Local Polymer Replacement for Neuron Patterning and in Situ Neurite

May 21, 2014 - and axonal outgrowth of embryonic hippocampal neurons in situ. Cultures of ... concept allows for building complex neuron networks acti...
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Local Polymer Replacement for Neuron Patterning and in Situ Neurite Guidance Harald Dermutz, Raphael R. Grüter, Anh Minh Truong, László Demkó, János Vörös,* and Tomaso Zambelli Laboratory of Biosensors and Bioelectronics, Institute for Biomedical Engineering, ETH Zurich, CH-8092 Zurich, Switzerland S Supporting Information *

ABSTRACT: By locally dispensing poly-L-lysine (PLL) molecules with a FluidFM onto a protein and cell resistant poly-Llysine-graft-polyethylene glycol (PLL-g-PEG) coated substrate, the antifouling layer can be replaced under the tip aperture by the cell adhesive PLL. We used this approach for guiding the adhesion and axonal outgrowth of embryonic hippocampal neurons in situ. Cultures of hippocampal neurons were chosen because they mostly contain pyramidal neurons. The hippocampus is known to be involved in memory formation, and the stages of network development are well characterized, which is an asset to fundamental research. After fabricating diffuse PLL spots with 10−250 μm diameter, seeded hippocampal cells stick preferentially onto the spots migrating toward the spot center along the PLL gradient. Cell clusters were formed depending on the lateral size of the PLL dots and the density of seeded cells. In a second step of this protocol, the FluidFM is used to connect in situ the obtained clusters. The outgrowth of neurites, which are known to grow preferentially on adhesive substrates, is tailored by writing PLL lines. Antibody staining confirms that the outgrowing neurites are mostly axons, while the activity of the neurons is assessed by a calcium indicator, proving cell viability. The calcium signal intensity of two actively interconnected clusters showed to be correlated, corroborating the formation of vectored and polarized interconnections.

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INTRODUCTION

After cell patterning the connections between neurons or clusters of neurons are formed spontaneously. However, the control of axon outgrowth is of main interest for many applications in neuroscience. Full knowledge of network connectivity makes it possible to study how single cells adapt to certain stimuli in far more detail than would be possible in random networks. Together with the possibility to study the activity of the whole network, such defined network cultures are a powerful tool to study the adaptation processes of neuronal networks. E-beam,13 photo,3,14 or scanning probe15 lithography was exploited to form surface patterns influencing the polarity of single neurons. Additionally, advances in microfluidics facilitate to test different axon guidance cues in cell stripe arrays.16−19 Besides these prepatterned “static” approaches, active surfaces were designed enabling a “dynamic” control over the cell adhesive properties20,21 for neuron manipulation. Either electrical impulses,22 UV irradiation,23 or laser pulses24 have been applied to remove the cell repellent coating and to direct the outgrowth of neurites. In another approach, the outgrowth of growth cones was induced by the hydrostatic force of a scanning ion conductance microscope.25 Even though these

Letourneau first described the effect of surface-adhesion properties on neuronal morphogenesis.1 Nowadays, this concept allows for building complex neuron networks acting as reliable logic devices2 or forming small neuron circuits of active, synaptically connected neurons.3 Thus, basic mechanisms of neuronal network behavior like synapse formation, synaptic plasticity, synaptic development, and functional properties can be addressed under defined conditions. Poly-L-lysine (PLL), a well-known “cell-adhesive” polymer, is often used for neuron patterning. Using standard techniques like photolithography4−6 or microcontact printing,7−9 the PLL layer and consequently the seeded neurons can be patterned. Hereby, the contrast may be improved by backfilling with “nonfouling” polymers. The activity and connections between the neurons can be recorded by deposition of the PLL pattern directly onto a multielectrode array5−7,9 or using the patchclamp technique.7,8 Besides these techniques, direct patterning methods have been conceived to avoid the photolithography step. Adapted microspotters and inkjet printers have been used for direct cell printing10 and quick production of surface patterns for cell adhesion.11 As an alternative, scanning probe lithography protocols like dip-pen lithography have been applied to deposit cell adhesive proteins locally on the surface.12 © 2014 American Chemical Society

Received: April 2, 2014 Revised: May 15, 2014 Published: May 21, 2014 7037

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(OW 2400; MicroVacuum Ltd.) were first cleaned with SDS (sodium dodecyl sulfate), and rinsed with ultrapure water (Millipore Corporation, Billerica, MA). Afterward, the waveguides were immersed in isopropanol, cleaned in an ultrasonic bath for 10 min, rinsed again with ultrapure water, and dried under a nitrogen stream. As last steps the waveguides were treated in an oxygen plasma cleaner (PDC-32G; Harrick Plasma, Ithaca, NY) at “high” power (18 W) for 2 min and mounted into the flow cell, and the flow cell was filled with PBS (phosphate-buffered saline, pH 7.4; Life Technologies Ltd., Paisley, United Kingdom). Before the measurement the flow cell was kept overnight to stabilize the baseline signal. Chemicals. PLL-g-PEG (PLL(20)-g[3.5]-PEG(2)) and fluorescent PLL-g-PEG-Atto (PLL(20)-g[3.5]-PEG(2)/Atto633) were purchased from Surface Solutions (SuSoS AG, Dübendorf, Switzerland) as powder. PLL (poly-L-lysine hydrobromide, MW 30−70 and 70−150 kDa) and fluorescent PLL-FITC (poly-L-lysine-FITC labeled, MW 30−70 kDa) were purchased from Sigma-Aldrich (St. Louis, MO) as a powder. Stock solutions of the polymers were done in HEPES II, at a concentration of 10 mg/mL, and stored at −20 °C. The polymers were dissolved in PBS (phosphate-buffered saline, pH 7.4; Life Technologies Ltd., Paisley, United Kingdom) at the desired concentration. Substrate Preparation. Glass dishes (WillCo-Dish; WillCo Wells B.V., Amsterdam, Netherlands) were first cleaned in oxygen plasma at high power (18 W) for 2 min and subsequently covered with filtered (0.2 μm) PLL-g-PEG solution for 45 min. Finally, they were rinsed with ultrapure water and dried under nitrogen stream. Primary Neuronal Culture. Animals. All experiments were performed with primary rat hippocampal cell cultures prepared from E17 embryos, taken from time-mated pregnant Wistar rats (Harlan Laboratories, Netherlands). The hippocampus was dissected on ice. Dissection of the animals was carried out by personnel of the Institute of Pharmacology and Toxicology - Morphological and Behavioral Neuroscience of the University of Zurich. Culture of Hippocampal Neurons. Mixed primary embryonic hippocampal neurons were counted (Countess Automated Cell Counter; Life Technologies Ltd.) and diluted to the desired concentration (from 10 000 to 100 000 cells/cm2) with serum containing media, further called “attachment medium”. Attachment Medium. High Glucose Dulbecco’s Modified Medium (DMEM, Life Technologies Ltd.) was mixed with 1% GlutaMAX (Catalog # 61965-026, Life Technologies Ltd.), 10% fetal bovine serum (Catalog # 10270-106, Life Technologies Ltd.), and 1% antibiotic-antimycotic (Catalog # 15240-062, Life Technologies Ltd.). Cell Culture Medium. After the attachment process the medium was exchanged with serum-free medium to suppress astroglia proliferation and optimal cell survival in the low-density cultures.34 This medium is composed of neurobasal medium (Catalog # 21103049, Life Technologies Ltd.) mixed with 2% B-27 serum-free supplement (Catalog # 17504-044, Life Technologies Ltd.), 1% penicillin−streptomycin (Catalog # 15140-148, Life Technologies Ltd.), and 1% GlutaMAX (Catalog # 61965-026, Life Technologies Ltd.). Primary Neuron Culture Treatment. Before cells were plated, the patterned dishes were filled with attachment medium and incubated at 37 °C, 5% CO2 for at least 15 min to adjust the pH. After cell plating, dishes were incubated for 30 min at 37 °C, 5% CO2 before they were intensively rinsed with cell culture medium several times to dispose of nonadhering neurons. Cultured dishes were incubated at 37 °C, 5% CO2. Half of the medium was exchanged three times a week. Calcium Imaging. Spontaneous activity of neurons was shown with a calcium indicator (Oregon Green 488 Bapta-1, AM; Life Technologies Ltd.). A stock solution of 50 μg of Oregon Green Bapta-1 dissolved in DMSO was prepared. After at least 14 days in vitro (DIV), stock solution of Oregon Green Bapta-1 was diluted to 6 μM in cell culture medium, added to the dish of interest, and incubated for 60 min at 37 °C, 5% CO2. Images were taken with an electron multiplier CCD camera (Hamamatsu C9100-13; Hamamatsu Photonics K. K.) connected to an AxioObserver-Z1 (Carl Zeiss AG)

techniques allow for a direct manipulation of neurons and axons, the controlled formation of an active synaptic connection has yet not been reported. Such artificially designed patterns of neuron networks are one approach to simplify the complexity of the neuronal systems in the mammalian brain. In a defined system, it becomes possible to study the electrical interplay and the underlying operational principles of basic neuronal circuits. The accessibility of such systems makes it possible to interact with single nodes and to study the influence of external stimuli on the output of the network. The activity of patterned networks was shown via a calcium sensitive dye2,26 or with patterned neuron clusters on top of a multielectrode array (MEA).5,27,28 In this work, we established a scanning probe lithographybased method allowing for both tasks: neuron patterning and in situ creation of connections between electrically active neuron clusters. The ability of a dynamic deposition in the presence of cells substantially differs from previous experiments using scanning probe lithography. For this, the FluidFM,29 which combines a microchanneled cantilever with a closed fluidic circuit and a pressure controller, is used as lithography tool in liquid environment.30,31 We utilized this technique to locally modify the surface properties not only by removing the low fouling background but also by simultaneously depositing PLL molecules without any additional fabrication steps. Fluorescently labeled polymers enabled an immediate visual control of the replacement process. Furthermore, it allowed the direct comparison between the fabricated polymer pattern and the dynamic process of patterned cluster formation of the seeded rat hippocampal neurons. Additionally, the FluidFM system allowed direct manipulation of the neuron clusters under physiological conditions and thereby actively control the network formation. Thus, PLL guiding lines were written in situ to stimulate the neurons to establish connections between the patterned clusters. Finally, the synchronized activity of connected clusters was recorded with calcium imaging.

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EXPERIMENTAL SECTION

FluidFM Probes. For all experiments tipless cantilevers fixed on a CYTOCLIP (Cytosurge AG, Zurich, Switzerland) were used. They were 200 μm long and 36 μm wide, having a 1 μm thick internal channel and a circular aperture of 2 μm diameter at the end of the cantilever (stiffness 3 N/m). FluidFM Equipment. The cantilevers were mounted on a standard atomic force microscope (JPK Instruments, Nanowizard I AFM) and connected to a pressure controller (Fluigent, MicroFluidics Control System), with an output pressure precision of better than 2.5%, to adjust the pressure in the liquid reservoir of the CYTOCLIP. For deposition experiments in the presence of neurons, the AFM head was set up onto a laser scanning microscope (LSM 510, Carl Zeiss AG) equipped with an incubation box assuring a controlled temperature of 37 °C. The samples were imaged using an excitation wavelength of 633 nm (HeNe laser) with a high pass (635 nm) and a low pass (650 nm) filter to detect the Atto dye and an excitation wavelength of 488 nm (argon laser) with a band-pass (505−550 nm) filter for the FITC dye. Optical Waveguide Lightmode Spectroscopy (OWLS) and Waveguide Preparation. In optical waveguide lightmode spectroscopy, the change in the refractive index within the evanescent field of a laser, coupled into a planar waveguide via an optical grating, is measured. Adsorption and desorption of molecules to and from the waveguide lead to a change of refractive index, which can be translated into a shift of the incoupling angle of the laser. The adsorbed mass can be determined using the formula of de Feijter.32 For PLL and PLL-gPEG, a dn/dc value of 0.169 was used.33 For the OWLS measurements (OWLS 120, MicroVacuum Ltd., Budapest, Hungary) the waveguides 7038

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with a Colibri fluorescence system. The dye was excited with a wavelength of 470 nm (LED) through a band-pass filter (470/40 nm). As emission filter, a band-pass filter (525/50 nm) was used. During the measurement, the incubator box around the microscope was regulated to maintain a temperature of 37 °C. Data Analysis. Videos were recorded at 30 and 65 frames/s and evaluated with ImageJ.35 The effect of bleaching has been approximated with exponential intensity decay and removed by the following steps. First the time constant of the decay was calculated from the intensity change of the pixels representing cells having no or minimal activity; then this time constant was used to fit the time course of every pixel individually. Finally, these fits were removed, keeping the baseline and neuronal activity only. Fluorescence signals were quantified by manually selecting a region of interest (ROI) over a region of neurons on the different spots. The mean pixel values of the ROI was calculated with ImageJ and plotted in Origin (Origin 8; OriginLab USA). Change of fluorescence I/I0 was computed, where I0 was calculated by calculating the mean over the last 100 data points of the signal, which corresponds to 7 s. The normalized cross-correlation was calculated by the corresponding algorithm in Origin. Immunocytochemistry. After calcium imaging, cells were fixed for 10 min with 4% paraformaldehyde (Sigma-Aldrich, Switzerland) and washed three times for 10 min in PBS (phosphate-buffered saline, pH 7.4; Life Technologies Ltd., Paisley, United Kingdom). For labeling, cells where permeabilized for 7 min with 0.1% Triton X-100 in PBS and washed three times for 5 min. To avoid nonspecific binding, the sample was incubated for 1 h at room temperature in 10% fetal bovine serum (FBS) (Sigma-Aldrich, Switzerland). The primary antibodies mouse anti-Pan-Axonal Neurofilament Marker (SMI 312) (Enzo Life Science, Switzerland) and rabbit antimicrotubule-associated protein 2 (MAP2) (Sigma-Aldrich, Switzerland) were diluted in 10% FBS and incubated overnight at 4 °C, followed by three 5 min PBS washing steps. Afterward, the samples were incubated with anti-mouse IgGAtto 633 antibody produced in goat (Sigma-Aldrich), anti-rabbit IgGAlexa Fluor 488 produced in goat (Life Technologies Ltd.), and DAPI (4′,6-diamidino-2-phenylindole dihydrochloride; Sigma-Aldrich).

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Figure 1. (a) Black curves: adsorbed mass for sequential adsorption of PLL-g-PEG and PLL involving (A) preadsorption of PLL-g-PEG layer after addition of PLL-g-PEG solution, (B) rinsing with ultrapure water and PBS, and (C) exposure of the layer to PLL. Red curves: adsorption of PLL onto uncoated substrate. (b−g) Confocal images of a glass bottom dish using Atto-settings (b−d) and FITC-settings (e− g) before coating (b, e), after immersion for 45 min in PLL-g-PEGAtto (0.1 mg/mL in PBS) solution (c, f), and after subsequent immersion for 45 min in PLL-FITC (40 μg/mL in PBS) solution (d, g).

RESULTS AND DISCUSSION Polymer Replacement at the Macroscopic Scale. The phenomenon at the basis of this study is the local polymer replacement of a nonfouling polymer (in our case PLL-gPEG36−38), with a cell adhesive polymer (in our case PLL, known in cell culture to promote neuron adhesion onto substrates39−42). The polymer exchange was reported first by Gon et al.43 Using optical reflectometry, they observed that PLL from solution can challenge a PLL-g-PEG layer, partially replacing it at the surface. This replacement process was found to be dependent on the grafting ratio g (number of lysine groups over number of PEG groups) and the chain length of the polymers. In order to evaluate the extent and kinetics of this exchange between the polymers at our disposal, OWLS and CLSM experiments were performed. First, the adsorbed mass of a PLL monolayer on the OWLS waveguide was determined by injecting PLL solution (40 μg/mL in PBS) into the flow cell (Figure 1a, red curves). The PLL concentration was selected based on toxicity tests exploring the effect of the PLL concentration onto the viability of the neurons (see Supporting Information, Figure SI1). All three measurements resulted in an adsorbed mass density of 136 ± 2 ng/cm2 45 min after the injection. Repeating the same protocol with PLL-g-PEG solution (0.1 mg/mL in PBS) resulted in a consistently thicker monolayer (Figure 1a, black curves). The adsorbed mass density was measured to be 184 ± 1 ng/cm2 45 min after injection for all three waveguides. Continuing the measurement by rinsing with water and PBS and subsequent injection of the

PLL solution, the replacement phenomenon was studied. After injection the mass signal instantly increased (overshoot) and then slowly decreased, reaching a value of 154 ± 6 ng/cm2 45 min after PLL injection. Thus, around 62 ± 13% of the PLL-gPEG layer was replaced by PLL. Gon et al.43 reported a complete replacement for PLL-g-PEG molecules with a smaller grafting ratio (2.7). For bigger grafting ratios the adhesion of the resulting PLL-g-PEG layer is stronger because of the higher amount of charges on the backbone and the lower steric hindrance between the PEG side chains.44 The overshoot right after injection is in good agreement with literature43 and is caused by a fast adsorption of PLL molecules, followed by a slower desorption of PLL-g-PEG molecules from the surface. The slow desorption of PLL-g-PEG is also apparent in the exponential decay (see Supporting Information, Figure SI2a) of the adsorbed mass during the exchange, which indicates a surface-limited process. Fu et al. 45 studied the same phenomenon showing that the chain length of exchanged homopolymers is influencing the kinetics of the replacement 7039

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Figure 2. Confocal images of a glass bottom dish using Atto-settings (a) and FITC-settings (b) before (left) and after (right) deposition of PLL dots with 500 mbar for 2 min. (c) Color-coded release of PLL-g-PEG; the image after deposition was subtracted from the image before deposition and normalized. (d) Color-coded deposition of PLL; the image after deposition was subtracted from the image before deposition and normalized. (e) Intensity profiles (along red line) using Atto settings (above) and FITC settings (below) at different times t0−t3. (f) Schematic illustration (top and side view) of a FluidFM probe during deposition in contact mode. (g) Fluorescence intensity over time for a deposition with 500 mbar for 1 min. The fluorescence intensity is measured for Atto and FITC settings at the red spot in the fluorescence images. Overpressure is applied at time point A and stopped at time point B when also the cantilever is removed from the surface.

conditions (more details in the Supporting Information). Fluorescent labeling allows to directly analyze the polymer replacement under the optical microscope. Local Polymer Replacement at the Microscopic Scale. Upon characterization of the system at the macroscopic scale, we transferred the polymer exchange protocol to the micrometer scale using the FluidFM. The FluidFM allows for the pressure-controlled local deposition in liquid environment. Both pressure and contact time influence the amount of dispensed volume and therefore the number of deposited molecules.31 The FluidFM microchannel was filled with PLLFITC solution (40 μg/mL in PBS) which was delivered onto the glass surface coated with PLL-g-PEG-Atto. Once the cantilever was approached at the surface using the AFM forcefeedback, an overpressure in the fluidic system led to a local dispensing of PLL-FITC molecules onto the surface. The surface was immersed in PBS. Applying a pressure of +500 mbar for 2 min led to the exchange of PLL-g-PEG for PLL under the aperture of the cantilever. Figures 2a,b are confocal images of the substrate before (t0) and after (t4) local polymer replacement at four chosen locations. Cell adhesion spots (PLL) of ∼200 μm diameter were written into a cell repulsive background (PLL-g-PEG). The extent of polymer exchange could be estimated relying on the fluorescence intensity measurements in Figure 1. The change in fluorescence for each pixel was translated into a color-coded scale, illustrating

process. Especially for small differences in the chain length, like in our case, the exchange is surface-limited. The doubleexponential fitting curves (R2 ≥ 0.99) resulted in a fast (t1 = 3.8 ± 1.1 min, A1 = 22.9 ± 0.6 ng/cm2) and a slow process (t2 = 17.7 ± 4.4 min, A2 = 27.5 ± 3.2 ng/cm2) for all three experiments. These kinetics can be explained by a fast desorption due to the additionally adsorbed PLL molecules during the overshoot (high polymer density) and a slower relaxation of the polymer layer after the overshoot. As a summary so far, the OWLS experiments confirmed the replacement process of PLL-g-PEG and PLL with minor kinetic discrepancies to literature, probably related to the differences in structural properties and concentrations of the polymers used in this study. Since we envisaged monitoring the exchange process in situ during the FluidFM lithography experiments, we additionally investigated the polymer replacement of fluorescently labeled PLL and PLL-g-PEG with a CLSM. By consecutive immersion of a glass dish into PLL-g-PEG-Atto and PLL-FITC, the fluorescence signal can be used to analyze the replacement phenomenon (Figure 1b−g). After immersion in PLL-FITC solution for 45 min, 82 ± 4% of the PLL-g-PEG-Atto molecules were desorbed from the substrate while 77 ± 9% of a PLLFITC monolayer was formed. These values were calculated by comparing them to the fluorescence intensity of the polymer layers that formed directly on a bare dish under the same 7040

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the amount of desorption of PLL-g-PEG and adsorption of PLL (Figures 2c,d). Thus, in the center of the dots (r ± 25 μm) 69 ± 7% of the PLL-g-PEG layer was replaced by PLL molecules, whereas outward the amount of the exchange gradually decreased. This gradient formation is shown in the intensity profiles (Figure 2e) along a line across the dots before deposition (t0), right after deposition at the right (t1) respectively left (t2) spot and 2 min (for the left spot) respectively 4 min (for the right spot) after deposition (t3). A Gaussian fit of the profiles at t3 results in a full width at halfmaximum (FWHM) of 122 ± 3 μm for both dots. While the intensity profile for PLL-FITC is unaltered after dispensing, the intensity of PLL-g-PEG-Atto clearly keeps decreasing. This relatively slow desorption process of the PLL-g-PEG molecules is consistent with the overshoot effect in the OWLS measurements. Once the equilibrium is reached, the spots show an asymmetric shape with a higher gradient to the right. This can be explained by an increased transport of PLL molecules to the left due to the 10° tilt angle of the FluidFM cantilever (i.e., the aperture edge is not completely in contact with the underlying substrate), illustrated in Figure 2f. To provide a deeper insight in the dynamics of the exchange process, fluorescence movies for both Atto- and FITC-settings were recorded during the FluidFM replacement experiment. Figure 2g shows the evolution of the intensity signals at the position marked with a red spot in the inset fluorescence images recorded before and after the measurement. In contrast to the OWLS experiments, this allows a clear distinction between the adsorption and the desorption signals. By applying a pressure (500 mbar), the PLL signal increased instantly whereas the PLL-g-PEG signal is affected only little. Afterward, the adsorbed PLL molecules slowly displaced the PLL-g-PEG from the surface. One minute later, the pressure was stopped and the cantilever was retracted from the surface. Even though this stopped the flux of PLL molecules to the surface, the desorption of PLL-g-PEG layer continued. In fact, about 50% of PLL-g-PEG was released after the flux of PLL was stopped, which again confirms that the polymer exchange is a surfacelimited process. The double-exponential fit (R2 ≥ 0.99, see Supporting Information, Figure SI2b) for the decay of the fluorescence signal resulted again in a fast (t1 = 1.05 ± 0.03 min) and a slow (t2 = 18.0 ± 2.7) desorption process. The time constant of the fast process t1 is consistent with the length of the pressure pulse (1 min) and represents the direct exchange during PLL adsorption. Since the PLL adsorption is stopped after the pressure pulse, t1 is consistently shorter compared to the OWLS experiment where the PLL adsorption is not restricted to a pulse. On the other hand, the time constant t2 of the slower process describes the kinetics of the polymer relaxation after adsorption of the PLL molecules, which is identical for both FluidFM and OWLS experiments. Cell Patterning on the PLL Spots and Cell Migration along the PLL Gradient. Using the approach described above, we generated cell-adhesive PLL spots of tunable size (5−200 μm) and separation (20−400 μm) in an antifouling PLL-g-PEG background. The Petri dish was immersed in attachment medium. After the preparation of the spots, dissociated rat embryonic hippocampal neurons were seeded onto the glass dish. The cells were incubated for 30 min in serum containing media. Nonadherent cells were rinsed off immediately afterward with serum free medium. As described also by Yavin et al.,39 rat embryonic neurons show a fast adhesion time onto PLL layers. Figures 3a,b show fluorescence

Figure 3. (a−d) Left: confocal image using Atto settings showing the pattern of the PLL-g-PEG layer. Right: corresponding bright field images of the neurons at 1, 4, and 6 days in vitro (DIV). It can be seen that the neurons migrate into the center of the deposited spots after days, forming clusters. Depending on the spot size and distance, these clusters formed spontaneous interconnections or stayed separated.

images of the patterns and bright field images of the neurons right after seeding and after 4 days in vitro (DIV). Whereas the distribution of attached neurons correlates only little after 1 DIV, the neurons are able to “feel” the presence of the PLL gradient. They gradually migrate along it toward the center of the PLL spots, where they build clusters after 4 DIV. This observed soma migration is consistent with results from other reports,46−48 where hippocampal neurons were seeded onto a predefined “static” surface pattern of cell adhesive coatings. Yet, in these experiments, migration was only observed on homogeneous coatings with a sharp border between the less and more adhesive coating and not on an inverse gradient between antifouling PLL-g-PEG and adhesive PLL like in our case. The interplay of the decreasing concentration of PLL-gPEG and the increase of PLL probably favors a preferred polarization of focal adhesion, resulting in the consequent observed directional migration. Neuron migration toward areas with an increased local adhesivity around the soma is supported by the findings of Corey et al.46 By means of laser ablation, they fabricated a poly-D-lysine (PDL) grid pattern with bigger circles (nodes) at the intersections and showed that hippocampal neurons migrate from the thinner paths of PDL toward the bigger PDL nodes. This is dependent on the distance and ratio 7041

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Figure 4. (a, b) Bright field image of neuron clusters before (left) and 2 respectively 11 days after PLL line writing (right). The in situ deposited patterns are highlighted in the confocal image using Atto settings (middle). It can be clearly seen that the neurites reacted to the deposited PLL lines.

cones which already grew by chance with the right direction toward another cluster are stimulated to grow over the nonadhesive gap via chemical attraction as proposed by Staii et al. In Situ Guided Connection of Two Neuron Clusters. Whereas the neuron polarity in the spontaneously built connections is random, we envisaged the possibility to actively influence the creation of connections having a specific polarity. The advantage of the presented method is the ability to modify the surface in situ, not only before cell seeding but also in the presence of already seeded cells. In particular, a PLL line can be written into the region between two neuron clusters in order to trigger the outgrowth of already formed neurites toward the other existing clusters (“guided” connection). We previously spotted a pattern so that the separation between the clusters exceeded the mean length of neurites normally formed on the patterned circular spots after 6 DIV. Since the neurites outgrowing on the line will be much longer than the already existing neurites, only axons should bridge the distance between the two spots as foreseen by Dotti and Banker,50 Goslin and Banker,51 and more recently by Yamamoto et al.52 All these works agree that the longest neurite will become the axon. For example, if an already emerged axon is cut on purpose, the longest remaining neurite forms the new axon.50 Although the size of the spots investigated in this work probably allows for the existence of long neuronal processes also inside individual spots, the distance between the spots increases the probability that only axons will reach the adjacent spot. After 6 DIV PLL lines were written between unconnected neuron clusters using a pressure of 5 mbar while the tip was moved over the surface with a velocity of 2 μm s−1. In Figure 4, confocal images of the resulting removal of the PLL-g-PEG layer along the PLL line are shown for two different areas. In both cases the neurons clearly react to the deposited lines and grew neurites along the PLL lines. However, the timing of the outgrowth can differ strongly from cluster to cluster (2−12 days) which is probably dependent on the PLL density around the neuron cluster and the exact shape of the deposited PLL line. To determine the polarization of the formed neuronal connections, immunostaining of the samples was carried out using pan-axonal neurofilament marker SMI312 and MAP2,

between the path and the node diameter. In our work, the amount of cells in the clusters is dependent on the size of the PLL spots and on the cell density during seeding. In Figure 3a, a 200 μm PLL-spot with a seeding density of 100 000 cells/cm2 induced the formation of a cluster containing around 100 neurons. Smaller spots (60 μm diameter) and seeding densities (50 000 cells/cm2) led to the formation of clusters with around 10 neurons (Figure 3b). By reducing the size of the spots down to around 10 μm, single cell resolution could be reached (see Supporting Information, Figure SI3). However, at this scale, placing the cells by adhesion out of suspension without any further guidance (e.g., with help of microfluidic tools) is not reliable enough to achieve reproducible patterns. Furthermore, the survival rate of not connected neurons is much lower. This could eventually be improved by adding feeder layers with high density cell culture areas in close proximity to the pattern.49 We noticed that spontaneous interconnections between the neuron clusters were formed for specific size and separation distance of the PLL spots (Figure 3c,d). In the same glass bottom dish two areas with three PLL spots in a row were prepared. While the distance between the spots was kept the same (∼250 μm), the amount of adsorbed PLL was adjusted by different contact times of the cantilever. At spots with longer contact times (Figure 3c, 3 min, corresponding to a FWHM 213 ± 2 μm size) the neuron clusters built connections, whereas for shorter contact times (Figure 3d, 1 min, corresponding to a FWHM 164 ± 2 μm size) the clusters did not connect at all. Longer contact times result in more PLL diffusion into the area between the PLL dots and therefore wider PLL gradients. Considering the findings of Staii et al.,15 the connection between two spots over such a long distance (>35 μm) should only occur, if an adhesive path is accessible to the outgrowing processes pointing toward the adjacent spot. Staii and co-workers produced the patterns of adhesive squares and thinner adhesive lines by scratching a PEG monolayer with a standard AFM cantilever from a gold surface. Afterward, they backfilled with PDL and plated the cells. By changing the distance between the adhesive squares, they could investigate the optimal distance over which the cells sitting on the squares could still interconnect. Since in our work the FluidFM dispensing leads to approximately concentric PLL spots, growth 7042

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Figure 5. (a) Bright field image of neuron clusters before PLL line writing. (b) Confocal image using FITC settings highlighting the in situ written pattern. (c) Confocal image showing the fluorescently labeled dendrites (MAP2). (d) Confocal image showing the fluorescently labeled axons. The outgrowth of a single axon from cluster C along the deposited PLL line is shown by the axon specific antibody SMI-312. Strong attachment of the axon to the deposited PLL line is evident in the shape of the axon.

cluster activity of around 0.6−0.75 Hz corresponds to the results shown by Jun et al. for a density of 100−200 cells/ mm2.5,27,53 However, due to the limited time resolution of the camera, conclusions over the single spiking frequency cannot be made. To analyze the connections between the clusters, the cross-correlation between the fluorescent signals of connected clusters was determined (Figure 6b).54,55 Between clusters Aguide and Bguide a correlation coefficient of around 0.6 was observed. This is consistent with the work of Liu et al., reporting correlation coefficients in the range of 0.55−0.69 for neurons, which were connected to the rest of the network.56 The time shift of the correlation shown in Figure 6b shows a time delay of around 50 ms between clusters Aguide and Bguide. Although the temporal resolution of the calcium indicator recording is limited to 30 Hz, which makes it impossible to resolve single synaptic delays,57 these results indicate a synchronized bursting activity between clusters Aguide and Bguide. Considering the conduction velocities in unmyelinated neurons reported in the literature, the delay due to traveling of the action potential would lead to a delay of around 830 μs. This indicates that a time delay of 50 ms between clusters Aguide and Bguide cannot be explained by one direct connection between the two clusters, but rather shows that the recorded signals are caused by synchronized bursting activity of the

which labels the axon and somatodentric regions, respectively. In Figure 5, fluorescence images are shown after 20 DIV for three clusters connected both spontaneously (clusters A and B) and guided (clusters B and C; the PLL line was written after 12 DIV as shown in Figure 5b). The spontaneous connection is bulky and contains several dendrites (Figure 5c) and axons (Figure 5d), whereas the deliberate connection consists of a single axon. Thus, writing a line of adsorption molecules in situ can guide the outgrowth of a single axon. Although a specific staining of the clusters for excitatory and inhibitory neurons was not made, the findings by Sun et al. suggest that they mostly consist of excitatory neurons.48 Indeed, Sun et al. created different adhesive surfaces with spot sizes similar to those of our work by microcontact printing and found that the clusters consist of excitatory neurons while inhibitory neurons were observed only in the periphery of the geometric constraint away from the clusters. Activity of Interconnected Neuron Clusters. As last issue, the vitality and activity of two neuron cluster (Aguide, Bguide) with guided connections (Figure 6a) were shown using a calcium indicator. Binding of intracellular Ca2+ to the dye molecule generates a fluorescence signal, which allows to study the activity of neurons. The fluorescence signal of the clusters was recorded for 120 s (Figure 6b). The observed rate of 7043

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Figure 6. (a) From left to right: bright field imaging of the neuron clusters Aguide and Bguide before PLL line writing, fluorescence imaging of the neuron clusters Aguide and Bguide using FITC settings after PLL line writing and fluorescence imaging using a calcium indicator. (b) Measured relative change in intensity over time for neuron clusters Aguide and Bguide. The clusters show regular bursting frequency over the whole recording time. Inset: magnified region of the intensity change over 10 s and the normalized correlation factor between the two connected clusters.

clusters.58 The fact that the action potential of cluster Aguide occurs always prior to Bguide shows that the induced connection is unidirectional. However, due to the temporal resolution of the signal, a clear analysis is difficult. Adaption of the system toward a multielectrode array will bring better temporal resolution in the future.

of highly defined neuron networks. The presented technique is a powerful tool for future applications in neurosciences, in particular for the assembly of neuron networks of high complexity in a straightforward process.

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ASSOCIATED CONTENT

S Supporting Information *

Experimental details; Figures SI1−SI3. This material is available free of charge via the Internet at http://pubs.acs.org.

CONCLUSIONS We defined a method to locally replace an antifouling PLL-gPEG with cell-adhesive PLL polymer by releasing the PLL from the aperture of a FluidFM cantilever. We used it to fabricate cell-adhesive spots into a nonadhesive background to pattern hippocampal neurons in a desired topology. Furthermore, the FluidFM technique was used to write PLL lines in situ between the neuron clusters. It was shown that the neurons react to these guiding lines by the outgrowth of axons. One week after the in situ polymer deposition the activity of the neurons was monitored with a calcium sensitive dye. The signal propagating between connected clusters was assessed. The size, number, and position of the neuron clusters can easily be adapted, and the resulting differences in signal propagation can be analyzed. This will provide further insights into the adaptation processes

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected] (J.V.). Author Contributions

H.D. and R.R.G. equally contributed to this work. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by the Swiss Nano-Tera initiative, by EuroBIOSAS ICS Initiative, the Swiss National Science Foundation, and the 3DNeuroN project in the European 7044

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Union’s Seventh Framework Programme, Future and Emerging Technologies, grant agreement no. 296590. We are grateful to Stephen Wheeler and Martin Lanz (LBB) for their technical support. Special thanks also goes to Prof. Dr. Jean-Marc Fritschy and especially Giovanna Bosshard from Institute of Pharmacology and Toxicology - Morphological and Behavioral Neuroscience at the University of Zurich for kindly providing us with dissected rat hippocampi and sharing their knowledge of primary neuron cultures.

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