glucose deprivation in hiPS-CMs

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Live cell imaging in microfluidic device proves resistance to oxygen/glucose deprivation in hiPS-CMs Sebastian Martewicz, Giulia Gabrel, Marika Campesan, Marcella Canton, Fabio Di Lisa, and Nicola Elvassore Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b05347 • Publication Date (Web): 29 Mar 2018 Downloaded from http://pubs.acs.org on March 29, 2018

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Analytical Chemistry

Live cell imaging in microfluidic device proves resistance to oxygen/glucose deprivation in hiPS-CMs Sebastian Martewicz1,2,3, Giulia Gabrel2, Marika Campesan4, Marcella Canton4, Fabio Di Lisa4, Nicola Elvassore1,2,3,5*. 1

Shanghai Institute for Advanced Immunochemical Studies (SIAIS), ShanghaiTech University, Shanghai, China. Department of Industrial Engineering, University of Padova, via Marzolo 9, 35131 Padova, Italy. 3Venetian Institute of Molecular Medicine, via Orus 2, 35129 Padova, Italy. 4Department of Biomedical Sciences, University of Padova, via Bassi 58/B, 35121 Padova, Italy. 5Stem Cells & Regenerative Medicine Section, UCL Great Ormond Street Institute of Child Health, 30 Guilford Street, London WC1N 1EH, UK. *Corresponding author at: [email protected], tel: +39 049 8275469 2

ABSTRACT: Analyses of cellular responses to fast oxygen dynamics are challenging and require ad hoc technological solutions, especially when decoupling from liquid media composition is required. In this work, we present a microfluidic device specifically designed for culture analyses with high resolution and magnification objectives, providing full optical access to the cell culture chamber. This feature allows fluorescence-based assays, photo-activated surface chemistry and live cell imaging under tightly controlled pO2 environments. The device has a simple design, accommodates three independent cell cultures and can be employed by users with basic cell culture training in studies requiring fast oxygen dynamics, defined media composition and in-line data acquisition with optical molecular probes. We apply this technology to produce an oxygen/glucose deprived (OGD) environment and analyze cell mortality in murine and human cardiac cultures. Neonatal rat ventricular cardiomyocytes show an OGD time-dependent sensitivity, resulting in a robust and reproducible 66±5% death rate after 3 hours of stress. Applying an equivalent stress to human induced pluripotent stem cell-derived cardiomyocytes (hiPS-CMs) provides direct experimental evidence for fetal-like OGDresistant phenotype. Investigation on the nature of such phenotype exposed large glycogen deposits. We propose a culture strategy aimed at depleting these intracellular energy stores and concurrently activate positive regulation of aerobic metabolic molecular markers. The observed process, however, is not sufficient to induce an OGD-sensitive phenotype in hiPS-CMs, highlighting defective development of mature aerobic metabolism in vitro.

Oxygen concentrations in biological samples are major determinants of cell behavior and fate. In vivo, cells experience different oxygen partial pressures (pO2) ranging between 1% and 13% pO21,2, thus making pO2 a de facto characterizing feature of each cellular niche. In vitro, cell cultures are continuously exposed to non-physiological hyperoxia3 but retain oxygen-level sensitivity. Cellular responses to fast and dynamic pO2 changes are of particular interest: in vivo such conditions are often associated with pathological events such as ischemia, and recreating these conditions in a controlled experimental setting allows valuable insight for improving therapeutic approaches4. Ischemia is a multifactorial condition, in which arrested blood supply generates changes in both cellular liquid and gaseous environments, with concentrations of glucose, pH, O2 and CO2 changing rapidly and dynamically5. The heart is an organ that deeply relies on blood supply for its survival and function, and possesses extremely limited energy-stores in physiological conditions6, rapidly depleting its glycogen stores upon birth when it switches from the prenatal hypoxic environment to the oxygen-rich newborn one7. While retaining some substrate adaptability8, adult cardiomyocytes generate energy mainly by mitochondrial oxidation of fattyacids9 and acute ischemic events produce severe repercussions on cardiac tissue function and survival10. Observing real-time cellular responses to such rapid variations in environmental

conditions and understanding the relative contributions of single factors of ischemia can provide insight into cardiacprotective therapeutic solutions. Taken together, these considerations urge for the development of experimental setups aimed not only at precise control over O2 and nutrients independently, but also allowing easy coupling with unspecialized analytical tools with unrestricted range for optical detail. Animal cardiac cells are widely employed as in vitro models, but not always represent to full extent human physiology11. The advent of human pluripotent stem cells (hPSCs) and their differentiation into cardiac lineage provides a good cellular substrate to conduct studies in a human physiological background12. Nevertheless, cardiomyocytes derived from hPSCs (hPSC-CMs) display immature fetal-like phenotypes, which extend to their metabolic profile and energy substrate preference13. While often reported in comparison with their fetal counterparts14, the hypoxia-response of hPSC-CMs has never been experimentally assessed. In particular, despite one recent study reporting the effect of maintaining hPSC-CMs in a hypoxic chamber for days15, there are no available data describing live changes in cell behavior during fast pO2 variations. Oxygen control over cell cultures has been traditionally performed by regulating macroenvironments in airtight incubators, hypoxic workstation or flow-chambers1, providing an

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indirect and imprecise control over cellular microenvironments, regulated by slow diffusive processes5,16. In these setups, administration of fast and sharp stimuli requires cumbersome equipment and hours-long pre-equilibration periods17. The extreme reduction of handled liquid volumes and operation in laminar flow-regimes allow microfluidic-based technologies to easily overcome these major limitations18. In these settings, pO2–control methods range from oxygen-scavenging substrates19-22 to physical isolation of oxygen-consuming cultures with gas-impermeable materials23,24. Most commonly, microfluidic devices take advantage of the high permeability to gasses of polydimethylsiloxane (PDMS) to promote oxygen exchange between the culture chamber and either liquid-25,26 or gas-perfused27-31 microchannels, which geometrical complexity can generate spatially homogeneous or gradient-like O2 concentrations at cell culture level22,26,32-35. While these technological solutions provide robust control over cellular microenvironments and are in principle suitable for coupling with optical imaging setups, there are few examples of devices specifically designed for in-line data acquisition during hypoxic stimuli. In most cases, coupling with a microscope is limited to cell culture monitoring or cell tracking27,35-42. Live imaging of biological responses to hypoxia in terms of intracellular Ca2+ dynamics and mitochondrial membrane polarization has been reported in large organoid structures (pancreatic islets) in microfluidic devices characterized by a gas-controlled bottom layer, which does not allow more detailed observations43,44. To our knowledge, at the moment of the preparation of this manuscript, a part from our previous setup45 there are only two other reports of live imaging assessing hypoxia-responses at cellular level for an ongoing stimulus46,47. Khanal et al. take advantage of microfluidic to reduce media volumes and feed the cultures, however they still rely on slow gas diffusion in a hypoxic chamber for pO2 control47. In a recent paper, Byrne et al. show changes in mitochondrial redox state upon delivery of a hypoxic stimulus employing an open-well, gas-controlled bottom layer microfluidic setup46. While open-well systems can robustly generate pO2-controlled environments, they are (by design) impossible to completely isolate from external conditions and the positioning of the oxygen-control layer underneath the cell culture can hinder imaging at higher magnifications than low NA air objectives. Our previous bimodular system45 required steady and continuous flow of medium to maintain hypoxic conditions, preventing long-term cultures48 and exposing the cells to constant and potentially detrimental shear stress. Here, we present a microfluidic device specifically designed for single-cell live imaging at high resolution and magnification with confocal-grade optics. The system is based on a noflow, periodic-perfusion strategy allowing long-term culture and reducing shear stress to minimum levels during rapid media changes. The device can be lodged easily in a multiwell plate and allows even untrained users straightforward coupling with epifluorescence or confocal imaging systems. The equilibration kinetics for %pO2 at cell culture-level are in the order of tens of seconds until reaching equilibrium for any desired O2 concentration. We provide proof-of-concept for full optical accessibility to the cell culture chamber, opening the device to any kind of histochemical or immunofluorescent assays, surface photochemistry and direct cell live imaging. The device allows separate control over gaseous and liquid components of cell cultures. We employ it to assess the response to oxygen-, glucose- and oxygen/glucose-deprivation (OGD) in murine

cardiac cultures, providing a very precise time window to study cardiomyocyte viability in ischemic-like conditions. Finally, comparison between murine primary cultures and hiPS-CMs subjected to an equivalent OGD stress experimentally proves their hypoxia-resistant fetal-like phenotype. We show how this hiPS-CM feature is independent of their abnormal glycogen stores, as glycogen depletion and concurrent positive regulation of fatty acid metabolism markers by culturing cells in a “cardiac medium” does not promote a hypoxiasensitive phenotype.

EXPERIMENTAL SECTION Device fabrication. The microfluidic device was made by standard soft-lithography technique49 on the design reported in Fig. S1A&B. Further details on the manufacturing process are included in the Supporting Information section. Oxygen measurement. The oxygen measurements were carried out as previously described45. A detailed description of the procedure is available in the Supporting Information section. Cell Cultures. Human fibroblasts: HFF-1 cell line was purchased from ATCC and cultured according to suppliers’ instructions. hiPSCs: A previously reported hiPS cell line generated by mmRNA protocol was used50. hiPS-CMs: Cardiac differentiation was performed according to protocols described by Lian et al.51 and Burridge et al.52 and schematically shown in Fig. S5A&B. Glucose-reduced media were obtained by mixing at proper ratios B27-supplemented RPMI w/ or w/o glucose (ThermoFisher Scientific, USA). NRVMs: Neonatal rat ventricular myocytes were obtained from Sprague-Dawley 2–3 days old pups according to a published protocol53. Detailed information about subculturing, differentiation, dissociation of hiPS-CMs and isolation of NRVMs are available in the Supporting Information section. Acrylamide photopatterning. The glass surface of the culture channel was functionalized with 1% 3-(trimethoxysilyl)propyl methacrylate in 94% ethanol and 5% glacial acetic acid for 4 minutes. After extensive washing with ethanol and milliQ water the channel was airdried and filled with prepolymer solution: 8% monomeric acrylamide in milliQ water and 50 mM HEPES pH 8. Prior to injection the solution was completed 1:10 with photo-initiator IRGACURE 2959 (Ciba) prediluted at 350 mg/ml in methanol. Photopolymerization was achieved by exposure to 365 nm UV light source (DYMAX Corporation, USA) through a highdefinition acetate printed mask with the desired geometry for 2–5 min. The excess of not polymerized solution was washed away, the surface was sterilized by ethanol washing and coated with MRGF. All chemicals were purchased from SigmaAldrich, USA if not otherwise stated. Immunofluorescence. The list of antibodies employed in this study, inclusive of fixation and incubation information is available in the Supporting Information section. Live imaging. Confocal calcium and membrane potential measurements were performed as reported previously54. VF2.1Cl voltage sensitive dye was kindly provided by Prof. Roger Tsien55. pHyPer-cyto was purchased from Evrogen, Russia. All acquisition parameters along with a modified version of Lipofectamine 2000 transfection protocol (ThermoFisher Scientific, USA) and the composition of the recording saline solution are described in the Supporting Information section. Live&Dead Assay. Live&Dead assay was carried according to manufacturer’s instructions (cat#L3224, ThermoFisher Scientific, USA). Periodic-Schiff Acid staining. PAS staining was performed according to manufacturer’s instructions (cat#395B, Sigma-Aldrich, USA). Real-Time


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Analytical Chemistry PCR. RNA was isolated with TRIzol Reagent (ThermoFisher Scientific, USA) followed by purification with RNeasy Mini

Kit

(Qiagen,

USA).

RNA

was

retro-

Figure 1 Microfluidic device design and validation (A) Real-life picture of the device: cell culture channels are filled with red dye, gas chamber is filled with blue dye. (B) %pO2 plot show equilibration times for different oxygen levels. (C) A step-like variation in the %pO2 in the gas chamber, rapidly matched by dropping %pO2 in the cell culture chamber.

-transcribed with High Capacity cDNA Reverse Transcription Kit (ThermoFisher Scientific, USA). Primer sequences for are reported in Table S1 in the Supporting Information section. Western Blot analyses. Cells were lysed with sodium deoxycolate 5% (Sigma-Aldrich, USA) with complete protease/phosphatase inhibitor cocktail (Sigma-Aldrich, USA) and resolved in NuPAGE 4-12% Bis-Tris precast gels (ThermoFisher Scientific, USA). The list of antibodies employed is available in the Supporting Information section. Statistical analysis. Numerical data are presented as means ± standard error of means (SEM). Data pairs were compared by nondirectional Student’s t-test. All data computation was performed with Origin 8.1 software.

RESULTS AND DISCUSSION Microfluidic device design and validation. The device is designed to allow three parallel and independent cultures within the same microfluidic chip, completely isolated from atmospheric conditions by a common gas chamber directly above culture microchannels (Fig. S1A). The entire system fits on top of standard 25 mm microscopy coverslips and can be handled in sterile conditions within a 6-well multiwell plate (Fig. 1A). pO2-control requires only an additional pre-mixed gas tank at the desired pO2 to be plugged-in through a flowmeter into the inlet/outlet gas ports. We maintained the same parameters for gas-exchange surface as our previous design45, with a 50 µm thick PDMS-membrane separating medium and gas chamber (Fig. S1B) allowing very short O2 equilibration periods within the 375 nl of media volume in the cell culture chamber. Fig. S1C shows fluorescence at different %pO2 of the oxygen-sensitive Ru(ddp) dye inside the microfluidic device. The %pO2 in the culture medium matches the gas mixture flowing in the top chamber within 30 seconds (Fig. 1B). Once reached the pre-set value, %pO2 remains stable until further modification. Upon subjecting the device to a step-like change in oxygen levels from atmospheric conditions to complete anoxia, %pO2 at cell culture level reaches a steady-state of 0% pO2 within 60 seconds without reoxygenation until the flow of 95% N2, 5% CO2 is maintained (Fig. 1C). Dehydration of the cell culture chamber during prolonged experiments is avoided by bubbling gas mixtures through a previously oxygen-stripped milliQ water glass vessel28,56,57. Gas flow-rate was kept at 100 ml/min for all described experiments, but sta-

ble %pO2 conditions could be achieved for flow-rates as low as 35 ml/min (data not shown). In summary, we here validate a novel microfluidic device designed for long-term, no-flow cultures under user-defined %pO2. It allows tight and independent control over the liquid and gaseous components of cell cultures, with oxygen variation kinetics in the order of tens of seconds with stable steadystate until further user input. In-chip photopatterning and cell live imaging. We designed the microfluidic device to be fully optically accessible despite the isolation from atmospheric conditions of the cell cultured microchambers. To validate the cellular viability inside the device, we cultured several cell types in the microchambers, such as neonatal rat ventricular myocytes (NRVMs) shown in Fig. 2A. NRVMs were maintained for up to 7 days in the device in normoxic conditions on a periodic-perfusion regime of two complete medium changes every 12 hours. The cultures displayed healthy morphology and sustained contractile activity throughout the whole duration of culture, both when seeded as compact monolayer (Movie S1) or sparse single-cell distribution (Movie S2). Immunofluorescence assays can be easily performed and the design allows for sample analysis with a broad range of detail, up to high NA oil immersion objectives (Fig. 2A inset) and confocal-grade optics. Cellular niches are characterized by both physical properties of the microenvironment and soluble cues58, and geometrical constrains in culture have been widely reported to affect cell behaviour59. Being able to couple surface-modifying techniques with oxygen control could be instrumental in providing more detailed insight in biological processes in which both features have been shown to play pivotal roles, as in earlyembryonic zonation60,61 or cardiac-chamber in vitro selforganization59,62. The optical accessibility of the microfluidic device allows employment of photo-activated surface chemistry, as proven by the human fibroblast culture (HFF-1) seeded inside a microchannel patterned for a linear geometry (Fig. 2B). The HFF-1 cells are precluded adhesion to glass surfaces coated by photopolymerized linear acrylamide, and this capability of surface modification can be extended to other 2D geometries or even more complex 2-photon generated 3D structures, as we recently reported63. When challenged with an anoxic stimulus for 1 hour, HFF-1 displayed a clear and clas-



sical hypoxic-response, by rapidly accumulating and translocating to the nucleus the hypoxia inducible factor 1α (HIF1α) (Fig. S2A&B). HIF1α is a master regulator and one of the best-characterized elements of the early hypoxic response30, thus providing proof of a correctly functioning device from a biological standpoint.

Figure 2 Optical accessibility of the microfluidic device. (A) NRVMs cultured in the microfluidic channel and stained for cTnT by immunofluorescence (bar = 500 µm, inset represents a field of

the same sample acquired with oil-immersion 63x confocal-grade objective, bar = 25 µm). (B) HFF-1 cultured in the microfluidic channel after linear photopatterning of the culture surface (bar = 500 µm, bar inset = 50 µm). (C) NRVMs transfected inside the microfluidic channel with a ROS-sensitive green-fluorescent HyPer probe (bar = 100 µm). (D) Live imaging of HyPer fluorescent response to H2O2 stimuli in the microfluidic device (fluorecence a.u.).

Direct analyses of cellular responses to ongoing stimuli provide detailed insight in the molecular mechanisms involved in cellular processes64. The molecular tools employed in such studies vary from loaded cellular dyes to genetically encoded probes, which require optimized delivery in microfluidic settings. We optimized a transfection protocol for NRVMs with liposomes carrying the reactive oxygen species (ROS)sensitive probe HyPer (Fig. 2C). Intracellular ROS generated during ischemic events in vivo are thought to be the main determinant of acute cardiomyocytes death65 responsible for cardiac dysfunction in later stages of the pathology. As proof of concept for the feasibility of in-line acquisition of intracellular ROS in our device, we challenged transfected NRVMs with 1 mM H2O2. In a series of experiments, we transiently perfused beating cardiomyocytes with puffs of 1 mM H2O2, acquiring fluorescence intensity data (Fig. 2D). The cells showed a corresponding rapid and transient increase in fluorescence upon oxidative stress, displaying sensitivity to multiple stimuli, and proving the viability of our experimental setting for live imaging. These proof-of-concept examples of applications for our microfluidic device show its complete optical accessibility, both for cell culture analyses (either on fixed samples or by live cell imaging concurrent with microenvironment modification) and for culture substrate modification. This feature makes the device suitable for a broad range of studies, in which precise control over the gaseous environment is desirable. OGD results in NRVMs death in a time-dependant manner. Next, we decided to test the microfluidic device inducing an ischemic-like stress to NRVM cultures, with a fastgenerated and stably-maintained oxygen-glucose deprived (OGD) environment. Ischemia occurs when blood flow is blocked in parts of an organ, cutting oxygen and nutrient supply and allowing toxic waste to build-up. Traditionally, OGD experiments are performed placing culture vessels in anaerobic chambers or hypoxic incubators66,67 with little control over actual oxygen levels and slow dynamics. Cellular responses to OD and OGD are extremely fast and targeted experiments, although not impossible to perform, require cumbersome equipment with hours-long equilibration times17 or specialized instruments without in-line monitoring capabilities68. In our experimental setting, oxygen and glucose variations can be generated by a rapid medium change to serum- and glucosefree conditions, while stripping oxygen by maintaining a steady flow of 95% N2, 5% CO2 in the gas chamber. The design of the device offers independent control over the two parameters, thus allowing studies decoupling the effects of any combination of gaseous and soluble factors. We evaluated cell mortality in actively beating NRVMs by means of fluorescent dye intake assay (Live&Dead, calcein/di-ethidium bromide assay). Similarly to other reports17, depriving cells just of one between glucose (Fig. S3A) or oxygen (Fig. S3B) did not affect significantly cell viability in the time span taken into account (Fig. 3B). In vivo, the duration of an ischemic event


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Analytical Chemistry plays a pivotal role in the severity of the ischemic insult10. In our experimental setup, time extent of OGD proved to be similarly important, as exposure to OGD for 1 hour and 2 hours did not produce significant cell death (Fig. 3A&B). Cell-death magnitude increased dramatically after 3 hours of OGD with massive di-ethidium bromide staining marking nuclei of dead

cells (Fig. 3A). Adverse effects on cell viability due to gasflow in the top chamber can be excluded, as control NRVMs subjected only to glucose-deprivation (GD) were still maintained under steady flow of 20% O2, 75% N2 and 5% CO2 (Fig. S3A).

Figure 3 NRVMs are OGD-sensitive in a time dependent manner. (A) NRVMs exposed to oxygen/glucose-deprivation under 95% N2 flow (bar = 100 µm). (B) Quantification of cell viability by calcein/di-ethidium bromide staining in different experimental conditions.

In both cases, after recovering basal culture medium and atmospheric O2 levels, surviving cardiomyocytes kept displaying contractile activity (Movies S3&S4). Repeated experiments proved robustness of cell mortality magnitude after 3 hours of OGD producing a 66±5% mortality (Fig. 3B). During an ischemic event, reperfusion and the associated ROS production are mostly cited as the major cause of cell mortality65. Nevertheless, in our experimental setup, time extent of OGD alone was responsible for cell mortality, as shown by in-line cell morphology monitoring during a continuous OGD without reoxygenation or medium replacement (Fig. S3C). 2 hour-long OGD seemed to represent a boundary condition for NRVMs, as cellular morphology started to appear abnormal but no diethidium bromide staining was apparent (Fig. S3D). At this point, perfusion with complete and oxygenated medium allowed recovery of cellular activity and did not produce significant cell mortality (Fig. 3A&B). This result correlates perfectly with previous reports on the timing of apoptosis activation in NRVMs exposed to oxygen- and serum-deprivation69, and highlights the ability of a microfluidic system to define a more accurate timeframe for the cellular response than one only estimated in a diffusion-based hypoxic chamber. Wholemicrochannel imaging revealed the remarkable spatial definition of the anoxic condition and proved the actual isolation of the cell culture chamber from external environment: the boundary of the top gas chamber above the culture microchannel almost perfectly demarks the spatial distribution of dead cardiomyocytes (Fig. S3E). More importantly, Fig. S3E (insets) shows that the device offers a valuable additional internal control on the culture conditions between oxygendeprived and normoxic culture areas inside the very same microfluidic channel, thus distinguishing between oxygenation effects and external- or medium change-related effects In summary, we delivered a fast ischemic-like OGD stress to neonatal rat ventricular myocytes. By live monitoring the cells during the stimulus, we were able to define a precise timeframe for the onset of cell mortality, with a boundary condition for cell survival after 2 hours of OGD. The spatial distribution of dead cardiomyocytes after 3 hours of OGD provides an excellent proof of precision for delivered O2 drop,

setting a viable internal control for the cell cultures protected by the PDMS layer. hiPS-CMs are OGD-resistant. The optimized OGD protocol proved robust in affecting viability in bona fide neonatal cardiomyocytes, thus providing a tool for assessing oxygensensitivity of human cardiomyocytes differentiated from induced pluripotent stem cells (hiPS-CMs). These cells hold considerable potential in becoming an excellent in vitro cell model for cardiac diseases with a human genetic and physiological background70, but display an immature phenotype limiting their application14. Starting from hiPS cells reprogrammed with non-integrative mmRNA technology (Fig. S4A)50, we obtained hiPS-CMs expressing proper cardiacspecific molecular markers (Fig. S4B) and displaying cardiaclike functional features including contractility associated with membrane depolarization during action potential (Fig. S4C) and calcium transients (Fig. S4D). Nevertheless, when exposed to an equivalent OGD stress as the NRVMs, hiPS-CMs showed a remarkable viability even to an extended 4 hourlong OGD (Fig. 4A). While often inferred from their similarity to fetal cardiomyocytes14, to our knowledge hypoxiaresistance in human iPS-derived cardiomyocytes was experimentally observed only recently, proving to go beyond 24 hours in presence of media with complete nutrient supply, thus allowing glycolytic energy production15. We here show experimental evidence for prolonged OGD-resistance in hiPS-CMs, marked by high resilience and sustained contractile activity even under harsh conditions where hiPS-CMs were devoided of any external nutrient supply (3 hours OGD in PBS with Ca2+ and Mg2+) (data not shown). This apparent energetic selfsufficiency prompted us to investigate the presence of intracellular energy-reserves that could support the high energydemanding contractile activity. hiPS-CMs differentiated in well were tested for glycogen presence by periodic-acid Schiff (PAS) staining, and proved positive, identifying specifically only the beating cTnT+ areas of differentiated monolayers (Fig. 4B). After enzymatic digestion and single-cell replating, a more precise picture of the magnitude of these intracellular stores was apparent, with PAS staining marking a significant



portion of intracellular hiPS-CM-volume (Fig. 4C) and precisely identifying cardiomyocytes71. In summary, we show how hiPS-CMs display a highly OGD-resistant phenotype, strikingly differing from their neonatal murine counterparts. We provided evidence of prenatalor pathological-like glycogen intracellular stores, developmentally lost in more mature cells6,9,72. Glycogen depletion does not promote OGD-sensitivity in hiPS-CMs. Mature adult cardiac myocytes do not possess intracellular glycogen stores and positive PAS staining is usually associated with pathological and degenerative events6,72, in which adult CMs dedifferentiate reactivating fetal-like expression profiles6, matching the phenotypic description of hiPS-CMs. We sought to improve the features of hiPS-CMs in culture

Figure 4 hiPS-CMs are OGD-resistant and display extensive glycogen stores. (A) hiPS-CMs exposed to 3 and 4 hours of OGD in the microfluidic channel (bar = 100 µm). (B) hiPS-CMs differentiated in well assayed for cTnT expression by immunofluorescence and glycogen content by PAS staining (bar = 5 mm). (C) PAS staining showing glycogen stores in single-cell cultures of hiPS-CMs.

by optimizing a protocol aimed at depleting intracellular energy reserves. The basal cardiac differentiation medium is composed by RPMI supplemented with complete B27 nutrient mix, providing a rich environment for differentiating hiPS cells51.We asked if the rich B27-supplemented medium employed for differentiation could be accountable for glycogen accumulation during small-molecule cardiac differentiation of hiPSCs or it is an intrinsic feature of cardiac differentiation process. Applying a second differentiation protocol based on a supplement-free basal medium52 produced a similarly intense PAS staining in a cardiomyocyte-specific fashion (Fig. S5B), thus suggesting a B27-indipendent process. Reducing the concentration of only one component (glucose) of the B27supplemented medium proportionally decreased the intensity of PAS staining in hiPS-CM differentiated in well, reaching a stable level within 5 days after the medium switch (Fig. 5A). Nevertheless, glucose is an essential component of the medium, as prolonged complete withdrawal of this carbon source

resulted in cell death within the following days, even in presence of all the other B27 components (data not shown). In a manner similar to what performed during NRVMs isolation, when FBS is rapidly withdrawn from the culture medium after cell adhesion and recovery, we hypothesized that switching to a more essential medium after completing differentiation could be beneficial to hiPS-CM phenotype. Cultures maintained in a “cardiac medium” composed of serum-free αMEM supplemented with ITS and NEAA showed a significant reduction in PAS staining already within 5 days (Fig. 5B), without compromising cell viability for an extended culture up to 1 month. We then asked if culturing cells in the “cardiac medium” had any effects on gene expression in hiPS-CMs, by considering transcription and protein expression of metabolism-related markers. In general, we observed a trend towards a more metabolically aerobic expression profile, with transcriptional downregulation of glucose metabolism-related hexokinases (with a more marked reduction for the fetal isozyme HK-2) and an upregulation of fatty-acid metabolismrelated transcripts, such as mitochondrial very long-chain specific acyl-CoA dehydrogenase (ACADVL) and carnitinepalmitoyl transferase IB (CPT1b) (Fig. 5C). At protein level, transcriptional data for HK-2 were confirmed by drastic reduction of its protein product (Fig. 5D). Moreover, antagonist metabolic-regulators acetyl-CoA carboxylase beta (ACCβ) and malonyl-CoA decarboxylase (MCD) showed antithetic behaviours, with MCD transcripts being upregulated and ACCβ mRNA levels mostly unchanged (Fig. 5C), but showing marked increase in post-translational inhibitory phosphorylation (Fig. 5D. Nevertheless, despite intracellular glycogen depletion and the molecular hints for a starting aerobic metabolic activation, hiPS-CMs retained an OGD-resistant phenotype. Dissociated hiPS-CMs cultures were maintained in culture inside the microfluidic device for 7 days and fed with either RPMI + B27 or “cardiac medium”, but both conditions showed high cellular viability after 3 hours of OGD (Fig. 5E, suggesting a still immature phenotype for both conditions. In summary, monolayer small molecule-based cardiac differentiation protocols produce cardiomyocytes characterized by extensive glycogen stores, observed after PAS staining. The glycogen deposits represent considerable intracellular energy-stores and can be reduced by modulating glucose concentration in the culture medium. Switching culture medium to a more essential composition after completion of differentiation allows long-term culture, while depleting glycogen content and activating a more β-oxidation leaning expression pattern. Nevertheless, this process does not confer hiPS-CMs an OGD-sensitive phenotype.

CONCLUSIONS In this study, we developed a microfluidic device aimed at fast generation of stable environments at precisely defined O2 partial pressures, while maintaining independent control over cell medium composition. The device is aimed at non-expert users with basic cell culture training, and is based on a no-flow, periodic perfusion strategy that allows long-term cell cultures, including human fibroblasts, primary murine cardiomyocytes and human cardiomyocytes derived from induced pluripotent stem cells. The simple design allows easy sterile procedures in standard laboratory cultureware and provides full optical accessibility to the cell culture chamber. Proof-of-concept applications of this specific feature are shown by performing high


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Analytical Chemistry magnification immunofluorescent assays, surface photochemistry for cell culture geometrical patterning and singlecell live imaging with a fluorescent molecular probe for ROS detection, all in-chip. The experimental setting was employed to generate oxygen-, glucose- or oxygen/glucose-deprived environments for cardiac cultures, both murine and human. We show how deprivation of just one component is not sufficient to induce cell death in murine cardiac cultures, while complete OGD results in robust and reproducible mortality

rate in a time dependant fashion. Human cardiomyocytes derived from hiPS cells on the other hand, are resistant to equivalent stimuli. In this study, we provide experimental evidence of the hypoxia-resistant phenotype associated with hiPS-CMs and further describe fetal-like features like extensive intracellular glycogen stores. Aiming at glycogen depletion to deprive hiPS-CMs of OGD-protective energy reserves, we develop a culture protocol that concurrently positively regulates genes related to aerobic fatty-acid metabolism.

Figure 5 Glycogen depletion and metabolic-marker induction in hiPS-CMs. (A) Reducing glucose concentration after completion of differentiation proportionally decreases glycogen content in hiPS-CMs. (B) An essential “cardiac medium” induces a more robust glycogen depletion. (C) Real-time PCR analyses for metabolic-marker transcripts (n=2) (D) Western blot for metabolic regulators and relative quantification of band intensity (n=2). (E) hiPS-CMs cultured either in B27-supplemented medium (top) or essential “cardiac medium” (bottom) display OGD-resistance after 3 hours of stress.

ASSOCIATED CONTENT Supporting Information An extended version of the Experimental Section along with 5 supporting figures are available as Supporting Information (PDF file). We include Movies S1-S6 as .avi files in the online version of the manuscript.

AUTHOR INFORMATION

REFERENCES

Corresponding Author * Department of Industrial Engineering, University of Padova, Via Marzolo, 9. 35131 Padova, Padova, Italy. E-mail: [email protected], Tel.: +39 049 8275469

Author Contributions SM performed most of the experimental work; GG performed oxygen measures and HyPer experiments; MC, MC and FdL provided primary cardiomyocyte cultures and discussed the research activity; SM and NE designed the research and wrote the manuscript.

ACKNOWLEDGMENT

This research was supported by Progetti di Eccellenza CaRiPaRo (to N.E.), Oak Foundation Award (to N.E. Grant#W1095/OCAY14-191), Fondazione Città della Speranza (to S.M.) and TRANSAC Progetto Strategico Università di Padova (to N.E.). We thank Lia Prevedello for her assistance in lithography and microfabrication. We are grateful to Prof. Roger Tsien (Howard Hughes Medical Institute) for providing the membrane voltage sensitive dye.

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