WAT’s up!? – Organ-on-a-chip integrating human mature white adipose tissues for mechanistic research and pharmaceutical applications

Obesity and its numerous adverse health consequences have taken on global, pandemic proportions. White adipose tissue (WAT) – a key contributor in many metabolic diseases – contributes about one fourth of a healthy human’s body mass. Despite its significance, many WAT-related pathophysiogical mechanisms in humans are still not understood, largely due to the reliance on non-human animal models. In recent years, Organ-on-a-chip (OoC) platforms have developed into promising alternatives for animal models; these systems integrate engineered human tissues into physiological microenvironment supplied by a vasculature-like microfluidic perfusion. Here, we report the development of a novel OoC that integrates functional mature human WAT. The WAT-on-a-chip is a multilayer device that features tissue chambers tailored specifically for the maintenance of 3D tissues based on human primary adipocytes, with supporting nourishment provided through perfused media channels. The platform’s capability to maintain long-term viability and functionality of WAT was confirmed by real-time monitoring of fatty acid uptake, by quantification of metabolite release into the effluent media as well as by an intact responsiveness to a therapeutic compound. The novel system provides a promising tool for wide-ranging applications in mechanistic research of WAT-related biology, in studying of pathophysiological mechanisms in obesity and diabetes, and in R&D of pharmaceutical industry.


Introduction
The global obesity pandemic poses one of today's biggest challenges to public health. Each year, 2.8 million people die from causes related to overweight or obesity. (1) Since the 1970s, the worldwide prevalence of obesity has nearly tripled, also leading to an upsurge in associated comorbidities such as type 2 diabetes (figure 1a).(2) Future projections reveal the gravity of this public health crisis: the prevalence rates of childhood obesity are increasing at an alarming pace (3), and by 2030, more than 50% of U.S. adults are predicted to be obese.(4) White adipose tissue (WAT) is the principal organ in obesity. In healthy human adults, WAT comprises approximately 20-25% of the total body mass, thus constituting the second largest organ, after the skin. In obese individuals, WAT's contribution to the total body mass may become as high as 50% (figure 1b). (5) WAT is tightly involved in the two most important functions of an organism -energy homeostasis and reproduction. (6) In energy homeostasis, not only does WAT act as the main storage site of excess dietary energy (figure 1c), it also performs crucial endocrine and metabolic functions (figure 1d). (7,8) WAT can sense the body's energy status, and respond appropriately, either by storing fuel, in the form of triacylglycerides, or by releasing it as glycerol and fatty acids, for ultimate delivery to organs in need. While this classically described role of WAT already entailed extensive crosstalk between WAT and other organs, its inter-organ communications extend beyond simple feedback loops activated by fed-or fasted states.
Endocrine functions of white adipocytes, and other WAT-resident cells in the stromal vascular fraction, are performed by the release of a variety of adipokines (adipose-associated cytokines) which affect the functioning of the brain, liver, pancreas and immune system.(9) Besides managing nutritional homeostasis, WAT contributes to the regulation of the hypothalamicpituitary-gonadal axis by secreting and metabolizing sex steroids.(7) Especially adipose tissue intracrinology, i.e., the modulation of sex steroid levels by significant numbers of locally expressed enzymes that activate, convert, or inactivate circulating steroid hormones, plays important roles in human reproductive function. (7,10,11) Given its prominent endocrine functions and extensive cross-talk with other organs, it comes as little surprise that abnormal amounts, or altered functioning of WAT may result in wide-ranging disorders, including hepatic and cardiovascular diseases, (12,13) diabetes and cancer. (14)(15)(16) In line with its important roles in metabolism, inflammation and cancer, WAT has emerged as a drug target with major therapeutic potential for a variety of diseases. (17)(18)(19) Additionally, storage metabolism by WAT can have a major impact on the efficacy of drug therapies, such as those for cardiovascular diseases and cancer. (20,21) For instance, adipocytes may metabolize and inactivate the chemotherapeutic, Daunorubicin, (21) which strongly affects the efficacy of this anticancer therapeutic. Furthermore, the capacity of WAT to sequester hydrophobic compounds, gives the tissue a prominent role in absorption, distribution, metabolism, and excretion (ADME) processes.
Even though WAT plays such a significant role in many diseases, surprisingly little is known about pathophysiological processes of WAT, especially when considering the ever-increasing prevalence of obesity and its co-morbidities. An important reason for this lacking insight is that mechanistic studies in humans often involve unacceptable health risks. Hence, most research depends on clinical observation, genome-wide association studies (GWAS) as well as animal studies, all with limited success. (22) Although animal models have led to many insights in obesity and diabetes, they often are lacking in predictive validity for human body functioning, first because of important species differences in nutrition and metabolism, and second because the distinct, unique physiological and pathophysiological roles of WAT in humans (23).
Alternatively, conventional, cell culture-based in vitro models have been widely utilized and are significantly less controversial ethically. Typically, researchers have used a variety of cell sources, ranging from (immortalized) murine, to primary human (pre-)adipocytes, each featuring distinct advantages and limitations. (24) In vitro differentiation of adipogenic progenitor cells, i.e., either pre-adipocytes or multipotent stem cell lines, or (induced) pluripotent stem cells, is often the method of choice due to the availability of donor-specific information, and the capacity of these cells to undergo expansion and cryopreservation while retaining their characteristic fat depot. Yet, the in vitro use of differentiated adipocytes has two major drawbacks: unlike their in situ matured congeners, (i) their lipid contents never reach a state of unilocularity, but remain distributed among multiple small lipid vacuoles, and (ii) their secretome differs immensely in the relative proportions of adipose-associated hormones (25,26). Both phenomena reflect a pre-mature state of the in vitro differentiated adipocytes, which strongly suggests that mature human adipocytes provide the best recapitulation of a mature human adipocyte physiology. Yet, the ex vivo culture of primary human adipocytes is extremely challenging, due to issues with the cells' buoyancy, fragility and de-differentiation, which so far have hindered development of robust protocols for longterm culturing. Additionally, inter-individual variability complicates the interpretation of study results based on samples from different donors.
Overall, it is of upmost importance to develop microscale platforms that provide microphysiological environments for the long-term culture of white adipocytes in structures that may recapitulate in vivo physiology, and functionality based on a minimal amount of cells. By combining modern techniques in microfabrication, biomaterials and tissue engineering, organon-a-chip technology has enabled the construction of promising platforms for mechanistic and pharmaceutical studies, that have great potential for disease modeling as well as the optimization of personalized medical treatments of obesity and diabetes. (27)(28)(29)(30) The integration of tissues with in vivo-like structure and functionality in perfused microenvironments is of particular interest when studying endocrine tissues and multi-factorial diseases, due to the possibility to combine individual chips into multi-organ systems. (31,32) However, although organ-on-a-chip research has burgeoned in recent years, and numerous platforms have been developed for many organs and tissues, WAT appears to have been largely overlooked, and only a few relevant efforts have been undertaken. (25) Several research groups injected preadipocytes from murine (33,34) or human (35,36) sources into microfluidic chambers, and were able to subsequently induce adipogenesis. Others directly introduced primary adipocytes from mice into mesoscopic, perfused reservoirs.(37) All of these innovative approaches, however, did not achieve the goal of generating human white adipose tissue structures with in vivo-like physiology.
Here, we present the first OoC platform that integrates human primary mature adipocytes into a perfused microfluidic chip. The human WAT-on-a-chip consists of multiple, tissue-specific chambers that are fluidically connected to a vasculature-like microchannel, while being shielded from the shear forces of the perfusion fluid. Using specifically tailored isolation and injection protocols, 3D microtissues based on freshly isolated adipocytes are generated inside a series of individual chambers. Thus, a large number of independent replicate cell cultures from individual donors can be produced and kept viable -as well as functional -for over one month. By analyzing media effluents and taking advantage of the optical accessibility of the tissue chambers, the patency of cells as well as key cell-physiological aspects, e.g., fatty acid metabolism and drug responsiveness, could be monitored successfully. Fig. 1 Relevance of research on WAT. a. The worldwide prevalence of obesity has nearly tripled since 1975; in 2016, almost 40% of adults aged 18 and over could be classified as overweight, 13% of them were even obese. This developments has coincided with a rising prevalence of diabetes, which in 2016 affected approximately 9% of adults worldwide. T2DM -the most prevalent form of diabetes -often develops during chronic positive energy balance, e.g., by a combination of excessive energy intake and physical inactivity.(1,2) b. WAT's contribution to body mass is 20-25% in healthy individuals, and up to 50% in the obese.(5) c. Traditional view of WAT as an organ primarily for energy storage. d. Additional, modern-day concepts of WAT functionality, including extensive endocrine functions.

Results and Discussion
Concept and characterization of the microfluidic platform Fig. 2 Concept of the WATmicrophysiological platform and microfluidic perfusion. a. Schematic of the chip platform featuring two independent systems with eight tissue chambers each. As shown in the cross-sectional zoom-in, tissue chambers, which feature tissue channel inlets at the bottom, are located right below the perfusable media channels, and separated from them by a microporous membrane. b. During flow, the chip's circuitry imposes a separation of the two transport processes, convection and diffusion. Convective flow is confined to the vasculature-like media channels, as confirmed by computational modeling (i). The tissue chambers are supplied by a diffusive transport as confirmed by observable diffusion of a fluorescent dye from the media channel into a hydrogel-filled tissue chamber (ii), as well as by computational modeling (iii).
In order to generate a microphysiological environment capable of integrating human WAT, and maintaining tissue viability and functionality during long-term culture, we developed a specifically tailored microfluidic platform featuring a footprint comparable to the size of a stamp (2 x 2 cm). The essential features of the platform, depicted in figure 2a, are 3D-tissue chambers and perfusable microchannels that are separated by an isoporous membrane. Each chip platform houses two identical, independent systems each featuring eight individual tissue chambers. All tissue chambers are located at the end of individual branches of a main channel, which can be accessed via a common inlet for cell injection. The tissue chambers are specifically designed to accommodate adipose tissue, particularly its large-sized, fragile and buoyant adipocytes: they are cylindrical structures with a diameter of 720 µm and a height of 200 µm. The tissue-chamber microstructures are encased by transparent glass (cover slip or microscope slide) at the bottom, enabling easy visual inspection of the tissues with most current types of microscope, and by an isoporous membrane on the top side. The membrane was specifically functionalized (supplement S2) to ensure a tight sealing of chip components, and to provide a porous barrier to the adjacent media microchannels. The channels can be connected to external devices (e.g. syringe-or peristaltic pumps), to achieve a vasculaturelike perfusion with media. This allows for a precisely controllable convective transport of nutrients, metabolites, and other dissolved molecules towards, as well as away from, the tissue chambers, mimicking the in vivo circulation of blood. It also opens up the possibility to administer compounds with high temporal resolution. The membrane further ensures that convective transport is restricted to the media channels, thereby shielding the tissue chambers Due to the microscale footprints of chambers and platform, only a small number of cells are required per chamber. This is of particular importance, as the tissues are based on primary, non-proliferating cells, and multiple chips (independent replicates) can be loaded with cells from a single biopsy, enabling testing under multiple experimental and control conditions of microtissues from the same donor. This is of crucial advantage when dealing with inter-donor variability, while it also enables patient-specific experiments. Online monitoring of fatty acid transport on day 6. Monitoring of fatty acid uptake and accumulation was realized by the addition of a fluorescent fatty acid analog (4 µM BODIPY™ 500/510 C1, C12) to the perfused medium for one hour, followed by measurement of the mean fluorescence intensity. Upon removal of the fatty acid analog from the medium, fatty acid release could be observed (n = 47 individual tissue chambers). Scale bar equals 150 µm. c. Evaluation of fatty acid metabolites in media effluents. Using colorimetric assays, glycerol and oleic acid (representative of non-esterified fatty acids) concentrations were found to be stable for the first three days of on-chip culture (n = 8 systems with 8 tissue chambers each). d. Upon addition of 0.2% w/v human serum albumin (HSA) to the culture medium for 24 h, fatty acid release significantly increased compared to that during previous days (p < 0.005; n =11 systems with eight tissue chambers each).

Structural analysis of WAT on-chip
To assess the capability of the WAT-on-a-chip system to maintain viability and functionality of the integrated tissues, we deployed a comprehensive toolbox of chip-specific, functional readout methods. First, the viability and integrity of the vast majority of cells after eight days of on-chip culture was confirmed using a FDA/PI-based live/dead-staining protocol tailored specifically for the chip's configuration (figure 4a). In addition, to verify that the objective of any microphysiological OoC system -the recapitulation of in vivo functionality -was indeed attained, we examined multiple functional endpoints. Taking advantage of the optical accessibility and continuous medium perfusion through our system, we performed noninvasive imaging and analysis of media effluents under several conditions.
To monitor fatty acid metabolism in real-time, we added the fluorescently tagged fatty dodecanoic acid (C12H24O2) analog (BODIPY™ 500/510 C1, C12) to the perfusion medium, and characterized the dynamics of fatty acid influx into, and efflux from, the WAT. This assay represents a powerful tool that allows non-invasive assessment of the cellular uptake of fatty acids, their accumulation in intracellular lipid vacuoles, and their release from adipocytes when the fatty-acid analog is removed from the medium. Utilizing time-resolved fluorescence microscopy, we were able to observe these characteristic uptake and secretion kinetics in the WAT cultured on-chip (figure 4b).
The continuous, convective transport by the perfusion flow, of metabolized and secreted factors away from the tissues, provides opportunities for effluent sampling in a time-resolved manner, allowing the dynamics of e.g. metabolism, secretion, and endocrine functionality to be measured. Using colorimetric assays, we characterized the levels of oleic acid, a representative of non-esterified fatty acids (NEFAs), and of glycerol in the effluent medium, and found that their concentrations remained very stable for several days of culturing (figure 4c). The measured NEFA and glycerol levels were comparable to levels from human subcutaneous adipose tissue explants immediately after biopsy (41). In a similar approach, we examined the impact on fatty acid release of a 0.2 (w/v) % supplementation of the perfusion medium with human serum albumin (HSA). After 24 h of perfusion, we observed a significant increase of NEFA levels in the effluent (figure 4d). Thus, in our system, effluent sampling provided a powerful, non-invasively obtained readout of secretome and metabolome dynamics with high temporal resolution (depending on the sensitivity of the employed assay). This constitutes an effective link for the cross-correlation with clinical data and biomarker development.
Long-term functionality of integrated white adipose tissue One of the main challenges of conventional adipocyte in vitro culture is the long-term maintenance of functional stability. As discussed above, issues related to buoyancy, fragility and de-differentiation of adipocytes, typically limit the duration of integrity of in vitro cultures to a couple of days at most (41,42). Our structural and functional characterization of the WAT-ona-chip indicated that this microphysiological platform is capable of maintaining stable culture conditions over prolonged time periods. However, to investigate the potential of the platform to support a robust, long-term culture, we performed fatty acid uptake assays on the tissues after 6 days, as well as after 36 days of on-chip culture (figure 5). We observed that even after more than a month of in vitro culture, adipocytes still showed functional uptake and accumulation of fatty acids. Interestingly, a difference in initial uptake rates emerged, which might be related to size limitation of the "long-term-fed" cells on day 36. Still, these findings highlight that the WAT-on-a-chip is indeed capable of maintaining the functionality of adipose tissue for much longer periods than are conventional methods. This brings within reach a variety of novel opportunities for in vitro studies, e.g. of long-term effects of nutrition, repeated exposure of potential toxicants, or of long-term endocrine dynamics. (43) Applicability of WAT-on-a-chip for drug testing Fig. 6 Applicability for drug testing illustrated by the effect of isoproterenol on fatty acid uptake and release. Normalized mean fluorescence intensity obtained by time-lapse fluorescence microscopy of WAT-on-a-chip systems perfused with medium supplemented with the fluorescently tagged fatty acid analog (4 µM BODIPY™ 500/510 C1, C12) during the first 60 minutes. Kinetics in systems exposed to 10 µM Isoproterenol (red circles) show significant differences to non-treated controls (grey polygons) (n = 7 and n = 8 individual tissue chambers; mean fluorescence intensities ± SEM are presented).
Since an important area of application for the organ-on-a-chip technology is drug development, we conducted a proof of concept compound test to assess the applicability of our WAT-on-achip for pharmaceutical research. After 6 days of on-chip culture, we exposed the WAT to the β-adrenergic agonist, isoproterenol, which is known to induce lipolysis. By additionally supplementing the medium with the fluorescently tagged fatty acid analog (BODIPY™ 500/510 C1, C12) for 60 minutes, we were able to monitor isoproterenol-related effects on fatty acid uptake and release by WAT, using standard fluorescence microscopy ( figure 6). While media with fluorescently tagged fatty acids was perfused, the normalized fluorescent intensity in the isoproterenol-treated systems increased significantly more slowly than in non-treated control systems. After switching to media without the fluorescently tagged fatty acid analog, the fluorescent intensity decreased much quicker in the isoproterenol-treated systems. Taken together, this means that the isoproterenol exposure induced a reduction of the net uptake rate of fatty acids and an increase of their release rate. Both findings are in line with the expected lipolysis-inducing effect of isoproterenol.
Evidently, the used approach based on standard microscopy provides facile and non-invasive read-outs for drug screening, with a very high time-resolution, which is amenable for massive parallelization and automation. One noteworthy limitation is the requirement for normalization of readings from individual tissue chambers. Absolute quantitative values would provide even more information and potential for cross-correlation; it is, therefore, planned to address this in future generations of chips and readout-infrastructure. Overall, the successful proof of concept highlights the applicability of the WAT-on-a-chip system for screening purposes and the potential the system can have for drug development purposes.

Conclusion
Due to the lack of suitable human model systems, research in adipose tissue biology and obesity has so far relied mostly on animal models, non-physiological in vitro systems, GWAS studies and (clinical) epidemiology. The first two approaches have major limitations in terms of their translatability to humans; the latter two require complex statistical analyses of large data sets and do not permit strong conclusions on pathophysiological mechanisms or efficacy of therapeutic interventions in individuals. The human WAT-on-a-chip platform presented here provides a novel tool that enables the maintenance, monitoring and manipulation of human adipose tissue in a durably stable, microphysiological environment. The vasculature-like perfusion provides opportunities for biomarker evaluation and "liquid biopsies" that can be cross-correlated with clinical endpoints, as well as a potential connection link for integration with other organ-on-a-chip platforms.(32) Multi-organ-chips integrating WAT are likely to be of major interest e.g. for ADMET, diabetes or non-alcoholic steatohepatitis (NASH) studies.
Overall, the WAT-on-a-chip system presented here, offers new perspectives and opportunities in mechanistic studies, pharmaceutical development and testing, as well as in personalized medicine.

Chip fabrication by soft lithography and replica molding
The microfluidic platform is a custom-designed three-layered hybrid device featuring two micro-patterned polydimethylsiloxane (PDMS; Sylgard 184, Dow Corning, USA) layers, which are separated by an isoporous semipermeable membrane. The media channel and tissue chamber microstructures in the PDMS slabs were generated using two differently patterned master wafers that served as positive molding templates. The intricately structured masters were fabricated by commonly used photolithographic processes described previously.(34) The chips' PDMS structures were created using two different molding techniques: standard molding to obtain thicker slabs with closed structures, and exclusion molding to obtain thin layers with open structures (cf. "Replica molding of PDMS parts" in the supplements section). To prepare the semipermeable membranes, commercially available polyethylene terephthalate (PET) membranes (rP =3 µm; ρP = 8 × 10 5 pores per cm 2 ; TRAKETCH® PET 3.0 p S210x300, SABEU GmbH & Co. KG, Northeim, Germany) were functionalized by a plasma-enhanced, chemical vapor deposition (PECVD) process (cf. "Membrane functionalization" in the supplements section). In a final step, chips were assembled in three subsequent O2-plasma activation (15 s, 50 W; Diener Zepto, Diener electronic GmbH + Co. KG, Ebhausen, Germany) and bonding steps, followed by an overnight exposure to 60 °C for bonding enhancement (cf. "Chip assembly" in the supplements section).

Chip preparation for experiments
On the day before cell injection, chips were O2-plasma sterilized (60 s, 50 W) and subsequently All primary mature adipocytes were isolated from biopsies that were taken from female, preobese donors (BMI 25.0 -29.9, as per the WHO classification), aged 45 to 55.

Isolation of primary human adipocytes
Primary mature adipocytes were isolated from subcutaneous adipose tissue samples. The isolation protocol was performed as previously described (44,45), with slight modifications. In

Injection of human mature adipocytes into the microfluidic platform
Immediately after the isolation of white adipocytes from tissue samples, the adipocytes were prepared for injection into the microfluidic platforms. Sixty µl of densely packed adipocytes were mixed with 24 µl of a dispersion of 10 mg/ml collagen type 1 (from rat tail, provided by

Responsiveness to β-adrenoreceptor agonistic drugs
To assess the effect of β-adrenergic agonist drugs on fatty acid metabolism, AM-1 medium was supplemented with 10 µM isoproterenol hydrochloride (I6504, Sigma-Aldrich Chemie GmbH). Fluorescence intensity was measured for 60 min with, and then for 60 min without addition of 4 µM BODIPY™ 500/510 C1, C12 to the conditioned medium at 37°C with an inverted fluorescence microscope (LEICA DMi8, LEICA Microsystems GmbH) to assess the dynamics of fatty acid uptake and release. Measurement and analysis of fluorescence was performed as described above.

Analysis of adipose-associated metabolites
The non-esterified fatty acid (NEFA) contents of medium effluents were determined by enzymatic analysis, using the ACS-ACOD-MEHA method (NEFA-HR(2) Assay, FUJIFILM Wako Chemicals Europe GmbH, Neuss, Germany). Effluents were thawed and centrifuged at As another readout for lipolysis, glycerol concentrations were determined using a colorimetric assay. Media effluents were thawed and centrifuged at 10,000 x g for 10 min. Then, 40 µl of sample [i.e. the effluents' supernatants, or AM-1 as blank, or Glycerol Standard Solution for a standard curve (G7793, Sigma-Aldrich Chemie GmbH)] were supplemented with 60 µl of the Free Glycerol Reagent (F6428, Sigma-Aldrich Chemie GmbH). After a 5-min incubation at 37°C, absorbance at 540 nm was recorded.

Statistical analysis
All graphs show raw data means ± standard deviation (unless otherwise indicated). We annotated or sample size n as follows: -For analyses pertaining to data at tissue chamber level (e.g. from fatty acid transport assays), n denotes the number of tested tissue chambers.
-For analyses pertaining on tissue system level only, i.e., the collectivity of eight chambers connected via one media channel, (e.g. metabolite measurements), n denotes the number of tested tissue systems.
For statistical analysis of differences, we performed independent two-sample t-tests using  PDMS slabs featuring media channel microstructures were produced by a standard molding process. First, 25 g of a PDMS mixture (containing a 10:1 w/w ratio of PDMS pre-polymer and curing agent) was poured over the media wafer. Then, after curing overnight at 60 °C, the PDMS mold was peeled off the wafer. b. PDMS layers featuring the tissue chamber systems were fabricated by exclusion molding. For this procedure, only ~ 1 g of the PDMS mixture was poured over the microstructured wafer. Then, a release liner, consisting of a polyester film, coated with fluoropolymer, was placed on top of the PDMS mixture, with the coated side facing the PDMS. A blank wafer was pressed against the other side of the foil to create open PDMS structures. After peel-off, PDMS stuck to the foil and PDMS residues were gently removed to uncover the intended open structures.

Membrane functionalization
Commercially available PET-membranes (TRAKETCH® PET 3.0 p S210x300, SABEU GmbH & Co. KG, Northeim, Germany) were modified using a plasma-enhanced chemical vapor deposition (PECVD) process. The PECVD, carried out by a low-pressure plasma, was performed in a custom-built plate reactor. The plasma was generated using a high-frequency United States). Before initiation of the coating process, the reactor chamber was cleaned using isopropanol, and all gas lines were evacuated to remove any residual gas. The PETmembranes, sized 210 × 300 mm², were placed inside the reactor chamber on top of a piece of protection paper of the same size, provided by the supplier. The plasma reactor was closed and evacuated until a pressure of approximately 1 -2 µbar was reached. Before starting the PECVD, the membranes were pre-treated by a 30-s exposure to an oxygen plasma to clean and activate the substrate. The process parameters are listed in table S1. Immediately after the pre-treatment, the membranes were coated by PECVD for 40 sec, using the parameters in table S1. The pre-treatment and coating process were applied on both sides of the membranes. After the membrane functionalization, the membranes were cut to the desired size, using a CO2-laser-cutter (VLS2.30, Universal Laser Systems). (2), and finally, the top layer and membrane were bonded to the bottom layer on the glass slide (3). Alignment of structures in top and bottom layer was conducted under visual guidance of a stereomicroscope. To enhance bonding, the chip was firmly pressed together between two microscope slides using foldback clips and kept overnight in an oven at 60°C. To verify the sealing tightness and fluidic connectivity of the system, tissue chambers and media channels were perfused with 70% ethanol.

Influence of human serum albumin presence on non-esterified fatty acid analysis
To ensure that increases in the measured NEFA concentrations could not be attributed to the presence of HSA during assay performance, we performed NEFA-measurements of the oleic acid standard with and without HSA supplemented medium. Oleic acid standard (100 µM) was added to AM-1 only or to AM-1 supplemented with 0.2% (w/v) HSA, and the NEFA assay was performed as described. No differences were observed in the measured absorbances between the two conditions ( figure S4).