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. 2022 May 5;16(3):034103. doi: 10.1063/5.0087532

Development of a microfluidic platform to maintain viability of micro-dissected tumor slices in culture

Maryam Parsian 1, Pelin Mutlu 2,a), Ender Yildirim 3, Can Ildiz 4, Can Ozen 1, Ufuk Gunduz 1,4,1,4
PMCID: PMC9076128  PMID: 35547184

Abstract

One of the issues limiting the development of personalized medicine is the absence of realistic models that reflect the nature and complexity of tumor tissues. We described a new tissue culture approach that combines a microfluidic chip with the microdissected breast cancer tumor. “Tumor-on-a-chip” devices are suitable for precision medicine since the viability of tissue samples is maintained during the culture period by continuously feeding fresh media and eliminating metabolic wastes from the tissue. However, the mass transport of oxygen, which arguably is the most critical nutrient, is rarely assessed. According to our results, transportation of oxygen provides satisfactory in vivo oxygenation within the system. A high level of dissolved oxygen, around 98%–100% for every 24 h, was measurable in the outlet medium. The microfluidic chip system developed within the scope of this study allows living and testing tumor tissues under laboratory conditions. In this study, tumors were generated in CD-1 mice using MDA-MB-231 and SKBR-3 cell lines. Microdissected tumor tissues were cultured both in the newly developed microfluidic chip system and in conventional 24-well culture plates. Two systems were compared for two different types of tumors. The confocal microscopy analyses, lactate dehydrogenase release, and glucose consumption values showed that the tissues in the microfluidic system remained more viable with respect to the conventional well plate culturing method, up to 96 h. The new culturing technique described here may be superior to conventional culturing techniques for developing new treatment strategies, such as testing chemotherapeutics on tumor samples from individual patients.

I. INTRODUCTION

Recent statistics from the World Health Organization (WHO) report indicate that cancer is the leading cause of death, with an estimated rate of around 10 × 106 deaths per year worldwide.1 During the significant progress in cancer treatment over the past few decades, the need for personalized therapies gained much attention.2 Anticancer drug candidates have a very high attrition rate, and only about 5% of drugs successfully complete phase III clinical trials.3 Besides these problems, a significant number of cancer patients are non-responsive to regular chemotherapy treatment. Furthermore, the adverse side effects of chemotherapy with low clinical benefits are seen in many patients.4 The lack of relevant models with appropriate human cancer nature and complexity is one of the problems that reduce the development of anticancer therapies. Therefore, developing novel drug testing tools and technologies besides new strategies for efficient monitoring of therapeutic agents is attractive to enhance treatment outcomes and specificity while minimizing toxicity.5–7

Recently, studies on spheroids produced by 3D cell culture and organoid models have been increasing. It begins to bridge the gap between the traditional 2D cell culture and the tissue culture. However, spheroids have a limited ability to mimic the complex tissue architecture and cell composition of human tumors. From this point of view, they cannot represent the real tumor tissue that is taken from patients in terms of different genetic makeup, tumor grade, or responsiveness to therapy.4,8,9 This has triggered the search for alternative or complementary therapeutic strategies with traditional tissue culture methods. Ex vivo cultures obtained from xenografts generated by tumor cells in immunodeficient mice constitute a structure much closer to the tumor microenvironment.10 Ex vivo cultures can supply suitable environments for experimental studies for the actual testing of therapeutics on real tissues from patients.11 However, this method has some significant problems, such as the inability to maintain viability and metabolic activity over a sufficient number of days for different analytical purposes. Since they lack any functional vasculature ex vivo primary tissues, they are prone to premature death and leave no time to test neither therapies nor obtain a reasonable output.12 Solid tumors are characterized by hypoxic and necrotic areas resulting from the high metabolic activity of tumor tissue and lack of oxygen supply due to insufficient vasculature. Hypoxic areas of solid tumors have an increased resistance to chemotherapy and radiation compared to better-oxygenated tumor tissue.13 In addition, the metabolic waste content of tumor sites is generally higher than normal tissue's microenvironment.14 Hypoxemia can be reduced by slicing the tissue into thin slices (250–500 μm) of relatively large (∼4 mm) diameter, with the smaller dimensions facilitating the transport of nutrients and oxygen to the center.5–15 Since the sections taken in this tissue culture method are very thin, there is no oxygen and nutrient deficiency, and the viability of the cells continues in the culture medium for a week. In addition, recently, researchers have successfully extended the culture time of tissues or cell aggregates in their microfabrication device or biochips by several weeks.11

Microfluidic devices have become very useful for the study of biological tissues and for simulating tissues and organs. These devices better mimic the in vivo environment than traditional in vitro methods, and research is moving toward developing a “human-on-a-chip” that enables fluid contact between tissues or organ-mimicking cell structures on a microfluidic platform.11–16 Tissue-on-a-chip devices developed for culture-on-chip tissue biopsy, toxicity studies, separation and detection of antibody-labeled cells from tumor cells, perfusion of living heart tissue by electrochemical monitoring, and examination of radiation-induced cell death in tissue.11,15,9–17 Thus, the importance of long-term culture with sufficient nutrients, oxygen, and better handling of waste products in microfluidic devices is beginning to be recognized.15,18–20 A microfabricated chip made of polydimethylsiloxane (PDMS) and containing an array of microchannels etched or molded into it can act as a culture vessel for cells and easily accommodate the medium flow or circulation. In addition, tissue pieces, such as tumor slices, were cultured in microfluidic devices to better preserve their function than traditional culture methods.21,22

A promising alternative would be directly testing alternative drugs on small amounts of cancer tissue taken from patients inside microfluidic biochips. Consider the following examples, patient derived liver and colorectal carcinoma models are used for drug screening processes by cancer-on-a-chip platforms.23,24 In addition, developing detection methods to measure the viability of tissues and chemotherapy testing simultaneously are novel approaches in a 3D ex vivo tissue culture from patients.9–25 We propose a new tissue culture method combining the microfluidic biochip with a microdissected breast cancer tumor to overcome these problems. This new strategy is an ideal model for tissue viability testing, new drug, drug life cycle from preclinical to patient use in terms of most suitable treatment regimen selection.

II. MATERIAL AND METHODS

A. The microfluidic chip

The microfluidic chip illustrated in Figs. 1(a) and 1(c) consists of three polydimethylsiloxane (PDMS) layers. The middle layer involves the chambers to accommodate the tissue slices. The top and the bottom layers contain the inlet and outlet channels, respectively. Polycarbonate filters [polycarbonate (PCTE) Membrane Filters, 8.0 Micron, Sterlitech, Auburn, WA, USA] were placed on the top and at the bottom of each chamber, both to preserve the tissue slices in the chamber and to ensure a more homogeneous flow profile in the chamber. Figure 1(b) compares the flow profile in one tissue chamber with filters to that without filters simulated by using COMSOL Multiphysics. PCTE filters and the tumor slice were modeled as porous media with the Darcian flow. Based on the manufacturer's data, the porosity and the permeability of the filter were set as 0.05 and 4 × 10−14 m2. Similarly, the porosity and the permeability of the tumor slice were set as 0.82 and 3.10 × 10−17 m2 by referring to Ref. 26. During the operation, flow rate across the microfluidics chip was measured as 20 ml/day. Assuming that the flow rates across four tissue chambers are equal to each other, the 5 ml/day flow rate was set at the inlet in the simulation model. The outlet was set to zero pressure. Navier–Stokes equations were solved assuming the laminar flow with the stated boundary conditions. The simulations resulted in that the hydrodynamic resistance of the tissue chamber with the filters and the tumor slice was in the order of 1015 Pa s/m3. This resistance was significantly greater than the hydraulic resistances of the channels, which were calculated to be in the order of 1011 Pa s/m3. The observation that the hydraulic resistance of the microfluidic network was dominated by the chamber resistances justified the assumption of equality of the flow rates across the tissue chambers.

FIG. 1.

FIG. 1.

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Overview of microfluidic device design and fabrication. (a) Schematic view of the micro-chamber sections used in the tumor viability analysis. (b) Flow profiles were obtained using COMSOL Multiphysics. Laminar flow physics was utilized with a constant flow rate of 5 ml/day at the inlet and zero pressure at the outlet. The PCTE filter and the tumor slice were modeled as porous media with the Darcian flow. (a) Unfiltered chamber geometry, (b) filtered chamber geometry, (c) flow profile observed in the chamber in the unfiltered condition, and (d) flow profile observed in the chamber in the filtered condition. (c) Micro-channel system. I. Bottom layer (outlet layer); II. Middle layer (tissue chamber); and III. Upper layer (inlet layer). (d) Photograph of the PDMS chip sandwiched by PMMA slide layers and binder clips.

The layers were sandwiched between two polymethylmethacrylate (PMMA) plates by using binder clips [Fig. 1(d)]. This non-permanent assembly scheme allows the tumor slices to be removed from the chip for further investigation during the experiments.

The top PDMS layer was fabricated as a thin layer (2 mm) to allow oxygen and carbon dioxide regulation within the tissue incubation chamber during testing. In order to facilitate permeation, air inlets were drilled on the top PMMA plate. The layers were fabricated by the standard PDMS molding process. The molds were fabricated as pockets on brass plates by milling (Proxxon FF500/BL-CNC Milling Machine), such that the thickness of the layers was directly controlled by the depth of the pockets.

During the operation, the medium is directed to chambers via the inlet channel posed in the top PDMS layer. The liquid is perfused through the tissue in the chamber and then collected via the exit channel at the bottom PDMS layer. However, since multiple chambers are fed in parallel, it is possible to have bubbles trapped in the inlet channel at the beginning of the operation. In order to prevent air bubbling problems, PDMS layers were pre-treated with 100% ethanol, and after drying, they were sterilized under UV light. However, bubbles were observed especially during the priming of the channels. To capture these air bubbles, a dummy outlet was located at the end of the inlet channel. The bubbles in the inlet channel were captured through this dummy outlet by using a syringe. Once the bubbles were completely removed, this dummy outlet was plugged and kept closed during the operation. For daily examinations, the media were collected from the outlet on the bottom PDMS layer and used in the biochemical analysis.

The amount of dissolved oxygen (DO) and pH were measured from the outlet medium of the non-tissue chip to check gas diffusion through the PDMS layers. The DO was measured by a micro-DO-electrode (YSI, 556MPS), and pH was measured by a pH-meter (WTW InoLab 720 pH/Cond Meter) from 20 ml outlet medium after 24 h. In addition, the pH of the outlet medium from the microdissected tissue cultured chip was measured during culture time, every 24 h.

III. EX VIVO BREAST CANCER MODELS

Female CD-1 athymic (without thymus gland) nude mouse models weighing ∼25 g were obtained from Kobay A.S. (Ankara, Turkey). The mice were maintained under a 12 h light/dark cycle in steel micro-isolator cages in a 22 ± 2 °C temperature and 45%–55% humidity-controlled room with free access to autoclaved water and food (SDS/RM3, DU, IRR) and handled only under sterile conditions. The use of animals for these experiments was approved by the Animal Ethics Committee of KOBAY DHL A.S. (local ethical comity-02.01.2017-#208).

1. Preparation of tumors and microdissected tumor slices

Tumors were generated in mice by using MDA-MB-231 and SKBR-3 cell lines. Mice were monitored for the development of primary xenograft tumors and weighted at least once a week. Tumors were measured weekly with a vernier caliper, and once they reached roughly 1 × 1 cm in size, they were removed and the animal was sacrificed. All of the surgical procedures began after placing the animals under general anesthesia. Following a subcutaneous incision, the tumor was exposed, measured, and placed in an ice-cold (1–2 °C) Hank’s medium (HBSS, #311-516-CL, Wisent Inc., Saint-Bruno-de-Montarville, Canada) supplemented with 1% Pen-strep (HyColone-Thermo Scientific). To produce the microdissected tumor slices, the tumor tissue was embedded into 4% low melting point agarose [SIGMA (A-9539), USA]. The agarose was solidified on ice for at least 10 min, thereby creating a supporting structure around the embedded tissue. A traditional Vibratome (The Campden Instrument LTD,1990) was used to produce 350 μm thick slices of the whole tumor inside a 10 °C bath containing Hank’s buffered saline solution supplemented with 1% Pen-Strep. The produced slices were kept in the same solution and finally further cut into disk-like tissues using a 3 mm diameter tissue punch inside Hank's buffer. The final product was a cylindrical microdissected tumor of approximately 350 μm in height and 3 mm in diameter and had a wet weight of about 3.4 μg. The representative scheme of the ex vivo culture system is given in Fig. 2.

FIG. 2.

FIG. 2.

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Representative scheme of the ex vivo culture arrangement. Preparation of tumors and microdissection of the tumor for culturing at two systems.

2. Culture of microdissected tissues

Immediately after slicing, the tissue slices were incubated by two different methods. The microfluidic chip and 24-well plates with a permeable polycarbonate membrane polystyrene transwell plate (Corning, 6.5 mm diameter, 8.0 μm pore size) were used. Every week one tumor was removed from one animal and incubated in the same conditions for microfluidic chip and well-plate experiments.

a. Incubation system

A: Microfluidic chip

Four microdissected tissues were placed in four chambers in the middle layer of the chip that had been attached by inlet and outlet layers. PDMS layers were fixed by two PMMA layers and placed in an incubator set at 37 °C for 96 h (from day 1 to day 4) [Figs. 3(a) and 3(b)].

FIG. 3.

FIG. 3.

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Placement of tumor sections in a microfluidic chip (a) and (b). Overview of incubators and placement of chips inside incubators. (c) Schematic view of a system and (d) real view of microfluidic chips’ incubation systems.

Sterile glass containers (100 ml) containing the medium were closed with two-ports microfluidic reservoir caps. Each container was connected to one chip so that many microfluidic chips could be operated in parallel. The containers were pressurized through the tubing and a manifold connected to a microfluidic autonomous pressure pump (Elvesys Elveflow AF1, Paris, France) [Figs. 3(c) and 3(d)].

The pressure provided by the microfluidic pressure pump was adjusted to allow ∼20 ml of medium per day to pass through each chip in parallel. The dummy outlet on the top PDMS layer was plugged with a 5 ml syringe. This outlet was used to capture the air bubbles present in the inlet flow channel. The media from the outlet on the bottom PDMS layer were collected daily in 100 ml sterile glass containers with special four-port lids. The flow rate was monitored by measuring the volume of the medium collected in the glass container after one day of operation by using the graduations on the container.

B. : Well plates

Tissue slices were incubated in a 24-well plate with a transwell permeable plate in a 1 ml RPMI 1460 medium and incubated at 37 °C for 96 h. The medium was removed for the metabolic activity test and replaced by a fresh medium every 24 h (Fig. 4).

FIG. 4.

FIG. 4.

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Well-plate culture system. Tumor slice cultured in the upper chamber of the transwell permeable plate.

b. Hematoxylin–eosin staining

After 6 weeks, H&E staining of the tumors obtained from MDA-MB-231 and SKBR-3 cells grown in CD1 mice was performed. In addition, pathological analyses were done on the first day of tumor removal to prove the development of tumors from tissue samples.

c. Tissue viability assessment with fluorescein diacetate and propidium iodide dyes by confocal microscopy

As a way to compare the viability properties of microfluidic chips and well-plate culture systems, microdissected MDA-MB-231 and SKBR-3 tumor slices from the same tumor were stained on the first and fourth day of cultivation. Following vibratome sectioning, the 350 μm slice was carefully punched to 3 mm and then washed with PBS before being incubated in a microfluidic system and well-plate system.10 μl fluorescein diacetate (FDA) (Sigma-Aldrich, USA) and 5 μl propidium iodide (PI) (Sigma-Aldrich, USA) were dissolved at 5 ml PBS. On the first day, slices were incubated at 24 well-plates with 1 ml/well PBS containing FDA and PI and rotated for 1 h. After the incubation time, the microdissected tissues were washed with PBS and imaged. Layer-by-layer photos of each microdissected tumor were captured by confocal microscopy on the first day immediately after segmentation and 96 h later from two cultivated method tumors. Two replicate samples were used to capture 10 sequential images using a 20× lens to test each cultivation method. The contrast of images was improved by using deconvolution software with Huygens essential software (SVI, Hilversum, the Netherlands). Totally six different images were taken from each group (two replicated slices and three shots from different positions of a slice). The intensity sum (px intensity) of fluorescence was calculated by using the MIPAR software.27

d. Daily analysis of the supernatant

The metabolic activity was detected daily from the outlet fractions of the microfluidic device and well-plate system by using an automated biochemistry autoanalyzer (Beckman Coulter DXC800, USA). A supernatant volume of 1 ml/day was used to assess the lactate dehydrogenase (LDH) and glucose content of the output media from slices in both systems.

IV. RESULTS AND DISCUSSION

Tumor slices are cultured in a newly designed microfluidic chip system and compared with traditional culture methods. One of the microfluidic chip's primary purposes is to keep the tumor fragments alive in culture systems for anticancer drug testing on real tumor tissues from patients in the future. The continuous supply of fresh media to the tissue samples and removing metabolic wastes maintain viability during the culture period. Many conventional culture methods implicate various drawbacks, including maintaining viability over a sufficient number of days for various analytical purposes. Furthermore, without adequate feed and oxygen, tumor slices die prematurely, leaving little or no time to test therapeutics for guiding the selection of an optimal treatment regimen.

A. Ex vivo culture studies

In the ex vivo culture studies, tumors were developed in CD-1 nude mice with SKBR-3 and MDA-MB-231 cells by subcutaneous injection. In each experiment, when the tumor reached around 1 × 1 cm, the mouse was anesthetized and the tumor was removed. After the operation, the removed tumor was immediately placed in a 50 ml cold Hank's buffer and transferred in a cold chain. The tumor was dissected within 60 min into 3 mm × 350 μm pieces and placed into the well-plate and microfluidic systems.

In the microfluidic system, four microdissected tumor slices were placed in the chambers of the chip, between two layers of polycarbonate filters, and sealed with the perforated PMMA layers. The tumor chip was connected to a container containing the appropriate media and attached to a calibrated pressure pump within an in-house incubator with an installed CO2 and O2 sensor. The pressure pump was set to ∼19 mbar, so the medium was infused continuously into the chip at a rate of 14 μl/min for 96 h. Fresh media were provided to the tissue samples for 96 h uninterruptedly, and the media coming out of the system were collected every 24 h. Infusions were stopped after 96 h, and microdissected tumor samples were removed from the device for post-chip analyses.

B. Tumor tissue analyses by hematoxylin–eosin staining

Tumors that were derived from MDA-MB-231 and SKBR-3 breast cancer cell lines in CD1 mice were visualized on a stereomicroscope for the identification of tumor size, morphology, and angiogenesis [Fig. 5(a)]. Then, further pathological analyses were done by the hematoxylin–eosin staining method. Hematoxylin–eosin staining of tumor sections that were derived from MDA-MB-231 cells indicated that cells preferentially grew in ducts to form ductal papillae being the main characteristic of MDA-MB-231 originated tumors [Fig. 5(b)].

FIG. 5.

FIG. 5.

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(a) MDA-MB-231 tumor image under a stereo microscope and (b) representative H&E images of MDA-MB-231 tumor slice sections on the first day.

C. Assessment of cell viability of microdissected tissue cultures

Following segmentation of the MDA-MB-231 and SKBR-3 tumor tissue, fluorescein diacetate (FDA) and propidium iodide (PI) dyes were used to assess the viability of microdissected tissues before and after 4 days of cultivation. Various methods can be used to measure the survival of cells within tissue slices. Confocal laser scanning microscopy eases access for living cell imaging in the 3D tissue. By using confocal microscopy, it is possible to obtain stratified images through the depth of the 3D tissue sample, allowing for information on each focal plane to be obtained. A computer-controlled confocal microscopy creates digital images that are compliant with image and data processing.28 FDA has been used for the viability assessment of a wide variety of cells and tissue types.29,30 The FDA assay does not have any significant toxicity, making this assay suitable for the cell and tissue culture analysis. FDA is not a fluorescent molecule, and by the opportunity of bipolar side chains, it penetrates into living cells. FDA remains colorless until esterases convert it to fluorescein inside the metabolically active cells. Under the light of microscopy, the intensity of fluorescence was seen; it gave the proportion of live cells in the layers of tissue.30,31

The maximum intensity projection (MIP) and the fluorescence intensity histogram for the green (live) and red (dead) were subtracted from the 3D image [Figs. 6(b), 6(d), and 6(f)]. The MIP visualized the volume of live and dead cells, and the intensity histogram of MIP demonstrated fluorescent intensity to the number of voxels in each 3D image. MIP images with low green intensity and high red intensity were seen in an image from well-plate system slices [Fig. 6(f)]. However, imaging from the microfluidic slice showed that red fluorescence was strongly diminished, whereas green cells remained brightly fluorescent [Fig. 6(d)].

FIG. 6.

FIG. 6.

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Confocal imaging and maximum intensity projection of MDA-MB-231 microdissected tissues. Layer-by-layer images indicated the differences between the amount of live and dead cells in microdissected tissues on the first day immediately after operation (a) and (b) and after 4 days of cultivation at the microfluidic chip (c) and (d) and the well-plate system (e) and (f). Live and dead maximum intensity projections from a representative MDA-MB-231 slice of confocal image stacks from the first day and after 96 h from the microfluidic and well-plate system (by Huygens Essential software) (b), (d), and (f).

Figure 7 shows the observed difference between average red and green fluorescence intensity from six images. The sum intensity for each layer of images is calculated separately. The loss of red fluorescence intensity was more pronounced in whole tumor (MDA-MB-231 and SKBR-3) images from the microfluidic chip. In comparison, green fluorescence remained bright, even more than first-day green intensity.

FIG. 7.

FIG. 7.

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The mean sum intensities of MDA-MB-231 (a) and SKBR-3 (b) tumors from three different positions of two slices of microdissected tissues (slice number: n = 2, image number: n = 6). The sum intensity for live and dead images is calculated by the MIPAR program for each layer-by-layer image. The average of sum intensities shows the difference between the first and fourth days of different cultivation methods.

D. Mass transfer analysis of oxygen and carbon dioxide

The addition of dry ice to the incubator periodically provides the required amount of CO2, as measured by a CO2 sensor. By virtue of PDMS gas permeability, it allows CO2 and oxygen to diffuse into the chamber from the ambient air. Therefore, controlling carbon dioxide gas concentration in a surrounding area could provide the optimal pH for tissue viability.

During the setting up in the first step, the stability of pH values was measured every day in a non-tissue installed microfluidic chip for 96 h. The pH value was measured in the supernatant medium of the chip and compared with CO2 concentration in the incubator (ppm/min) to determine how much dry ice was needed to be added to the incubator during the day. A small butterfly valve controlled the amount of exiting CO2 from the reservoir of the dry ice container. By this method, the microfluidic chip outlet medium pH was measured as ∼7.4 ± 0.05 after 24 h, indicating equilibrium with 5% CO2 at 37 °C.32 Furthermore, the pH value of the tissue-containing device perfusion experiments was also constant and suitable for tissue viability during 4 days. The pH value of the outlet medium of the micro-device was measured every 24 h, and results were the same as a non-tissue-containing device, between 7.4 and 7.6 ± 0.05. For the well-plate system, the media of the tissue were changed every day; the pH value of media was around 7.6–7.8 ± 0.05. In addition, we used a medium (RPMI1640) containing phenol red, which is a pH indicator. No color change and, as a consequence, no pH change were observed in the outlet medium of tissue containing chips during all experiments, which indicated that the CO2 level was sufficient over 24 h. However, a significant color change could be seen in the medium of control (non-tissue well) and tissue-contained wells at well-plates experiments.

The concentration of dissolved oxygen (DO) in the outlet medium of the non-tissue-cultured device was also very high and stable, around 98%–100% (9.38–9.43 mg/l) on a supernatant every 24 h. The concentration of DO in the outlet medium in the presence of tissue was also checked at the end of 96 h and found as 100% (9.43 mg/l). The O2 permeability of PDMS and the oxygen concentration in the inlet medium are enough since the medium is initially saturated with oxygen during the pumping process. Therefore, a constant flow of the oxygenated medium through the chambers offers the appropriate amount of oxygen for the micro-tissues during culture time in the microfluidic system.32,33 Misra et al. showed that 350 μm thick precision-cut slices represent the maximum width for the diffusion of oxygen and substrates into the deepest cell layers of the tumor slice. Cultured tissue slices were metabolically active for 96 h at their well-plate culture system. Also, they indicated that oxygen tension did not affect the proliferative activity of the tumor.33 Monitoring O2 levels is crucial for assessing tissue cell viability and metabolic activity in microfluidic systems. As a result of recent advances in microfluidics, O2 sensing has become an essential feature for organ-on-a-chip devices for various biomedical applications. Therefore, various optical and electrochemical on-chip O2 sensors were integrated with the organ-on-a-chip devices.21 The low concentration of O2 inside many microfluidic chips requires precise and selective quantification for confined settings. Based on COMSOL simulations, the suitability of different tumor slices for oxygen mass transport within microfluidic devices was evaluated, leading to the development of an optimized design for achieving defined in vivo oxygenation within an organ-on-a-chip system.18

E. Microdissected tumor slice viability analysis

Christensen et al. show the oxygen transport efficiencies in the precision-cut liver slice fabricated device by the LDH assay.18 The viability of the tumor tissue samples was evaluated by lactate dehydrogenase (LDH) release and the glucose consumption analysis. LDH is a cytoplasmic enzyme constitutively expressed in most mammalian cells. Determining the amount of LDH released into the medium is widely used to assess plasma membrane integrity. Excessive release of LDH enzyme into the medium indicates that the cells are dead, while the consumption of glucose from the medium shows that the cells are alive for determining the micro-slice viability in the microfluidic chip and well-plate incubation systems, leakage of LDH and glucose consumption amount were measured every 24 h for 96 h. In each experiment, the tumor of the same mouse was incubated in the two incubation systems.

1. Lactate dehydrogenase analysis

In order to confirm that the tumor micro-slices were viable in the micro-chamber, the viability was examined using LDH release over a 96 h period and compared to traditional well-plate systems.

All experiments were performed with four tumor slices per mouse for five mice for SKBR-3 and four mice for MDA-MB-231. Due to the use of one tissue in the well-plate system, the amounts of LDH in the microfluidic system were also calculated for one tissue. A supernatant volume of 1 ml was used to assess the LDH content of the slice. First and foremost, the initial peak of LDH releases at the beginning of the experiment (first 24 h) was found in all mice trials in the two incubation systems. It is probably due to the damage caused to the surface of the tissue in harvesting, handling, and slicing procedures, causing LDH enzymes’ leakage in the first hours of incubation. Van Midwoud et al. (2009) and Olubajo et al. (2020) also were found a similarly high percentage of LDH release from liver and glioblastoma slices in the first hours.32–5

Figure 8 demonstrates the level of LDH released from MDA-MB-231 tumors slices decreased around 50% in the second 24 h in both incubation systems (Average: 6.1 MU/ml for MF system and 52.0 MU/ml for well-plate). After 72 h, the rate of decrease remained low in both systems (2.6 MU/ml for MF system and 39.0 MU/ml for well-plate). As shown in Fig. 8, at 96 h measurements, the amount of release increased around 50% at the well-plate system (62.3 MU/ml), however at microfluidic systems, the LDH leakage remained stable during the last 24 h (3.4 MU/ml).

FIG. 8.

FIG. 8.

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Comparison of the amount of LDH released from MDA-MB-231 tissues in the microfluidic and well-plate system over four days (a: 24 h, b: 48 h, c: 72 h, and d: 96 h). Results are mean ± standard error of the mean (SEM) of four mice, one slice per incubation system per experiment. This difference was statistically significant when tested with a student t-test (*p < 0.05, **p < 0.01, and ***p < 0.001).

The pattern of LDH release into the effluent from SKBR-3 tumors was distinctly similar to MDA-MB-231 tumors release in both systems. LDH release profiles for SKBR-3 tumors slices in the microfluidic and well-plate systems every 24 h during four days were 38.7, 5.4, 3.5, 2.6 MU/ml and 214.3, 110.7, 44.7, 86.3 MU/ml, respectively. By comparing two systems in both tumors, LDH release results indicated that the tissues in the microfluidic chip system were not damaged by the flow and remained viable during 96 h, despite the well-plate culture method (Fig. 9).

FIG. 9.

FIG. 9.

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LDH release from SKBR-3 tissues at microfluidic and well-plate systems (a: 24 h, b: 48 h, c: 72 h, and d: 96 h). Results are mean ± standard error of the mean (SEM) of five mice, one slice per incubation system per experiment. This difference was statistically significant when tested with a student t-test (*p < 0.05, **p < 0.01, and ***p < 0.001).

Van Midwoud et al. (2010) showed high viability in the micro-chamber and well-plate system. In both systems, the LDH leakage patterns are similar during 24 h. However, the main differences between our system and this published experiment are the thickness of their tissues (100 μm) and experiment time (24 h). By making thinner slice, the diffusion distance of nutrients and oxygen toward the inner cells of the slice is minimized; so during 24 h in the well-plate system, the amount of dead cells is similar to the micro-chamber system. Also, the Van Midwoud group showed that because PDMS is a relatively hydrophobic polymer, protein LDH was not adsorbing onto the PDMS and all LDH was released to the supernatant.32

2. Metabolic activity

Glucose levels were measured in the two culture methods every 24 h (Fig. 10). In contrast to LDH release, the glucose consumption of the microfluidic chip increases during this period, which shows the groups are alive inside the micro-chamber during 96 h.

FIG. 10.

FIG. 10.

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Comparison of the microfluidic and well-plate systems metabolic activity over 4 days. (a) Glucose consumption in MDA-MB-231 tissues at the microfluidic and well-plate systems. (b) Glucose consumption in SKBR-3 tissues at the microfluidic and well-plate system tissues.

The glucose consumption amount of the well-plate measured for one slice inside 1 ml medium/day, 1 ml fresh medium added to each well every day. RPMI-1640 medium contained ∼2 mg/ml glucose, which is sufficient for a slice of tissue culture. As shown in Fig. 10, the metabolic activity is stable for three days, while the glucose level decreased more than fourfolds in the MDA-MB-231 tumor slice and around 1.5-folds in the SKBR-3 tumor slice at last 24 h.

At the microfluidic chip, four slices were cultured on one device by the flow rate of ∼20 ml/day. 40 mg glucose is available at this culture medium for 24 h, sufficient for four slices of tissue culture. The quantity of glucose used is determined by subtracting the amount of glucose at fresh medium from the amount of glucose in the supernatant. The amount of glucose consumption is calculated for one slice in the microfluidic system. Comparing the results of the two culture methods demonstrated that the tumor slice used a remarkably high glucose amount in a microfluidic device during the incubation time. Furthermore, this result shows that the tissue was alive and metabolically active in the microfluidic system even after 96 h.

V. CONCLUSION

In conclusion, we successfully developed a new PDMS microfluidic device. Three thin polymer layers with polycarbonate filters at the bottom and top of the microchamber were used. Filters were necessary to ensure an equal distribution of the medium flow across the entire chamber; this caused the same pressure over the entire surface of the tissue and prevented the curling up of the slices in the microchamber. When slices are folded or curled up, diffusion distances to inner cells are increased and affect the nutrient and oxygen delivery to inner cells. The characterization reveals a versatile device, which can operate as a suitable platform for tumor slice cultivation with an appropriate amount of oxygen and carbon dioxide. Consequently, our study showed the most intensive metabolic activity observed in tumor slices cultivated in a microfluidic system for up to 96 h compared with a well-plate system. Using microdissected tumor tissue in microfluidic system will allow for the study of the therapeutic response in the tumor tissue much more closely than in vivo studies. In addition, it may significantly benefit the development of testing of the biopsy samples from patients rapidly for the effectiveness of different chemotherapeutic agents under laboratory conditions in the future.

ACKNOWLEDGMENTS

The authors thank Ali Can Atik, Dr. Nusret Taheri, and Nalan Kamalı for technical assistance. This study was supported by The Scientific and Technological Research Council of Turkey (TUBITAK 1001 Project No.117Z092).

AUTHOR DECLARATIONS

Conflict of Interest

The authors have no conflicts to disclose.

Ethics Approval

Ethics approval has been obtained by the Animal Ethics Committee of KOBAY DHL A.S. (local ethical commity-02.01.2017-#208).

DATA AVAILABILITY

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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Associated Data

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Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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