system, but suffer from unreliability in chronic settings. This decline in chronic device performance correlates with a reactive response of brain tissue (Vetter et al., 2004). Designing therapeutic approaches to counter this decline in device performance is complicated by the lack of detailed mechanistic understanding of the progression of the reactive tissue. Dural and vascular damage appear to be major factors contributing to the reactive tissue response (Karumbaiah et al., 2013; Saxena et al., 2013). Using novel device capture techniques (Woolley et al., 2011, 2013a,b), this reactive tissue response has been shown to be non-uniform and depth dependent, with stronger scarring closer to the surface of the brain (Woolley et al., 2013c). Transdural implants elicit a much greater response than implants dwelling completely within the brain (Markwardt et al., 2013). These findings collectively suggest that the introduction of non-native cellular and molecular components into the brain amplifies inflammatory pathway activation, and that this activation is strongest at the site of injury to respective structures. Recently, potential therapeutic targets such as reactive oxygen species and toll-like receptor 4 (TL4) have been identified (Potter et al., 2013; Ravikumar et al., 2014), but the complexity underlying *in vivo* conditions can obscure investigations of biological mechanisms.

These obstacles can be somewhat overcome by studying simpler models, such as *in vitro* cell cultures. The most widely used model, first described by Polikov et al. (2006, 2009), presents microscale foreign bodies to primary mixed neural cultures., and has been applied to test biocompatibility of various materials as neural interfaces (Achyuta et al., 2010; Tien et al., 2013). This model requires the modification of the culture media to achieve a globally elevated activation state. We posit that a more localized inflammatory microenvironment may better represent the nonuniform reactive tissue response, and propose a modification to the model whereby the foreign objects are dip-coated in lipopolysaccharide (LPS) to simulate a localized inflammatory microenvironment. LPS is a known upregulator of microglial activation through TL4 binding (Lehnardt et al., 2003; Tzeng et al., 2005), and as such is an attractive option for modifying the Polikov model to test cellular responses to localized targeting of TL4 receptors. In contrast to the previous model, the creation of a localized inflammatory microenvironment also enables the analysis of neuronal responses.

Previous research in the neurotrauma field has also found that, due its surfactant properties, soluble polyethylene glycol (PEG) can induce membrane sealing of damaged cells and reduce edema (Borgens et al., 2002). This effect significantly improves recovery from both spinal cord and traumatic brain injuries by inducing cellular and behavioral recovery (Koob et al., 2005, 2008; Koob and Borgens, 2006). Additionally, we have recently showed that a dip-coated PEG film can modulate impedance changes caused by non-cellular components both *in vitro* and *in vivo* (Sommakia et al., 2014). In this regard, a non-grafted dip-coated PEG film is a technically and economically attractive option to achieve both antifouling and membrane sealing. Our hypothesis is that a dip-coated layer of high molecular weight PEG will exhibit sufficient short term stability to modulate cellular responses to microelectrodes *in vitro*. Given the importance of the early stages of the injury response in shaping the later chronic stages, this approach might prove highly beneficial *in vivo*.

In this work we test our PEG hypothesis using the local inflammation-modified Polikov model. We show that, as expected, coating segments of microwire with LPS results in an increase in microglial activation at distances up to 150 µm, and, importantly, co-depositing LPS with a PEG solution prevents observed increases in microglial activation. We also observe a slight increase in astrocyte activation in response to LPS-coated microwire, but not at the same magnitude or spatial distribution as microglia. Interestingly, neuronal responses in this *in vitro* paradigm do not appear to be influenced by corresponding glial responses.

#### **MATERIALS AND METHODS**

#### **CELL CULTURE AND MICROWIRE PLACEMENT**

The experimental procedures complied with the Guide for the Care and Use of Laboratory Animals and were approved by The Purdue Animal Care and Use Committee (PACUC). Forebrains from E17 embryonic rat pups were received suspended in 5 ml of Solution 1 (NaCl 7.24 g/L; KCl 0.4 g/L; NaH2PO4 0.14 g/L; Glucose 2.61 g/L; HEPES 5.96 g/L; MgSO4 0.295 g/L; Bovine Serum Albumin 3 g/L) in a 50 ml centrifuge tube. Under sterile conditions, the tissue was gently triturated with an added 18 µl of trypsin solution (Sigma-Aldrich, St. Louis, MO) (7.5 mg/ml in 0.9% saline) and incubated for 20 min in a 37◦C water bath. Following the incubation step, 100 µl of trypsin inhibitor/DNAase solution (Sigma-Aldrich, St. Louis, MO) (2.5 mg/ml trypsin inhibitor, 400 µg/ml DNAase in 0.9% saline) was added and tissue was again gently triturated. The tissue was then centrifuged at 1,000 rpm for 5 min at room temperature and supernatant was poured off. Cells were re-suspended in 16 ml of Hibernate-E (Brainbits, Springfield, IL) and triturated once again. Cells were filtered through a 70 µm cell strainer (Fisher Scientific) and centrifuged at 1,400 rpm for 5 min at room temperature. Supernatant was poured off and cells were re-suspended in a culture medium consisting of Dulbecco's modified Eagle's Medium (DMEM) with 10% Fetal Bovine Serum (FBS) and 10% horse serum (HS). The cells were then seeded in 96 well plates at a density of 500,000 cells/cm2 , and cultured for 7 days at 37◦C and 5% CO2, with the cell media being replaced every 48 h. At day 7 *in vitro*, lengths of 50 µm-diameter tungsten microwire (California Fine Wire Co., Grover Beach, CA) were autoclaved then cut into small segments of 5–7 mm in length using carbide scissors. The microwire segments were treated by dip coating with one of four treatments: LPS (50 ng/ml) only, PEG (20% aqueous solution, 4000 MW) only, a 1:1 mixture of LPS and PEG, or uncoated. A relatively low LPS concentration was chosen based on reported literature values (Das et al., 1995; Wang et al., 2005) in order to achieve localized activation of microglia, but prevent a generalized activation that might result from a higher concentration of LPS diffusing rapidly throughout the well. PEG concentration is based on our previous work demonstrating a proof of concept for using PEG to modulate impedance changes to neural microelectrodes (Sommakia et al., 2014). In each well, one segment of microwire was dropped into the medium and allowed to sink to the bottom of the well. The plates were then placed in the incubator for an additional 7 days.

#### **CELL FIXING AND LABELING**

At day 14 *in vitro*, the cultures were fixed with 4% paraformaldehyde for 10 min, rinsed 3× with HEPES Buffered Hank's saline (HBHS) (in g/L; 7.5 g NaCl, 0.3 g KCl, 0.06 g KH2PO4, 0.13 g Na2HPO4, 2 g Glucose, 2.4 g HEPES, 0.05 g MgCl2:6H2O, 0.05 g MgSO4:7H2O, 0.165 g CaCl2, 90 mg NaN3, at pH 7.4), then permeabilized with 0.2% Triton-X (Sigma-Aldrich, St. Louis, MO). The cultures were then blocked with 10% normal goat serum (Jackson Immunoresearch, West Grove, PA) for 1 h, after which primary antibodies to beta-3-tubulin (β-3-tub) (Covance, Princeton, NJ), which labels neurons; Glial Fibrillary Acidic Protein (GFAP) (Millipore, Billerica, MA), which labels astrocytes; and Ionized Calcium binding adaptor molecule 1 (Iba1) (Wako, Osaka, Japan), which labels microglia, were added, and the cultures incubated in a 4◦C refrigerator overnight. The wells were then aspirated, rinsed in HBHS 3×, and the following secondary antibodies were added: Alexa Fluor 488 Goat anti-mouse, Alexa Fluor 555 Goat anti-chicken, and Alexa Fluor 635 Goat antirabbit (Invitrogen, Carlsbad, CA). After a 2 h incubation at room temperature, the secondary antibodies were rinsed 3× with HBHS, and a final volume of 100 µl of HBHS was left in the wells for imaging. Special care was taken to ensure the microwire segments remained attached to the bottom of the wells.

#### **IMAGE ACQUISITION AND ANALYSIS**

Fluorescent images (512 × 512 pixels) were obtained on a confocal microscope fitted with a long working distance 10× air objective using Fluoview software (Olympus, Center Valley, PA). The different channels were imaged sequentially, and noise reduction was achieved by applying a Kalman filter built into the acquisition software to 3 scans for each channel. Each plate was imaged using the same set of imaging parameters (laser power, aperture, acquisition time) to ensure uniformity. Source images were imported into ImageJ (ImageJ, U. S. National Institutes of Health, Bethesda, MD), visually inspected and rotated to place the microwire in a vertical orientation. When possible, two adjacent rectangular selections, 480 pixels high by 240 pixels wide (equivalent to 994 µm by 496 µm), were made with the long edge running on the center of the wire. If that was not possible due to excessive proximity to wall of the well, only a single rectangular selection was made facing the interior of the well. Each of these selections was considered a single sample for analysis purposes. From these selections, intensity profiles of average brightness of each vertical line were generated, as shown in **Figure 1C**. Microwire segments were also imaged in three empty wells, and an average intensity profile was obtained and subtracted from the intensity profile generated from cell-containing wells. One response index (RI) per cell type was obtained for each region by summing the area underneath the intensity profile line between the distance points corresponding to the region boundaries and dividing by 10000. Statistical analysis was performed using the SAS 9.3 statistical package (SAS Institute Inc., Cary, NC). A general linear model (GLM) procedure was used perform to a one way ANOVA with block, to remove the effects of variations between the plates by treating the plates as a statistical block. *Post hoc* Tukey tests were used to determine statistical significance between the treatment groups at a significance level of α = 0.05. The error bars plotted

channel. The fluorescent image is pseudocolored to show neurons in red, astrocytes in green, and microglia in blue. Scale bar is 50 µm. For each

under the graph corresponding to the chosen distances and dividing by 10000.

represent the standard error of the means. *P*-values less than 0.05 are denoted in the figures by a single asterisk, while *p*-values less than 0.001 are denoted by double asterisks. Plots were generated using MATLAB (The MathWorks Inc., Natick, MA).

#### **RESULTS**

**Figure 1** shows an overview of the methodology employed to analyze the cellular responses to microwire segments. Microwire segments placed in the wells (**Figure 1A**) were imaged, resulting in sets of images such as the one shown in **Figure 1B**. Intensity profiles (**Figure 1C**) of areas of various widths were analyzed to obtain the results described below.

#### **MICROGLIA**

**Figure 2** shows the different levels of aggregate microglial response in interface areas of different sizes. In the interface area containing only the microwire (i.e., 25 µm), the only significant difference in the microglial RI was between the PEG coated microwire and LPS coated microwire (RI = 1.37 vs. 2.2, *p* = 0.007). For the interface area containing the wire and extending over an adjacent 25 µm, significant pairwise differences were observed between the LPS coated wire (RI = 5.84) and the other wires (uncoated RI = 4.92, *p* = 0.041; PEG RI = 4.26, *p* < 0.0001; LPS + PEG RI = 4.82, *p* = 0.022). For a wider interface area containing the microwire and extending over the adjacent 50 µm, these pairwise differences get stronger between the LPS coated wire (RI = 8.27) and the other wires (uncoated RI = 6.58, *p* = 0.0007; PEG RI = 5.8, *p* < 0.0001; LPS + PEG RI = 6.4, *p* = 0.0002). Notably, the relative pattern of the microglia response indices for the different conditions is the same for all three regions. Only the overall magnitude of the responses increase as the anti-Iba1 fluorescence is summed over larger areas. This indicates that all three size regions up to the wire plus 50 microns are representative of interface. In all three cases LPS induced microglia activation rises to a level statistical significance. As more distance is included in the interface measurement, the ability of the PEG to relieve microglia activation by LPS rises to statistical significance.

**Figure 3** shows the microglial responses at more distant regions. For the closest distant bin extending from 50 to 150 µm from edge of wire, the RI for LPS coated wire (RI = 5.62) was significantly higher than all the other treatments (uncoated RI = 4.21, *p* = 0.0001; PEG RI = 3.71, *p* < 0.0001; LPS + PEG RI = 3.91, *p* < 0.0001). For the next three distant 100 µm wide bins, the only significant difference observed was between LPS coated wire and PEG coated wire in all 3 bins. These calculated RI are as follows: for the bin extending from 150 to 250 µm from edge of wire: LPS RI = 4.5 vs. PEG 3.12, *p* = 0.0001; for the bin extending 250–350 µm from edge of wire: RI = 5.12 vs. 3.8, *p* = 0.0003; for the bin extending 350–450 µm from edge of wire): RI = 4.98 vs. 3.9, *p* = 0.01. Again the pattern of relative RI's is consistent at all distances in the distant regions and matches that of the interface region. In this case the size of each measurement area is the same across cases and is always 100 microns. Again LPS induces an increased RI in all regions reaching statistical significance when comparing LPS to the PEG coated wire and in the most near distant region (50–150 microns) when comparing LPS to an uncoated wire. The wire likely induces some basal level of microglial attachment or activation, which is reduced by PEG alone, therefore the effect of LPS is most pronounced when we compare the PEG wire. These results are consistent with a diffusion based model whereby the effect of LPS will decrease with increasing distance from the wire.

#### **ASTROCYTES**

**Figure 4** shows the astrocyte RI at interface areas. For the interface area containing only the microwire, the astrocyte RI for LPS

**FIGURE 3 | Microglial response to LPS-coated microwire compared to all other treatments is elevated at distances up to 150** µ**m, after which a tiered response is observed**.

coated wire (RI = 3.33) was significantly higher than PEG coated and LPS + PEG coated wire (PEG RI = 2.59, *p* = 0.015; LPS + PEG RI = 2.63, *p* = 0.02). For the interface area containing the wire and extending an adjacent 25 µm, the same pairwise difference were observed, but with a stronger difference between the LPS coated wire (RI = 6.7) and the LPS + PEG coated wire (PEG RI = 5.75, *p* = 0.012; LPS + PEG RI = 5.64, *p* = 0.0045). For the interface area containing the microwire and extending an adjacent 50 µm, the same observation of the LPS astrocyte RI being higher than both PEG and LPS + PEG was noticed (LPS RI = 7.54, PEG RI = 6.49, *p* = 0.02; LPS + PEG RI = 6.19, *p* = 0.002). Overall the astrocytes show a similar pattern in the interface as the microglia, but to a lesser extent. Importantly, for all three interface sizes (at the wire, within 25 µm of the wire, and within 50 µm of the wire), the PEG coating is able to significantly reduce the LPS-induced astrocyte response.

**Figure 5** shows the astrocyte RI at distant areas. No significant differences were observed between the different treatments for the closest distant bin extending from 50 to 150 µm from edge of microwire. For the middle two distant bins, a slightly significant difference was observed between LPS coated wire and LPS + PEG coated wire [bin 2 (150–250 µm from edge of wire): LPS RI = 2.31, LPS + PEG RI = 1.37, *p* = 0.012; bin 3 (250–350 µm from edge of wire): LPS RI = 2.73, LPS + PEG RI = 1.73, *p* = 0.03].

#### **NEURONS**

**Figures 6**, **7** show the neuron RI in interface and distant regions respectively. No significant differences in the neuron response were found between any of the treatment conditions in either interface or distant region. In contrast to microglia and astrocytes, where the RI was higher in distant areas in comparison to the widest interface area examined, the neuron RI in distant areas was roughly equal to that in the widest interface area examined.

#### **DISCUSSION**

#### **VALIDITY OF MODEL SYSTEM**

To test the effects of a dip coated PEG film on the cellular responses to implanted electrodes, we modified a robust and frequently replicated *in vitro* mixed cortical culture model pioneered by Polikov et al. (2006, 2009, 2010; Achyuta et al., 2010; Tien et al., 2013). The anatomical and physiological complexity *in vivo* obscures mechanistic investigations into the reactive tissue response to implanted microelectrodes. Using an *in vitro* model allows us to simplify the biological system under study, and isolate particular components of interest. The challenge with *in vitro* models is to simulate physiological conditions in the absence of particular anatomical structures. In this particular model of primary cortical cell cultures, the cells exist in isolation from supporting vasculature, structural extra-cellular matrix components, and meninges. These aforementioned structures are heavily damaged during microelectrode insertion, which has been shown to strongly affect the chronic response of the brain to implanted microelectrodes (Karumbaiah et al., 2013; Markwardt et al., 2013; Saxena et al., 2013). The original model (Polikov et al., 2006) did not elicit a consistent glial scar, and it was necessary to alter the composition of the culture media to place all glial cells in the culture in an elevated reactive state, thereby ensuring a consistent glial scar (Polikov et al., 2009). By coating LPS directly onto microwire, we are able to create a localized inflammatory microenvironment that more closely mimics the reality of an indwelling cortical implant, rather than placing the glial cells in the culture in a globally activated state. This localized inflammatory microenvironment enables us to examine distance related effects on the cultured cells. For the LPS + PEG condition, concerns about cross contamination and the potential to disrupt the dip-coated PEG film led to the decision to co-deposit PEG and LPS via dip-coating from a single pot. While polymeric films containing PEG have the potential for prolonged drug release, they are typically crosslinked to form hydrogels (Peppas, 1997; Lin and Anseth, 2009) or composites (Ramakrishna et al., 2001). Dipcoated films of a pure hydrophilic polymer, such as PEG, are rarely used for prolonged drug release due to their burst release characteristics and potential for dissolution over timescales shorter than is therapeutically beneficial (Acharya and Park, 2006). PEG, in various conformations, has been shown to accelerate the release of small hydrophobic molecules similar to LPS (Ooya et al., 2003; Kang et al., 2007). For these aforementioned reasons, we were confident that our codeposition of PEG and LPS would not hinder the exposure of the cells to LPS.

To examine microglial response, we chose to quantify Iba1 fluorescence across relatively wide bins. The choice of Iba1 was due to its high specificity to the microglia/macrophage cell type. The function and level of Iba1 expression is directly related to the classic morphological changes associated with microglial activation (Ito et al., 1998). Iba1 crosslinks actin and is involved in the formation of membrane ruffles and rapid motility (Sasaki et al., 2001). Additionally, Iba1 levels correlate directly with morphological feature changes associated with microglial activation (Kozlowski and Weimer, 2012). While current morphological analysis methods have been found lacking (Beynon and Walker,

2012), the method of quantification we employ is not without its shortcomings. Given our bin sizing, an increase in fluorescence in a bin could be attributed to either an increase in microglial cell numbers, or a higher level of Iba1 expression, or both. However, proliferation, migration, and morphological changes are all important components of microglial activation. Quantification of Iba1 fluorescence in a given area can therefore capture an aggregate of these aspects of microglial activation, but cannot distinguish between the individual components. We chose our method of quantification of Iba1fluorescence using bin sizes of up to 100 µm as an indicator of microglial response because we were most interested in quantifying gross activation across an extended distance from the foreign body. This resulted in a tradeoff against smaller bin sizes and higher magnification examination of individual microglia. Similar image analysis approaches quantifying fluorescence levels have been used *in vitro* (Polikov et al., 2009, 2010; Achyuta et al., 2010; Tien et al., 2013) and *in vivo* (Azemi et al., 2011; Potter et al., 2013, 2014) to analyze responses to microelectrodes and microscale foreign bodies., while presenting similar shortcomings in terms of elucidating separate aspects of microglial activation. Additional markers of microglial activation, such as secreted cytokines, are also a major factor of interest when studying microglial responses. Commercially available biochemical assays are not sensitive enough to detect secreted cytokines in this particular *in vitro* injury model. Future studies should examine improved experimental and analysis methodologies to combine gross microglial responses with morphological changes and biochemical expression patterns.

#### **ANALYSIS OF CELLULAR RESPONSES Microglia**

The microglial response in a narrow interface region comprising only the area under the microwire exhibits a three tiered response where a significant difference exists between the LPS only and the PEG only treatments, but not between the other conditions. This tiered response might be attributed to the difference between increased activation caused by the LPS and reduced cellular adhesion caused by PEG. The three data sets from the interfacial region included in **Figure 2** (wire only, wire + 25 µm, wire +50 µm) examine the RI of the microglia near the wire by summing the fluorescence over progressively increasing areas. We note that all three sets have the same relative trend when we compare each condition (bare wire, PEG only, LPS, LPS + PEG), only the magnitudes increase as the sets progress because the summation area increases. We observe a microglial monolayer forming at the surface of the wire, explaining the lack of a significant difference between the different treatments. The difference becomes significant only at the wire +50 µm because the magnitudes of the fluorescence become large enough to detect the differences given the variance of the assay. Even though this effect is not statistically significant when we analyze the wire only and wire +25 µm, the trend is consistent. The same is true for LPS vs. a bare wire, although this effect becomes significant at both wire +25 µm and wire +50 µm. We therefore think that the effect of LPS to locally activate microglia and mitigate that activation by a PEG coating is happening throughout the entire interfacial area. This increase in microglial response might be explained by elevated activation of microglia through amplification of inflammatory pathways precipitated by TL4 binding, leading to an increase in microglial response at distance. The observed elevation of Iba1 fluorescence persists in the next 100 µm wide distant region, again indicating an extended inflammatory response, potentially mediated by secreted cytokines produced by activated microglia but dissipates in further distant regions, reverting to a tiered response, where the only significant pairwise difference is between LPS and PEG. This tiered response can again be attributed to distinct pathway amplification between the two treatments; the difference appearing only between the increased upregulation of microglial activation due to LPS and the reduced microglial activation due to PEG.

#### **Astrocytes**

In interface regions of varying width, the astrocyte response also exhibits a three tiered response, where an elevated astrocyte response is observed with LPS, and a reduction occurs with both PEG conditions (with LPS and without). In the distant regions, the first and fourth 100 µm wide distant bin do not exhibit any differences between the different treatments, but we observe a difference between LPS and LPS + PEG in the middle two 100 µm wide bins, but surprisingly no difference between LPS and PEG in these distant areas. A potential explanation is that the astrocytes are exhibiting a dose dependent response to LPS. Under this explanation, the increased activation in the interface area for the LPS only treatment results in both astrocyte migration from distant regions and increased overall proliferation; delivering the LPS with PEG results in astrocyte migration without an accompanying equivalent increase in proliferation, resulting in a depletion of distant astrocytes; while PEG only results in even less astrocyte activation in interface areas, which in turn does not signal migration of distant astrocytes. Because we did not directly test for whether the LPS was acting through direct binding to receptors on astrocyte surfaces, we are merely discussing correlative effects. It is unclear whether the astrocyte response is due to direct action by LPS, or if it they are reacting to cytokines and chemokines secreted by microglia. While astrocytes are not typically thought to express TL4 receptors, there is some evidence to the contrary (Bowman et al., 2003). Additionally, while GFAP-positive astrocytes are observed in primary cultures, a considerable portion of them differentiate *in vitro* from astrocyte precursors (Abney et al., 1981). It is possible that due to these culture conditions that astrocyte response is altered from normally developing astrocytes *in vivo*.

#### **Neurons**

No significant differences were detected in neuron response to any of the treatments, in either interface or distant regions. While not statistically significant, a coupling between neuron and astrocyte response can be noticed, where slightly higher (but not significantly different) neuron growth was observed for the LPS treatment. Neuronal growth has been consistently shown to occur on a supporting substrate of astrocytes (Noble et al., 1984; Tomaselli et al., 1988). In contrast to the microglia and astrocytes, where the response in the widest interface bin was considerably higher than the first adjacent distant bin, the neuron RI in the first distant bin was comparable to the neuron RI in the wide interface bin, and we did not observe a decline in neuron density over distance. One explanation mirrors the concern expressed earlier about the maturity of the astrocytes, where immature astrocytes in culture provided a better substrate for neuron outgrowth compared to mature astrocytes (Smith et al., 1990). An alternative explanation is that elevated glial activation is not in and of itself neurotoxic or neurodegenerative within a foreign body reactive tissue response paradigm. If the latter explanation is correct, then the loss of neural density *in vivo* following implantation of a microelectrode might be better explained by displacement of neurons following insertion trauma and edema which fail to reoccupy depleted zones because of the glial scar formation, or that *in vivo* neurotoxicity occurs due to direct contact between neurons and extrabrain components.

#### **CONCLUSIONS**

We have shown that microglial response in a primary mixed cortical culture can be manipulated by dip-coated treatments. Microglial response can be increased by coating the surface of the foreign body with LPS, and this increase can be prevented by codepositing LPS and PEG. We hypothesize that the film of high molecular weight PEG, while allowing for LPS release, presents a hydrated physical barrier that disrupts cytokine, chemokine and adsorbed protein gradients that typically guide pathological responses. Astrocyte response also increased for LPS coated foreign bodies, but it is unclear whether this response is directly mediated by LPS or whether it is caused by other microgliasecreted factors. Neuron response was not negatively correlated with microglial response, suggesting mechanisms other than glial activation causing *in vivo* neuronal density loss. Our results highlight the importance of considering the *in vivo* chronic foreign body response as a complex phenomenon with multiple, interconnected yet parallel processes. Attempts to target an individual brain cell type to reduce the overall chronic response are unlikely to be successful. The differences between our findings and typical *in vivo* responses indicate the importance of components other than native brain cells in the progression of the reactive tissue response. Our findings additionally point to a viable alternative hypothesis regarding neuronal density depletion following microelectrode implantation in the brain.

#### **ACKNOWLEDGMENTS**

Funding for this research was provided by the Purdue Research Foundation, the Indiana Spinal Cord and Brain Injury Research Grant Program (Fund # 00015115), and the Defense Advanced Research Projects Agency (DARPA) Microsystems Technology Office (MTO), under the auspices of Dr. Jack W. Judy<EMAIL_ADDRESS>and Dr. Doug Weber<EMAIL_ADDRESS>as part of the Reliable Neural Technology Program, through the Space and Naval Warfare Systems Command (SPAWAR) Systems Center (SSC) Pacific grant No. N66001- 11-1-4013. Thanks to members of the Neuroprostheses Research Laboratory for feedback on the manuscript.

#### **REFERENCES**

contamination on the neuroinflammatory response to sterilized intracortical microelectrodes. *J. Mater. Chem. B Mater. Biol. Med.* 2, 2517–2529. doi: 10. 1039/c3tb21453b

**Conflict of Interest Statement**: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 01 May 2014; accepted: 28 October 2014; published online: 14 November 2014*.

*Citation: Sommakia S, Rickus JL and Otto KJ (2014) Glial cells, but not neurons, exhibit a controllable response to a localized inflammatory microenvironment in vitro. Front. Neuroeng. 7:41. doi: 10.3389/fneng.2014.00041*

*This article was submitted to the journal Frontiers in Neuroengineering*.

*Copyright © 2014 Sommakia, Rickus and Otto. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution and reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms*.

### Bio-inspired hybrid microelectrodes: a hybrid solution to improve long-term performance of chronic intracortical implants

#### *Sara De Faveri 1,2 \*, Emma Maggiolini 1, Ermanno Miele3 , Francesco De Angelis 3 , Fabrizia Cesca2 , Fabio Benfenati 2,4 † and Luciano Fadiga1,5 †*

1 Department of Robotics, Brain and Cognitive Science, Istituto Italiano di Tecnologia, Genova, Italy

2 Department of Neuroscience and Brain Technologies, Istituto Italiano di Tecnologia, Genova, Italy

3 Department of Nanostructures, Istituto Italiano di Tecnologia, Genova, Italy

4 Department of Experimental Medicine, University of Genova, Genova, Italy

5 Section of Human Physiology, University of Ferrara, Ferrara, Italy

#### *Edited by:*

Ulrich G. Hofmann, Albert-Ludwigs-University Freiburg, Germany

#### *Reviewed by:*

Christian K. E. Moll, University Clinic Hamburg-Eppendorf, Germany Jürgen Krüger, University of Freibourg, Germany

#### *\*Correspondence:*

Sara De Faveri, Department of Robotics, Brain and Cognitive Science, Istituto Italiano di Tecnologia, via Morego 30, Genova 16163, Italy e-mail<EMAIL_ADDRESS>
†Fabio Benfenati and Luciano Fadiga have contributed equally to this work. The use of implants that allow chronic electrical stimulation and recording in the brain of human patients is currently limited by a series of events that cause the deterioration over time of both the electrode surface and the surrounding tissue. The main reason of failure is the tissue inflammatory reaction that eventually causes neuronal loss and glial encapsulation, resulting in a progressive increase of the electrode-electrolyte impedance. Here, we describe a new method to create bio-inspired electrodes to mimic the mechanical properties and biological composition of the host tissue. This combination has a great potential to increase the implant lifetime by reducing tissue reaction and improving electrical coupling. Our method implies coating the electrode with reprogrammed neural or glial cells encapsulated within a hydrogel layer. We chose fibrin as a hydrogel and primary hippocampal neurons or astrocytes from rat brain as cellular layer. We demonstrate that fibrin coating is highly biocompatible, forms uniform coatings of controllable thickness, does not alter the electrochemical properties of the microelectrode and allows good quality recordings. Moreover, it reduces the amount of host reactive astrocytes – over time – compared to a bare wire and is fully reabsorbed by the surrounding tissue within 7 days after implantation, avoiding the common problem of hydrogels swelling. Both astrocytes and neurons could be successfully grown onto the electrode surface within the fibrin hydrogel without altering the electrochemical properties of the microelectrode.This bio-hybrid device has therefore a good potential to improve the electrical integration at the neuron-electrode interface and support the long-term success of neural prostheses.

**Keywords: gliosis, intracortical microelectrodes, fibrin, bio-inspired,** *in-vivo***, chronic implants**

#### **INTRODUCTION**

Patients with various neurological disorders could benefit from the stereotaxic implant of chronic intracerebral microelectrodes to record endogenous activity and/or to electrically stimulate specific brain areas (Donoghue et al., 2007; Larson, 2008; Truccolo et al., 2008; Velliste et al., 2008; Benabid et al., 2009; Normann et al., 2009; Hemm and Wårdell, 2010). To be functional and safe, the implant should keep a stable electric interface with the tissue, enabling an efficient recording and/or activation of neurons over time. However, the long-term functionality of implanted electrodes has been very critical, because of the progressive deterioration of both the electrode surface and the surrounding neural tissue.

Electrode insertion is a traumatic event that induces acute injury to the brain tissue (Polikov et al., 2005; Grill et al., 2009; Marin and Fernández, 2010). The tissue is cut, stretched or compressed by the penetration of the electrode and neurons and glial cells within and around the electrode path are killed or severely injured during insertion. Since the first day of implantation, microglial cells -that are extremely sensitive to even small pathological changes- start to be activated to scavenge damaged neurons and cell debris. Moreover, activated microglia also releases proinflammatory cytokines that propagate the inflammatory reaction. The deepening of the electrode into the brain pushes aside the tissue and increases the local pressure of the surrounding tissue. Blood vessels and capillaries are disrupted and the blood– brain barrier is compromised (Bjornsson et al., 2006), leading to plasma extravasation and local edema, that further increase the intracranial pressure (Unterberg et al., 2004).

This initial acute tissue reaction triggers, and is followed by, a more adverse long-term response mainly operated by activated astrocytes that is dependent on the extension of the injury and ends up in the generation of a glial scar, neuronal loss and impairment of the regrowth of neuronal processes (Fawcett and Asher, 1999; Polikov et al., 2005; Grill et al., 2009; Marin and Fernández, 2010). The glial encapsulation surrounding the electrode becomes denser and organized over time pushing away neurons from the tip of the probe (the so-called "kill zone"). This leads to

a progressive increase of the electrode-electrolyte impedance and to a deterioration of the recordings. All these events reduce the long-term performance of the electrodes, making the long-term application of this type of technology to human subjects almost unfeasible.

The physical properties of the electrodes such as surface composition, size, shape and stiffness, the mechanical connection between the probe and the recording system, the functionalization of the electrode and the implantation method all have some impact on the intensity of the tissue reaction (Edell et al., 1992). Different approaches have been proposed to solve or reduce the inflammatory reaction. One is the reduction of the electrode size. Minimizing the device footprint is important to reduce the insertion trauma. A smaller device induces a much smaller increase in the local pressure and minimizes tissue displacement (Kozai et al., 2012). However, miniaturization of the electrode worsens its electrochemical characteristics in terms of contact impedance and signal-to-noise ratio (SNR). To circumvent this effect, various groups have used conductive polymers such as poly(3,4-ethylenedioxythiophene, PEDOT) or nanostructured materials (e.g., carbon nanotubes) to reduce the impedance of the recording site (Richardson-Burns et al., 2007; Kim et al., 2010b; Ansaldo et al., 2011; Baranauskas et al., 2011).

Other solutions employing untethered or flexible probes (Rousche et al., 2001; Takeuchi et al., 2005; Kim et al., 2010a; Castagnola et al., 2013; Zhang et al., 2013) have been adopted to decrease the formation of the glial scar. Indeed, when an electrode is tethered to the skull or the device has a different compliance with respect to the adjacent brain tissue, the brain, subject to normal movement induced by respiration and circulation, moves around the probe causing chronic trauma as well as instability of the recording (Kim et al., 2004b; Biran et al., 2007; Thelin et al., 2011).

Approaches aimed at reducing the mechanical mismatch at the interface between the stiff intracortical electrode and the soft brain tissue have been proposed. Intracortical wires were covered with soft materials (Kim et al., 2010b) or with polymeric nano-composite materials that switch from a stiff to a compliant state when inside the cerebral tissue (Harris et al., 2011). The soft coating, matching the host tissue softness, was shown to play an important role in limiting the severity of the inflammatory reaction over time in terms of the amount of reactive astrocytes, neuronal death, and density of active microglial cells around the probe (Harris et al., 2011). The use of hydrogel scaffolds has also been introduced to contrast the consequences due to the inflammatory and immune responses by astrocytes and oligodendrocytes involved in the assembly of the glial scar. Both biologically derived hydrogels (e.g., alginate, Frampton et al., 2011; hyaluronic acid, Hou et al., 2005) and synthetic hydrogels (e.g., polyethylene glycol Bjugstad et al., 2010) have provided promising results in terms of repair and regeneration after surgical implants (Nisbet et al., 2008; Green et al., 2010). Bioactive materials such as L1, growth factors, dextran or laminin, have also been used to functionalize the probe improving its tolerability (He et al., 2006; Klaver and Caplan, 2007; Azemi et al., 2011). Furthermore, cell-coated interfaces were proposed to

increase the biocompatibility of the device and minimize the reaction of the host tissue (Purcell et al., 2009; Richter et al., 2011).

Although all the above-mentioned approaches may lead to potentially interesting results, the preservation of the performance of intracortical devices still remains an unresolved issue. To contribute to this challenge, we developed a strategy to improve the biocompatibility of the implant by coating microelectrodes with soft hydrogel and live cells to mimic both the mechanical properties and the biological composition of the host tissue. We used fibrin as hydrogel and either glial cells or neurons to immunologically hide the foreign object to the host tissue. By using such approach we obtained: (i) a strong adhesion of hydrogel and cells to the microelectrode without altering its electrochemical properties; (ii) a small and transient tissue reaction to the implant; and (iii) high quality of electrophysiological recordings.

### **MATERIALS AND METHODS**

#### **IMPLANTABLE WIRES** *Quartz-coated metal microelectrodes*

Single core quartz-insulated metal microelectrodes (Thomas Recording, Giessen, Germany) were prepared in-house by mechanically grinding 95% platinum, 5% tungsten microwires of 20 μm diameter coated with 30 μm of quartz (see **Figure 1A**). The measured impedance of these electrodes, measured in saline at 1 kHz, was typically between 500 and 700 k- (Ansaldo et al., 2011).

This probe was used as a seeding substrate and to evaluate the grip of the hydrogel to the electrode surface during intra-cortical insertion.

#### *Black-lead sticks*

Black-lead sticks were 350 μm in diameter and 3 mm long (see **Figure 1B**). The tip was mechanically ground. This probe was used to evaluate the time of hydrogel in-vivo reabsorption and the extent of tissue reaction to the implant without removing it from the brain, thanks to the possibility to cut the stick inserted in the brain with the microtome.

#### **FIBRIN HYDROGEL COATING**

Human fibrin (TISSEEL, Baxter, Deerfield, IL, USA) from pooled human plasma was used. It was composed by two sterile components, namely the sealer protein solution (aprotinin and fibrinogen) and thrombin solution (thrombin, CaCl2 and factor XIII). To form a cross-linked fibrin gel, fibrinogen was dipped in the thrombin solution to generate fibrin monomers that were subsequently cross-linked by factor XIII (soluble transglutaminase). Wires were coated by alternate dippings in the two components of the fibrin sealant. The resulting electrode coating was uniform and its thickness was controllable by the number of dippings (Kim et al., 2010b). The radial thickness of the coating was evaluated by measuring the hemi-diameter of the covered probe in five points and subtracting the probe diameter. Thin films of fibrin were prepared by dropping 50 μl of the first component of the fibrin sealant and 50 μl of the second component over sterile glass slides. The drops broadened and they were left drying in air for 15 min in a laminar hood.

#### **EVALUATION OF THE ELECTROCHEMICAL PROPERTIES OF THE FIBRIN-COATED DEVICE**

The electrochemical properties of the microelectrode were studied via electrochemical impedance spectroscopy (EIS) in a physiological aqueous solution (NaCl 0.9%). For the measurements, a sine wave (10 mV root mean square amplitude) was superimposed on the open circuit potential while varying the frequency from 1 Hz to 100 kHz. The electrochemical characterization was carried out using a potentiostat/galvanostat/ZRA (Reference 600, Gamry Instruments, Philadelphia, PA, USA) connected to a standard electrochemical cell used in a three-electrode configuration with a platinum counter-electrode (35 mm2 area) and an Ag/AgCl reference electrode.

#### **MODIFICATION OF THE MICROELECTRODE QUARTZ SURFACE**

The roughness of the microelectrode quartz surface was manipulated by using an argon ion plasma etching. Sputter-coated and annealed nanometric gold islands acted as masks for quartz surface milling. The ions hit all the surface, etching both quartz and gold but where quartz was not covered, the generated holes were deeper than where it was covered by gold. The process stopped when all the gold coating was removed. Thanks to this etching, the peak-to-peak average distance was about 500 nm (Goyal et al., 2006).

#### **EXPERIMENTAL ANIMALS**

Adult Sprague-Dawley rats (from Charles River, Calco, Italy) and Long-Evans rats (born in house) were used. All experiments were carried out in accordance with the guidelines established by the European Communities Council (Directive 2010/63/EU of September 22nd, 2010), and were approved by the Italian Ministry of Health.

#### *Neurons*

Neurons were isolated from E18 rat embryos. Briefly, embryos were removed, placed in cold Hank's balanced salts solution (HBSS) and hippocampi were dissected under sterile conditions. Subsequently, the tissue was enzymatically digested with 0.125% Trypsin for 20 min at 37◦C and disaggregated with a pipette. The cells were incubated with 1% Glutamax, penicillin-streptomycin, 2% B-27 supplemented Neurobasal medium in a humidified 5% CO2 atmosphere at 37◦C. Half of the medium was changed every week. All culture media and reagents were from Invitrogen (Milano, Italy).

#### *Glial cells*

Glial cells were isolated from the cerebral cortex of P0 rat pups. Cortices were dissected in cold HBSS, broken into small fragments by pipetting and enzymatically digested with 0.25% Trypsin – 1 mg/ml DNase for 15 min at 37◦C. After centrifugation at 1200 rpm for 5 min at 8◦C, the digestion medium was removed and the pellet was suspended in 10% horse serum, penicillin-streptomycin and 33 mM glucose-supplemented minimum essential medium. Cells were plated in culture flasks and half of the medium was changed every week.

#### **SEEDING OF CELLS OVER THE MICROELECTRODES**

Microelectrodes were fixed in a silicon cap by the end opposite to the tip and coated with 0.1% poly-D-lysine, which created a positively charged layer enhancing cell adhesion. Microelectrodes were dipped inside a vial containing the cells suspended in the medium (300000 cells/500 μl). Half of the medium was harvested from a high concentration cell culture, the so-called conditioned medium. The vials were kept in slow and continuous rotation in an incubator (2.31 rpm; relative centrifugal force 0.013 × *g*).

#### **EVALUATION OF THE BIOCOMPATIBILITY OF THE FIBRIN FILM**

Approximately 5000 astrocytes or neurons in 100 μl of medium were plated over a thin layer of fibrin. After the cells attached to the surface, the coverslip was covered by culture medium and the cell viability was analyzed at 1, 5, 7, and 11 days. Viability was evaluated by rating the surviving cells over the total number of cells. Samples were incubated for 3 min at room temperature in Ringer-Locke solution containing propidium iodide (PI; Sigma-Aldrich, Milano, Italy; 15 μg/ml), fluorescein diacetate (FDA; Sigma–Aldrich, Milano, Italy; 5 mg/ml) and Hoechst-33342 (Sigma-Aldrich, Milano, Italy; 3.3 μg/ml). After incubation, samples were washed once in Ringer-Locke solution and immediately imaged. At least 15 different fields of view were acquired for each sample. For each field, the ratio of PI-positive cells over the total number of nuclei, identified by Hoechst-33342, was calculated (Limongi et al., 2013).

#### **ANIMAL SURGERY**

Male Long Evans rats were anesthetized with a cocktail of Zoletil (Virbac, Carros cedex, France; 30 mg/kg) and Xylazine (Bayer, Leverkusen, Germany; 5 mg/kg) administered intraperitoneally. Their heads were then shaved and swabbed with ethanol. The anesthetized animal was placed in a stereotaxic apparatus (Myneurolab, St Louis, MO, USA). Lacrigel (Farmigea, Pisa, Italy) was distributed over the eyes to avoid dryness. An approximately 2 cm incision was made along the midline of the cranium. The underlying muscle and connective tissue were retracted to expose the skull. A craniotomy (5 mm × 3mm) was made in the parietal bone to expose the somatosensory cortex. Sterile saline was applied during pauses while drilling to help dissipate any local heating and to clean the surfaces. The exposed dura mater was wetted with saline, and carefully incised using surgical micro-scissors. Pia mater was incised too. Implantable wires were lowered perpendicularly through the cortical surface using a hydraulic microdrive (David Kopf Instruments, Tujunga, CA, USA) to 2800 μm from the surface of the brain (layer V). To minimize the variability associated with the surgery, all implants – both quartz-coated metal microelectrodes for acute experiments and black-lead sticks for chronic experiments – were performed by the same subject. During the surgery, the depth of anesthesia was monitored by testing for the absence of hindlimb withdrawal reflex and was maintained by additional doses of anesthetic (i.p. or i.m.), while the rat body temperature was kept at 36–38◦C by positioning the animal onto a thermostatic heating pad.

#### **ACUTE NEURAL RECORDING**

Acute recordings in layer V of rat cortex were performed to characterize *in vivo* the electrical performance of the fibrin-coated microelectrodes. Recordings were acquired for 20 min using a custom-made, low-power recording system (Bonfanti et al., 2008; Borghi et al., 2008) composed of a compact headstage (amplification gain = 62 dB) and a control unit for data acquisition. Raw traces were band-pass filtered between 3 Hz (first-order Butterworth) and 8 kHz (fourth-order Butterworth). The amplified signals were multiplexed, digitally converted (10-bit, 40 k sample/s) and sent via USB to a PC for subsequent analysis. LabView-based (National Instruments, Austin, TX, USA) acquisition software was used for real-time data visualization and storage. All experimental sessions were conducted inside a Faraday cage to reduce the electromagnetic noise interference.

#### **EVALUATION OF THE SIGNAL-TO-NOISE RATIO**

To evaluate the SNR, each acquired trace was high-pass digitally filtered above 300 Hz. The signal was wavelet decomposed and thresholded to 3.5 standard deviations (SD) above and below the mean of the sample distribution to discriminate signal from noise. Waveforms were clustered using Run-Valley seeking sort method (Offline-Sorter; Plexon Inc., Dallas, TX, USA). Signal amplitude was defined as the peak-to-peak amplitude of the mean waveform for each cluster. Noise was defined as two times the calculated rms noise voltage. The SNR was calculated as follows:

$$\text{SNR} = \frac{\text{S}}{2\,\text{V}rm\text{s}\_n} \tag{1}$$

where *S* is the signal amplitude and Vrmsn is the voltage root mean square of the noise. SNR values were sorted into categories (Ludwig et al., 2006, 2009; Abidian et al., 2009; Kim et al., 2010b). Clusters with a SNR greater than 4 were categorized as high-quality units, while clusters with a SNR between 3 and 4 were categorized as moderate units.

#### **IMMUNOHISTOCHEMICAL AND IMMUNOCYTOCHEMICAL TECHNIQUES**