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. 2025 Aug 26;5(5):636–646. doi: 10.1021/acsmeasuresciau.5c00028

Bubble Perfusion Brain Slice Culture with Single-Droplet Stimulus Delivery in a 3D Printed Microfluidic Device

Genoveve G Gutierrez †, Richard J Ortiz ‡, Victoria Norman §, Rebecca A Prosser §, Christopher A Baker †,*
PMCID: PMC12532049  PMID: 41113152

Abstract

Ex vivo tissue culture can model tissue physiology under well-controlled conditions and is especially promising for understanding the complex mechanisms of the brain. Three-dimensional (3D) printing has immense potential to accelerate microfluidic technology development, especially for ex vivo tissue culture devices where miniaturization is ultimately limited by the physical dimensions of tissue explants. Here we describe the development of a 3D printed microfluidic perfusion device for ex vivo brain slices that utilizes media droplets segmented by oxygen bubbles, a perfusion technique we call “bubble perfusion”. Device design considerations are described, including materials property challenges associated with 3D printed plastic, such as wetting behavior and thermal conductivity challenges. Integrating a heated water circulation chamber and media prewarming chambers yielded media droplets delivered to brain slice explants at a temperature of 36.8 ± 0.13 °C, with tissue experiencing a temperature drift of 0.5 ± 0.09 °C over the course of a 60 s media droplet exposure. Murine brain tissue explants containing the suprachiasmatic nucleus (SCN) or entorhinal cortex (EC) were observed to be viable within the perfusion system by fluorescence imaging of intracellular Ca2+ flux induced by single-droplet stimulus of 60 mM KCl. Robust Ca2+ flux was observed for perfusion experiments lasting up to 12 h, with sequential droplet observations indicating the temporal dynamics of Ca2+ responses. End-point propidium iodide staining was used to characterize the health of EC and SCN tissue, with ca. 60% of cells in both regions showing no sign of membrane damage after 12 h of perfusion. The utility of the perfusion system toward pharmacological studies was demonstrated by comparing the Ca2+ flux induced by stimulus with 50 μM cannabidiol (CBD) vs 50 μM anandamide (AEA). Interestingly, similar magnitude and temporal dynamics of Ca2+ flux were observed for both CBD and AEA stimuli despite differential proposed mechanisms of action with respect to the CB1 receptor. These studies demonstrate the utility of the 3D printed bubble perfusion system toward the study of receptor-binding ligands that induce relatively modest magnitudes of Ca2+ flux.

Keywords: microfluidic, 3D printing, ex vivo, brain slices, calcium imaging, cannabidiol, anandamide

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Introduction

Neurodegenerative diseases are characterized in part by dysfunction or loss of synapses and neurons, and formation of protein aggregates, which severely impact behavior and physiology. Alzheimer’s disease (AD) is the most common age-dependent neurodegenerative disease and is the most common cause of dementia in elderly people. An estimated 6.7 million Americans aged 65 and older are living with AD. By 2050, the number of people aged 65 or older living with AD is projected to grow to 12.7 million. Initial AD neuronal dysfunction and cell death occur in areas associated with memory and cognitive processing such as the entorhinal cortex (EC). AD patients commonly experience disruption in circadian rhythms, which influences various homeostatic functions, including disturbances with thermoregulation and sleep-awake cycles. − Additionally, circadian disruption becomes more severe as AD severity increases. ,

In animal models, accumulation of amyloid β (Aβ) interferes with the expression of circadian clock-related genes. − Specifically, Aβ accumulation has shown to alter expression patterns of the circadian clock genes Bmal1 and Per2. Therefore, age-related neuroendocrine degeneration and disruption of circadian rhythms have been proposed to have a consequential, bidirectional interaction , that require further examination.

Recent studies have shown that the G-protein coupled receptor cannabinoid receptor 1 (CB1) plays critical roles in various physiological and behavioral systems including memory formation, , nociception, and energy homeostasis. Centrally, CB1 is expressed in various areas of the brain, including the suprachiasmatic nucleus (SCN), which acts as the body’s central circadian timekeeper, and the EC, which is involved in time perception. , CB1 activation has been shown to have an excitatory effect on SCN neurons by reducing GABA release, affecting circadian light attenuation. Cannabidiol (CBD) is a phytogenic ligand of CB1 and an emerging pharmacotherapy for AD. CBD has been found to be neuroprotective, to prevent hippocampal and cortical degeneration, and to protect against Aβ neurotoxicity. ,

Modeling microphysiological environments with the precision flow control of microfluidics has provided new understanding of biochemical signaling processes and accelerated drug discovery. − Emerging microfluidic technologies are enabling various ex vivo organ-on-chip models, including brain, kidney, and heart tissue. Integrating microfluidic perfusion systems with ex vivo tissue slices enables precise delivery of nutrients and various chemical stimuli. Additionally, perfusate sampling facilitates the analysis of tissue-secreted factors such as hormones and neurotransmitters. , These techniques together are especially promising steps toward understanding complex signaling in neural tissue.

A key challenge for maintaining brain tissue samples ex vivo is meeting the high oxygen demand without intact vasculature. The rotary-tube culture method was an early approach to addressing this challenge, but while it provides adequate oxygen influx, the mechanics of rotary tube culture do not allow for real-time analysis by microscopy or electrophysiology. As such, rotary tube culture has largely been supplanted by membrane-interface culture methods, in which tissue slices are kept stationary on a porous membrane at a liquid–gas interface. Porous membranes are permeable to flow and thus present a difficulty for microfluidic flow control, and the challenge of miniaturizing ex vivo brain slice culture to create brain-on-chip models remains an active area of research.

Our laboratories previously developed a brain-on-chip microfluidic device that mimicked the alternating oxygen and media delivery of rotary tube culture by delivering segmented flow of media droplets and oxygen bubbles in a method we called bubble perfusion. The original bubble perfusion technology was limited by a need for specialized microfabrication, and by a continuous flow operation that increased total perfusion volumes despite the small volume of individual droplets. Here, we describe the development of a bubble perfusion system fabricated using accessible, consumer grade three-dimensional (3D) printing technology, and designed for droplet-on-demand fluid delivery that reduces overall perfusion volumes. Single-droplet stimulus delivery was demonstrated in various experiments delivering KCl, anandamide (AEA), or CBD. Tissue responses to stimuli were observed via fluorescence imaging of Ca2+ flux in either SCN or EC tissue, and tissue health was assessed via end-point propidium iodide (PI) staining. Robust Ca2+ flux was observed in SCN and EC tissue for the full duration of perfusion experiments lasting up to 12 h.

Materials and Methods

Design and Operation of the Perfusion System

Device architectures were designed in AutoCAD 2023 (Autodesk, San Francisco, CA). Devices were 3D printed using BioMed Clear resin on a Form 3B SLA printer (Formlabs, Somerville, MA). Cytocompatibility of the device was improved by a postprint procedure including rinsing in isopropyl alcohol, and postprint curing by baking and additional ultraviolet (UV) exposure. These procedures are known to improve cytocompatibility of structures printed in Formlabs BioMed resin. − Specifically, the printed device was sonicated for 20 min submerged in isopropyl alcohol, followed by flushing the interior of the device with several mL of fresh isopropyl alcohol. The rinsed devices were air-dried, then baked at 60 °C for 1 h before additional UV irradiation for 30 min.

The tissue chamber was sealed using a 25 × 20 mm2 coverslip cut from GelBond film (Lonza, Basel, Switzerland) adhered to the device using a double-sided adhesive gasket cut from 300LSE tape (3M, St. Paul, MN). The tissue chamber had a designed internal volume of 19 μL, with tissue slices occluding approximately 6 μL (ca. 3.5 mm × 3.5 mm × 0.5 mm) of the chamber volume. Droplets delivered to the tissue chamber had a volume of 30 μL, the volume reduction of which was limited by dead volume within the device and by the tissue chamber dimensions. Further optimization of these parameters may be warranted in future revisions of this system.

On-demand delivery of aqueous droplets and oxygen bubbles was achieved using a custom-built valving system (see Supporting Information, Figure S1). Briefly, 3D printed two-point pinch valves were supplied with pressurized oxygen and placed upstream of the perfusion device. For bubble delivery, the pinch valve was connected directly to the perfusion device. For droplet delivery, a sealed 50 mL centrifuge tube was placed in-line between the pinch valve and the perfusion device. Metered volume delivery was accomplished by entrapping a defined volume of pressurized oxygen between the dual pinch points of the valve, then releasing this entrapped pressurized volume to flow toward the perfusion device. The metered volume was defined by the over pressure of the entrapped volume relative to atmospheric pressure and the volume entrapped between the two pinch points of each valve.

Temperature Characterization

Temperature characterization evaluated three factors: prewarming chamber volume, droplet hold time, and circulating water temperature. Prewarming chamber volume was evaluated at 19.5, and 156 μL termed 1X, and 8X, respectively, for their relative volume to the tissue chamber. Droplet hold time was evaluated at 1 and 30 min. Circulating water temperature was evaluated at 37 and 45 °C. Experiments lasted approximately 2.5 h allowing for both 1 and 30 min droplet delivery times (n = 57 and n = 4, respectively). A miniature thermistor (TH10K, THORLABS, Newton, NJ) was sealed into the tissue chamber to record the response of temperature vs time.

Hydrogel Glue Preparation

Hydrogel glue was synthesized following a protocol developed by Balakrishnan and colleagues, and was used to immobilize tissue slices on the GelBond coverslips. The two-part glue was comprised of Chitosan hydrochloride (ChitHCl) and Dextran dialdehyde (DDA).

ChitHCl Preparation

5 g of high molecular weight chitosan was mixed into 60 mL of 12 M HCl and 40 mL of ethanol and allowed to stir at room temperature for 3 h. The ChitHCl salt was filtered and washed with a 3:1 mixture of acetone–water and dialyzed against ultrapure water until no precipitate appeared when 1 wt % silver nitrate was added to the dialysate dropwise. The ChitHCl solution was dried to a pellet by vacuum centrifuge and stored at −20 °C until needed.

DDA Preparation

Five g of dextran and 4.5 g of sodium metaperiodate were mixed with 100 mL of ultrapure water for 6 h while protected from light. Five mL of the solution was dialyzed against milli-Q water until no precipitate appeared when 1 wt % silver nitrate was added to the dialysate dropwise. The DDA solution was dried to a pellet by vacuum centrifuge and stored at −20 °C until needed.

Tissue Slice Preparation

Coronal brain slices of 400 μm (Baker Lab) or 500 μm (Prosser Lab) thickness containing the SCN and surrounding anterior hypothalamus or the EC were collected from adult male C57BL/6 mice (Prosser Lab) or 10–12 week-old female Swiss Webster mice (Baker Lab). All animals were housed in 12 h light: 12 h dark conditions with food and water provided ad libitum. Tissue slices were initially placed in an interface brain slice chamber with Hank’s Balanced Salt Solution (HBSS, pH:7.4, Baker Lab) or Earle’s Balanced Salt Solution (EBSS, pH:7.4, Prosser Lab) and aerated with 100% O2 (Baker Lab) or 95%O2/5%CO2 (Prosser Lab) for 30 min.

Prior to transferring to the perfusion device, tissue slices were incubated in balanced salt solution supplemented with 20 μM (Baker Lab) or 50 μM (Prosser Lab) Calbryte 520 AM (AAT Bioquest, Pleasanton, CA) and 1% pluronic acid, then protected from light for 45–60 min. The dye-loaded tissue was washed by incubation in fresh balanced salt solution prior to loading into the perfusion device.

Working solutions of 10 wt % DDA in phosphate buffered saline and 5 wt % of ChitHCl in ultrapure water were prepared. Four μL of each working solution were combined into a single droplet on the GelBond coverslip, and the tissue slice was immediately placed onto the glue droplet. The coverslip was sealed to the perfusion device to load the tissue into the tissue chamber.

Drug-Stimulated Ca2+ Response Measurements

Experiments evaluating SCN response to drug stimuli were conducted in the Baker Bioanalysis Lab. Alternating oxygen bubbles and aqueous droplets containing HBSS were delivered to the tissue chamber, with each bubble or droplet held stationary on the tissue for 60 s before being evacuated by the subsequent droplet or bubble. This pattern of media and oxygen delivery was maintained for the full duration of each perfusion experiment (8–12 h). Once per hour a single stimulus droplet was delivered which was comprised of HBSS supplemented with 60 mM KCl, 50 μM AEA, or 50 μM CBD.

Fluorescence image sequences were collected using an upright fluorescence microscope (BX41, Olympus, Tokyo, Japan) outfitted with a 488 nm laser excitation source (Sapphire-100, Coherent Corp., Saxonburg, PA) and sCMOS camera (Kiralux, Thorlabs, Newton, NJ). Image sequences were collected for 6 HBSS droplets prior to the stimulus droplet (“pre-stimulus”), the stimulus droplet, and 4 HBSS droplets following the stimulus droplet (“post-stimulus”). Images were collected with a field of view of several hundred μm2, giving physiological-level, but not cellular-level, spatial resolution. Quantitative processing of fluorescence images is illustrated in the Supporting Information (Figure S2). Briefly, fluorescence vs time profiles were extracted from image sequences using ImageJ to define a region of interest (ROI) corresponding to the tissue region of study, keeping only image data collected during the interval when an aqueous droplet filled the tissue chamber. Nonlinear background drift resulting from photobleaching and dye leakage was fit to a single-phase exponential decay, and the fluorescence vs time profile of the full droplet sequence was divided by the exponential decay fit to give the signal-to-background ratio profile for each droplet. − The mean profile for all prestimulus droplets was calculated to give the intradroplet baseline profile. Finally, the intradroplet baseline profile was subtracted from each droplet’s signal-to-background ratio profile to get the baseline-subtracted signal-to-background ratio profile of each droplet.

Comparison of EC and SCN Tissue

Comparisons of EC and SCN tissue were conducted by the Prosser Lab. Alternating gas bubbles and aqueous droplets were delivered consecutively to the tissue chamber using various droplet pause times but maintaining the 60 s pause time for oxygen bubbles. Preliminary experiments indicated that droplet pause times up to 8 min maintained robust Ca2+ responses for at least 12 h; thus, tissue comparison experiments used 8 min droplet pause times for droplets delivered between stimulus sequences.

Tissue slices were stimulated with 60 mM KCl at 1, 6, and 12 h of perfusion time. Each stimulus sequence included observations for 4 prestimulus, 1 stimulus, and 4 poststimulus droplets. During the stimulus sequence, droplet pause times were decreased to 1 min, and a series of time-lapsed images, one every 5 s, was taken of the tissue. Fluorescence image sequences were collected using an upright fluorescence microscope (BH-2, Olympus, Tokyo, Japan) outfitted with a 488 nm laser excitation source (GH04850B2G, Sharp Corp., Osaka, Japan) and sCMOS camera (FMA050, AmScope, Irvine, CA).

After 12 h of perfusion, slices were transferred to an interface brain slice chamber and perfused for 2 h with EBSS containing 4.6 μg mL–1 PI. Following PI loading, slices were fixed in 4% PFA for 10 min then transferred to 30% sucrose overnight at 4 °C. Slices were embedded in optimal cutting temperature (OTC) and stored at −80 °C. 25 μm sections were cut using a cryostat and mounted to gelatin subbed slides and stored at −80 °C. Slides were thawed at room temperature and washed three times for 5 min each with PBS. Coverslip media containing DAPI was added to the surface of slides before they were coverslipped and sealed with nail polish.

Fluorescence images were acquired via 10× objective using a Leica DM6000B microscope and analyzed using ImageJ software. One image was analyzed for each tissue slice, and a ROI was drawn to contain either the entire SCN or a portion of the EC within that section. The images were split into blue and red channels to isolate DAPI-stained cells and PI-stained cells, respectively. Within each ROI, the total number of cells were counted using the DAPI image, and the PI was assessed in the same ROI. The ImageJ thresholding feature was used to differentiate between high PI intensity cells corresponding to dead cells, moderate PI intensity indicating damaged cells, and cells with minimal PI staining, considered healthy cells. −

Results and Discussion

Device Design

The final design of the 3D printed bubble perfusion device is shown in Figure A. Transitioning from microfabrication in PDMS/glass to SLA 3D printing presented challenges related to materials properties. First, we anticipated a substantial reduction in thermal conductivity for SLA printed plastics as compared to the previously employed glass substrates. Whereas glass devices can rely on simple resistive stage heaters, the 3D printed perfusion system integrated a 10.6 mL water circulation chamber for on-device thermal management. This approach utilized the thermal mass of circulating water to achieve a stable temperature equilibrium within the culture volume. Temperature characterization using this strategy is discussed below.

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Perfusion Device Design. (A) A photograph illustrating the final perfusion device design including water circulation chamber­(i), internal structural supports (ii), media prewarming chambers (iii), tissue culture chamber (iv), finger tight threaded port for gas inlet (v), and barbed connectors for two media/stimulus inflows (vi), perfusate outflow (vii), and heated water circulation (viii). (B) Micrograph illustrating uncleared droplet solution remaining in the chamber for square (i) and 45 rotated (ii) tissue chambers. Minimal uncleared droplet solution was observed for the lemon geometry (iii), which mimicked the spontaneous wetting behavior observed in (ii). A CAD illustration of the device geometry and a and schematic diagram of the microscopy-integrated perfusion system are provided in the Supporting Information.

Another key challenge was the difference in surface energy between the 3D printed plastic substrate and the previously employed PDMS/glass materials, which ultimately influences wetting behavior within the tissue culture chamber. Complete clearing of the tissue chamber of each media droplet by the subsequent gas bubble is important to ensure removal of metabolic waste products, and to maintain temporal sampling fidelity in downstream droplet analyses. In our previous work, we achieved effective tissue chamber clearance by identifying several empirical “best practices” for tissue chamber design, but we were unable to identify a specific model to relate design parameters (such as geometry and material) to droplet clearance behavior. In the 3D printed device, the previously identified best practices did not translate to the plastic substrate material. Figure B shows the evolution of tissue chamber design, highlighting the retained volume of a previous droplet after a bubble had filled the chamber. The initial chamber design (Figure B,i) was identical in geometry to that employed in our previous work, however here we observed retention of a substantial fraction of the droplet volume. Rotating the chamber geometry by 45° (Figure B,ii) showed little improvement. The chamber geometry was then modified to mimic the inherent wetting behavior observed in the suboptimal geometries, resulting in what we called the lemon geometry (Figure B,iii), which showed effective droplet clearance. One drawback to this approach was the need to scale the lemon geometry up slightly to facilitate the dimensions of a typical tissue slice. This resulted in increased chamber volume, from 12 μL for the square geometries to 19.5 μL for the lemon geometry, although this volume does not consider the volume occluded by tissue slice samples housed within the chamber. Future work may address large chamber volumes with refinements to the tissue preparation procedure, but we anticipate that tissue dimensions will remain the limiting factor for volume reduction in this and similar perfusion systems for brain slice culture.

Temperature Characterization

Figure shows the temperature profiles of droplets delivered to the tissue chamber and held static within the chamber for 60 s, with circulating water heated to 37 °C. Our initial perfusion design delivered room temperature perfusion media, which warmed only as it transited through the delivery channels prior to entering the tissue chamber. In that design, media droplets largely came to temperature within the tissue chamber, showing a profile of steadily increasing temperature spanning 2.3 ± 0.21 °C (Figure A, green trace). To reduce temperature variation within the tissue chamber, media holding volumes, referred to as prewarming chambers, were added to the device prior to the tissue chamber. Prewarming chambers were evaluated at equal volume (Figure A, red trace, “1X”) and eight times the volume (Figure A, blue trace, “8X”) of the tissue culture chamber. The range of temperature variation was reduced with prewarming chambers to 0.4 ± 0.05 °C for the 1X prewarming chamber, and 0.7 ± 0.04 °C for the 8X prewarming chamber.

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Temperature control characterization. (A) Representative temperature vs time profiles measured within droplets delivered to the tissue culture chamber in device designs with no media prewarming chamber (green), and prewarming chambers with volumes of 1X (red) and 8X (blue) that of the tissue culture chamber. Profiles were measures with a circulating water temperature of 37 °C. (B) Mean temperature of the 60 s droplet temperature profile for 1X and 8X designs vs the temperature of water circulating through the integrated water circulation chamber. Profiles were measured for droplets delivered after 1 min holding time in the prewarming chamber (n = 57 for each condition), and after 30 min holding time (n = 4 for each condition). Boxes indicate the central 50th percentile of observed values, whiskers indicate the full range.

Importantly, circulating 37 °C did not provide the desired chamber temperature of 37 °C, and subsequent calibration (data not shown) revealed that a circulating water temperature of 45 °C was required to reach 37 °C within the tissue chamber. Operating with circulating water >37 °C presented the potential of overheating the media in prewarming chambers, which was of particular concern when delivering stimulus droplets after lengthy residence times in the prewarming chamber. Therefore, we evaluated the 60 s temperature profiles of droplets with 1 and 30 min residence times in the prewarming chambers (Figure B).

With circulating water heated to 37 °C, droplets with 1 min residence time in the prewarming chamber were measured as 31.3 ± 0.06 and 31.6 ± 0.08 °C within the tissue chamber, for 1X and 8X prewarming chambers respectively (n = 57). Droplets with 30 min prewarming residence times yielded 31.6 ± 0.08 °C, 0.10 °C for both prewarming chamber volumes (n = 4). With circulating water heated to 45 °C, 1 min prewarming residence times yielded 36.8 ± 0.13 and 37.3 ± 0.08 °C, for 1X and 8X designs respectively (n = 57). 30 min residence times yielded 36.9 ± 0.09 °C and 37.0 ± 0.13 °C for 1X and 8X designs respectively (n = 4).

Intradroplet temperature stability was characterized with 45 °C circulating water by observing the trend of temperature versus time for droplets held in the tissue chamber for 60 s. Temperature stability was summarized by the range of temperature drift over the course of 60 s (Figure ). With 1 min prewarming residence time, temperature ranges were 0.5 ± 0.09 and 1.5 ± 0.14 °C for 1X and 8X designs, respectively (n = 57). Temperature range for 30 min prewarming residence time were 0.7 ± 0.01 and 1.4 ± 0.13 °C for 1X and 8X designs, respectively (n = 4).

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Temperature stability characterization. (A) The magnitude of the temperature drift over 60s (“Temperature Range”) for droplets delivered to the tissue culture chamber from 1X and 8X prewarming chambers with circulating water set at 45 °C. Holding times in the prewarming chamber are indicated. n = 57 for conditions with 1 min holding time, n = 4 for conditions with 30 min holding time. Boxes indicate the central 50th percentile of observed values, whiskers indicate the full range. (B) Representative temperature vs time profiles for droplets delivered to the tissue culture chamber with a circulating water temperature of 45 °C.

With 45 °C circulating water, the 1X prewarming chamber design showed no statistical difference in either mean temperature or range of temperature drift between prewarming residence times of 1 or 30 min. This suggests that with the 1X prewarming chamber and water circulation at 45 °C, droplets delivered to the tissue chamber were at appropriate physiological temperature and were near thermal equilibrium. Thus, the risks of overheating or thermal shock to the tissue slices were low under these conditions. Importantly, this result illustrates that media droplets delivered with relatively short prewarming residence time (i.e., 1–2 min) and stimulus droplets with much longer prewarming residence times (i.e., 30–60 min) can be prewarmed in chambers of identical volume, simplifying device design and operation.

The 8X prewarming chamber design showed reduced temperature stability compared to the 1X design. This may result from increased interfacial area between the prewarming chamber and water circulation chamber, which would alter the thermal equilibrium, although further work would be needed for such a conclusion. Still, no benefit was evident to indicate the use of an 8X prewarming chamber design, and the 1X chambers were incorporated into the final device design. The final design of this perfusion device can be accessed via the NIH 3D print exchange under model number 3DPX-021779.

Evaluation of Tissue Viability

To evaluate the efficacy of the perfusion system for maintaining viable brain tissue slices, intracellular calcium changes were measured in response to a depolarizing chemical stimulus. Tissue slices containing the SCN were loaded with the fluorescent calcium indicator Calbryte 520 AM prior to 10–12-h perfusion experiments. Tissue was perfused with alternating HBSS droplets and oxygen bubbles. Depolarization was induced by delivering a single droplet containing HBSS supplemented with 60 mM KCl. Fluorescence intensity vs time was monitored within the SCN tissue for six droplets prior to KCl stimulation (Figure , Prestimulus), the single stimulus droplet (Figure , Stimulus), and four droplets after stimulation (Figure , Poststimulus). Fluorescence profiles observed after 30 min (Figure A) and 12 h (Figure B) of perfusion indicated that tissue remained responsive to the depolarizing stimulus for at least 12 h in the device. We have shown previously that robust KCl-induced intracellular Ca2+ increases are well-correlated with end-point PI staining to indicate a low level of cell death in the calcium responsive tissue.

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KCl-induced Ca2+ response in SCN tissue slices. (A) Baseline subtracted fluorescence of Calbryte 520-loaded SCN tissue vs time for 6 prestimulus droplets (HBSS), 1 stimulus droplet (HBSS + 60 mM KCl), and 4 poststimulus droplets (HBSS) delivered after 30 min of tissue perfusion. (B) The same experiment described for panel A, but for stimulus delivered after 12 h of perfusion. (C) Summary of KCl-induced Ca2+ response from 23 stimulus deliveries performed across 3 tissue samples, with each perfusion experiment lasting 10–12 h. Only the final prestimulus droplet is shown in the summary for figure clarity. Boxes indicate the central 50th percentile of observed values and whiskers indicate the full range. **** p < 0.0001.

Figure C shows the change in baseline subtracted fluorescence for the droplet sequence starting with the final prestimulus droplet for 6 stimulus experiments across 3 tissue preparations. Since droplets are delivered at regular intervals of once every 120 s, the sequential droplet comparison illustrates the temporal dynamics of intracellular Ca2+ levels in the SCN tissue slices, with a return to baseline [Ca2+] within 3–4 poststimulus droplets.

Comparison of EC and SCN Tissue Health

Tissue slices containing the SCN or EC were loaded with Calbryte 520 AM prior to 12-h perfusion experiments. Depolarization was induced by delivering a single droplet of EBSS supplemented with 60 mM KCl at 1, 6, and 12 h. Fluorescence intensity vs time was monitored for four prestimulus droplets, the single stimulus droplet, and four poststimulus droplets for 3 tissue preparations of each EC and SCN tissue (Figure A and B, respectively). Interestingly, although in all cases the KCl induced significant intracellular Ca2+ flux, differences in the magnitude and duration of the changes are apparent between the SCN and EC, and between the SCN tissue studied in the Prosser lab vs the Baker lab. The reasons for these differences are unclear, although between-lab differences may be influenced by differences in optical efficiency of the relative imaging systems, differences in perfusion media used, or biological differences between the tissue donor animals. Further investigation may be warranted. To corroborate evidence of tissue health observed by fluorescence Ca2+ imaging, end-point PI staining was performed following 12-h perfusion experiments. Slices were removed from the tissue chamber and perfused with PI to identify dead and damaged cells. After fixation, tissues were imaged, and PI-stained cells were counted in defined ROIs corresponding to the tissue regions of interest. Figure shows the quantitation of healthy, damaged, and dead cells for the EC and SCN regions. Importantly, end-point measurements cannot resolve tissue damage occurring in the tissue preparation (i.e., dissection and sectioning) from tissue damage occurring in the perfusion system. Thus, this data gives evidence that a nonzero fraction of undamaged cells persisted throughout the time course of these perfusion experiments, but it cannot attribute cause or mechanism to the observed cell damage.

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Comparison of Ca2+ response in EC and SCN tissue. (A) Summary of Ca2+ response in EC tissue induced by delivery of 60 mM KCl after 1 h (i), 6 h (ii), and 12 h (iii) of perfusion. Each data set pools observations from three tissue slices. (B) The analysis is identical to A, but summarizing Ca2+ response in SCN tissue. ***p < 0.001, ****p < 0.0001, ns = p > 0.05.

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End point health of EC and SCN tissue (A) Representative fluorescence images showing typical ROIs (shown in yellow) used for EC (i) and SCN (ii) analyses. Tissue samples were PI stained and PFA-fixed after 12-h perfusion experiments, then imaged by fluorescence microscopy with additional DAPI staining. (B) PI intensity analysis in conjunction with PI/DAPI ratio revealed the percentage of healthy, damaged, and dead cells in the tissue region of interest. Data represents pooled observations from 3 tissue samples for each region.

Observations of depolarization-induced Ca2+ flux in the SCN and EC provide three important insights. First, the ability of SCN and EC brain slice tissues to maintain robust depolarization-induced Ca2+ flux demonstrates that the 3D printed bubble perfusion device is suitable for maintaining viable brain slices from different brain regions for at least 12 h. The histological data using PI staining confirms the robust health of the tissues at the end of the 12-h perfusion experiments, with EC tissue and SCN tissue showing comparable distributions of healthy, damaged, and dead cells. Combining both assessments, we observed no indicators that 12 h represents a physiological limit, but rather a pragmatic limit of perfusion experiments that were not designed around unsupervised automation. Therefore, we anticipate that future work with this system will readily extend into assaying important physiological phenomena in ex vivo tissues, such as those investigating neural plasticity responses of the EC or circadian functions of the SCN on 12–24-h time scales. Second, the ability for on-demand delivery of individual stimulus droplets enabled a time-resolved picture of depolarization-induced changes in intracellular [Ca2+]. We anticipate this capability will be useful for evaluating other dynamic responses in brain tissues, such as time course profiles of short duration drug stimuli. Third, the extended droplet pause times utilized when comparing EC to SCN slices may offer a straightforward approach to enriching the concentration of cellular secretions in future applications focused on downstream analysis of droplet contents.

Drug Stimulus Studies

KCl-induced depolarization studies demonstrated the efficacy of the perfusion system for maintaining viable brain slices and established a method to analyze temporally resolved measures of stimulus-induced Ca2+ flux in SCN tissue. To demonstrate the utility of the perfusion and imaging system toward more selective mechanisms of chemical stimulus, such as via receptor-mediated cell activation, we applied the same methodology described for KCl stimulus to compare the Ca2+ response induced by CB1 receptor ligand CBD to that of the endogenous CB1 ligand AEA.

Figure shows the Ca2+ response of SCN tissue stimulated with 50 μM AEA or CBD. Each tissue slice preparation was stimulated with both AEA and CBD, at alternating 60 min intervals, to prevent aliasing slice-to-slice variability with real differences in stimulus response. For both CB1 ligands, measurable Ca2+ flux was observed in SCN tissue. A comparison of the sequential droplet profile of AEA and CBD-induced activation (Figure C) shows similar dynamics both in terms of the magnitude of stimulus-induced Ca2+ increase and the time-course of recovery.

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Comparison of Ca2+ response induced by AEA and CBD. Representative fluorescence vs time traces illustrate Ca2+ response in SCN tissue induced by delivery of (A) 50 μM AEA; or (B) 50 μM CBD. (C) Summary of AEA and CBD-stimulated Ca2+ responses overserved across 3 tissue perfusion experiments, with each tissue sample receiving both AEA and CBD stimuli. n = 9 for AEA stimulus; n = 8 for CBD stimulus.

The similarities in Ca2+ response between AEA and CBD is curious, since AEA is a CB1 receptor agonist while CBD has been shown to be a CB1 receptor antagonist. This may suggest that the dominant mechanism of action of CBD within SCN tissue is not via CB1 antagonism. We hypothesize that the observed CBD-induced Ca2+ flux may be via TRPV1 receptor activation, − but further work beyond the scope of this technology development report is needed to resolve differential mechanisms of CBD and AEA activation in the SCN.

As an initial step toward elucidating differential mechanisms of action, we evaluated the response of SCN tissue explants to stimulus by the selective TRPV1 inhibitor A784168 (50 nM) in the absence of AEA or CBD. Figure A shows an overall decrease in calcium flux with TRPV1 inhibition. This suggests that osmo- or mechanoreception generates background Ca2+ influx even in the absence of other chemical stimuli. Inhibiting such background effects may prove beneficial in future studies for characterizing low levels of cellular activation such as by AEA or CBD stimulation. In contrast, inhibition of TRPV3 by administration of 60 μM dyclonine hydrocholoride shows a marked increase in Ca2+ influx (Figure B). We hypothesize that this activation is via the known interaction between dyclonine and the transcription factor NRF2, although further work is needed to resolve specific mechanisms.

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Ca2+ response induced by TRPV1 and TRPV3 inhibition. Box plots summarizing the Calbryte 520 fluorescence response to stimulus with (A) 50 nM A784168, a selective TRPV1 inhibitor; and (B) 60 μM dyclonine, an inhibitor of TRPV3. Concentrations were selected to be twice the known IC50 for each drug. Boxes represent the summary of 4 replicate droplets in each condition, with one tissue explant evaluated for each drug.

Taken together, these drug stimulus studies highlight the complexity of evaluating cellular activation by Ca2+ fluorescent imaging in response to drug stimuli. This may motivate the use of alternative imaging modes in future studies, such as voltage sensitive dyes or genetically encoded fluorescent reporters. Still, these studies highlight the utility of the microscopy-integrated perfusion platform for characterizing dynamic responses in living tissue explants.

Conclusions

This work demonstrates the development of a perfusion system for ex vivo brain slice culture fabricated by widely accessible consumer-grade 3D printing, which enables well-controlled delivery of chemical stimuli and straightforward integration with fluorescence microscopy. Two challenges were addressed relating to fabrication by 3D printing. First, the wetting properties of 3D printed plastic were addressed by modifying device geometry to achieve effective clearance of droplets from the tissue chamber. Second, the reduced thermal conductivity of plastic as compared to conventional glass microfabrication substrates was addressed by integrating circulating heated water. Thermal characterization achieved appropriate temperature and temperature stability for mammalian tissue culture.

The perfusion system was demonstrated to maintain Ca2+ responsive brain tissue for at least 12 h of perfusion via KCl-stimulated depolarization and fluorescence imaging of Ca2+ responses. Two distinct brain regions, the SCN and EC, were studied in these experiments. Observing the trend of sequential droplets revealed time-resolved dynamics of these changes, demonstrating the suitability of the system for characterizing dynamic processes on minute time scales. Tissue health after 12 h was verified using PI histological analysis. Finally, system utility was demonstrated by observing drug-induced activation in SCN tissue. Specifically, we observed similar activation profiles for SCN stimulated by AEA and CBD, while observing overall reduced activation with selective TRPV1 inhibition, and increased activation with TRPV3 inhibition by dyclonine. Further work is needed to expand these studies toward elucidating specific mechanisms of CBD-induced activation in the SCN.

This work was motivated to advance our previous proof-of-principle bubble perfusion technology, with an emphasis on enabling wider access to the technology for researchers who may not have access or expertise in precision microfabrication. The 3D printed fabrication strategy enables broad access and adoption of this technology and facilitates further customization and scalability to meet specific research goals. Work remains to improve the performance, and to better understand biological implications, of this mode of tissue perfusion. For example, volume reduction is hindered by the dimensions of explanted tissue slices, which will introduce dilution that negatively impacts the analysis of secreted factors. In addition, the present work did not elucidate the mechanisms of observed cellular damage, nor quantify any potential for hypoxic conditions within the tissue. Further studies into mechanisms of cellular damage in bubble perfusion are warranted. Still, the bubble perfusion system has demonstrated utility, and offers a platform technology with value for applications in neuroscience, toxicology, pharmacology, and other areas that may benefit from organ-on-chip modeling or ex vivo tissue culture.

Supplementary Material

tg5c00028_si_001.pdf (428.2KB, pdf)

Acknowledgments

This work was supported by the National Institute of General Medical Sciences of the National Institutes of Health under award number R35GM138173 and by startup funds provided by New Mexico State University.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsmeasuresciau.5c00028.

  • Schematic diagrams illustrating the microfluidic valving system, device integration with the epifluorescence microscope, and the data processing workflow, and CAD illustrations of device geometry (PDF)

∥.

G.G.G. and R.J.O. contributed equally to this work. CRediT: Genoveve G Gutierrez data curation, formal analysis, investigation, visualization, writing - original draft; Richard Joaquin Ortiz data curation, formal analysis, investigation, methodology, supervision, visualization, writing - original draft; Victoria Norman data curation, formal analysis, investigation, visualization; Rebecca A Prosser methodology, project administration, resources, supervision, writing - review & editing; Christopher A. Baker conceptualization, data curation, funding acquisition, methodology, project administration, resources, supervision, visualization, writing - original draft, writing - review & editing.

The authors declare no competing financial interest.

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Supplementary Materials

tg5c00028_si_001.pdf (428.2KB, pdf)

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