Incubator-independent cell-culture perfusion platform for continuous long-term microelectrode array electrophysiology and time-lapse imaging

Most in vitro electrophysiology studies extract information and draw conclusions from representative, temporally limited snapshot experiments. This approach bears the risk of missing decisive moments that may make a difference in our understanding of physiological events. This feasibility study presents a simple benchtop cell-culture perfusion system adapted to commercial microelectrode arrays (MEAs), multichannel electrophysiology equipment and common inverted microscopy stages for simultaneous and uninterrupted extracellular electrophysiology and time-lapse imaging at ambient CO₂ levels. The concept relies on a transparent, replica-casted polydimethylsiloxane perfusion cap, gravity- or syringe-pump-driven perfusion and preconditioning of pH-buffered serum-free cell-culture medium to ambient CO₂ levels at physiological temperatures. The low-cost microfluidic in vitro enabling platform, which allows us to image cultures immediately after cell plating, is easy to reproduce and is adaptable to the geometries of different cell-culture containers. It permits the continuous and simultaneous multimodal long-term acquisition or manipulation of optical and electrophysiological parameter sets, thereby considerably widening the range of experimental possibilities. Two exemplary proof-of-concept long-term MEA studies on hippocampal networks illustrate system performance. Continuous extracellular recordings over a period of up to 70 days revealed details on both sudden and gradual neural activity changes in maturing cell ensembles with large intra-day fluctuations. Correlated time-lapse imaging unveiled rather static macroscopic network architectures with previously unreported local morphological oscillations on the timescale of minutes.

1. Introduction

Unlike cells within living organisms, cells in vitro lack a supporting body infrastructure. They are not protected by the immune system or other basic system-wide regulatory mechanisms that control the temperature, pH and the turnover of nutrients, metabolites and signalling factors. Nevertheless, they are considered meaningful model systems for investigating functional physiological subsets or for testing in vivo-like constructs [1–5]. However, the cell-culture infrastructure (e.g. incubator, sterile hood) required to keep cultures alive is largely incompatible with most screening tools (e.g. microscopy, electrophysiology) and thus limits experimental possibilities. For this reason, most non-sacrificial cell-culture experiments are performed at ambient conditions. To prevent drifts in temperature, pH and osmolality, they rely on taking exemplary, quasi-static data snapshots over short periods only. This practice biases insights into physiological events and their temporal correlation. Typically, the data are collected at different basal activity levels. These independent and fragmented datasets may furthermore be distorted by both environmental and handling artefacts. For instance, manual medium exchange and variations in cell-culture handling during culture transfer from the humidified CO₂ incubator to the dry experimental set-up are acting as non-reproducible stimuli that modulate cell physiology. By contrast, an incubator-independent and automated perfusion system would stabilize physiological conditions on the experimental set-up and thus allow for the functional separation of environmental variability from physiological fluctuations. It would also extend the experimental time frame. In the best case, a culture system could reach, if not supersede, the natural lifetime of the donor animal that provided the tissue. Various custom-made [6,7] or commercial microscope incubators have been demonstrated to stabilize the pH, temperature and humidity (for a list see Nikon MicroscopyU). In general, they copy the elements and working principle of an incubator in a miniaturized, platform-adapted implementation. Therefore, they share some of the limitations of classical incubators. For instance, without further shielding the exposed electronic circuits or connectors against humidity, a highly humid atmosphere can lead to corrosion and functional failure of electronic data acquisition components. For imaging, optical windows need to be heated to prevent water condensation along temperature gradients. Furthermore, the majority lack an automated medium perfusion function. This shortcoming is addressed by a plethora of cell-culture perfusion and microchannel devices in different experimental contexts, as recently reviewed in general [8–14] and with a special focus on applications in neuroscience [15–17]. However, the majority still depends on classical incubation schemes to keep the cells viable, thereby limiting the experimental possibilities. To get around above described constraints, our two main goals were the decoupling of cell cultures from standard cell-culture infrastructure accompanied by the automation of cell culturing tasks. To this end, we devised a replica-casted perfusion cap and paired it with a simple multi-purpose benchtop perfusion scheme for timed or very slow-flow operation to reduce manually induced interference artefacts. In combination with a chemically buffered medium and a temperature controller, this platform allows for unsupervised multimodal long-term imaging and extracellular electrophysiology studies at ambient conditions immediately after cell seeding that are unbiased by physical and chemical handling artefacts.

2. Material and methods

2.1 Polydimethylsiloxane cap moulding master templates

Perfusion cap templates were designed (Alibre Design) to fit the standard outer diameter (OD) of 24 mm glass or polymer rings that are usually found on commercial microelectrode arrays (MEAs). The polymethyl methacrylate moulding template consisted of five slidable parts made for the arbitrary definition of vertical cap geometries (electronic supplementary material, figure S1). These were held in place by M4 polymer screws. Part 1 determined the OD of the polydimethylsiloxane (PDMS) lid; parts 2 and 5 determined its total height. Part 3 defined the inner cap diameter, which equalled the OD of the culturing dish; in this case, this was the OD of the Ø 24 mm glass ring. A smooth groove slightly below its upper edge added a sealing O-ring feature to the PDMS cap. Two holes (Ø 2.2 mm) centred at opposite sides of the upper edge of part 3 (or part 4, not shown) provided relative positioning and temporary fixation of the polytetrafluoroethylene (PTFE) inlet and outlet tubes. The shape of the inner cylinder (part 4) defined the slope of the cap ceiling. If the bottom of part 4 was levelled with those of parts 1–3, a casted cap would abut on the edge of the glass ring of the MEA. The thickness of the cap ceiling could reach several millimetres, depending on the bending radius of the somewhat stiff PTFE tubing. Part 5, the top cylinder, was affixed slightly above the tubing to fully embed the tubing in the PDMS ceiling. There was a vertical slit with dimensions of 10×2 mm on the top of part 1 to allow the passage of the PTFE tubing. Part 5 had a vertical groove to allow the escape of excess PDMS precursor mix or air when finalizing the assembly after filling the template. Depending on the resulting membrane thickness, each cap consumed 4.5 ml to 7 ml of the PDMS precursor mix.

2.2 Polydimethylsiloxane cap fabrication

PDMS properties and the cap fabrication procedure have been described in detail in a previous publication on caps without tubing [18]. We therefore summarize the main steps and only add detail for the tubing-specific differences. PDMS (Dow Corning Sylgard 184, 50 Shore A) prepolymer and the curing agent were thoroughly mixed with a metal spatula at a 10 : 1 ratio (v/v). The uniform distribution of trapped air bubbles indicated the sufficient homogeneity of the mixture, which was then degassed in a desiccator in which the vacuum was periodically broken to rupture any surfaced bubbles. PTFE tubing (OD 2.1 mm, inside diameter (ID) 1.5 mm, Supelco 20531) was reversibly clogged at one end with fishing line to prevent the entry of PDMS and was inserted into the guiding holes of part 3 (or part 4, not shown) of the moulding template (electronic supplementary material, figure S1). A small polymer half-sphere with a 2.5 mm hole was slipped onto the outlet tube to create a dome-shaped cavity in the resulting cap (figure 1, inset). To not obstruct the central optical path, both tubes were slightly bent towards the template wall and guided through the slit of part 5. The remaining slit opening was sealed with Parafilm to prevent PDMS outflow. The PDMS mixture was poured into the cavities of the assembled moulding template and cured within 24 h at room temperature, within 2 h at 60°C, or within half an hour at 85°C. After curing, the cap with its two PTFE tubes was released from the template at room temperature by pushing the slidable template components upward with the help of a rubber bottle stopper placed underneath and with 96% ethanol as a release agent. The tube endings were fitted with male Luer lock injection sites (WPI, 14034-40) with replaceable turnover flange stoppers (Carl Roth, EE00.1, Ø7.1 mm) via short pieces of Viton tubing (Cole-Parmer, 06435-01).

2.3 Preparation and plating of rat hippocampal cell suspensions

Where not stated otherwise, chemicals were bought from Life Technologies. Pregnant Sprague Dawley rats (CD IGS, Charles River) were anesthetized and sacrificed by cervical dislocation 18 days after conception. Following standard tissue dissociation protocols [19], their embryos (E17/E18) were harvested, put on ice in Hank's balanced salt solution (HBSS) and decapitated. After removing the meninges, the hippocampi were extracted, minced and transferred to fresh HBSS and dissociated into single cells using 0.25% (w/v) trypsin in HBSS buffer. After incubation for 10 min at 37°C, the trypsin was deactivated by 0.25 mg ml⁻¹(final concentration) soya bean trypsin inhibitor along with 0.01% (w/v) DNase (Sigma). Cell suspensions were prepared by sequential trituration (15–20 times) using three fire-polished Pasteur pipettes with decreasing diameters. Cells were then centrifuged at 200g for 5 min and the pellets were resuspended in Neurobasal medium (NBM) containing 2% B-27 serum-free supplement, 1 mM penicillin/streptomycin and 2 mM Glutamax. MEAs (30/200iR, Multi Channel Systems) carrying an OD 24 mm glass ring as a culture medium container were autoclaved and subsequently hydrophilized beforehand by a short O₂ plasma treatment (0.3 mbar, 1 min, 60 W, 2.45 GHz, Diener plasma GmbH) and thereafter were coated with a 10 μl drop of a poly-d-lysine (0.1 mg ml⁻¹) and laminin (5 μg ml⁻¹) mix in ultrapure sterile water. Drops were allowed to dry in the vacuum of the plasma chamber. Soluble coating components were thoroughly rinsed with ultrapure sterile water. The MEA was dried again in the vacuum of the plasma chamber before plating the cells at a final density of approximately 60 000 cells mm⁻². The cells were protected against medium evaporation by PDMS caps without perfusion functionality [18] and were allowed to settle for less than 10 min in a humidified (92–95% relative humidity (RH)) incubator at 37°C in a 5% CO₂ in air atmosphere before the pre-warmed cell-culture medium was added.

2.4 Culture and perfusion medium

Cultures were plated and grown in the above-mentioned NBM. For the benchtop experiments and control cultures, the medium contained a suitable pH buffer (l-histidine, Fluka 53370; 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), Sigma H0887). In the first experiment, the pH of the buffered media was adjusted to a physiological pH of 7.4 at room temperature without its preconditioning to ambient CO₂ levels (0.038%). To avoid pH drift in the second experiment, the medium was transferred to a glass beaker sealed by Parafilm and shaken for at least 4 h in a 37°C water bath to precondition the medium to ambient CO₂ at physiological temperature. Then, HEPES was mixed in at a final concentration of 10 mM. Owing to the temperature dependency of the pK_a of most buffers [20], the pH was titrated to physiological pH at 37°C with 1 M HCl (8 μl ml⁻¹) by comparing the colour of the medium (w/phenol red) with that of a control medium in a sealed flask at the same temperature. The osmolality was controlled with a vapour pressure osmometer (Vapro 5520, Wescor) before and after the pH adjustment and found to stay in a physiologically acceptable range between 220 and 260 mOsmol kg⁻¹. The pH-adjusted media stocks were sterile-filtered through 0.2 μm syringe filters and stored in a standard refrigerator in airtight containers (e.g. Falcon tubes, capped syringes) for later use. The buffers were found to stabilize the pH equally well at ambient CO₂ levels and in the control cultures that were kept in a humidified 5% CO₂ incubator. l-histidine was used for the entire perfusion period of the first hippocampal culture, HEPES for the second.

2.5 Culture maintenance

Control cultures and some of the perfusion cultures were kept in a humidified (92–95% RH) incubator at 37°C in a 5% CO₂ in air atmosphere prior to their insertion into the perfusion platform or their placement on the heating pad. Where necessary, non-buffered medium was replaced by one of the chemically buffered media before transfer. All MEAs that were stored in the incubator either carried a gas-permeable PDMS cap without tubing [18] or a perfusion cap. The caps were tightly seated around the MEA glass rings to prevent contamination and evaporation. Depending on the colour of the phenol red pH indicator, up to half of their medium (500 μl) was exchanged once a week, either by temporarily removing the caps without tubing or by manual syringe-driven perfusion through the tubing of perfusion caps.

2.6 Perfusion parameters

For MEAs with 7 mm high and ID 20 mm glass rings, the total cell-culture volume after cap placement varied for different inner cap geometries between 1.6 ml and 2.2 ml. Approximately, 88 μl had to be added for every 50 mm of the ID 1.5 mm PTFE tubing. Where necessary, the materials were handled and assembled on a sterile workbench to avoid contamination. For the long-term recording and imaging experiments, the tubeless PDMS cap was replaced by a PDMS perfusion cap. During cap placement and chamber filling, the Luer lock injection sites were temporarily removed from both tube ends to avoid pressure build-up and air bubble entrapment, which tended to slow down or entirely obstruct the fluid flow. Both the chamber and the tubing were filled through the inlet tube with buffered medium from a syringe before reattaching the outlet and then the inlet injection sites.

In the first perfusion experiment, a short (less than 10 mm) silver wire was inserted into the outlet septum holder to reduce the contamination risk should medium backflow from the non-sterile waste container (e.g. 15 ml Falcon tube). To prevent Ag-ion-related cytotoxicity [21], the wire was replaced with an ultraviolet A light emitting diode (UVA LED) (405 nm, 500 mcd, Conrad Electronics) in follow-up experiments [22]. This was attached by a short piece of thick tubing with a Ø 5 mm hole that was wrapped around the PTFE tubing behind the outlet septum (figure 3).

All perfusion cultures were kept at 35.5–36.5°C with an uncontrolled, but constant vertical T-gradient through the culture dish. They were either perfused automatically every eight hours (first hippocampal culture, 7 DIV+) or continuously (second hippocampal culture, 3 DIV+). With few exceptions, the exchanged volume of the above-mentioned chemically buffered media did not exceed 500 μl per day. If possible, the fluid level of the waste was kept level with the embedded end of the outlet tubing in the perfusion cap to avoid the build-up of negative pressure in the culture chamber, which could drain it in the case of any upstream leakage.

For the first experiment, the perfusion medium was stored at room temperature in a 100 ml glass bottle with a PDMS membrane (Ø 30 mm, 3 mm thick) in its cap to allow for the pH adjustment of the medium at ambient conditions without risking contamination. The flow pressure of the gravity-driven flow was adjusted by hanging the supply bottle approximately 5–20 cm above the MEA. The same type of PTFE tubing used for the perfusion caps was moulded into the bottle caps with PDMS membranes as supply lines. PTFE tubing also led from the perfusion cap outlet to the waste container (e.g. a Falcon tube). It was dipped into the waste medium to prevent clogging. Its position was slightly below the MEA surface to keep negative pressure on the outlet low.

In all cases, the PTFE tubes were interconnected with a combination of autoclavable Viton adaptor tubing (Cole-Parmer, 06435-01), polypropylene Luer fittings and syringe needles (OD 0.9 mm, G20) that pierced the cap silicone septa. This needle diameter allowed for sufficient flow while causing the least damage to the septa, thereby lowering the risk of leakage at the penetration site.

The first hippocampal culture was exposed to timed perfusion using a normally closed, computer-controlled (Velleman relay card K8056, Abacom Profilab) solenoid pinch valve (Model 360P071-21, NResearch Inc.). A short piece of soft silicone tubing was inserted into the PTFE tubing endings between the cap outlet and the waste. Placing the pinch valve at the outlet tubing (figure 2d) maintained positive pressure within the culture chamber. This helped in limiting the formation of bubbles and driving out any trapped bubbles at the risk of not being able to stop upstream medium leakage. Owing to such upstream leakage problems, the pinch valve was positioned between the medium supply and the cap inlet during the second half of the timed perfusion experiment (figure 2c). Volume replacement in the culture compartment was controlled by a stop-go flow. Flow rates were dictated by the opening time of the valve, the ID of the tubing and the relative height of the supply medium level with respect to the cell culture. They were empirically set to approximately 200 μl per opening of the pinch valve.

The constant perfusion experiment was based on a custom-made stepper motor-driven [23] 5 ml syringe pump. A micrometre screw gauge (MW-Import, 10-000-500-100-P) attached to a custom-made syringe holder for 0.5–5 ml syringes (inspired by World Precision Instruments, model MMP) was actuated by a stepper motor (1.8°/step). Stepping intervals were set by a timer circuit (based on National Semiconductor, LM555), which triggered the input of a stepper motor controller integrated circuit (National Semiconductor, SAA1042). Perfusion rates were set to less than or equal to 250 μl per day. An additional Luer T-piece was inserted directly behind the inlet septum holder as a bubble trap (figure 2a).

2.7 Multichannel microelectrode array electrophysiology and spike train analysis

Extracellular signals were recorded and processed by a commercial 60-channel, 1 Hz–3 kHz bandpass filter-amplifier data acquisition system (25 kHz sampling rate per channel) (Multi Channel Systems, MEA60-Up). The MEA socket in the base plate featured a resistive heating element and a Pt-100 temperature sensor. An external T-control unit (Multi Channel Systems, HC-1) kept the temperature of the socket surface at less than or equal to 36.5°C. The amplifier was mounted on a fixed, custom-made stage of an inverted microscope (Zeiss, Axiovert 200). This allowed the firm positioning of the amplifier with respect to the microscope optics to stabilize the region of interest (ROI) over time. To avoid the need to lift the cap during the MEA insertion, a custom-made Al spacer was placed between the base plate and the amplifier stage to create a gap for the free passage of the cap tubing (electronic supplementary material, figure S2). For the same reason, the lower right corner of each MEA was diagonally removed with a diamond pen. This strategy allowed the independent handling of the amplifier without lifting the cap or disconnecting the perfusion line. A grounded metal cap with central hole for illumination was placed onto the amplifier as a Faraday shield. Additionally, two grounded microclips (Conrad, 102407) were connected to both metal syringe needles at the inlet and outlet septa. A grounded wire mesh sock surrounding the tubing reduced the level of noise picked up by the fluid lines.