Human vascularised synovium-on-a-chip: a mechanically stimulated, microfluidic model to investigate synovial inflammation and monocyte recruitment

Healthy human fibroblast like synoviocytes (hFLS) from 3 male donors aged 32, 39 and 75 were purchased from a commercial source (Articular Engineering, Illinois, USA) and used from passage 1-6 for experiments. Cells were cultured in monolayer in Dulbecco's modified eagle medium: nutrient mixture F-12 (DMEM/F12; 1:1) supplemented with 2 mM penicillin/streptomycin and 10% (v/v) foetal bovine serum (FBS; all Gibco, Loughborough, UK). hFLS were cultured to 80% confluence prior to passaging and seeding for experiments. HUVECs (Lonza, Cambridge, UK) were used from passage 3–7 and cultured in endothelial growth medium-2 (EGM-2; Lonza). THP-1 monocytes (Invivogen, Toulouse, France) were cultured in Roswell Park Memorial Institute (RPMI) medium (Gibco) supplemented with 2 mM penicillin/streptomycin and 10% (v/v) FBS. All cells were cultured in a humidified incubator at 37 °C with 5% CO₂.

The Chip-S1^® (Emulate Inc., Boston, USA) was used (figure 1(B)) to generate the vascularised synovium-on-a-chip model. This PDMS based chip comprises two overlapping channels separated by a thin (50 µm), porous (7 µm pore diameter) membrane. The channels are 1 mm in width and the channel height is 0.2 mm and 1 mm for the bottom and top channels respectively. Each channel is connected to its own medium reservoir using the Pod^™ system (Emulate) such that each cell population can be supplied individually with required media formulations, and fluid shear can be applied to each channel independently. Two vacuum channels either side of the culture channels are used to apply uniaxial cyclic tensile strain across the membrane (figure 1(B)). A commercially available organ-chip system was selected for use in the present study to allow for the wider research community to adopt the optimised model without the need for specialist knowledge of chip fabrication techniques.

Prior to cell seeding, the inner surfaces of the channel were activated according to standard protocols (emulatebio.com; 'Basic Research Kit Protocol: EP223). In brief, 0.5 mg ml⁻¹ ER-1 solution (Emulate) was added to each channel and the chips were then treated with UV light for 10 min. The channels were aspirated, and the process repeated with additional ER-1 solution. Channels were then washed with ER-2 solution (Emulate) followed by phosphate buffered saline (PBS; Gibco). Extracellular matrix, in the form of 500 µg ml⁻¹ collagen Type I (Corning, Arizona, USA), was added to the top channel, and a mix of 250 µg ml⁻¹ collagen type 1 and 25 µg ml⁻¹ fibronectin (Thermo Fisher Scientific, Massachusetts, USA) added to the bottom channel. Chips were incubated overnight at 4 °C, followed by 1 h at 37 °C.

In the final chip configuration, hFLS were cultured in the top channel and HUVECs in the bottom channel (see schematic representation in figure 1(C)). Following the matrix coating of the two channels, chips were washed with pre-warmed culture medium and HUVECs seeded to the bottom channel at a concentration of 6 × 10⁶ cells ml⁻¹. Chips were inverted for 2 h to encourage attachment to the membrane, then righted and cultured for a further 24–48 h to encourage formation of a robust monolayer with associated tight junctions. hFLS were then seeded to the top channel at a concentration of 1 × 10⁶cells ml⁻¹ and allowed to attach for 4 h prior to gentle washing. After a further 24 h, chips were connected to the automated culture module (ZÖE; Emulate) and cultured with their respective media flowing at a rate of 30 µl h⁻¹ for 7 d to encourage deposition of hFLS-derived matrix before the experimental parameters, optimised as described subsequently, were applied.

Several experiments were initially carried out on 2D monocultures of hFLS to optimise experimental parameters, which were subsequently applied to the organ-chip system. Firstly, an interleukin−1β (IL−1β) dose response experiment was performed. hFLS were subjected to an IL−1β (Peprotech, London, UK) dose of 0, 0.1, 1 or 10 ng ml⁻¹ for 24 h to identify an optimal dose for use in subsequent studies, with culture supernatant examined for the presence of inflammatory signalling molecules (NO, PGE₂, interleukin-6 (IL−6) and matrix metalloproteinase 1 (MMP-1)). Based on results from these experiments (detailed in the results section) a dose of 1 ng ml⁻¹ of IL−1β was used for subsequent experiments.

Mechanical strain regimes were also initially optimised off-chip. Cyclic tensile strain (CTS) was applied using a commercially available stretch system (Flexcell; Flexcell^® International Corporation, North Carolina, USA). hFLS were seeded on Collagen Type I coated Uniflex plates (UF4001-C, Dunn Labortechnik, Thelenberg, Germany). Plates were additionally coated overnight with 50 µg ml⁻¹ Collagen type 1 (354249; Corning, Arizona, USA) at 4 °C followed by incubation for 1 h at 37 °C and washing with culture medium. hFLS were seeded at a density of 5000 cells cm⁻² and cultured until confluent (approx. 7 d). hFLS were subjected to a strain regime which comprised 22 h of unloading immediately followed by 2 h of loading, at 2% or 12% strain at a frequency of 0.2 Hz using a sinusoidal waveform. These experiments were performed in the presence or absence of 1 ng ml⁻¹ IL−1β.

The optimised mechanical regime was applied to the chips from day 7, following the initial culture period, such that chips were unstretched for 22 h then subjected to cyclic stretch for 2 h at 12% strain, 0.2 Hz using a sinusoidal waveform. This was repeated for 3 d (figure 1(D)). At the same time, inflammation was induced in the synovium-on-a-chip through addition of the optimised dose of 1 ng ml⁻¹ IL−1β in serum-free hFLS media to the top channel inlet media reservoir at day 7. IL−1β containing media was then flowed through the top channel at a rate of 60 µl h⁻¹, for 3 d (figure 1(D)).

On day 10, THP-1 monocytes were labelled with 2 µM cell tracker dye (CMRA orange, Thermo Fisher Scientific) for 15 min in serum-free medium at 37 °C. Cells were then centrifuged at 200 ×g for 8 min to form a pellet, washed with serum-free medium then resuspended to a concentration of 4 × 10⁶ cells ml⁻¹. Cells were subsequently mixed 1:1 with an equal volume of buoyancy medium (1.6% v/v gelzan in Percoll; Sigma-Aldrich, Dorset, UK) and added to the inlet reservoir of the bottom channel of the Pod^™. Fluorescently labelled monocytes were flushed through the bottom channel at 1000 µl h⁻¹ for 10 min then incubated under static conditions for 3 h. The channel was then flushed with fresh EGM-2 for 10 min to removed unattached cells and chips were imaged to assess monocyte recruitment using confocal microscopy. Labelled monocytes were counted using Image J (Image J 1.53c, National Institutes of Health, USA) for 15 fields of view per chip.

2.6.1. Sample collection

2.6.2. RNA isolation, cDNA synthesis and qRT-PCR

2.6.3. Effluent analysis

Nitric oxide (NO) release was monitored indirectly using the Greiss assay, a widely used test which measures the levels of nitrite (NO₂), a stable metabolite of NO. The nitrite content of culture media was assayed in triplicate against a sodium nitrite standard curve. Commercially available enzyme-linked immunosorbent assays (ELISAs) were used to examine the release of PGE₂ (KGE004B), MMP-1 (DY901B), IL−6 (D6050) and hyaluronan (HA; DHYAL0; all R&D systems, Abingdon, UK). Optimisation was performed to determine suitable dilutions for each analyte to fit within the recommended range of the corresponding assay and samples were diluted as required in culture media and assay diluent before being analysed in duplicate. For all analytes, the detection limit (DL) was determined [21]. The DL was based on the standard deviation (SD) (σ) of the response, and the slope (S) and can be expressed as follows:

$$\begin{equation*}{\text{DL}} = \frac{{3.3\sigma }}{S}.\end{equation*}$$

For all analytes, values below the DL were replaced with the DL value to enable statistical analysis, as is common practice.

To assess barrier function in monoculture and co-culture chips, fluorescent tracer dye was added to the culture media in the top channel only throughout the duration of the experiment. To examine transport of differently sized molecules, cascade blue conjugated 3 kDa dextran (Thermo Fisher Scientific) and FITC-conjugated 70 kDa dextran (Sigma-Aldrich) were used at a concentration of 0.1 mg ml⁻¹. In brief, the synovium-on-a-chip was disconnected from the automated culture module, ZÖE (Emulate), and 100 µl media removed from each reservoir. Fluorescence intensity was measured at 355 nm and 485 nm and the concentration of dye was determined for the dosing channel (top channel) and receiving channel (bottom channel) using a standard curve. This data was used to assess the apparent permeability (P_app) according to the following calculation:

$$\begin{align*}{P_{{\text{app}}}} & = \, - \frac{{{Q_{\text{R}}}*\,{Q_{\text{D}}}}}{{{\text{SA}}*\left( {{Q_{\text{R}}} + \,{Q_{\text{D}}}} \right)}}\nonumber\\ & \quad\,\, *{\text{In}}\left[ {1 - \frac{{{C_{{\text{R,0}}}}*\left( {{Q_{\text{R}}} + {Q_{\text{D}}}} \right)}}{{{(Q_{\text{R}}}*{C_{{\text{R,0}}}} + {Q_{\text{D}}}*{C_{{\text{D}}{\text{.0}}}})}}} \right]\end{align*}$$

where P_app is the apparent permeability in units of cm s⁻¹, SA is the surface area of the co-culture channel (0.17 cm²), Q_R and Q_D are the fluid flow rates in the receiving and dosing channels, respectively, in units of cm³ s⁻¹, and C_R,0 and C_D,0 are the recovered concentrations in the receiving and dosing channels respectively (Emulate).

While all cells used in the present study were commercially sourced, we used primary synovial tissue collected from patients undergoing treatment for OA in the knee (collected for a separate ongoing study) to generate the representative images of healthy and inflamed synovium shown in figure 1(A) only. All patients provided written informed consent, approved by the National Research Ethics Committee in 11/NW/0875. Samples were formalin fixed and paraffin embedded before being sectioned at a thickness of 5 µm and stained with haematoxylin and eosin. Images were captured using a DS-FiL camera (Nikon Corporation, Tokyo, Japan).

Immunocytochemistry of hFLS monolayers and synovium chips was performed according to established protocols. In brief, samples were fixed with 4% PFA and then washed with PBS. This was followed by permeabilisation with 0.5% triton X-100 for 5 min, blocking with 5% donkey serum for 1 h at room temperature and the application of primary antibodies overnight at 4 °C in 0.1% bovine serum albumin (BSA; Sigma-Aldrich)/PBS. Primary antibodies used in this study are as follows: fibronectin-1 (ab2413, Abcam, Cambridge, UK), cadherin-11 (5B2H5, Invitrogen, Loughborough, UK), lubricin (ab28484, Abcam), VE-cadherin (sc-9989, Santa Cruz Biotechnology, California, USA), platelet endothelial cell adhesion molecule-1 (PECAM-1)/CD31 (ab9498, Abcam), intercellular adhesion molecule-1 (ICAM-1) (ab2213, Abcam). After 24 h samples were washed with 0.1% BSA/PBS and incubated with appropriate Alexa Fluor-conjugated secondary antibodies (Molecular Probes, Massachusetts, USA) for 1 h at a dilution of 1:500. Nuclei were counterstained with DAPI (D3571, Thermo Fisher Scientific) and actin labelled with Alexa Fluor 555-conjugated Phalloidin (Santa Cruz Biotechnology) for 1 h at room temperature. Primary antibodies were diluted at 1:200, secondary antibodies at 1:500 and DAPI was used at 1 ug ml⁻¹. Membranes were mounted cell side up with Fluoromount G (00-4958-02, Thermo Fisher Scientific). Samples were imaged using a Nikon CSU-W1 SoRa Spinning Disk Confocal (Nikon) using a CFI Plan Fluor 10× objective (0.3 NA). Isotope controls are not included for immunocytochemistry as our group has extensive experience in the use of these optimised antibodies which have limited non-specific activity and label characteristic structures which are extremely unlikely to be shown by non-specific binding.

Statistical analysis was performed using GraphPad Prism version 9.5 for Windows (GraphPad Software, San Diego, California USA). Prior to statistical analysis, all data was assessed for normality (Shapiro–Wilks test). With normal distribution confirmed, statistical significance was assessed by Students t-test, one-way, two-way or three-way ANOVA followed by Sidak's multiple comparisons test, as highlighted in the individual figure legends. The threshold for statistical significance was set at 0.05. Data is represented as mean ± SD. Statistical significance is displayed as p < 0.05 = *, p < 0.01 = **, p < 0.001 = ***, p < 0.0001 = ****. Data from multiple donors are displayed in unique colours which remain consistent throughout all figures.

HUVECs were seeded into the bottom channel of the chip S1^® and cultured for 48 h prior to the addition of hFLS. HUVECs attached well to ECM coated surfaces, forming an even monolayer through the length of the channel (figure 2(A)). An observable permeability barrier was present from day 3 in culture following connection to the automated culture module and application of fluid flow, which acted to reduce diffusion between the channels (figures 2(C) and (D)). Tight junction formation was observed with VE-Cadherin labelling (figures 2(E) and (F)) and cells labelled positive for PECAM-1; figure 2(F). HUVECs grew on all inner surfaces of the bottom channel forming a tubule-like structure with a hollow lumen (figure 2(E)).

Zoom In Zoom Out Reset image size

Following HUVECs pre-culture, healthy hFLS were seeded into the top channel of the chip S1^® (figure 3(A)). hFLS attached well along the length of the channel (figure 3(A)) and could be successfully cultured for 24 h without media refreshment with no negative effects on cell viability. Matrigel^™ has commonly been used for the generation of synovial organoids [8, 22, 23]. However, within the chip environment, hFLS were found to exhibit a more invasive phenotype when cultured on Matrigel. Cells migrated through the membrane pores within 24 h of seeding and grew on the opposing side of the membrane (figures S1(B) and (C)). Therefore, initial studies explored the use of alternative ECM coatings to limit migration, with membrane coatings optimised using transwell cultures (figure S2), in which collagen type I was found to be most effective at limiting/delaying this behaviour (figures S2(B) and (C)).

Zoom In Zoom Out Reset image size

On collagen type I coated surfaces, hFLS formed a loose network at the surface of the membrane (figures 3(E) and (F)). Co-culture chips formed a robust permeability barrier to 70 kDa dextran but were found to be more permeable to 3 kDa dextran than monoculture chips (figures 3(C) and (D)). Deposition of fibronectin-1 (FN-1) was observed in the synovial compartment (figure 3(E)) and hFLS exhibited positive labelling for PRG4 (lubricin), which was found distributed throughout the cell cytoplasm (figure 3(G)). Furthermore, expression of the synovial marker cadherin-11 could be observed at points of cell–cell contact, overlapping with actin labelling (figure 3(F)). In addition, hFLS-secreted HA was detected in the culture media (figure 6(F)).

FLS exhibit mechanosensitive gene expression of HA synthase enzymes (HAS-1, 2 and 3) and production of HA both in vivo [24, 25] and in vitro [26]. However, many of these studies use cells from animal models, cell lines or cells isolated from diseased tissue. We therefore examined this response in a 2D stretch system on healthy human FLS, prior to testing in the chip environment. Isolated hFLS were cultured on collagen type I coated elastomeric membranes and subjected to cyclic tensile strain at 2% or 12% strain for 2–24 h. We found that after 24 h synoviocytes with strain began to detach from the membrane even at the lowest strain levels (data not shown), therefore a shorter loading regime of 2 h was used. Actin stress fibre formation was observed in hFLS subjected to 12% strain while only faint actin labelling was observed under both control (0% strain) and 2% strain conditions (figure 4(A)). Significant upregulation of HAS-1 (p = 0.0087) and HAS-3 (p = 0.0112) were observed in response to 12% strain only (figures 4(B) and (D)). HAS-2 gene expression was found to be significantly downregulated by 2% strain (p = 0.0013), but not 12% strain (figure 4(C)). Gene expression of HAS-2 and PRG-4 (encoding lubricin) were not significantly altered by the application of strain (figures 4(C) and (E)).

Zoom In Zoom Out Reset image size

We then applied these findings to co-culture organ-chips for further optimisation. In unstrained chips, actin labelling of hFLS was weak, with little definition of stress fibre formation, and cells had a strong tendency to align along the length of the channel within a loose network (figure 4(F)). Upon the addition of 12% strain, this cell alignment was disrupted, and cells stained more intensely for actin, although stress fibre formation was not easily observable under the culture conditions (figure 4(F)). There was no significant difference in barrier permeability to 3 kDa or 70 kDa dextran in response to strain (figures 4(G) and (H)). Consistent with our observations of increased HAS gene expression in response to stretch, we observed a trend towards an increase in HA concentration in the conditioned media from the synovium channel in response to strain, but this was not statistically significant (data not shown).

The application of IL−1β to hFLS was used to simulate synovial inflammation. The response to IL−1β was initially examined in 2D on tissue culture plastic and, subsequently, in the presence of strain. Isolated hFLS were subjected to IL−1β treatment 0–10 ng ml⁻¹ for 24 h to identify an optimal dose for use in subsequent studies, and culture supernatant examined for the presence of inflammatory signalling molecules. No significant alterations in the release of nitric oxide (NO) were observed in response to IL−1 (figure S3(A)). However, significant release of PGE₂ was observed at all concentrations of IL−1β compared to the control condition, and release of IL−6 observed from 1 ng ml⁻¹ (figures S3(B) and (C)). Significant release of the catabolic enzyme MMP-1 was also observed from 1 ng ml⁻¹ IL−1β (figure S3(D)). No significant effects on cell viability were observed but the FLS began to look stressed morphologically upon the addition of 10 ng ml⁻¹ (data not shown). For this reason, an IL−1β dose of 1 ng ml⁻¹ was used in subsequent experiments. Gene expression was also analysed following the addition 1 ng ml⁻¹ IL−1β to 2D hFLS cultures. There was a trend towards increased expression of catabolic markers ADAMTS-4, ADAMTS-5, and MMP-3 while MMP-1 expression was significantly increased (p = 0.0006, figure 5(B)). Moreover, expression of MMP-13 was observed in cells treated with IL−1β but not untreated cultures (not shown).

Zoom In Zoom Out Reset image size

The hFLS response to 1 ng ml⁻¹ IL−1β was then explored in the presence of 12% cyclic tensile strain, using the optimised regime based on the previously mentioned experiment. There was a significant upregulation of PGE₂ in the presence of IL−1β in both the unstrained (+IL−1β: 2834.59 pg ml⁻¹ ± 3272.13_SD vs −IL−1β: 22.83 pg ml⁻¹ ± 19.72_SD; p < 0.0001) and strained (+IL−1β: 2871.57 pg ml⁻¹ ± 2706.58_SD vs −IL−1β: 42.96 pg ml⁻¹ ± 56.81_SD; p < 0.0001) conditions (figure 5(G)). Similar trends were observed in the secretion of IL−6, in response to IL−1β addition, in the strained (+IL−1β: 17 774.44 pg ml⁻¹ ± 9469.82_SD vs −IL−1β: 118.48 pg ml⁻¹ ± 3.14_SD; p < 0.0001) and unstrained conditions (+IL−1β: 17 472.40 pg ml⁻¹ ± 9603.14_SD vs −IL−1β: 114.43 pg ml⁻¹ ± 0_SD; p < 0.0001; figure 5(H)). MMP-1 was similarly upregulated in the presence of IL−1β in the strained (+IL−1β: 3914.25 pg ml⁻¹ ± 4007.83_SD vs −IL−1β: 31.15 pg ml⁻¹ ± 32.12_SD; p < 0.0001) and unstrained conditions (+IL−1β: 3079.46 pg ml ⁻¹ ± 3270.75_SD vs −IL−1β: 71.55 pg ml⁻¹ ± 107.40_SD; p < 0.0001; figure 5(I)). Interestingly, MMP-1 secretion was further upregulated in the presence of strain and IL−1β, compared to IL−1β alone (+IL−1β, +strain: 3914.25 pg ml⁻¹ ± 4007.83_SD vs +IL−1β, −strain: 3079.46 pg ml⁻¹ ± 3270.75_SD; p = 0.0465; figure 5(I)). NO secretion was significantly greater with the application of strain, both in the presence of IL−1β (+IL−1β, +strain: 9.00 µM ± 12.83_SD vs +IL−1β, −strain: 2.40 µM ± 5.45_SD; p = 0.0207) and in the absence of IL−1β (−IL−1β, +strain: 17.89 µM ± 24.78_SD vs −IL−1β, −strain: 9.28 µM ± 11.46_SD; p = 0.0256),although this was largely due to the increased secretion detected in the hFLS of one of the three donors (red; figure 5(F)). Gene expression analysis of hFLS in the presence of CTS also revealed similar trends following the addition of IL−1β, with upregulation ADAMTS-4 and -5, MMP-3 and MMP-1, in which the difference was statistically significant (p = 0.0023; figure 5(B)).

The response of hFLS to inflammatory stimuli in the vascularised synovium-on-a-chip was then examined by the application of 1 ng ml⁻¹ IL−1β to the top channel for three days from day 7. There was no observable difference in cell distribution or actin structure after 72 h of IL−1β treatment (figure 6(A)). After 24 h, there was a significant increase in barrier permeability to 3 kDa dextran. Studies in monoculture chips (hFLS only or HUVECs only) suggested that this response was due largely to alterations in the vascular channel, with a similar reduction in barrier function observed in HUVECs monoculture chips at 24 h, which then recovered (figure S4). Consistent with this finding, VE cadherin labelling was found to be more variable in some regions after 72 h of IL−1β treatment, and cell number was reduced. Interestingly, permeability to 70 kDa dextran was also increased and remained high at 48 h post stimulation with IL−1β. However, this response was not seen in HUVECs monoculture chips, suggesting that the effect was mediated, at least in part, by the hFLS. A functional barrier was restored by 48 h with respect to 3 kDa dextran, however chips remained permeable to 70 kDa dextran until 72 h post application of IL−1β.

Zoom In Zoom Out Reset image size

The secretion of the aforementioned pro-inflammatory and catabolic mediators, as well as the SF constituent, HA, were assessed in the conditioned media collected from the hFLS channel at 24 h, 48 h and 72 h after the addition of IL−1β. There were no significant differences observed in terms of NO release in the presence or absence of IL−1β (figure 6(B)). At all three timepoints, increased secretion of PGE₂ was observed in the presence of inflammatory stimuli, which was statistically significant at 72 h (+IL−1β: 11 207.86 pg ml⁻¹ ± 4427.86_SD vs −IL−1β: 22.52 pg ml⁻¹ ± 26.05_SD; p = 0.0039; figure 6(C)). The secretion of IL−6 was significantly higher in the IL−1β treated chips compared to the control chips at 24 h (+IL−1β: 13 735.70 pg ml⁻¹ ± 7205_SD vs −IL−1β: 177.34 pg ml⁻¹ ± 136.92_SD; p = 0.0004), 48 h (+IL−1β: 12 677.76 pg ml⁻¹ ± 7555.83_SD vs −IL−1β: 160.3 pg ml⁻¹ ± 44.05_SD; p = 0.0009) and 72 h (+IL−1β: 9156.36 pg ml⁻¹ ± 4626.15_SD vs −IL−1β: 153.49 pg ml⁻¹ ± 0_SD; p = 0.0318). At 24 h, the secretion of MMP-1 was significantly lower in the control chips (2347.33 pg ml⁻¹ ± 1648.66_SD) compared to in those to which IL−1β was added (3792.27 pg ml⁻¹ ± 1551.08_SD; p = 0.0176), but this difference was lost at the later timepoints (figure 6(E)). The secretion of HA was also stimulated in the synovium chips by the addition of IL−1β and was found to be higher in the IL−1β stimulated chips compared to the control chips at all three timepoints, with statistical significance at 48 h (+IL−1β: 1820.36 ng ml⁻¹ ± 936.06_SD vs −IL−1β: 377.86 ng ml⁻¹ ± 0_SD; p= 0.0249) and 72 h (+IL−1β: 2019.56 ng ml⁻¹ ± 831.01_SD vs −IL−1β: 377.86 ng ml⁻¹ ± 0_SD; p = 0.01; figure 6(F)).

The expression of ICAM-1 was increased within the endothelial channel of synovium-on-a-chip, and labelling intensity was significantly higher, in IL−1β treated chips versus untreated control chips (figures 7(E) and (F)). No positive labelling of ICAM-1 was observed in the monoculture inlet region of the chip where there was no mixing between synovium and vascular channels (figure S5), suggesting that IL−1β, or one of its downstream mediators is likely responsible for this effect.

Zoom In Zoom Out Reset image size

To determine if the activated endothelium (figures 7(E) and S5) is functionally relevant in synovium-on-a-chip, THP-1 monocytes were introduced to the bottom channel of the chip. Monocytes were labelled with cell tracker and incubated in the bottom channel under static conditions for 3 h before the channel was flushed with fresh medium at 1000 µl h⁻¹. Few cells attached to the surface of the endothelial channel in untreated chips, however in IL−1β treated chips, attachment was significantly increased by 16-fold (p < 0.0001; figures 7(G) and (H)).