Rapid manufacturing of flexible microstructured PDMS substrates, using 3D DLP printing technique, for flexible pressure sensors

Rapid manufacturing of flexible microstructured pdms substrates, using 3d DLP printing technique, for flexible pressure sensors

Florian PISTRITU , Mihaela CARP , Violeta DEDIU , Catalin PARVULESCU , Marian VLADESCU6, Paul SCHIOPU

In this work, we have carried out research on the microstructured substrates obtained with molds made by the 3D DPL printing technique, in order to obtain a microstructured substrate with maximum displacement. Microstructured PDMS and PDMS/aerogel substrates were tested. Compression tests were performed at 80N, 100N and 120N force The PDMS pyramid-type microstructured substrates, having the side of the base and the height of the pyramid of 1500µm x 1060µm, respectively 2000µm x 1414µm, obtained the best value for displacement. The better results obtained make the PDMS/aerogel composition appropriate to be used as sensitive elements/membranes in pressure sensors.

 Keywords: flexible substrates, microstructured PDMS, 3D printing technique, rapid manufacturing, flexible pressure sensor

We kindly thank the researchers at IMT Bucharest for this collaboration, and for sharing the results obtained with their system.

Introduction

Flexible pressure sensors are mainly used in robotics and medicine. These sensors have found applicability in various fields such as displays [1], robotics [2, 3, 4, 5], human pulse waveform [6,7,8, 9], very sensitive pressure detection [3, 8, 10], voice recognition [10], gas flow monitoring [3,8,10, 11], human-machine interface technologies [3,4,5,9], foot pressure [3]. In the field of medicine, the most used detection methods, of a pressure sensor, are based on the piezoresistive and capacitive effect [12]. The piezoresistive detection method is based on the piezoresistive effect and consists in the conversion of the deformation of a material into a variation of the resistivity, which can be measured. Depending on the application chosen for the pressure sensor, the most important parameters of the pressure sensor is also established, such as response time, sensitivity, measurement range, elasticity, bending resistance, transparency, and cost.

For a low-price method of obtaining a pressure sensor, inkjet printing technology can be used. A pressure sensor made through this technology involves the integration of a flexible substrate with an elastomeric substrate. The flexible substrate can be PET, Kapton or something similar on which it is deposited by inkjet printing, a resistor. Among the materials used for the production of the elastomeric substrate are polydimethylsiloxane (PDMS) [7, 13] and Ecoflex [14], polyethylene terephthalate (PET) [15], polyethylene (PE), polyurethane (PU), polyimide (PI) [16], and others.

In this work, we conducted research on the microstructured substrates obtained with molds made by the 3D printing technique. The duration to obtain a microstructured substrate, through this method, is relatively short (from the 3D CAD modeling to obtaining the microstructured model from PDMS: 4 hours), offering the possibility of rapid modification of microstructure configurations. The purpose of this research is to obtain a microstructured substrate with as much displacement as possible. Integrating this microstructured substrate into a pressure sensor, we have the possibility to measure high pressures. For the realization of the microstructured substrate, Fig. 1 shows the stages from concept to test bench. In the first stage, E-I, we must have software for creating 3D CAD mold models, installed on a PC. Stage II, E-II, includes the creation of 3D CAD models of the molds. The transfer of these 3D CAD models of the molds to the 3D printer and their printing represents Stage III, E-III. After 3D printing, in Stage IV, E-IV, we have the treatments applied to the obtained 3D molds. In Stage V, E-V, we obtain the microstructures from PDMS using the molds obtained in the previous stages. In the last stage, Stage VI, E-VI, we will test these structures with the Mecmesin MultiTest 2.5 i device. When creating the 3D CAD model of the mold, we took into account the type of printer used, CADworks3D µMicrofluidics M50. The resin used to make the molds is Master Mold Resin for PDMS devices. Thus, having the printed mold, we investigated the displacement for microstructured structures from PDMS, but also PDMS/aerogel in two ratios.

Materials and Methods

 2.1 Molds fabrication - Desing and printing of 3D molds

The elastomeric layer was made from PDMS, this being a silicone rubber used in the field of electronic devices. PDMS is a low-cost, low hardness elastomer, it’s a biocompatible material [17, 18], preferred for medical applications.

Several studies on the geometric variation of the microstructured PDMS layer are published in the specialized literature [20]. This microstructured layer is based in most cases on micro-cylinders, micro-pyramids, micro-domes [8]. The analyzes carried out by several authors, on different types of structural models, have shown superiority in terms of sensitivity [19] of micro-domes and micropyramid structures in relation to other structures.

From the more detailed analysis carried out in 2018 by Shuangping Liu and Monica Olvera de la Cruz [22], we observe that the thickness of the base of the micropyramids in the elastomeric layer does not have a significant influence, so we eliminated this parameter from the analysis of the structural models.

Seven 3D CAD mold models were made for the fabrication of pyramidtype microstructured substrates, and one mold model for obtaining a parallelepiped-type microstructured substrate. The general rule for all 8 types of molds was that the size of the base of the microstructure should be equal to the distance between the microstructures in all directions. Pyramid microstructures were made with base side x height of: 200µm x 141µm (P200), 350µm x 247µm (P350), 500µm x 353µm (P500), 750µm x 530µm (P750), 1000µm x 707µm (P1000), 1500µm x 1500µm (P1500), 2000µm x 1414µm (P2000). The microstructures of the parallelepiped type had a base side of 500 µm and a height of 353 µm (D500). Fig. 2 shows the 3D CAD models of the molds.

The stages of making the 3D mold are: 1) Design a 3D CAD model of the mold. The software used is FreeCAD. Execution time: 20 minutes; 2) Preparing the 3D printer for making the mold. The 3D printer used is CADWORKS3D - µMicrofluidics M50; 3) Transferring the file with the mold model from the PC to the 3D printer. The software used for the transfer is Utility 6.0. 4) Making a 3D mold. The resin was used: Master Mold for PDMS Devices – 3D Printing Resin Photopolymer Resin (Composition: Methacrylated oligomer, Methacrylated monomer, Photoinitiator & Additives). Time needed to print: approximately 15 minutes; 5) UV treatment for the printed model. After printing, the mold was cleaned in isopropyl alcohol (Isopropyl Alcohol- IPA, concentration 99.9%) twice for one minute. After cleaning in alcohol, they were dried. Then a UV treatment was performed for 20 minutes for each of the 2 faces.

2.2 Realization of the microstructured layer from PDMS

After obtaining the 3D CAD molds, the next import step is the choice of the elastomeric material to be poured into the obtained molds. Two types of PDMS were used: Sylgard 184 from DOW Chemical Company and KER 4690 from Shin-Etsu Chemical Company. These two types of PDMS are different both in terms of the modulus of elasticity and the method of exposure for curing. The steps for obtaining the microstructured PDMS substrate are: 1) mold cleaning with isopropyl alcohol IPA and drying with a nitrogen gun; 2) obtaining PDMS mixture. To prepare Sylgard 184, the mixing ratio between polymer and hardening agent is 10:1 [21], being one of the best mixing ratios. Homogenization time: 20 minutes. To prepare KER 4690, I mixed the two components KER 4690- A and KER 4690-B in a 1:1 ratio for 20 minutes.; 3) degassing PDMS mixture: for 45 min; 4) PDMS deposition on the mold; 5) Treatment for curing.

The curing treatment for Sylgard 184 consisted of exposure to a temperature of 100 C for 50 minutes, and for KER 4690 it consisted of exposure to UV radiation.

The 3D CAD models of the pyramid-type and parallelepiped-type microstructured substrate can be seen in Fig. 4 and Fig. 5

2.3 Realization of the microstructured layer from PDMS/aerogel

The PDMS used was Sylgard 184 from DOW Chemical Company. The aerogel used is powder aerogel (Powder aerogel <0.125 mm - Green Earth Aerogels).

The steps for obtaining the microstructured substrate from PDMS/aerogel are: 1) mold cleaning with IPA isopropyl alcohol and drying with a nitrogen gun; 2) obtaining the PDMS mixture. For the preparation of Sylgard 184, the mixing ratio between polymer and hardening agent is 10:1, for 20 minutes; 3) Adding to the obtained PDMS a quantity of 5% or 10% of powder aerogel, and mixing for homogenization for 15 minutes; 4) degassing PDMS mixture for 45 min; 5) depositing the PDMS/aerogel mixture on the mold; 5) Treatment for hardening at a temperature of 100°C for 50 minutes.

Materials

Master Mold Resin

M Series

Results

14 types of microstructured PDMS substrates and 4 types of microstructured PDMS/aerogel substrates were tested. Compression tests were performed at 80N, 100N and 120N. In Table 1, you can see a comparison of several parameters for the two types of PDMS used.

The characterization of all microstructured substrates was carried out on the Mecmesin MultiTest 2.5i device. With the help of this device, we analyzed the compression of the microstructured substrates when applying a force of 80N, 100N, and 120N, at a compression speed of 1mm/min. Compression tests were performed using Emperor Force software. At the end of the test, the software generates an analysis report.

 In Table 2 you can see the results when applying a force on a type of microstructured substrate and the displacement obtained.

In Tables 3 and 4 you can see the results obtained for different types of micro-pyramidal substrates when applying a force of 100N and 120N.

In Table 5, the results obtained from the compression of the pyramidaltype microstructured substrates made of both PMDS and PMDS/aerogel are presented.

The tests performed at a compression of up to 80N showed that the microstructured substrates made of PDMS Sylgard 184 have a greater displacement than those made of PDMS KER 4690

The highest displacement values were obtained for the micromicrostructured P1000, P1500 and P2000 substrates, which is why we performed the tests by increasing the compression force to 100N. The displacement obtained when applying a force of 100N on the substrates P1000, P1500 and P2000, can be seen in the diagram from Fig. 6.

According to the obtained results, it appears that the micromicrostructured substrates P1500 and P2000 have the best movement. Additional tests were performed only with these two types of structures, at a compression force of 120N. The results are roughly equal, with a slight edge for the P2000. By introducing aerogel into the PDMS composition, the results showed higher obtained values. The displacement obtained during compression for the microstructured substrates made of PDMS and PDMS/aerogel can be seen in the diagram in Fig. 7.

Conclusion

We conducted research on microstructured substrates obtained with molds made by the 3D DLP printing technique, to show if the obtained substrates are suitable for use as sensitive elements/membranes in pressure sensors. Making the molds according to their 3D CAD models showed the simplicity and speed in obtaining the molds. With these molds we made 4 types of microstructured substrate. The best results were obtained for the substrates obtained from PDMS/aerogel. The large difference between the displacements occurring upon the addition of aerogel makes the obtained pyramid-type microstructured structures suitable for use as sensitive elements/membranes in pressure sensors.

In the future, we will use the obtained substrates to make a flexible pressure sensor

Controlling bead and cell mobility in a recirculating hanging-drop network

Controlling bead and cell mobility in a recirculating hanging-drop network

Nassim Rousset , Martina de Geus , Vittoria Chimisso , Alicia J. Kaestli , Andreas Hierlemann  and Christian Lohasz

Integrating flowing cells, such as immune cells or circulating tumour cells, within a microphysiological system is crucial for body-on-a-chip applications. However, ensuring unimpeded recirculation of cells is a significant challenge. Closed microfluidic devices have a no-slip boundary condition along channel walls and a defined chip geometry (laminar flow) that hinders the ability to freely control cell flow. Open microfluidic devices, where the bottom device boundary is an air–liquid interface (ALI), e.g., hanging drop networks (HDNs), offer the advantage of an easily-actuatable fluid-phase geometry, where cells can either flow or stagnate. In this paper, we optimized a hanging-drop-integrated pneumatic-pump system for closed-loop recirculation of particles (i.e., beads or cells). Experiments with both beads and cells in cell culture medium initially resulted in particle stagnation, which was suggestive of a pseudo-no-slip boundary condition at the ALI. Transmission electron microscopy and dynamic light scattering measurements of the ALI suggested that aggregation of submicron-scale cell-culture-medium components is the cause of the pseudo-no-slip boundary condition. We used the finite element method to study the forces on particles at the ALI and to optimize HDN design (drop aperture) and operation (drop height) parameters. Based on this analysis, we report a phase diagram delineating the conditions for free flow or stagnation of particles at the ALI of hanging drops. Using our experimental setup with 3.5 mm drop apertures, we conducted particle flow experiments while actuating drop heights. We confirmed the ability to control the flow or stagnation of particles by actuating the height of hanging drops: a drop height over 300 μm led to particle stagnation and a drop height under 300 μm allowed for particle flow. This particle-flow control, combined with the ease of integrating scaffold-free organ models (microtissues or organoids) in HDNs, constitutes the basis for an experimental setup enabling the control of the residence time of single cells around 3D organ models.

We kindly thank the researchers at ETH Zürich for this collaboration, and for sharing the results obtained with their system.

Introduction

Designing microfluidic devices for cell culturing, especially multi-tissue cultures, has led to approaches that interconnect 2D or 3D cultures of different cell types (tumour, brain, liver, heart, etc.) through microchannels in a physiologically relevant combination and ratio.1 These microphysiological systems (MPSs) are often considered the next step in preclinical research toward more comprehensive and physiologically relevant in vitro testing systems.2,3 The interest in MPSs is mainly based on their potential to better predict the effect of compounds on processes in the human body4,5 and to better understand – in a more systemic way – how different healthy and diseased organs interact with each other3,6,7 when compared to traditional preclinical in vitro models. The potential applications of MPSs include pharmaceutical research and compound testing,8 basic research on tissue and cell interaction,9 and disease progression studies.10

One of the current challenges for MPS applications is the interaction between solid tissues and suspended cells, e.g., circulating tumour cells or immune cells. Such interaction studies are particularly interesting to, for example, mimic immunotherapeutic approaches,9 and recapitulate the interaction dynamics of circulating tumour cells11 and immune cells12 with other organs. Some strategies rely on a static interaction between cell suspensions surrounding solid tissues,13 ignoring the physiological behaviour of immune cells that migrate toward and around their target.14 Hydrogel-based approaches can generate stable signalling gradients that may guide the migration of immune or tumour cells.15 However, these approaches ignore the circulatory nature of immune cells that move around due to blood flow.

To enable the interaction between suspension cells and a series of immobilized tissue constructs, a liquid-phase transport system is needed. However, emulating a circulatory system with microfluidics is not trivial, as it requires flowing single cells that interact with a static organotypic tissue model over several days. Furthermore, a closed-loop recirculation of cells is crucial for the build-up of relevant concentrations of signalling molecules, e.g., cytokines and chemokines, and appropriate tissue/suspension cell interaction.9 Recent advances in achieving cell recirculation have been demonstrated,9,16 but have yet to meet the requirement of maintaining stable and long-term cell recirculation. The requirements of efficient cell recirculation are (i) minimizing cell/microfluidic structure interactions, (ii) minimizing cell stagnation, and (iii) minimizing cell agglomeration in larger chambers that are used to host tissue models.

Open microfluidic systems – such as hanging-drop networks (HDNs) – are particularly suited to meet the requirements detailed above.17 HDNs feature hanging drops, interconnected through microfluidic channels (Fig. 1a). Tissue models can be immobilized and cultivated within the individual drops, while fluid flow through the channels is used to establish inter-tissue communication through various signalling molecules. A key feature of such open systems is an air–liquid interface (ALI). ALI in this manuscript does not refer to epithelial cells exposed to air but to the interface between an air and a liquid phase. The ALI largely reduces the interaction between cells and microfluidic channel structures – e.g., SU-8 or polydimethylsiloxane (PDMS) – and that allows for direct optical access to the tissue and cell models with an inverted microscope (Fig. 1b). Additionally, the ALI in open microfluidic systems provides ample oxygenation, which reduces the risk of hypoxia-related cytolytic and migratory activity of immune cells, as well as cell death.18 An open microfluidic system also enables free liquid flow, where no stress is present at the ALI. The no-stress ALI boundary gives full control over the drop height (Fig. 1b) during an experiment. Free liquid flow also ensures continuous cell flow through the system due to the slip boundary condition (Fig. 1c) at the ALI. In contrast, free cell flow is not guaranteed within closed microfluidic systems, where the no-slip boundary condition at channel walls and rigid structures reduces the flow velocity (flow velocity is null at the walls), which gives only little freedom with chip operation and causes cell aggregation upon recirculating cells (Fig. 1d).

Fig. 1 Cell-mobility control scheme. (a) Schematic representation of a hanging-drop network (HDN) featuring four hanging-drop structures with in- and outlet drops at either end. (b) Colinear-to-flow cross-section of a typical hanging-drop compartment unit of an HDN. A hanging drop is connected to the network through microfluidic channels. The aperture (2a) of the drop is a design-defined constant. The height (h) of the drop can be controlled during an experiment. The air–liquid interface (ALI) is the key feature of HDNs, giving a slip boundary condition. (c) Visualization of the flow-velocity profile through an open microfluidic system. No stress is present at the ALI, which results in a slip boundary condition, where the flow velocity is maximal at the ALI. The slip boundary condition allows for unimpeded cell (orange) flow over time, even as cells settle due to gravity. (d) Visualization of the flow velocity profile through a closed microfluidic system. The no-slip boundary condition, caused by stiction of the outermost liquid-phase layers to the channel boundaries, sets the flow speed to zero. This may cause cells to stagnate and stick to the channel wall surface, particularly at the bottom where they settle due to gravity.

An HDN enabling closed-loop recirculation of fluids has been developed and validated in our laboratory.19 A unidirectional flow was achieved with an integrated-pump concept developed specifically for microfluidic HDNs. Here, we show a novel iteration of the device, which is aimed at flowing cells in a closed loop and controlling their interaction time with microtissues.

Preliminary tests with beads showed an unexpected behaviour. The slip boundary condition allowed for successful bead recirculation within de-ionized water. However, we observed a no-slip-like stagnation of particles (cells and beads) during recirculation within cell culture medium.

Mathematical modelling of the ALI as a slip or no-slip boundary with the finite element method (FEM) helped to explain this unexpected no-slip-like stagnation. The FEM is a widely used tool to model, predict, and characterize fluid dynamics within microfluidic chips. This modelling technique allows for computing hydrodynamic forces on spherical objects.20–22 We computed the forces on particles at the ALI, while varying operational parameters – e.g., drop height – and design parameters – e.g., drop aperture – for a set of defined experimental conditions.

The aim of this study was to investigate the forces on and behaviour of beads at the ALI as a surrogate for cell behaviour, which we thereafter confirmed experimentally with cells. Our goal was twofold: on the one hand, we wanted to find conditions where particles can freely recirculate within our device, despite the unexpected stagnation; on the other hand, we wanted to control particle stagnation in order to modify particle residence time in the hanging drop at will. Our theoretical and experimental findings suggest that, although the no-slip behaviour of liquid at the ALI is anomalous in cell culture media, careful experimental design can still enable unimpeded particle flow.

Experimental

 Flowing particles

The flowing beads were 8.0 ± 0.1 μm-diameter and 1.05 g cm−3-density polystyrene beads (Sigma-Aldrich, Buchs, Switzerland). They were suspended in de-ionized water or cell culture medium depending on the experiment.

The flowing-cell model used here was THP-1 (TIB-202; ATCC, Manassas, VA, USA), a human acute monocytic leukaemia cell line. THP-1 cells were cultured according to ATCC protocols and maintained in RPMI-1640 (PAN-Biotech GmbH, Aidenbach, Germany), supplemented with 10% foetal bovine serum (Sigma-Aldrich, Buchs, Switzerland) and 1% penicillin and streptomycin (Sigma-Aldrich, Buchs, Switzerland). The cell culture medium was filtered through a 0.2 μm-pore-sized filter (Thermo Fisher Scientific, Waltham, MA, USA) to ensure fibre-, aggregate-, and contaminant-free culture and microfluidic HDN operation. Cells were cultured in non-adherent flasks (Greiner Bio-One, Frickenhausen, Germany) at 37 °C, 5% CO2, and 95% humidity. Cells were subcultured every 2 to 3 days at a ratio of 1[thin space (1/6-em)]:[thin space (1/6-em)]5 to maintain a density of 0.2–1 × 106 cells per mL. Cell-culturing density was kept at these levels to ensure spherical cell morphology, because higher densities were shown to alter cell morphology.23

 Device fabrication

The fabrication process of the microfluidic device was identical to that of our previously published device19 with two PDMS (Sylgard 184, Dow Corning GmbH, Wiesbaden, Germany) layers – a microfluidic and a pneumatic layer – and a glass substrate to ensure device stability. A micrograph of the fabricated device is shown in Fig. 2a and b. A schematic illustration of the chip layout is shown in Fig. 2c–e.

Fig. 2 Hanging-drop network microphysiological system with on-chip pump drops for particle recirculation.

Picture of the recirculating hanging-drop network (HDN) filled with blue ink showing (a) the air–liquid interface and (b) the transparent slide with inlets and a digitally inserted highlight of pneumatic chambers. The fabricated chip consists of 2 PDMS layers: (c) a 750 μm-thick microfluidic network layer and (d) a 5 mm-thick pneumatic channel layer. (c) Schematic of the microfluidic layer of the recirculating HDN. A 2 × 3 on-chip pump drop setup, highlighted in orange at the top, increased the flow rate within the device. Four culture drops, highlighted in green at the bottom, enabled flowing-cell analysis and could accommodate tissue co-culturing. The sample inlet enabled the introduction of a cell suspension to the device with minimal chip handling. The height-control inlet enabled the in- and outflow of liquid with a syringe pump to maintain the drop height or actuate it to the desired value. (d) Schematic of the pneumatic layer of the recirculating HDN. The heigh-control, sample, and pneumatic inlets are 0.75 mm-diameter holes. The height-control and sample inlets were punched through to the microfluidic layer. Three pneumatic inlets control three separate pairs of on-chip pump drops (highlighted in white and red in b and in orange in c). (e) Side-view cross-section A (dashed line in c and d) of the device with a hanging drop (dimensions in μm). The side view shows that a 250 μm-thick PDMS layer (part of the microfluidic layer) seals the pneumatic layer. The microfluidic and pneumatic layers were plasma-bonded together (A-labelled dashed line in e). The pneumatic layer was bonded onto a transparent slide (not shown on cross-section) to ensure chip rigidity and optical access to the drops. Upon pressurizing the pneumatic chambers (red-labelled height), the 250 μm thick PDMS layer (black-labelled height) expands into the volume of the drop below. The red-coloured pneumatic chambers were actuated simultaneously and in an alternating pattern with the white-coloured pneumatic chambers. Combined with the integrated valves (yellow structures in c and e), the pumping produced a unidirectional flow (blue arrows in c). The positions of the integrated valves defined the flow direction. All interconnecting channels were 300 μm wide. All microfluidic structures were 500 μm high (blue-labelled height in e).

For the microfluidic layer (Fig. 2c), two masks were used to generate an SU-8 (Microchem Corp., Newton, MA, USA) microfluidic-channel master mould on a silicon wafer. We used a 7[thin space (1/6-em)]:[thin space (1/6-em)]1 PDMS-to-curing agent ratio to ensure stiff and reliable valve operation. For the pneumatic layer (Fig. 2d), one mask was used to generate the cavity master mould on a silicon wafer. We used a normal 10[thin space (1/6-em)]:[thin space (1/6-em)]1 PDMS-to-curing agent ratio for the pneumatic layer. Alternatively, with the aim to simplify the fabrication process, we 3D-printed a mould design identical to the one previously fabricated by photolithographic processes. We used a 3D printer designed for microfluidics (PR110-385, CADWorks3D, Toronto, Canada) with the dedicated “PDMS Mastermold” resin (CADWorks3D, Toronto, Canada) to 3D-print the mould. We used a high-intensity UV post-curing solution (Professional CureZone, Creative CADWorks, Concord, ON, Canada) to ensure proper curing of the resin.

For chip assembly, 0.75 mm diameter inlets were first punched in the pneumatic layer. After aligning and bonding both PDMS layers, 0.75 mm diameter inlets were punched for the sample and height-control drops through to the microfluidic layer. One of two processes was used to bond the resulting PDMS chip onto a supporting slide with 1.2 mm-diameter holes matching the chip inlets, which provided a stiff substrate to suspend the chip in a hanging-drop configuration. Process 1: following plasma activation, we aligned and bonded the PDMS chip onto a glass slide with drilled holes. Process 2: using double-sided tape, we bonded the PDMS chip onto an acrylic slide with laser-cut holes.

Design changes with respect to our previously published device19 included (i) the reduced pitch between culture drops to minimize travel time of the cells and (ii) the addition of height-monitoring drops (Fig. 2c). Two parallel and 3-pumping-drop configurations were designed to increase the flow rate in the device.

Materials

Clear Microfluidics Resin V7.0a

H Series

Pr Series

 Flow-rate measurement

Measuring the flow rate was done by imaging the circulating beads in the channel between the height-monitoring drops and the height-control-inlet. Videos were taken at 5 frames per second, and particles were traced manually. The flow rate was calculated by multiplying the average velocity of particles in the channel by the channel cross-section of 0.3 × 0.5 mm2. All experimental images, time lapses and videos were acquired using an inverted wide-field microscope (Leica DMI6000B, Leica Microsystems, Switzerland) with a 10× lens.

 Mathematical models

The finite-element method software, COMSOL Multiphysics® v. 5.4 (COMSOL AB, Stockholm, Sweden) was used to model a single hanging drop (Fig. 1b). A full 3D model of the liquid phase of a hanging drop was established using the laminar-flow module to solve the Navier–Stokes equation. The built-in physical properties of water were used for the liquid phase. The boundary conditions were set to “no slip” for the PDMS-water interfaces, constant “inflow” for the channel inlet, constant null “pressure” for the channel outlet, and “slip” (Fig. 1c) or “no slip” (Fig. 1d) for the ALI. We placed and moved, via a parameter sweep, a small particle at the ALI. Parametric sweeps were used to find the flow velocities and pressures for every modelled condition. We elaborate on the sweeping strategy in the “Particle-flow modelling strategy” section of the results. The hydrodynamic force on spherical particles at the ALI was computed by using the built-in reaction force (reacf) operator within COMSOL and summing it over the surface area of the spherical particle.

 Transmission electron microscopy

Transmission electron microscopy (TEM) was used to image the nanostructures present at the ALI and in the medium. TEM images were acquired with a Philips Morgagni 268D microscope. Each sample was deposited on a carbon-coated copper grid by gently touching the ALI with the flat side of the TEM grid. The samples were blotted, washed with MilliQ water and negatively stained with 1% phosphotungstic acid (PTA). TEM images were acquired with an acceleration of 90 kV, and each grid was imaged at three different sites.

 Dynamic light scattering

Dynamic light scattering (DLS) was used to determine the hydrodynamic radius (DH) of the particles present in the cell medium. The DH was calculated from scattering data collected using the Zetasizer Nano ZSP DLS measurement system (Malvern Panalytical, Volketswil, Switzerland).

The samples were prepared by transferring the content of four hanging drops to 100 μl of MilliQ water, and then analysed. Measurements were performed at 20 °C with non-invasive backscattering (NIBS) technology. Results were directly processed and displayed by the built-in software (Zetasizer software).

Results

 Particle-flow characterization

To achieve free flow of suspended particles through an open microfluidic system, we used an HDN composed of a sequence of drops, connected by channels, where each drop acted as a potential tissue compartment (Fig. 2a and b). Perfusion in a closed-loop system was induced by pneumatically actuating integrated on-chip pump drops (Fig. 2c–e) developed in our laboratory.19 Unidirectionality of the flow was achieved with integrated valves actuated through pneumatic inlets (Fig. 2c). This strategy reduced the large dead volume of – and prevented repeated squeezing of cells by – the peristaltic pump mechanism. Such integrated pneumatic pumps have a maximum possible flow rate that is defined by the microfluidic network design.19 A bead or cell suspension can be loaded into the system through the sample inlet in Fig. 2c to be recirculated within the HDN. The drop height and device level can be precisely monitored through height-monitoring drops (Fig. 2c) that were loaded with thin rings.24,25 Height-monitoring drops are positioned at the periphery of the device and fluidic system, thereby allowing to fine-tune the drop height at micrometre-precision prior to and during an experiment. Continuous drop-height control was achieved with an open-source “Droplet Based Microfluidics” software package implemented in YouScope,26 where the focal position of the aforementioned thin ring was kept constant by infusing or removing medium through the height-control inlet (Fig. 2c).

The chip shown in Fig. 2 was used to assess bead and cell mobility in the HDN. The experiments were conducted on an inverted microscope with an incubation box keeping the setup at 37 °C, 5% CO2, and 95% humidity. The chip was prefilled with either de-ionized water or culture medium. The pneumatic lines, set to an air pressure of 40 kPa, were connected to the pneumatic inlets (Fig. 2d) actuating the on-chip pump drops (Fig. 2c). The pump actuation protocol consisted of alternatively actuating the pneumatic inlets, highlighted in red and white (Fig. 2d), and was run for the duration of the experiment. The timing of the pump actuation protocol is ton = 200 ms, toff = 200 ms with a 200 ms delay between the red and white pneumatic valves. This scheme was previously optimized to achieve the highest flow rate possible of 1 μL min−1.19 The drop height of the height-monitoring drops was maintained at 550 μm, per previous protocols,19 by connecting a syringe of de-ionized water to the height-control inlet and by using our feedback controller26 as previously described.24 De-ionized water instead of cell culture medium for height-control was used to avoid changes in medium osmolality through evaporation of water and up-concentration of salts, proteins, and other molecules in the medium. At the start of every experiment, 10 μL of either polystyrene beads or THP-1 acute monocytic leukaemia cells were added to the sample inlet (Fig. 2c) with a seeding density of 0.7–7 × 106 particles mL−1. The effective flow rate of 1 μL min−1 was determined by measuring the average speed of single beads or cells through the channel connecting the height-monitoring drop and the height-control inlet.

First results shown in Fig. 3a indicated that beads in de-ionized water circulated continuously over the course of the experiment. The slip boundary condition at the ALI (Fig. 1b) allowed for rapid recirculation. However, cells and beads in cell culture medium settled to the bottom of the drop and stagnated within a few hours (Fig. 3b). This observed stagnation would be typical of a no-slip boundary condition (Fig. 1c). This behaviour was unexpected, considering that the density and viscosity of our cell culture medium did not deviate significantly from that of de-ionized water.27 This observed stagnation led to the hypothesis that the complex cell culture medium formulation (containing RPMI 1640 and 10% Foetal Bovine Serum – FBS), which includes various salts, amino acids, vitamins, nutrients, and other additives, may change the boundary conditions at the ALI to exhibit a behaviour featuring aspects of a no-slip boundary condition, i.e., a pseudo-no-slip boundary condition. As ALIs have been shown to accelerate protein crystallisation,28 we hypothesized that protein denaturation due to shear and the ALI29 may have caused this unusual hydrodynamic behaviour.

 Fig. 3 Observation of cells and beads within the microfluidic hanging drops at a recirculating flow rate of 1 μL min−1.

The drop aperture diameter was 3.5 mm, and the drop height was 550 μm. We highlight 4 particles in each snapshot (red, green, blue, and yellow dots) with their displacement over one frame marked up with arrows. (a) De-ionized water featured rapid bead movement at the ALI over 6 seconds. The snapshots of the bead flow were taken from a 16 h time lapse video. (b) and (c) Cell culture medium featured comparatively slow particle movement over 5 minutes as well as cell (b) and bead (c) stagnation at the bottom of the hanging drop. The snapshots of cell and bead stagnation were taken a few hours into a 19 h time lapse video, which suggested a significantly different hydrodynamic behaviour within the cell culture medium when compared to that in de-ionized water. The time difference between the first and second row was chosen to highlight a comparable displacement. The time difference of 6 seconds for the de-ionized water condition compared to 5 minutes for the cell culture medium condition indicates that particles at the bottom of the hanging drop have a slower movement, and a stronger tendency to form stagnation areas in cell culture medium than in de-ionized water, despite operating the chip with the same recirculating flow rate.

 Particle-flow modelling strategy

To investigate the pseudo-no-slip boundary condition, we modelled the forces acting on spherical particles (i.e., cells or beads) at the ALI. Hydrodynamic forces are the only ones that vary between flow in de-ionized water vs. cell culture medium. Therefore, computing the hydrodynamic forces on particles was key to studying cell and bead stagnation. To do so, we computed the flow in the hanging drop with a particle at a given location (Fig. 4a). We used the solution on the particle surface (Fig. 4b) to compute the hydrodynamic force.

 

 Fig. 4 Particle-flow modelling strategy. (a) and (b) show COMSOL simulation results with a slip boundary condition at the ALI. (a) Flow velocity profile (surface plot) and streamline solutions of the flow in the colinear-to-flow middle cut plane of the hanging drop. (b) Zoomed-in view of the square box in figure (a) showing a modelled particle of 8 μm diameter at the ALI. The hydrodynamic force is computed by summing the reaction forces of the flow on the surface (computed with the built-in reacf operator in COMSOL). (c) Schematic of the forces acting on particles at the ALI and (d) superimposition of the modelled net tangential force (white vector field) and its magnitude (coloured contour map) on the experimentally observed stagnation of beads at the bottom of the drop from Fig. 3c. The diameter of the drop in the image is 3.5 mm. The experimental drop height is 550 μm. The experimental flow rate is 1 μL min−1.

Particle free-body diagram. The free-body diagram of a particle (Fig. 4c) shows the forces that come into play when analysing free flow of particles at the ALI of a hanging drop. The externally acting forces are hydrodynamic, gravitational, and normal forces. The hydrodynamic force (Fhyd, green dashed vector in Fig. 4c) is caused by differentials in fluid pressure around the particle caused by the moving flow. The gravitational force (Fg, orange dashed vector in Fig. 4c) includes the weight of the particle and the buoyant force in medium. The normal force (F⊥, purple dashed vector in Fig. 4c) is derived from Newton's third law of motion and is the normal force of the ALI acting on the particle, stopping it from crossing the ALI.

We modelled Fhyd numerically by assuming a worst-case no-slip ALI boundary condition for the experimental results and conditions displayed in Fig. 3b and c. Fg was computed using the density of the particle and the density of the medium. The normal force of the particle on the ALI was computed by projecting the sum of hydrodynamic and gravitational forces on the normal vector of the particle-ALI contact point. F⊥ of the ALI on the particle is the negative of the previously computed force. F⊥ only exists if forces push the particle toward the ALI and is due to surface tension exceeding hydrodynamic forces. The net tangential force (F∥, black vector in Fig. 4c) is computed by adding all three external forces acting on the particle and is the force translating the particle along the hanging-drop surface.

The resulting net tangential force vectors (F∥, white vector field in Fig. 4d) and their magnitudes (coloured contour map in Fig. 4d) were superimposed on an experimental snapshot of the stagnation area observed for the case of polystyrene beads in cell culture medium within a hanging drop (Fig. 3c).

We estimated the minimum net tangential force required to ensure free flow of beads when assuming a no-slip boundary condition at the ALI (minimum F∥ for flow). To do this, we modelled higher flow rates, where a stagnation area is no longer present (ESI† Fig. S1, no-slip boundary condition). With a flow rate of 6 μL min−1, we estimated the minimum F∥ for flow at ∼100 fN.

Alternatively, we modelled a slip boundary condition at the ALI. This alternative did not yield a condition in which particles would stagnate, except for flow rates 20-fold lower than what was experimentally measured (ESI† Fig. S1, slip boundary condition). The use of a slip boundary condition did not yield the observed experimental result in Fig. 3b and c and 4d.

Therefore, with F∥ magnitudes under 100 fN, Fig. 4d supports our hypothesis that modelling a no-slip boundary condition at the ALI is a successful approach to predict the experimental stagnation area.

 Parametric sweep. The numerical model, supported by the experimental observation of stagnation in Fig. 4d, allowed us to perform a broad parametric sweep on design (drop aperture) and operation (bead position and drop height) parameters of hanging drops for a working flow rate of 1 μL min−1.

The upper limit for the size of a hanging drop for the parametric sweep was found by the capillary length (l) in eqn (2).30 This size was found via the Eötvös or Bond number, which establishes the ratio of gravitational forces, pulling the drop down, to capillary forces, hanging the drop up.

Here, γ is the water surface tension, 0.0727 N m−1 at 20 °C or 0.0709 N m−1 at 37 °C,31ρ is the water density, 998.2 kg m−3 at 20 °C or 993.3 kg m−3 at 37 °C,32 and g is the gravitational acceleration, 9.8 m s−2. These values yield a capillary length of 2.73 mm at 20 °C or 2.70 at 37 °C. This capillary length is calculated for the case where the surface tension is highest, i.e., with de-ionized water, which gives a maximum size value. A hanging drop with a radius equal to the capillary length (5.4 mm in diameter) will have a perfect equilibrium between its weight and surface tension. However, designing such a drop in a microfluidic network would be very unstable, since any hydrodynamic force on the interface would cause it to burst.33 Through previous experiments (data not shown), we have established that the practical upper limit for the diameter of a hanging drop within an HDN is 4 mm. In practice, the smaller the diameter of the hanging drop, the more stable it is.

Ultimately, modifying the design is laborious and requires optimizing the microfluidic network, device fabrication, and validating its functionality. On the other hand, modifying the drop height is doable during device operation by simply adding or removing liquid from one of the inlets of the system (Fig. 2c). Hence, for the purpose of this work, we wanted to find an aperture that enabled both, particle stagnation and flow.

Therefore, we carried out a parametric sweep on the drop aperture diameter (2a) from 0.5 mm, which is approximately twice the width of our channels (Fig. 2c), to 4 mm, which is the practical upper limit. A scheme of varying drop aperture radii (a) is shown in Fig. 5a. We also carried out a parametric sweep on the drop height (h) as seen in Fig. 1b relative to the drop aperture radius (Fig. 5b). The h/a parametric sweep varied from a flat drop at h/a = 0 to a hemispherical drop at h/a = 1.

Fig. 5 Parameter sweep results. (a–c) Schematics showing the parametric sweep methodology. (a) Schematic of the parametric sweep over the drop aperture radius (a) for a constant drop-height-over-aperture (h/a) ratio. (b) Schematic of the parametric sweep over the drop height (a), varying the h/a ratio for a constant drop aperture. A schematic representation of a bed of particles, constituting the stagnation area, is shown in green at the ALI. (c) Schematic of the parametric sweep over the particle's x position relative to the drop aperture radius, taken at the worst-case symmetry plane of the ALI. A schematic representation of particles (green circles) at various positions is shown at the ALI. (d) Plot of the h/a ratio required to expel a bead from a hanging drop at each bead position for various drop apertures. The stagnation area is delineated for a given h/a ratio of 0.6 (dashed red line) and a drop aperture diameter (2a) of 1 mm. This stagnation area implies that, if a particle settles at the ALI between x/a = 0.2 and 0.9, it will stagnate. The minimal h/a ratio (dash-dotted red line) is referred to as the “critical h/a ratio” under which no stagnation area is present. (e) Phase diagram showing the critical h/a ratio under which free bead or cell flow is allowed. Over a h/a ratio of 0.6, potential HDN instability can be observed and is further described in the discussion. The actual drop height in mm is shown for the given critical h/a ratio (dashed red line). The apparent local extrema in the dashed red line are numerical artefacts due to the parameter sweep steps of the critical h/a ratio.

Modelling particles at various positions (x) on the symmetry axis of the hanging drop (Fig. 5c) with a parametric sweep pointed out the worst-case scenario for particle stagnation. Since upstream particles (x/a < 0) are inevitably dragged down to the ALI due to gravity, it is sufficient to only model particles at positions from the bottom of the ALI to its apex downstream, as the particles need to be pushed out of the hanging-drop compartment by the hydrodynamic flow. Therefore, we modelled particle positions from x/a = 0 to x/a = 1.

 Modelling results

Our model provided an h/a ratio that is required to ensure that particles flow at each position in Fig. 5d for drop aperture diameters of 1, 2, 3 and 4 mm assuming a no-slip boundary condition at the ALI. Fig. 5d confirms that the centroid of the stagnation area is downstream of the hanging drop, as shown experimentally in Fig. 4d. As expected, larger drop apertures require smaller h/a ratios to ensure free flow of particles. In other words, the critical h/a ratio is inversely proportional to the drop aperture. For example, for a diameter of 1 mm, Fig. 5d shows that an h/a ratio under ∼0.43 allows for free flow of particles at the ALI. This data point and the absolute drop height (215 μm) were thereafter transferred to Fig. 5e. This process was repeated for every drop aperture, generating the black and red dashed lines of the phase diagram (Fig. 5e).

The phase diagram of Fig. 5e shows the critical h/a ratio to enable free particle flow in HDNs for a wide range of drop apertures. Fig. 5e essentially compares the operational parameter (drop height) on its y-axis to the design parameter (drop aperture) on its x-axis. The critical drop height in mm appears to plateau at 0.3 mm for drop aperture diameters larger than 2 mm. The optimal drop aperture then depends on the application and will be elaborated further in the discussion.

For our experiments, we used the existing on-chip pump and HDN design, which featured drop aperture diameters of 3.5 mm and still allowed for particle flow control.19 The model's findings are summarized in Fig. 6a for our 3.5 mm HDs and the particles of interest. The model predicts free particle flow for a drop height under 300 μm, i.e., at an h/a ratio under 0.17.

Fig. 6 Experimental validation of the phase diagram of Fig. 5e. a) Schematics of the various drop heights show the control over the h/a ratio. b) Images of the bead stagnation area (orange) for various drop heights. After waiting a few hours for bead stagnation, the drop height was actuated. The results show that, as expected, the stagnation area shrank before disappearing completely. ESI† Video S1 shows the videos from which the frames were taken. c) Images of the cell stagnation area (orange) and floating clusters (blue) for various drop heights. The noise in the background is due to the double-sided tape (process 2 described in the “device fabrication” section) used to bond the chip to the transparent slide support and does not affect the results otherwise. This double-sided tape was not used for the experiments in subpanel b. After waiting a few hours for cell stagnation, the drop height was actuated. The results show that, as expected, the stagnation area shrank before disappearing completely. ESI† Video S2 shows the videos from which the frames were taken.

In practice, since we are looking at phase diagrams, the h/a ratios in this study are rather indicative than decisive. There are two clear particle behaviours (phases) and a general transition (line) between these phases that is highly dependent on particle size, the effect of a stagnating pellet, static friction, adhesion, etc.

 Experimental particle-flow control

Using the same methodology as for the experiments in Fig. 3, we tested the model's findings (Fig. 6a) experimentally by flowing polystyrene beads in cell culture medium around the closed-loop HDN.

After a few hours of recirculation, bead stagnation was apparent, and we started actuating the drop height over several hours. With our existing drop aperture diameters of 3.5 mm and an experimentally measured flow rate of 1 μL min−1, the flow or stagnation of beads in cell culture medium could be successfully controlled by actuating the drop height (Fig. 6b).

With these findings for beads in cell culture medium, we repeated the experiment with THP-1 cells in cell culture medium. We found similar results for cells (Fig. 6c) when compared to beads flowing in cell culture medium. We noted that the visible background noise in Fig. 6c was caused by the double-sided tape used to bond the chip to its support slide in this experiment and did not affect flow results otherwise. We observed that the transition from a small stagnating particle bed (stagnation reduction in Fig. 6b and c) to unimpeded particle flow (particle flow in Fig. 6b and c) required gentle percussion, i.e., tapping the setup.

Microscopic investigation of the air–liquid interface

To investigate the changes that occur at the ALI and that generate the pseudo no-slip boundary condition, we performed transmission electron microscopy (TEM). Therefore, cell culture medium was recirculated through our HDN and was sampled after 0 h, 3 h, and 24 h (Fig. 7). The images obtained by TEM of stock cell culture medium (Fig. 7; first row) showed an abundance of micelles and small self-assembled structures (DH ∼ 20 nm and 100 nm respectively) and the presence of few large aggregates (>1 μm). TEM of the ALI after 3 h (Fig. 7; second row) showed an overall decrease in the number of single micelles on the grid, displaced by worm-like micelle aggregates, and larger aggregates resulting from them. TEM of the ALI after 24 hours (Fig. 7; third row) showed an abundance of the same micelle chains and structures deriving from their uncontrolled aggregation, as well as some salt precipitates. These results suggest that there is an induced aggregation of some medium components into larger aggregates, which could have an influence on the ALI's properties. Hence, the no-slip-like behaviour could be linked to the formation of aggregates over time.

Fig. 7 Transmission electron microscopy (TEM) and dynamic light scattering (DLS) results. Results are shown for stock cell culture medium (first row) and hanging-drop ALI after 3 (second row) and 24 (third row) hours. TEM images show several scales, from 5 μm down to 200 nm.TEM overview shows the increased prevalence of larger aggregates over time. TEM detail of larger nanostructures reveals the aggregation process. TEM detail of smaller nanostructures shows circular micelles. DLS results show the relative preponderance of nanostructures of scales from 10 to 10[thin space (1/6-em)]000 nm. The DLS measurements are qualitative, since the samples are heterogeneous and polydisperse. DLS curves are shown in triplicates, where the most representative data is drawn as a black line. 13° forward scatter DLS measurements: larger nanostructures primarily scatter light at forward angles. Therefore, we see the relative increase in micron-scale aggregates with time. 173° non-invasive back scatter DLS measurements: back scatter detection is less sensitive to the presence of large nanostructures. Therefore, we see a relatively constant signal from micelles and small self-assemblies over time. Repeating the TEM measurements by removing FBS from the cell culture medium showed a blank TEM image, where no large aggregates and mostly micelles could be seen.

To support these results, we performed dynamic light scattering (DLS) to qualify the change of the hydrodynamic radii of nanostructures at the ALI (Fig. 7; second and third columns). With the stochastic movement of aggregates in the polydisperse ALI samples and with the built-in DLS software peak smoothing, repeated measurements could not systematically find large aggregates and, therefore, DLS results should be considered qualitative. The highlighted DLS measurements confirmed an increased likelihood of uncontrolled aggregation (>1 μm) at the ALI (Fig. 7; second column) over longer periods of time and the presence of ∼20 nm nanostructures (Fig. 7; third column) throughout the experiment.

Discussion

With the experimental results displayed in Fig. 6, we validated a mathematical model of forces on polystyrene beads and cells settled at the ALI of a recirculating HDN. Here, we will elaborate on our insights in the fabrication and operation of HDNs designed for cell recirculation. We will also discuss how our results can be transferred to various applications. Finally, we will lay out the implications of our experimental observations and models on ALI modelling strategies.

 Insights on fabrication and operation

The phase diagram of Fig. 5e can be used to guide system design based on the application requirements (flow or stagnation of cells). However, even for a non-optimal system, the experimental conditions can be modulated by fine-tuning the operational parameters within specific boundaries.

If cell stagnation is desired through most of the operation of the HDN, a drop aperture of 3 mm or higher is a good choice. Choosing this aperture is preferable due to stagnation being present for h/a ratios above 0.2.

If free cell flow is desired through most of the operation of the HDN, a drop aperture of 1 mm or lower is preferable. Choosing this aperture is preferable due to free flow being present for h/a ratios under 0.4.

For a precise control over cell flow or stagnation, a drop aperture around 2 mm is preferable. This aperture will offer the maximum dynamic range for cell mobility control: working with an h/a ratio between 0 and 0.3 will allow for free cell flow, and an h/a ratio between 0.3 and 0.6 will entail cell stagnation.

Our model provides valuable aids in making design considerations before the fabrication of dedicated HDNs. Careful planning along these guidelines will save time and efforts by reducing the fabrication iterations needed to finalize chip designs through trial and error.

However, even when considering all these parameters, there are challenges that apply to HDNs in general.33 From a practical standpoint, potential fabrication inconsistencies and imperfect experimental chip levelling will have effects on hanging-drop stability. First, fabrication inconsistencies, especially when aligning and bonding multiple layers, could cause drop aperture variations throughout the device. Drop aperture variations could result in a “weak link” within the network, meaning that a misshaped hanging-drop compartment with a higher drop aperture diameter could cause the corresponding liquid drop to crash down. Second, an imperfect levelling could cause a variation in the hydrostatic pressure through the chip. Since the drops are fluidically interconnected within an HDN, pressure equilibrates through all drops. The drop at the lowest level will systematically crash if its h/a ratio reaches 1. Because of such experimental considerations, we recommend limiting the maximal prescribed h/a ratio to 0.6 (black dashed line in Fig. 5e).

 Impact on experimental applications

While the above listed recommendations hold for specific, established experimental conditions (8 μm diameter and 1.05 g cm−3-density particles recirculating at 1 μL min−1), our analysis can be repeated for particles of different diameters (d) and volumetric mass densities (ρ), or for different flow rates (Q). Larger particles would have a larger cross-section, leading to higher hydrodynamic forces proportional to their surface area (∝d2), increasing the force that pushes them out of the hanging drop. Larger particles, e.g., large cell aggregates or microtissues, would also have a larger volume, leading to higher gravitational forces proportional to their volumes (∝d3), pushing them to the bottom of the hanging drop. A cubic increase of gravitational forces trumps a quadratic increase of hydrodynamic forces. Therefore, an increase in particle size (for a particle with negative buoyancy) would lead to an increase in stagnation. The downstream location of the stagnation zone is constant for particles of a given size and density, subjected to a given flow rate, in a microfluidic chip of a given drop aperture (see Fig. 5d). However, increasing stagnation (Table 1 first row) would move the stagnation zone down, closer to the bottom of the hanging drop. Conversely, increasing circulation (Table 1 second row) would move the stagnation zone up, downstream in the hanging drop. Ultimately, increasing particle stagnation can be achieved by increasing particle size, particle volumetric mass density, drop height, or drop aperture, or by reducing the flow rate.

Table 1 summarizes how these various parameters affect particle stagnation and circulation. We also comment on the effort needed to change these parameters.

The preceding analysis can also be applied to particles with positive buoyancy, i.e., floating particles, by simply considering an inverted HDN, or standing-drop network.34,35 Neutrally buoyant particles, however, would only be affected by hydrodynamic forces.

 Usage in the context of microtissue and immune cell co-culture

During typical experiments, the stagnation area, as evidenced by Fig. 4d and ESI† Video S1, will steadily grow to its maximum steady-state size, as more particles are added into the system. The maximum size is highlighted for a h/a ratio of 0.6 and an aperture diameter of 1 mm in Fig. 5d. Once the stagnation area reaches its maximum size, particles will flow around the stagnation area and through the hanging-drop compartment, which effectively imposes a cap on the number of particles that can reside within the stagnation area. This cap on the number of particles enables simultaneous flow and stagnation of particles. Applying this knowledge to biological applications would allow for a precise control over the size of a stagnating-cell bed at the bottom of hanging-drop compartments. For example, by adding a microtissue in the system to study microtissue–immune cell interaction, the methodology outlined in this paper will enable to dynamically control the ratio of immune cells per microtissue in the system throughout the experiment.

However, our results do not directly translate to microtissue and immune cell co-cultures. The presence of a microtissue in the drop will reduce the flow speed near the bottom of the microtissue. In turn, this flow speed reduction will entail an increase of the stagnation area. Nevertheless, our modelling strategy provides a robust framework to recreate a flow-stagnation phase diagram (Fig. 5e) in the presence of microtissues and cells of various sizes. Such analysis was not conducted, as it was outside the scope of this publication.

Interpretation and context of pseudo-no slip boundaries

The no-slip-like fluidic behaviour observed in this study, as opposed to the expected slip behaviour (ESI† Fig. S1), suggests a fundamental change in how ALIs should be modeled.17,36,37 A medium-dependent boundary condition was not previously considered. The wrong boundary condition can lead to miscalculating flow rates by a factor of two to eight, which, in turn, can cause a large discrepancy between design and experimental operation of microfluidic chips. For microfluidic HDNs that are designed to be operated with cell culture medium, our results suggest verifying that there is a no-slip boundary condition at the ALI to ensure normal chip operation.

Although we show medium dependence of particle behaviour (Fig. 3), we show that particle circulation behaviour at the ALI does not significantly depend on whether the particles are cells or beads (Fig. 6). The similar performance is due to physical interactions (e.g., adsorption, aggregation, rolling, hydrodynamic push) prevailing over biological interactions (e.g., cell–wall interactions, etc.).

Effect of pseudo-no slip boundary on particles

Our TEM and particle flow results (Fig. 7) suggest that the pseudo-no-slip boundary is caused by the complex medium formulation of RPMI-1640, mixed with 10% FBS necessary for the culturing of our cell model. The composition of FBS is difficult to establish and, as our TEM measurements show, it contains several molecules that will aggregate and change ALI behaviour. Simpler medium formulations without a preponderance of micelles should, in principle, help to obviate the pseudo-no-slip boundary, if a slip boundary is required for the biological application.

Additionally, we observed a certain “stickiness” of beads at the ALI with cell culture medium. Combined with our TEM measurements, this observation suggests that, when beads stagnate at the ALI for a long time, they interact with proteins, molecular assemblies, and salts. This interaction leads to a stronger adhesion of beads to the ALI than if they were simply resting at the ALI. However, light tapping breaks this interaction, allowing beads and cells to simply rest at the ALI and follow the expected flow patterns.

Prevalence of the Marangoni effect

Surface tension gradients at an ALI will induce an interfacial flow from regions of low surface tension to regions of high surface tension; this is the so-called Marangoni effect. This interfacial flow can entrain a bulk liquid phase, leading to the more eye-catching examples of the Marangoni effect, e.g., tears of wine,38 or the reversal of coffee-ring deposition.39

Here, we estimate the scale of the Marangoni effect on particle displacement in our device.40 At 37 °C, the surface tension of de-ionized water is 70 mN m−1, whereas that of cell culture medium containing serum (e.g., 10% FBS) is 52 mN m−1.41 We examine the extreme case, where de-ionized water is inserted at the interface of a drop neighbouring a drop of cell culture medium (4.5 mm pitch). In this case, the surface tension gradient would generate a maximum and rapidly decreasing interfacial flow of 0.4 m s−1 from the culture medium to the de-ionized water interfaces.40 However, since we do not directly interact with the interface in the way described by this extreme case, this interfacial velocity is not possible in our system.

Surface tension gradients in our system can arise in two ways: (1) evaporation of the solvent (water) causing localized surfactant (e.g., micelles) upconcentration; and (2) advection and diffusion of surfactants. (1) Solvent evaporation is significantly mitigated by our experimental setup, which reduces the evaporation to less than 10 μL per hour while our system contains 100 to 300 μL. Additionally, solvent evaporation is uniform across the ALI surface. Therefore, evaporation does not induce surface tension gradients. (2) The hanging drop (at most 800 μm height) is hanging from a comparatively thick (500 μm height) bulk of liquid phase. Therefore, any local increase of surfactant concentration on the ALI surface is mitigated by the recirculating bulk of the liquid phase. Due to these mitigating factors, we can determine that surface tension gradients, i.e., the Marangoni effect, in our system are negligible.

Conclusion

We show that a judicious drop-height control is a viable way to counteract the unexpected effect of bead and cell stagnation when attempting particle recirculation in an HDN. However, the change of the ALI boundary condition from a slip to a no-slip condition is poorly defined. In this study, we achieved the transition from a slip to a no-slip condition by recirculating the medium in the HDN over several hours until a stagnating particle bed formed. Waiting for a particle bed to form ensures the ALI boundary reaches its steady-state and, therefore, a more predictable hydrodynamic behaviour within the recirculating HDN.

In practice, if the no-slip boundary condition can be reliably reproduced at the ALI in a sterile environment, the technique highlighted in this study will allow for a more precise prediction of and control over the flow and stagnation of cells than existing techniques.9 For a microtissue in such a system, a precise control over the residence time of flowing cells near the microtissue could be achieved simply by modifying the drop height. Such an approach would constitute an MPS that allows for studying the interaction between recirculating immune cells and various tissue or organ models without the need for tedious coating protocols as required for standard microfluidic devices.

Room temperature roll-to-roll additive manufacturing of polydimethylsiloxane-based centrifugal microfluidic device for on-site isolation of ribonucleic acid from whole blood

The rendered image of the room-temperature roll-to-roll additive manufacturing platform

Academic Article

Room temperature roll-to-roll additive manufacturing of polydimethylsiloxane-based centrifugal microfluidic device for on-site isolation of riboneucleic acid from whole blood

by Trung Hoang, Han Truong, Jiyeon Han, Saebom Lee, Jihyeong Lee, Sajjan Parajuli, Jinkee Lee and  Gyoujin Cho

Abstract: Polymer-based lab-on-a-disc (LoaD) devices for isolating ribonucleic acid (RNA) from whole blood samples have gained considerable attention for accurate biomedical analysis and point-of-care diagnostics. However, the mass production of these devices remains challenging in manufacturing cost and sustainability, primarily due to the utilization of a laser cutter or router computer numerical control (CNC) machine for engraving and cutting plastics in the conventional prototyping process. Herein, we reported the first energy-efficient room-temperature printing-imprinting integrated roll-to-roll manufacturing platform for mass production of a polydimethylsiloxane (PDMS)-based LoaD to on-site isolate ribonucleic acid (RNA) from undiluted blood samples. We significantly reduced energy consumption and eliminated thermal expansion variations between the mold, substrate, and resists by accelerating the PDMS curing time to less than 10 min at room temperature without using heat or ultraviolet radiation. The additive manufacturing technology was applied to fabricate a multi-depth flexible polymer mold that integrated macro (2 mm) and micro-sized (500 μm) features, which overcomes the economic and environmental challenges of conventional molding techniques. Our integrated R2R platform was enabled to print adhesion-promoting films at the first printing unit and continuously in-line imprint with a high replication accuracy (99%) for high-volume manufacturing of a new centrifugal microfluidic chip with an enhancement of mixing performance by integrating an efficient mixing chamber and serpentine micromixer. This research paved the way for scalable green manufacturing of large-volume polymer-based microfluidic devices, often required in real-world sample-driven analytical systems for clinical bioanalysis.

Keywords: room-temperature PDMS; centrifugal microfluidic; RNA extraction; roll-to-roll nanoimprint lithography; sustainable manufacturing

We kindly thank the researchers at Sungkyunkwan University for this collaboration, and for sharing the results obtained with their CADworks3D system.

The rendered image of the room-temperature roll-to-roll additive manufacturing platform

1. Introduction

In the last two decades, microfluidic systems have evolved rapidly for numerous chemical, biomedical, biological, and environmental applications [[1], [2], [3], [4]]. Among those devices, lab-on-a-disc (LoaD) platforms are gaining much attraction for biomedical applications due to the ability to integrate rapid sample preparation stages such as the isolation of nucleic acids from large-volume whole blood samples [5] with assays. The LoaD devices offer significant advantages over other microfluidic systems, especially a reliable, controllable, and compact pumping mechanism that enables efficient mixing of reagents [6], rapid response time, and enhanced assay sensitivity. In this platform, fluid flow is driven by centrifugal pumping which involves minimal instrumentation by using only a simple and compact motor to generate the force. This approach eliminates the need for external interconnects and syringe pumps, thereby preventing contamination of the sample by the surrounding environment [7]. By combining the benefits of both microfluidics and centrifugal forces in a single device, the centrifugal microfluidic technology has been identified as a standard tool for mainstream diagnostics especially point-of-care in vitro diagnostics (IVD), and achieved significant commercial success [8].


A typical LoaD consists of a multilayer platform made from thermoplastics, such as polycarbonate (PC), poly (methyl methacrylate) (PMMA), polystyrene (PS), cycloolefin polymer (COP), assembled by adhesive or through hot embossing and injection molding methods [[9], [10], [11]], which is cost-effective for high-volume manufacturing process. These devices can also be fabricated through a laser or router computer numerical control (CNC) machine for engraving and cutting plastics, in conjunction with the utilization of plotter machines to cut the adhesive film [[12], [13], [14]]. However, for CNC-manufactured microfluidic devices, the surface becomes extremely rough, leading to slow and inaccurate fluid flow and bonding inhibition as well as non-specific binding which can negatively impact the performance of microfluidic devices. Also, the utilization of these subtractive manufacturing systems has been constrained by the capability of the cutting tools, making the fabrication process of micro-scale features even more expensive, lengthy, and complex, limiting the fabrication throughput, and presenting challenges for mass production ability.


Among polymer-based materials, polydimethylsiloxane (PDMS) has been widely used to fabricate microfluidic devices via soft lithography [[15], [16], [17], [18], [19]]. The use of PDMS in manufacturing LoaD devices has been considered as an alternative strategy to the CNC-based one due to its capability to reduce production costs, increase flexibility, facilitate ease of fabrication, and permit rapid prototyping without the use of harmful etching chemicals. The precursors required for preparing PDMS, comprising prepolymers and curing agents, are notable for their cost-effectiveness and widespread commercial availability. The fabrication process of PDMS-based microfluidic devices can be executed without the utilization of specialized cleanroom facilities enabling rapid prototyping of devices at a lower cost than what is feasible using silicon technology [16]. Moreover, the surface properties of PDMS can be easily tuned, enabling the ability to bond with many materials like glass and PDMS itself [20]. The optical transparency and gas permeability of PDMS-based microfluidic devices are ideal for numerous biomedical applications, particularly in optical detection methods and cell culture. Therefore, large-scale manufacturing methods of PDMS-based microfluidic devices have recently gained much importance in various research areas of science and engineering to bring the usages of these devices to practical clinical applications.


Roll-to-roll (R2R) nanoimprint lithography is considered as an up-and-coming alternative to traditional manufacturing methodologies, owing to its ability to achieve high-throughput production and thus facilitate its application at an industrial scale [21]. PDMS was first demonstrated to be R2R processable by Ahn and Guo [22] (2008) for sub-micrometre test structures on a polyethylene terephthalate (PET) substrate. Later, Hiltunen proposed R2R fabrication of integrated PDMS-paper microfluidics for molecular diagnostics [19]. However, both techniques relied on thermal imprinting to cure PDMS resist by heating the imprinting roll to a high temperature with a large amount of energy. This requires roll modifications with complex and expensive auxiliary systems such as piping networks to circulate the heating liquids, the pre-heating and heating units, cooling system as well as temperature monitoring and control system, which increase waste heat and energy consumption, consequently causing greenhouse gas emissions. Additionally, the mismatch in thermal expansion coefficients between the mold and substrate results in the generation of lateral strain, which degrades the quality of the imprinted patterns and the lifetime of the mold [23]. Also, the molds used in these proposed methods were limited to a micrometre scale, making them impossible to apply for the fabrication of large-volume LoaD which requires millimetre-scale thickness.


To overcome those issues mentioned above, we developed the first room-temperature printing – imprinting integrated roll-to-roll (R2R) in-line manufacturing platform for mass production of a PDMS-based LoaD for on-site RNA isolation from whole blood samples with low cost, less energy consumption, and less by-products. In this work, multi-depth master stamps were first fabricated by using a 3D printer, overcoming the challenges of conventional CNC-based and photolithographic molding technology. The large area flexible polymer shim was fabricated by using a rubber sheet to stick PDMS molds which were replicated from 3D-printed stamps. For mass producing the large volume LoaD by R2R technology, the printing unit was integrated in-line with an imprinting platform for coating an adhesion promoter onto PET substrate to facilitate the demolding process. We incorporated well-defined compounds into commercial PDMS Sylgard 184 formulations to accelerate the curing time of PDMS at room temperature enabling the success of a low-temperature R2R imprinting process which helped to reduce heat waste and energy consumption. Finally, as a proof-of-concept study, novel LoaD devices with a high enhancement of mixing performance were sustainably manufactured by our green R2R platform. For the first time, the utility of these R2R-manufactured LoaD devices was demonstrated by isolating RNA from undiluted blood samples.

Apparatus Used

Master Mold for PDMS

Curezone

The CADworks3D Pr110 3D Printer with a 385nm wavelength projector

PR110
3D Printer

Legacy

2. Material and methods

2.1. Fabrication of negative 3D-printed mold

Our LoaD device comprises two layers: the designed patterns layer and a blank PDMS as a coverlid. The 3D model of the top part was designed by using SOLIDWORKS software, which was then exported as an STL file. We used the Utility Version 6.3.0t3 software to convert STL files into sliced files with appropriate layer heights. Printing options were optimized for high-resolution printing of mold features by using a 3D printer (PR110-385 from CADworks3D company) utilizing a 385 nm light source with a printing area of 110 × 62 × 120 mm and XY resolution of 40 μm. In this work, we printed all masters using the 3D printing protocol with the following parameters: a UV projection time per layer: 9.5 s and a z-axis resolution: 50 μm. It has been shown that 3D-printed templates interfere with the curing of PDMS due to uncured resin remaining on their surface [24]. Therefore, a post-print surface treatment technique is commonly necessary to overcome the aforementioned issues and prevent the adhesion of PDMS to the 3D-printed mold. A treatment method involving coating the template with a protective ink using airbrushing was reported by Comina et al. [25]. The group claimed this technique requires much effort for achieving optimal results. Ho et al. proposed another complicated surface treatment method for 3D printed templates, including heating, plasma treatment, and surface silanization [24]. However, cracks were formed in the template during the heating process. To avoid these time-consuming, nonreplicable, and ineffective surface treatment methods, we fabricated master molds directly via the DLP 3D printing method utilizing a novel resin formulated by Creative CADworks company (CCW Master Mold for PDMS devices), composed of methacrylated oligomers and monomers. After removing printed molds from the picker, they were subjected to a thorough rinsing procedure utilizing isopropanol. Subsequently, an air nozzle was employed to eliminate residual resin from edges and within extremely fine features. Finally, we postcured the molds by exposing each part to UV light at a wavelength of 405 ± 5 nm within a curing chamber, and then, the resulting molds were employed in our imprinting works.

 

2.2. Fabrication of positive PDMS mold

To fabricate positive PDMS mold, we prepared a mixture including PDMS base and curing agent in the ratio (W/W) of 10:1 (Sylgard 184 from Dow Corning, MI, USA). The mixture was mixed and defoamed with a centrifugal mixer (Thinky Mixer ARE-310) for 3 and 2 min, respectively, and then poured onto the 3D-printed molds without undergoing any surface treatment process. Later, we cured it in the oven for 4 h at 60 °C. Subsequently, the cured PDMS was carefully detached from the molds and cleaned thoroughly with isopropyl alcohol (IPA) and ethanol at least three times, followed by air drying between each wash. To be used as the mold for the R2R imprinting process, we applied a conformal coating of parylene C which serves as an anti-adhesion layer for demoulding these positive PDMS replicas [26]. Finally, a flexo plate with 5 mm thickness was cut with precise squares, used as the substrate for inserting PDMS positive molds, and wrapped to form a sleeve for preparing the imprinting cylinder.

 

2.3. PDMS formulations for R2R imprinting process

We formulated a new recipe for fast curable PDMS at room temperature (named Room-Temp PDMS in this paper) by adding compounds (the Ashby–Karstedt catalyst and tetrakis (dimethylsiloxy) silane) into commercial PDMS Sylgard 184 formulations [27]. SYLGARD™ 184 Silicone Elastomer and curing agent were supplied by Dow. Ashby–Karstedt catalyst (platinumcyclovinylmethyl-siloxane complex; 2 % Pt (0) in cyclomethylvinylsiloxanes) and Tetrakis (dimethylsiloxy) silane (TDS) were supplied by Gelest. We used a ratio of 10:1 (w/w) for the elastomer and curing agent to make the standard Sylgard 184 framework. In this study, these compounds were always added following the optimized recipe in exact order: (1) TDS 2 wt%; (2) Elastomer base 88.95 wt%; (3) Curing Agent 8.9 wt%; and (4) Ashby–Karstedt catalyst 0.15 wt%. After all the chemicals had been added, we placed the mixture in Reactor-Ready included circulator Huber Ministat 230, both were supplied by Radleys company (Fig. S8) for continuous mixing at 500 rpm and cooling at −15 °C to prolong the lifetime of PDMS for the imprinting process. After mixing for 1 h, we turned off the mixer and set the reactor at a vacuum of 0.2 psi for degassing in 1 h meanwhile maintaining cooling during the whole process. For comparison, we also prepared the standard PDMS Sylgard 184 as control samples (named Sylgard 184 in this paper) and the reference samples (named Vinyl-terminated PDMS in this paper) by optimizing the recipe of a fast, thermal-curable liquid resist [28] based on a modified PDMS. The formulation for this reference sample consists of three components: vinyl-terminated PDMS was supplied by Gelest, poly (dimethylsiloxane-co-methylhydrosiloxane) trimethylsilyl terminated which acts as a crosslinker and platinum (0)-1,3- divinyl-1,1,3,3-tetramethyldisiloxane complex solution as a catalyst, both were supplied by Sigma-Aldrich. In our optimized recipe, we mixed a ratio of 5:1 (w/w) for the vinyl-terminated PDMS and crosslinker, then added Pt catalyst into the mixer at a concentration of 470 ppm. This formulation effectively accelerated the curing time of this PDMS-based resist at room temperature, making it possible to be used for the R2R imprinting process.

 

2.4. Roll-to-roll imprinting process and chip assembly

The replication by the R2R imprinting process was conducted at room temperature through an imprinting unit as illustrated in Fig. S1. First, we treated a roll of 150 mm wide PET substrate with an adhesion promoter (Wacker Primer G790) by gravure printing method with a speed of 3 mm/s for five layers then dried in line at room temperature. The coated PET substrate remained as transparent as the original one, as shown in Fig. S2a. The premixed PDMS was deposited to the gap between the imprinting roll and pressure roll during the R2R imprinting process. We operated the process at room temperature with optimized pressure by testing 1 to 10 kgf/cm2 under an imprinting speed of 0.3 mm/s. After coming out from the imprinting unit, imprinted chips on the PET substrate were collected at the rewind roller and were applied to a cutting process for making single devices. The coverlids were simply fabricated by pouring our room-temperature PDMS into a 3D-printed square mold with a dimension of 55 × 55 mm, resulting in unstructured PDMS layers with a uniform thickness of 1 mm. The inlets and outlets for each chamber were manually punched. After that, a 3 mm diameter circular disc magnet which was supplied by First4Magnets, was preloaded into the mixing chamber of each device, and lids were then bonded onto the imprinted layer with a plasma treatment method.

 

2.5. Replication accuracy and material characterization

Replication accuracy. Dimensional analysis of the 3D printed mold, PDMS mold, and the R2R imprinted LoaD was performed with an industrial microscope Olympus BX53M.

 

Optical properties measurement. Autofluorescence measurements of three types of PDMS and glass were conducted by using a Nikon Eclipse Ti2 microscope with 4 different excitation wavelengths. The optical transmittance was measured on all three PDMS types: Sylgard 184, Modified Vinyl-terminated PDMS, and Modified Sylgard 184 using a complete setup of a 508 PV™ UV–visible–NIR Spectrophotometer integrated with a bright field microscope Olympus BX53M. For both measurements, we prepared 3 samples for each PDMS by pouring uncured PDMS into the circle Petri dish with a diameter of 55 mm with a uniform thickness of 2 mm and curing at 80 °C. All the samples were cleaned by sonicating in ethanol for 30 min and then dried with a nitrogen gun before analysis. To check the discoloration phenomenon of these materials, we prepared 5 samples for each type of PDMS in the same way as mentioned above, but all were cured at room temperature. After fully curing all samples, we aged them thermally in 5 isothermal ovens respectively at 20 °C, 40 °C, 60 °C, 80 °C, and 100 °C for 1 h before analyzing.

 

Contact angle measurements. We measured the static contact angle for each type of PDMS by using a Drop Shape Analyzer (DSA100) from Krüss. Measurements were conducted using an automated drop dispenser and deposition device to dispense a 2 μl drop of deionized (DI) water on the material surface. The static contact angle was calculated using computer software.

 

Mechanical characterization techniques. Tensile strength tests were conducted using the Instron Electropuls E3000 testing system. All samples for three types of PDMS were prepared following ASTM D-412 standards which were cut into dumbbell shapes, referred to as dogbone, using a type-D specimen die. The samples were put into the uniaxial grips and dragged at a speed of 3.84 mm/s until they reached a breakpoint [27]. We performed the tests on five trials for each sample then the engineering stress and strain were reported as averages and corresponding standard deviations.

 

2.6. Nucleic acid design and reagents

Complete genomes of SARS-CoV-2 Wuhan wildtypes (accession MN908947.3) were retrieved from NCBI (https://www.ncbi.-nlm.nih.gov/). Forward Primer -ACAGGTACGTTAATAGTTAATAGCGT and Reverse Primer -ATATTGCAGCAGTACGCACACA were purchased from Cosmogenetech Inc., (Seoul, Korea). The experiment used SARS-CoV-2 mRNA spiked in human whole blood (Innovative Research,USA) for further analysis. RNA purification reagents were purchased from MagListo 5 M viral DNA/RNA extraction (Bioneer). The buffer solutions (proteinase K, virus binding buffer (VB)), washing buffer (VWM1, RWA2, and WE buffer), elution buffer (ER buffer), magnetic nanobead (MNPs) were preloaded into the chip with the volume: 20 μl proteinase K, 200 μl VB, 400 μl absolute ethanol, 50 μl magnetic nanobead, 500 μl washing buffer, 100 μl ER buffer. 200 μl of blood was added at the beginning of the operation. Polymerase chain reaction (PCR) was performed using the following protocol: reverse transcription (42 °C for 5 min, 95 °C for 10 s), PCR test (40 cycles of 95 °C for 5 s, 60 °C for 30 s), melting curve analysis (95 °C for 0 s, 65 °C for 15 s, 95 °C for 0 s) with 20 μl of final PCR volume (One Step TB Green® PrimeScript™ RT-PCR Kit II (Perfect Real Time) (Takara Bio, Tokyo, Japan).

 

In our LoaD devices, we utilized ferro-wax valves which were made by mixing paraffin wax (327,204, Sigma-Aldrich, Inc., St. Louis, MO, USA) with Ferrofluid (EFH1, 60 cc, Ferrotec, Santa Clara, CA, USA) in the ratio of 1:1 b y mass and stirring the mixture at 65 °C on a hotplate for 12h.

 

3. Results

We developed the first sustainable R2R additive manufacturing platform for fabricating PDMS-based centrifugal microfluidic devices at room temperature without using heat and light sources that minimize the emission of greenhouse gas and hazardous by-products (Fig. 1). In this work, we used the continuous in-line printing and imprinting units of our R2R system, illustrated in Fig. 1a. By utilizing additive manufacturing, we introduced sustainable fabrication techniques for a deep-depth flexible shim that addressed the issues of traditional molding technologies (Fig. 1b). As illustrated in Fig. 1c, LoaD devices were successfully manufactured at ambient condition by using the new fast-room temperature-curing PDMS based on Sylgard 184 to eliminate the need for heat and light sources which are often used in conventional R2R imprinting process. The process of on-chip isolation of RNA from whole blood using our R2R imprinted LoaD was illustrated in Fig. 1d. Before the imprinting process, the PET substrate was uniformly treated by R2R gravure printing unit with an adhesion promoter that effectively assisted the demoulding of structures with a large thickness (∼2 mm) (Fig. 1e and f). Finally, we successfully demonstrated the high throughput and sustainable manufacturing platform for producing the LoaD devices (Fig. 1g).

 

Figure 1. Overview of the room-temperature roll-to-roll imprinted PDMS-based centrifugal microfluidic devices. (a) Roll-to-roll additive manufacturing platform; (b) Enlarged image of R2R imprint unit; (c) Rewinder unit for collecting imprinted LoaD devices; (d) Illustration of on-chip isolation of RNA from whole blood using our R2R imprinted LoaD device; (e) and (f) R2R gravure coating unit; (g) Mass production of PDMS-based centrifugal microfluidic devices on PET substrate.

3.1. Design and 3D printing of the centrifugal microfluidic devices

Before employing the R2R manufacturing system to mass-produce the LoaD devices, the 3D printed LoaD was first tested to prove the working concept. The prototype of LoaD was designed and fabricated using a 3D printer with a diameter of 55 mm, a channel depth and width of 500 μm, and all the chamber depths of 2 mm which enabled the device to process human blood volume of 150 μl (Fig. 2a). The LoaD was designed to have twelve liquid storage chambers connected by microchannels with ten 3D-printed screw valves, which consist of a head and a rod with a square hole of the channel size. The head has dimensions of 3 mm in diameter and 2 mm in height, while the rod has dimensions of 2 mm in diameter and 3 mm in height. The valve is opened by aligning the hole with the microchannel. To close the valve and stop fluid flow, it is rotated so that the hole is perpendicular to the microchannel [29]. In addition, a groove parallel to the microfluidic channel on the top head of the valve allows easy operation by using a screwdriver. Finally, the whole device was fabricated by the 3D printing method (Fig. 2b) to test the device’s performance. The structure of the valve and operating mechanism are shown in Fig. 2c. We conducted experiments on different geometries of the plasma separation chamber, such as a simple square chamber, a tilted square chamber [30], and a square chamber with tilted structures [31] to evaluate the separation efficiency (Fig. S3). Since the plasma separation efficiency was almost the same, we decided to choose the simple square chamber for easy fabrication. The mixing performance of our device was enhanced by integrating an S-shaped microchannel as a serpentine micromixer [32] and an optimized mixing chamber [33]. The whole process of RNA isolation from whole blood on this device consists of four steps: lysis step, binding step, washing step, and elution step. In Fig. 2d, we qualified the effectiveness of mixing performance and logical design of the 3D printed LoaD by using food dye color solutions. The solution flow direction is indicated by the red dashed line area to the light blue dashed line area. First, during the lysis step, when valves 1 and 2 were opened then the chip was centrifuged to transfer the solution (orange dye color) in three chambers (blood, proteinase k, and binding buffer chambers as named in Fig. 2a) to the mixing chamber. Second, in the second binding step, valve 2 was closed while valves 3 and 4 were opened sequentially to transfer the solution in the ethanol chamber and MNPs chamber respectively by centrifuging the device. After opening valve 5, the solution in the mixing chamber was transferred to the waste chamber. Third, in the washing step, valves 6,7, and 8 were opened serially to move the solution in VWM1, RWA2, and WE buffer chambers to the mixing chamber respectively, meanwhile, valve 5 was opened and closed alternatively to release the solution in the mixing chamber to the waste chamber. Finally, the elution step was performed by closing valve 5 and opening valve 9 to transfer the solution from the ER buffer chamber to the mixing chamber then valve 10 was opened to move the solution into the elution chamber to complete the sample preparation process. The eluted solution can be used for the diagnosis by extracting it from the elution chamber. Since our design has been proven to have efficient mixing and transferring performance, it can be further fabricated by the R2R additive manufacturing platform. After several flow tests on the 3D printed LoaD, the leakage at screw valves was observed because the micro-gap between the valve and the holding hole is inevitable. To solve this issue, we employed laser-actuated ferro-wax microvalves [34] for our R2R imprinted LoaD. Briefly, the working mechanism of this photonic valve is relied on the phase transition of the ferro-wax, actuated by using only a single laser diode instead of many microfabricated heaters and magnets. The valve can be opened by melting the ferro-wax plug in the channel with laser irradiation for a few seconds, allowing the melting wax to flow into two assistant chambers. The response time for the actuation of these photonic valves was accelerated due to the effective heating of iron oxide nanoparticles embedded in the paraffin wax matrix by the laser beam. The ferro-wax can be solidified rapidly at room temperature when we stop the laser illumination, enabling us to make a plug in the channel again as a closed valve. This simplifies the control of multiple microvalves. We demonstrated the operation of the ferro-wax microvalves in Fig. S4.

 

Figure 2. Design and 3D printing of the centrifugal microfluidic devices. (a) 3D model and detailed function of lab-on-a-disc (LoaD) device; (b) 3D printed LoaD device; (c) 3D design of screw valves; (d) Demonstration of device operation by food dyes, “v.1-10” stand for valve 1–10 and fluid flow sequences were indicated by dashed lines and yellow arrow, while red color circle represent closed valve and yellow one represents for opened valve.
Figure 2. Design and 3D printing of the centrifugal microfluidic devices. (a) 3D model and detailed function of lab-on-a-disc (LoaD) device; (b) 3D printed LoaD device; (c) 3D design of screw valves; (d) Demonstration of device operation by food dyes, “v.1-10” stand for valve 1–10 and fluid flow sequences were indicated by dashed lines and yellow arrow, while red color circle represent closed valve and yellow one represents for opened valve.

3.2. The multi-depth macro-to-micro flexible polymer shim

Due to the dramatic increase in complexity, more microfluidic devices require 3D structures, like multi-depth and layer channels. Moreover, microfluidic chips that combine micron-sized structures with large-volume liquid storage chambers are often required in real-world sample-driven analytical systems for clinical bioanalysis. The conventional way of using photolithography for fabricating these structures is time-consuming and labour-intensive, requiring a precise alignment process and extremely difficult to generate macro-sized features. By utilizing additive manufacturing technology, we developed a rapid and low-cost method for fabricating a multi-depth flexible polymer mold that overcomes the difficulties of traditional molding techniques, especially in integrating macro and micro-sized features. The whole process of fabricating this polymer shim is shown in Fig. 3a. We employed a commercially available resin from the Creative CADwork for the direct 3D printing of master molds that effectively addressed the current issues of time-consuming, nonreplicable, and ineffective surface treatment methods. A commercial flexoplate with low cost, flexible, and uniform thickness was used as the substrate for carrying the patterned molds. This method enabled to rapid manufacture of a large area flexible mold at the lab without using an industrial-scale high-resolution 3D printer. The multi-depth mold, which has a total thickness of 4 mm and consists of 2 mm in chamber depth with a channel depth of 500 μm was well fabricated with the dimension shown in Fig. 3d.

 

Figure 3. Fabrication of multi-depth flexible polymer shim. (a) Fabrication steps of polymer mold; (b) The complete large-area flexible polymer mold; (c) Wrapped polymer shim on imprinting roller; (d) Image of multi-depth macro-to-micro features of the mold; (e) Demonstration of effective anti-adhesive coating layer for long lifecycle of the mold by replicating master template M10 to 10 copies from C1 to C10.
Figure 3. Fabrication of multi-depth flexible polymer shim. (a) Fabrication steps of polymer mold; (b) The complete large-area flexible polymer mold; (c) Wrapped polymer shim on imprinting roller; (d) Image of multi-depth macro-to-micro features of the mold; (e) Demonstration of effective anti-adhesive coating layer for long lifecycle of the mold by replicating master template M10 to 10 copies from C1 to C10.

Before starting the R2R imprinting, we applied a conformal coating of parylene C served as surface anti-adhesion (Fig. 1a). This coating material is not only environment-friendly but also extremely effective to prolong the lifetime of the mold without any adhesion to the PDMS resist during the demolding process. To demonstrate that the mold treated with a single coating of parylene C can maintain its anti-adhesive property for a long lifecycle regardless of the number of replica molding cycles, we replicated 10 copies named from C1 to C10 from the master mold named M10. As shown in Fig. 3e, replicas remained high fidelity to the M10 without damaging the master mold. Finally, the flexible properties of this high-thickness polymer mold were demonstrated by wrapping on the imprinting roll with a conformal contact shown in Fig. 3c. This flexible polymer mold was found to be durable because of its capability to withstand high nip pressure (2MPa) for many imprinting cycles.

 

3.3. The fast-room temperature-curing PDMS

We effectively accelerated the curing time of our PDMS at room temperature by modifying Sylgard 184 formulation with Ashby-Karstedt catalyst and tetrakis (dimethylsoloxy) silane (TDS). As shown in Fig. 4a, the curing time at room temperature of standard Sylgard 184, our PDMS, and vinyl-terminated PDMS are 2 days (2880 min), 10 min, and 12 min, respectively. The addition of 0.1–0.3 wt% Ashby-Karstedt catalyst accelerated the curing time of Sylgard 184 at room temperature and improved its mechanical properties which were demonstrated by Murphy et al. [35]. Additionally, the incorporation of TDS can reduce the curing time significantly [27]. Aiming to reduce the heat waste and energy consumption of conventional R2R hot embossing methods, we optimized the concentration of Pt and TDS into Sylgard 184 formulation to make it possible for the R2R imprinting process at room temperature without using UV and thermal curing systems, which helps to reduce heat waste and energy consumption. In this work, we also modified vinyl-terminated PDMS to cure it rapidly at room temperature as a reference to compare with our PDMS.

 

Figure 4. Characterization of the fast-room temperature-curing PDMS. (a) Curing time of three different types of PDMS at room temperature; (b) Autofluorescence of all PDMS types and glass at four different excitations: 405 nm, 488 nm, 594 nm, and 647 nm; (c) Transmission spectra of PDMS samples; (d) Discoloration of all three PDMS formulations after 1 h of thermally accelerated aging; (e) Mechanical properties of Sylgard 184 and Room-temp PDMS.
Figure 4. Characterization of the fast-room temperature-curing PDMS. (a) Curing time of three different types of PDMS at room temperature; (b) Autofluorescence of all PDMS types and glass at four different excitations: 405 nm, 488 nm, 594 nm, and 647 nm; (c) Transmission spectra of PDMS samples; (d) Discoloration of all three PDMS formulations after 1 h of thermally accelerated aging; (e) Mechanical properties of Sylgard 184 and Room-temp PDMS.

The optical properties of three types of PDMS in this study were measured (Fig. 4b, c, and d). The autofluorescence intensities of three materials and glass substrate were measured by exciting light with four different wavelengths 405 nm, 488 nm, 594 nm, and 647 nm corresponding the excitation wavelength of DAPI, FITC, TRITC and Cy5. In Fig. 4b, the fluorescence intensity of 3 types of PDMS are almost the same in every excitation wavelength, while the glass showed a little decrease in fluorescence intensity. Overall, this confirmed the feasibility of using our room-temperature cured PDMS for biomedical devices at a good quality as the commercial PDMS Sylgard 184 and glass, which were used reference samples. The data about the fluorescence intensity is shown in Table S1. The transmittance of light through the microchannel, also referred to as optical transmittance, is a critical issue for a lab-on-chip (LOC) application since numerous analytical protocols employ visualization equipment operating within the visible wavelength range. For our devices, transmittance plays an important role in the efficiency of the laser-actuated ferro-wax valve. To verify the results, the optical transmittance was measured on all three PDMS samples after the thermal treatment, as shown in Fig. 4c. The transmittance of our room-temperature cured PDMS was lower than that of Sylgard 184 b y an amount of ∼3 %, and both samples exhibited an optical transmittance above 90 % for visible light. On the other hand, the vinyl-terminated PDMS showed poor transparency from 22 % to around 60 % for visible light, which causes difficulty for biomedical applications. This can be explained by the clear-to-yellow discoloration phenomenon of silicones caused by the interaction of platinum-complex [[36], [37], [38], [39]]. We observed the discoloration phenomena on all three types of PDMS by aging them for 1 h in a wide range of temperatures from 20 °C to 100 °C. As illustrated in Fig. 4d, the vinyl-terminated sample produced obvious color changes as temperatures increased. The significant discoloration of this sample can be explained due to the large concentration of platinum-complex by the addition of the Asbhy-Karstedt catalyst and the catalytic reaction of Pt was accelerated as temperature increased. The Sylgard 184 samples remained transparent because no additional platinum was added. Interestingly, even though platinum-complex was added in the formulation of our room-temperature cured PDMS, it remained almost transparent as Sylgard 184 for two reasons. First, the concentration of Pt added was small compared to the vinyl-terminated PDMS to be both cured rapidly at room temperature so that the discoloration level was significantly different between those samples. Second, the addition of TDS prevented discoloration in our PDMS [27].

 

The mechanical properties of our room-temperature cured PDMS and Sylgard 184 were measured by tensile testing on the dogbone specimens, as shown in Fig. 4e, confirming the quantitatively significant distinction between those two samples. The tensile test results revealed that the room-temp PDMS became harder and less flexible due to the addition of platinum-complex catalyst [35]. The maximum stress and strain of Sylgard 184 are 6.25 ± 0,83 MPa and 101.8 ± 7.02 %, while those of our room-temperature cured PDMS are 5.89 ± 0.98 MPa and 87.51 ± 9.64 %, respectively. We failed to measure the tensile strength of vinyl-terminated PDMS samples due to their extremely low modulus so they were broken during the gripping process before the measurement, as shown in Fig. S6. The water contact angle measurement results shown in Fig. S5 revealed that our room-temp PDMS has the same hydrophobicity property as Sylgard 184 (112.4°) in the meantime the vinyl-terminated PDMS showed a reduction to 105°, which caused adhesion problems during the imprinting process.

 

3.4. Roll-to-roll replication accuracy

Dimension analysis using the industrial microscope of imprinting tools and PDMS replica (Fig. 5) has shown that the master mold structures were transferred with high accuracy. The results (Fig. 5a) demonstrated that steep sidewalls could be fabricated by our R2R manufacturing platform with only slight bevelling. The cross-section images revealed that the multi-depth of the devices was successfully replicated with an accuracy of 99%. As shown in Fig. 5a, the imprinted chamber depth is 1.99 ± 0.011 mm, and the channel depth is 501.58 ± 1.36 μm while the CAD design were 2.00 mm and 500 μm, respectively. Since parameters such as roll temperature, applied nip pressure, and web transfer speed mainly influenced the quality of imprinted patterns, we optimized those parameters as shown in Table S1. In our developed platform, the operating temperature is low as room temperature, which is not only environmentally friendly but also addresses the common issue in the embossing process resulting from thermal expansion variations between the mold, substrate, and resists. The influence of nip pressure can be seen obviously in Fig. 5a. When the nip pressure increased from 1 kgf/cm2 to 10 kgf/cm2, the deformation of imprinted structures was generated and reduced the thickness of imprinted substrate resulting in failure products. Because the large volume of the devices required a large amount of dispensed PDMS, the imprinting speed was set at 0.3 mm/s to ensure the PDMS filled into the mold patterns with high fidelity and without generating bubbles. Therefore, the curing time of PDMS and printing parameters should be adjusted depending on the structures of the designs so that small devices could be manufactured more efficiently. The best condition for imprinting the LoaD is 1 kgf/cm2 and 0.3 mm/s as nip pressure and imprinting speed, respectively (Table S2). The dimension of three critical positions in the LoaD design (valve, inlet hole of each chamber, and S-shaped channel) of 3D printed mold, PDMS mold, and R2R imprinted LoaD were investigated as shown in Fig. 5b. The lowest variation of structural dimension between final products and computer aid design was in the range of ±2.7 μm, confirming the high replication accuracy of our R2R manufacturing method.

 

Figure 5. Roll-to-roll (R2R) replication accuracy. (a) Cross-sectional images with different magnifications from R2R imprinted samples under different operating nip pressure. Dashed areas on the left side images present the regions shown on the right; (b) Replication accuracy measuring at three positions: waste channel, S-shaped channel, and inlet hole on CAD design, 3D printed mold, PDMS mold, and R2R replicated LoaD with five samples per each.
Figure 5. Roll-to-roll (R2R) replication accuracy. (a) Cross-sectional images with different magnifications from R2R imprinted samples under different operating nip pressure. Dashed areas on the left side images present the regions shown on the right; (b) Replication accuracy measuring at three positions: waste channel, S-shaped channel, and inlet hole on CAD design, 3D printed mold, PDMS mold, and R2R replicated LoaD with five samples per each.

3.5. RNA extraction from whole blood on the R2R additive manufactured LoaD

To evaluate RNA extraction on our device, we implemented a design featuring ten preloaded liquid storage chambers that are separated by photonic valves. The complete protocol for extracting RNA from whole blood can be executed utilizing our LoaD (Fig. S7), comprising plasma separation, sample lysis, magnetic binding, washing, and elution which were designed based on prior literature [31]. The magnetic nanobeads were previously loaded into the mixing compartment and coupled with a small magnet (d = 3 mm) for binding to the intended RNA. All procedures were executed using a centrifuge machine manufactured by Optolane (Fig. S9).

 

 

As demonstrated in Fig. 6a, the whole chip process was conducted by using food dye for visualization of leakage testing and real blood samples for RNA extraction which shown in left and right images of each step, correspondingly. A variety of food dyes were pre-loaded onto the LoaD to illustrate each step and validate the functional capabilities of the LoaD in relation to leakage, separation, and mixing criteria (step 1) as following the same process in Fig. 2e. Under the centrifugal force of 2000 rpm per 1 min, no leakage was observed between compartments, and the buffer solutions were efficiently conveyed to the mixing compartment without any backflow to the primary channel. We proceeded with the whole operations (step 1–10) for evaluating all the compartments and obtained similar outcomes. This indicated that the whole functions of our R2R-manufactured devices were successfully tested. As a result, the LoaD was subsequently utilized for testing whole blood for the extraction and validation of RNA. The reagents and procedures are summarized in Table S3. 150 μl of whole blood (step 1) were loaded onto the LoaD and centrifuged at 2000 rpm in 3 min for successfully separating red blood cells (RBCs) and plasma (step 2). The simulation of this separation of plasma from whole blood was reported in our previous work [33]. The blood chamber was optimized on different designs for the easy fabrication and enhancement of the sedimentation rate of RBCs (Fig. S3). Furthermore, the connection channel to the mixing chamber is positioned at a higher elevation in the blood storage chamber to prevent the adhesion of RBCs on the connection channel, allowing the plasma to freely flow into the mixing chamber (step 3). After the plasma separation step, photonic valves 1 and 2 were activated by illuminating them with a laser (808 nm, 500 mW). The iron oxide nanoparticles embedded in the paraffin wax matrix (called ferro-wax) were heated by a laser beam, resulting in the melting of the ferro-wax and moving it from chamber 1 to chambers 2 and 3, thereby opening/closing the connection channel (as shown in Fig. 6b,c and Fig. S4). The chip was then centrifuged at 1000 rpm for 30 s which enabled the transfer of plasma, proteinase K, and VB buffer into the mixing chamber for lysing cell compartments (step 3). Subsequently, photonic valve 3 was opened to allow for the centrifugation-assisted transfer of ethanol to the mixing chamber which precipitated the DNA/RNA released from plasma (step 4) then rotating the devices at a mixing mode for 30 s and incubated at 60 °C for 10 min (step 5). After that, valve 4 was opened to transfer the magnetic nanobeads to the mixing chamber by spinning the chip at 1000 rpm for 15 s (step 6). The DNA/RNA is then attached to the MNPs and magnet in the mixing chamber by mixing and incubating at room temperature for 60 s. The magnet was placed in the mixing chamber from the beginning to reduce the number of steps as shown in Fig. 6d. For further improvement, we can preload magnetic nanobeads in the mixing chamber to avoid issues in improper movement of MNPs to the mixing chamber caused by resistance from PDMS walls. Following the binding process, photonic valve 5 was opened to transfer the aqueous part to the waste chamber. Next, valve 5 should be closed again to prevent waste solution from flowing back into the mixing chamber. Subsequently, photonic valve 6 was opened to release VWM1 and washout any impurities remaining on the mixing chamber and on the DNA/RNA (step 7). After washing, the VWM1 buffer was removed to the waste chamber by reopening valve 5 through centrifugation. This washing process was repeated twice with RWA2, and WE buffer to thoroughly washout all impurities (step 8 and 9). Finally, photonic valve 9 was opened to release ER buffer into the mixing chamber which detached the purified DNA/RNA from the magnet. The eluted DNA/RNA was then transferred to the eluent chamber through valve 10 (step 10). The eluent was then extracted for further analysis by RT-qPCR. The entire purification process of 150 μl blood on the chip could be completed within 30 min.

 

Figure 6. Room-temp PDMS-based LoaD operation. (a) Food dye visualization for testing leakage issues and photonic valves operation and snapshot images of the device for the whole process of RNA extraction from the whole blood, which were described by left and right images of each step correspondingly. The solution moves from the yellow dashed line area to the red dashed line area; (b) and (c) Photonic valve in close (left) and open (right) state. In the close state, ferro-wax was stored in chamber 1 and the connection channel, while assistant chambers 2 and 3 contained no wax (white dye color). In the open state, ferro-wax in the connection channel was melted by a laser and then moved to chambers 2 and 3 to open the channel enabling the transfer of liquid; (d) Magnification of magnet (d = 3 mm) in mixing chamber; (e) Gel electrophoresis of PCR result of on-chip extraction sample. Lane 1 represents the 50 bp ladder, lane 2 displays GAPDH gene in human, and lane 3 shows the amplification plot of SARS-CoV-2 (103 copies/μl) spiked in whole blood, and lane 4 is the SARS-CoV-2 (103 copies/μl) in whole blood and proceeded with conventional extraction method using the same kit as a positive control.
Figure 6. Room-temp PDMS-based LoaD operation. (a) Food dye visualization for testing leakage issues and photonic valves operation and snapshot images of the device for the whole process of RNA extraction from the whole blood, which were described by left and right images of each step correspondingly. The solution moves from the yellow dashed line area to the red dashed line area; (b) and (c) Photonic valve in close (left) and open (right) state. In the close state, ferro-wax was stored in chamber 1 and the connection channel, while assistant chambers 2 and 3 contained no wax (white dye color). In the open state, ferro-wax in the connection channel was melted by a laser and then moved to chambers 2 and 3 to open the channel enabling the transfer of liquid; (d) Magnification of magnet (d = 3 mm) in mixing chamber; (e) Gel electrophoresis of PCR result of on-chip extraction sample. Lane 1 represents the 50 bp ladder, lane 2 displays GAPDH gene in human, and lane 3 shows the amplification plot of SARS-CoV-2 (103 copies/μl) spiked in whole blood, and lane 4 is the SARS-CoV-2 (103 copies/μl) in whole blood and proceeded with conventional extraction method using the same kit as a positive control.

As a demonstration of the feasibility of our method, we performed RT-PCR to further confirm the extraction process by utilizing the LoaD. To validate the on-chip extraction and purification process, we included GAPDH gene primers as an internal control (IC) for house-keeping genes in human. The amplicons were visualized through gel electrophoresis after running a benchtop PCR. In Fig. 6e, lane 1 represents the 50 bp ladder, lane 2 clearly displays a strong band for the GAPDH gene in human, indicating that the plasma was successfully lysed and purified by our device. Next, lane 3 shows the amplification plot of SARS-CoV-2 (103 copies/μl) spiked in whole blood, and lane 4 is the SARS-CoV-2 (103 copies/μl) in whole blood and proceeded with conventional extraction method using same kit as a positive control for comparison with on-chip spiked samples. The intensity of COVID-19 in lane 3 exhibited adequate amplification efficiency compared to the positive sample in lane 4. However, it still demonstrated successful amplification on gel electrophoresis, indicating the extraction and purification of RNA from human whole blood. The weak amplitude may be caused by losing the spiked RNA during the centrifugation of whole blood in the plasma separation chamber. In summary, as a proof-of-concept test, we have successfully employed the R2R additive manufacturing platform to develop a whole blood extraction by utilizing the room-temperature cured PDMS chip and amplified both spiked SARS-CoV-2 and housekeeping gene (GAPDH) using our LoaD.

4. Discussion

The scope of this study was limited in terms of production rate compared to other techniques such as injection molding due to the lab-scale facilities. However, it is certainly possible to scale up this manufacturing process by increasing roll size with optimized mold design as well as reducing the PDMS curing time by adjusting the chemical composition. Firstly, we demonstrated that the production rate could be significantly enhanced approximately ∼7 times compared to the current one by simply optimizing the mold space with the current imprinting roll size (Fig. S10). Therefore, a larger imprinting roll can be employed to enable high throughput industrial-scale manufacturing process. Secondly, the curing time of PDMS is limited by 10 min due to the lack of an efficient dispensing unit that can continuously perform the mixing, degassing, and dispensing the proper amount of uncured PDMS in-line with the imprinting process. By developing this unit, a further study could assess the faster curing time at room temperature. As a result, the cooling condition (−15 °C) can be eliminated by simply adjusting the concentration of catalysts. Furthermore, a PDMS-tape bonding method can be employed for a rapid, simple, inexpensive, and energy efficient laminating method [40] which enable a greater degree of high throughput and sustainability for our proposed manufacturing process.

 

5. Conclusions

In conclusion, we demonstrated the printing-imprinting integrated R2R continuous in-line additive manufacturing platform, called as green R2R platform, for producing the PDMS-based LoaD with lower energy consumption and less by-products. To realize the green R2R platform, we addressed two main technological hurdles: multi-depth mold fabrication and the fast-room temperature-curing PDMS precursor, enabling a rapid imprinting process. Thus, we developed a rapid, cost-effective fabrication method of a multi-depth flexible polymer shim using 3D-printing technology, which overcomes the challenges in traditional molding techniques especially for integrating macro- and micro-sized features. In addition, we unveiled a novel PDMS formulation by utilizing Ashby–Karstedt catalyst that not only could cure quickly at room temperature, but also could gain better mechanical performance than Sylgard 184 standard. Finally, the resulting PDMS-based LoaD could be expandable for on-site RNA/DNA isolation from the large to a small sample size of whole blood (<150 μl). Our novel fabrication method operated at room temperature which eliminated energy consumption for UV light and heat source will pave the way for addressing the challenges in sustainable high-throughput manufacturing of PDMS-based microfluidic devices which have been highly demanded in the era of Coronavirus (COVID-19) pandemics.

 

Supplementary Materials

References

  1. X. Wei, W. Zhou, S.T. Sanjay, J. Zhang, Q. Jin, F. Xu, D.C. Dominguez, X. Li
    Multiplexed instrument-free bar-chart SpinChip integrated with nanoparticle-mediated magnetic aptasensors for visual quantitative detection of multiple pathogens
    Anal. Chem., 90 (2018), 10.1021/acs.analchem.8b02055
  2. M. Dou, S.T. Sanjay, D.C. Dominguez, S. Zhan, X. Li
    A paper/polymer hybrid CD-like microfluidic SpinChip integrated with DNA-functionalized graphene oxide nanosensors for multiplex qLAMP detection
    Chem. Commun., 53 (2017), 10.1039/c7cc03246c
  3. M. Dou, S.T. Sanjay, M. Benhabib, F. Xu, X.J. Li
    Low-cost bioanalysis on paper-based and its hybrid microfluidic platforms
    Talanta, 145 (2015), 10.1016/j.talanta.2015.04.068
  4. H. Tavakoli, W. Zhou, L. Ma, S. Perez, A. Ibarra, F. Xu, S. Zhan, X.J. Li
    Recent advances in microfluidic platforms for single-cell analysis in cancer biology, diagnosis and therapy
    TrAC, Trends Anal. Chem., 117 (2019), 10.1016/j.trac.2019.05.010
  5. F. Hu, J. Li, N. Peng, Z. Li, Z. Zhang, S. Zhao, M. Duan, H. Tian, L. Li, P. Zhang
    Rapid isolation of cfDNA from large-volume whole blood on a centrifugal microfluidic chip based on immiscible phase filtration
    Analyst, 144 (2019), pp. 4162-4174, 10.1039/C9AN00493A
  6. Z. Noroozi, H. Kido, R. Peytavi, R. Nakajima-Sasaki, A. Jasinskas, M. Micic, P.L. Felgner, M.J. Madou
    A multiplexed immunoassay system based upon reciprocating centrifugal microfluidics
    Rev. Sci. Instrum., 82 (2011), 10.1063/1.3597578
  7. R. Gorkin, J. Park, J. Siegrist, M. Amasia, B.S. Lee, J.M. Park, J. Kim, H. Kim, M. Madou, Y.K. Cho
    Centrifugal microfluidics for biomedical applications
    Lab Chip, 10 (2010), 10.1039/b924109d
  8. I.J. Michael, T.H. Kim, V. Sunkara, Y.K. Cho
    Challenges and opportunities of centrifugal microfluidics for extreme point-of-care testing
    Micromachines, 7 (2016), 10.3390/mi7020032
  9. T.H. Kim, V. Sunkara, J. Park, C.J. Kim, H.K. Woo, Y.K. Cho
    A lab-on-a-disc with reversible and thermally stable diaphragm valves
    Lab Chip, 16 (2016), 10.1039/c6lc00629a
  10. M.M. Aeinehvand, F. Ibrahim, S.W. Harun, A. Kazemzadeh, H.A. Rothan, R. Yusof, M. Madou
    Reversible thermo-pneumatic valves on centrifugal microfluidic platforms
    Lab Chip, 15 (2015), 10.1039/c5lc00634a
  11. S.M.A. Mortazavi, P. Tirandazi, M. Normandie, M.S. Saidi
    Efficient batch-mode mixing and flow patterns in a microfluidic centrifugal platform: a numerical and experimental study
    Microsyst. Technol., 23 (2017), 10.1007/s00542-016-3109-7
  12. B.L. Thompson, Y. Ouyang, G.R.M. Duarte, E. Carrilho, S.T. Krauss, J.P. Landers
    Inexpensive, rapid prototyping of microfluidic devices using overhead transparencies and a laser print, cut and laminate fabrication method
    Nat. Protoc., 10 (2015), 10.1038/nprot.2015.051
  13. S. Hosseini, M.M. Aeinehvand, S.M. Uddin, A. Benzina, H.A. Rothan, R. Yusof, L.H. Koole, M.J. Madou, I. Djordjevic, F. Ibrahim
    Microsphere integrated microfluidic disk: synergy of two techniques for rapid and ultrasensitive dengue detection
    Sci. Rep., 5 (2015), 10.1038/srep16485
  14. E. Duffy, R. Padovani, X. He, R. Gorkin, E. Vereshchagina, J. Ducrée, E. Nesterenko, P.N. Nesterenko, D. Brabazon, B. Paull, M. Vázquez
    New strategies for stationary phase integration within centrifugal microfluidic platforms for applications in sample preparation and pre-concentration
    Anal. Methods, 9 (2017), 10.1039/c7ay00127d
  15. Y. Xia, G.M. Whitesides
    Soft lithography
    Annu. Rev. Mater. Sci., 28 (1998), pp. 153-184, 10.1146/annurev.matsci.28.1.153
  16. G.M. Whitesides
    The origins and the future of microfluidics
    Nature, 442 (2006), pp. 368-373, 10.1038/nature05058
  17. L. Gervais, N. De Rooij, E. Delamarche
    Microfluidic chips for point-of-care immunodiagnostics
    Adv. Mater., 23 (2011), 10.1002/adma.201100464
  18. C.D. Chin, V. Linder, S.K. Sia
    Commercialization of microfluidic point-of-care diagnostic devices
    Lab Chip, 12 (2012), 10.1039/c2lc21204h
  19. J. Hiltunen, C. Liedert, M. Hiltunen, O.H. Huttunen, J. Hiitola-Keinänen, S. Aikio, M. Harjanne, M. Kurkinen, L. Hakalahti, L.P. Lee
    Roll-to-roll fabrication of integrated PDMS-paper microfluidics for nucleic acid amplification
    Lab Chip, 18 (2018), 10.1039/c8lc00269j
  20. E.K. Sackmann, A.L. Fulton, D.J. Beebe
    The present and future role of microfluidics in biomedical research
    Nature, 507 (2014), 10.1038/nature13118
  21. N. Kooy, K. Mohamed, L.T. Pin, O.S. Guan
    A review of roll-to-roll nanoimprint lithography | nanoscale research letters | full text
    Nanoscale Res. Lett., 9 (2014)
  22. S.H. Ahn, L.J. Guo
    High-speed roll-to-roll nanoimprint lithography on flexible plastic substrates
    Adv. Mater. (2008), p. 20, 10.1002/adma.200702650
  23. Y. Hirai, S. Yoshida, N. Takagi
    Defect analysis in thermal nanoimprint lithography
    J. Vac. Sci. Technol. B: Microelectronics and Nanometer Structures, 21 (2003), 10.1116/1.1629289
  24. H.N. Chan, Y. Chen, Y. Shu, Y. Chen, Q. Tian, H. Wu
    Direct, one-step molding of 3D-printed structures for convenient fabrication of truly 3D PDMS microfluidic chips
    Microfluid. Nanofluidics, 19 (2015), 10.1007/s10404-014-1542-4
  25. G. Comina, A. Suska, D. Filippini
    PDMS lab-on-a-chip fabrication using 3D printed templates
    Lab Chip, 14 (2014), 10.1039/c3lc50956g
  26. Y. Chen, W. Pei, R. Tang, S. Chen, H. Chen
    Conformal coating of parylene for surface anti-adhesion in polydimethylsiloxane (PDMS) double casting technique
    Sens Actuators A Phys, 189 (2013), 10.1016/j.sna.2012.09.024
  27. Z. Brounstein, J. Zhao, D. Geller, N. Gupta, A. Labouriau
    Long-term thermal aging of modified Sylgard 184 formulations
    Polymers, 13 (2021), 10.3390/polym13183125
  28. C. Pina-Hernandez, J.S. Kim, L.J. Guo, P.F. Fu
    High-throughput and etch-selective nanoimprinting and stamping based on fast-thermal-curing poly(dimethylsiloxane)s
    Adv. Mater., 19 (2007), 10.1002/adma.200601905
  29. M.T. Guler, P. Beyazkilic, C. Elbuken
    A versatile plug microvalve for microfluidic applications
    Sens Actuators A Phys, 265 (2017), pp. 224-230, 10.1016/j.sna.2017.09.001
  30. T.H. Kim, H. Hwang, R. Gorkin, M. Madou, Y.K. Cho
    Geometry effects on blood separation rate on a rotating disc
    Sensor. Actuator. B Chem., 178 (2013), 10.1016/j.snb.2013.01.011
  31. C.J. Kim, J. Park, V. Sunkara, T.H. Kim, Y. Lee, K. Lee, M.H. Kim, Y.K. Cho
    Fully automated, on-site isolation of cfDNA from whole blood for cancer therapy monitoring
    Lab Chip, 18 (2018), 10.1039/c8lc00165k
  32. J. Clark, M. Kaufman, P.S. Fodor
    Mixing enhancement in serpentine micromixers with a non-rectangular cross-section
    Micromachines, 9 (2018), 10.3390/mi9030107
  33. J. Lee, S. Lee, M. Lee, R. Prakash, H. Kim, G. Cho, J. Lee
    Enhancing mixing performance in a rotating disk mixing chamber: a quantitative investigation of the effect of euler and coriolis forces
    Micromachines, 13 (2022), 10.3390/mi13081218
  34. J.M. Park, Y.K. Cho, B.S. Lee, J.G. Lee, C. Ko
    Multifunctional microvalves control by optical illumination on nanoheaters and its application in centrifugal microfluidic devices
    Lab Chip, 7 (2007), 10.1039/b616112j
  35. E.C. Murphy, J.H. Dumont, C.H. Park, G. Kestell, K.S. Lee, A. Labouriau
    Tailoring properties and processing of Sylgard 184: curing time, adhesion, and water affinity
    J. Appl. Polym. Sci., 137 (2020), 10.1002/app.48530
  36. A.K. Antosik, P. Bednarczyk, Z. Czech
    Aging of silicone pressure-sensitive adhesives
    Polym. Bull., 75 (2018), 10.1007/s00289-017-2083-2
  37. K.R. McIntosh, J.N. Cotsell, J. Cumpston, A.W. Norris, N.E. Powell, B.M. Ketola
    The Effect of Accelerated Aging Tests on the Optical Properties of Silicone and EVA Encapsulants
  38. E.D. Cifter, M. Ozdemir-Karatas, A. Cinarli, E. Sancakli, A. Balik, G. Evlioglu
    In vitro study of effects of aging and processing conditions on colour change in maxillofacial silicone elastomers
    BMC Oral Health, 19 (2019), 10.1186/s12903-019-0798-1
  39. J.J. Gary, C.T. Smith
    Pigments and their application in maxillofacial elastomers: a literature review
    J. Prosthet. Dent, 80 (1998), 10.1016/S0022-3913(98)70111-8
  40. C.S. Thompson, A.R. Abate
    Adhesive-based bonding technique for PDMS microfluidic devices
    Lab Chip, 13 (2013), pp. 632-635, 10.1039/C2LC40978J

 

Portable impedance‑sensing device for microorganism characterization in the field

Portable impedance‑sensing device for microorganism characterization in the field

Karim Bouzid 1*, Jesse Greener 2, Sandro Carrara 3 & Benoit Gosselin

A variety of biosensors have been proposed to quickly detect and measure the properties of individual microorganisms among heterogeneous populations, but challenges related to cost, portability, stability, sensitivity, and power consumption limit their applicability. This study proposes a portable microfluidic device based on impedance flow-cytometry and electrical impedance spectroscopy that can detect and quantify the size of microparticles larger than 45 μm, such as algae and microplastics. The system is low cost ($300), portable (5 cm × 5 cm), low-power (1.2 W), and easily fabricated utilizing a 3D-printer and industrial printed circuit board technology. The main novelty we demonstrate is the use of square wave excitation signal for impedance measurements with quadrature phasesensitive detectors. A linked algorithm removes the errors associated to higher order harmonics. After validating the performance of the device for complex impedance models, we used it to detect and differentiate between polyethylene microbeads of sizes between 63 and 83 μm, and buccal cells between 45 and 70 μm. A precision of 3% is reported for the measured impedance and a minimum size of 45 μm is reported for the particle characterization.

Microorganisms are ubiquitous in nature, being found in environments such as lakes, soils, plants, and within animals. Some are involved in well-known bioprocesses such as fermentation in the food and drink industry, and more recently antibiotics and biofuels. New applications are currently researched in the field of biotechnology, with goals to degrade synthetic plastics1– 3, regularize emotions and stress responses using gut microorganisms4,5, monitor climate change and natural habitats6– 8, remediate nuclear wastes9, detect buried landmines10, or judge of the water quality of popular beaches based on the presence of large phytoplankton that produce neurotoxins such as Karenia brevis, Alexandrium fundyense, Dino-physis acuminata, and Pseudo-nitzschia11. However, despite their utmost importance and numerous applications to human and ecological activities, the vast majority of microorganisms are currently not catalogued, their existence having been only extrapolated from the results of recent phylogenetic studies and genomics12,13. Sophisticated sensors and equipment and a thorough understanding of physics, genomics, optics, taxonomy, and biology are necessary to test, characterize, and classify microorganisms, and a wide array of properties can be tested using different bioreceptors14– 16. Studying microorganisms is thus time-consuming and costly, added that microorganisms are too small to be studied with the bare eyes and mutate at a considerably faster rate than animals and plants, making it difficult to characterize them across time12,17,18. Moreover, replicating their heterogeneity, motility and unique behavior in laboratory settings is found to be challenging, especially considering their extreme sensitivity to their environment, where a minute variation in humidity, light intensity, pH, or temperature is enough to stunt the growth of entire populations19. The more resilient microorganisms are the ones most studied in the literature, the best example being the wellknown Escherichia coli.

Following these challenges, the objective of this study is to conceive a portable intelligent biosensor to characterize multiple properties of large microorganisms and microparticles autonomously and directly in their own natural habitat14. The device should be autonomous, requiring little to no supervision. Automated operations should include the retrieval of the important parameters of hundreds to thousands of microparticles per second. This will lead to a high-throughput technique to characterize and differentiate between microorganisms and microparticles polluting the ecosystems. A broad range of approaches currently exists for the characterization and study of microorganisms, including imaging and hyperspectral-based solutions20– 22, mass spectroscopy23, specialized biochemical sensors15,24, and flow cytometry25. Impedance-based measurements, especially when combined with electrical impedance spectroscopy (EIS)26,27 and impedance flow cytometry (IFC)28,29 seems especially promising.

The common way to monitor impedance is to use commercial benchtop instruments. However, those are generally too expensive and bulky for portable applications. Certain commercial LCR meters offer high precision impedance measurement with errors under 0.5%, but those units are costly, high power consuming, heavy, and bulky, which makes them unpractical for high-volume portable applications. Market-available impedance analyzers can be found in portable formats, but their prices are prohibitive for large scale deployment. As an alternative to these instruments, low-power low-cost integrated chips exist with impedance analyzer capabilities30– 33. These chips can be used as all-in-one-package solutions for low-cost impedance analysis, but are not as versatile as benchtop instruments and their excitation frequency often proves insufficient for microorganism characterization. Other portable impedance analyzers reported in Table 1 exist in the literature, based on techniques such as digital-signal-processing (DSP) sine-fitting34, direct digital synthesizer (DDS) EIS35, mixed analog/digital lock-in amplifier (LIA)36, indirect Kramers–Kronig transformation37, but none of these solutions is a perfect match for high-throughput microparticle characterization.

To fill this gap, we present here a low-cost portable impedance biosensor which improves the authors previous sensor design38,39 and concepts from printed circuit board integrated directly in a microfluidic device40. The presented device can autonomously monitor the impedance of large microorganisms at a high throughput directly in their own natural habitats without using any harmful chemicals, and determines their characteristics based on their impedance profile using EIS and IFC. The main novelty of the device is found in its square wave excitation signal and quadrature phase-sensitive detectors (PSDs). It is used with an algorithm to compensate for the high-level harmonics introduced with the square wave signal.

We kindly thank the researchers at Laval University for this collaboration, and for sharing the results obtained with their system.

Principles of impedance‑flow cytometry

The characterization of microorganisms can be performed using their impedance spectrum, which is a function of the resistivity, dielectric constant, and geometry of the substance under test (SUT). The resistivity and dielectric constant depends on the mobility and quantity of charge carriers in the material41. The impedance is defined from the complex Ohm’s law based on the ratio of a voltage signal to a current signal37, as shown in Eq. 1, where Z is the impedance, V is the applied (or measured) voltage, I is the measured (or applied) current, φ is the phase difference between the voltage and current, ω is the angular excitation frequency, and j is the imaginary unit value. EIS measurement consists in injecting an AC sinusoidal waveform of a known voltage or current to the SUT and measuring its respective output current or voltage response, for a certain number of frequency

IFC41,44,46,47 is the technique offering the best results in the literature for whole cell characterization so far. It is a label-free non-invasive impedimetric measurement technique based on the Coulter counter48 to measure the volume displacement of particles flowing in a fluid. The particles are detected using IFC by monitoring the impedance changes observed each time a particle passes over electrodes in a narrow channel. This is the case since the particles and fluid have different impedances. Figure 1 describes the principles of IFC. Electrodes are positioned on the walls of a microchannel and the impedance of a liquid flowing within is measured for several excitation frequency. A pulsed waveform resembling the one shown at the bottom of Fig. 1 is thus retrieved for each frequency when a cell flows in the channel. The shape of this pulse depends on the position and relative impedance of the microparticle and fluid in the channel, while its measured module and phase depends on the

Figure 1. Two microparticles named a and b submerged in a liquid flow from left to right at two different height in a microchannel. (a) Five-electrode configuration proposed by De Ninno51. (b) Observed current responses at the first and second pair of electrode based on the position of the two microparticles a and b, with which the impedance and permittivity can be calculated.

volume (or size) of the particle compared to the dimensions of the channel. When used as a spectroscopy with multiple excitation frequencies, it is possible to adequately characterize the dielectric properties of the cells flowing inside a microchannel, which can be used in association with their impedance models to retrieve important characteristics of microorganisms28,29,45,49. The impedance is generally converted to a complex permittivity since the geometrical parameters of the microchannel and electrodes are known, and used for the characterization. This is a simple and effective technique to count and characterize particles in a fluid, providing information in real-time for feedback control, or data for later analysis or post-processing50.

The high volume factor between the cell and detection area when using coplanar electrodes in IFC creates a fringing effect between the electrodes, which is difficult to modelize. A simple empirical equation is given in51 and52 as an alternative to estimate the size of microparticles when using IFC. The particle diameter D can be estimated using a fit to the cubic root of the combined measured impedance magnitude difference |Z1| and |Z2| observed when a particle passes the first and second electrode pairs in the channel, respectively. This is shown in Eq. 2, where G is a constant that accounts for all the parameters linked to the electronics and fluidics, such as the electrode configurations, magnitude and frequency of the excitation signal, filter bandwidth, channel depth and width, electrodes width, EDL capacitance Cedl , buffer conductivity, and electronics gains. The constant G can be determined empirically by testing the IFC system with circular beads of know diameters, then adjusting G until the estimated diameters match the effective beads diameters used during the test.

From the Randles53 model and from the impedance model of single-shelled cells42,43, it is possible to deduce the optimal excitation frequency range for microorganism characterization, which is found to be between 100 kHz and 10 MHz. For frequencies lower than 100 kHz, the sensibility of the sensor to microparticles is reduced considering that the electrical double-layer (EDL) and ionic diffusion from the Warburg element dominate the measured impedance38,44,46. Above 10 MHz, the PCB dielectric begins to shunt the channel impedance, and the parasitics of the measurement electronics significantly reduces the precision of the results. Above 1 GHz, the dionic reorientation of water molecules also affect the measured impedance. Calibration algorithms can be used to compensate for the errors obtained when using excitation frequencies outside of this range.

Materials

Master Mold Resin

H Series

Material and methods

The design and fabrication of the impedance-sensing system and microfluidic system will be described in this section based on the principles of IFC.

Impedance‑sensing system. The bloc-diagram of the impedance-sensing device is shown in Fig. 2. It is based on a LIA topology, and extracts the amplitude and phase of a high frequency input signal. It features two channels used to perform a differential analysis. A square waveform with frequency ranging from 200 kHz to 200 MHz is created by the clock-system (Si5351, Silicon Lab). This signal is sent to a quadrature generator to create two 90-degree phase-shifted square waveforms at half the clock signal frequency. The in-phase waveform is attenuated to 100 mVpp to keep a safe linear current response that will not harm the cells during the experiment, and then sent to the two differential electrode pairs of the microfluidics system. Two current responses are obtained, which are then amplified and converted to voltage signals by transimpedance amplifiers (TIAs). The outputs of the TIAs are mixed by two phase-sensitive detectors (PSDs). This yields four output signals representing the real and imaginary current responses of two electrode pairs, which can be sampled by the ADCs. The impedance magnitude and phase differences can be retrieved from the real and imaginary impedance components, and are processed by a square-to-sine spectroscopy algorithm to accurately retrieve the cells properties at a high throughput. The differential design is important to decrease the effects of the common noise and increase the sensitivity to the flowing particles46.

Square input signal. IFC and EIS systems are generally performed using sinewaves. The main advantage of square-waves over sinewaves is that they replace the complex hardware associated with conventional LIAs with much simpler clock-based circuitry54. For instance, the digital-to-analog converter (DAC), wave generator, and linear mixer can all be replaced by a simple clock-system and inexpensive controlled switches. This leads to a decrease of the system’s power-consumption and cost, which are of a prime importance in portable applications. The square wave input voltage Vin outputted by the clock-system and sent to the electrodes is defined in the time domain by Eq. (3). Square signals are multi-frequency signals. Looking at their Fourier transform, we see that a square wave is composed of a fundamental frequency followed by odd-frequency harmonics of decreasing amplitudes. An ideal symmetrical square wave of amplitude 2Vo with peaks at −Vo and +Vo follows the geometrical sum of Eq. (3).

An important point to note from Eq. (3) is that the amplitude of the fundamental differs from the one of a sinewave of amplitude 2Vo by a factor of 4 π . Such a difference biases the measurements and affects the precision but can be mostly corrected (up to a couple of percents) by following an algorithm proposed by Subhan54. Another consideration is the introduction of harmonics in the circuit, which raises the noise floor of the system.

Transimpedance amplifiers. Current responses are obtained from the two electrode pairs that follow Ohm’s law for complex impedance. Current responses being difficult to interact with, two transimpedance amplifiers (TIAs) are used to convert them to voltage signals. TIAs are current to voltage converters generally implemented using one operational amplifier, as shown in Fig. 3. The practical implementation also uses a capacitor for stability in parallel with the resistor in the feedback loop. In the simplest case for a square input signal, Eq. (4) represents the voltage output of the TIAs in time VTIA(t) , where |Z| and θ are respectively the impedance magnitude and phase of the SUT for any harmonic angular frequency nω0.

To accommodate for a wide range of input impedances, a programmable gain array (PGA) with a feedback resistor Rf and capacitor Cf is added to the TIAs to control the gain at will. The PGA is achieved using a multiplexer toggled by a microcontroller that can switch between different gain resistors in the feedback loop of the TIAs30. The feedback capacitor is needed by the TIAs to prevent high-frequency ringing. This can cause a limitation for high-frequency measurements since attenuation is expected at frequencies higher than a couple of megahertz because of the time constant of the RC network formed by Rf and Cf. However, the prototype can still be used at higher frequencies with adequate calibration, although with a reduced accuracy associated with the lessened measured signal amplitude. The trade-offs associated with TIAs are described in Orozco55

Phase‑sensitive detectors. The TIAs outputs cannot be sampled directly using an ADC since the frequency of the signals of interest is too high (the relevant harmonics can go as high as 110 MHz considering the five first harmonics of a 10 MHz square signal). A solution consists of using a mixing and filtering stage implemented from a phase-sensitive detector (PSDs). PSDs act as narrowband filter similarly to LIAs to precisely retrieve the amplitude and phase of a signal buried in noise41. PSDs use square signals and an inverter to switch between the original and inverted version of the signal of interest at the frequency of the square reference signal. This switching yields DC components proportional to the real and imaginary current of the SUT’s impedance. The behavior and implementation of the PSD is shown in Fig. 3. The DC values of the real and imaginary components of the current responses at the output of the PSDs are described by Eqs. (5) and (6).

Quadrature generator. Operating the PSDs mixers requires two square signals in quadrature. Those signals can be precisely obtained from a quadrature generator circuit using a comparator and two D-Flip-Flops, as shown in Fig. 3. This technique is ultrawideband and relatively simple to implement but can be used only for low-power binary signals since the current is sunk directly from the low-power flip-flops. Programmable delays are added in the path of the reference signals to compensate for the delays of the TIAs circuits. This way, the measured phase response from the PSDs is only affected by the SUT.

Square to sine spectroscopy algorithm. Now, as can be seen from Eqs. (5) and (6), it is not trivial to recover the impedance magnitude and phase of the fundamental when using PSDs, as is the case with LIAs. Indeed, harmonics of the square excitation signal are present at every odd frequency of the fundamental which adds a systematic error to the impedance measurement54,56. The harmonics present in the square signal are multiplied together by the mixer and pushed to DC along with the desired fundamental frequency. This systematic error is non-trivial as it depends on the impedance response of the SUT.

An algorithm inspired by Subhan54 can be used to cancel the systematic error. The values of square impedance at the harmonics can be subtracted or added to the fundamental impedance following a certain set of rules described in Subhan54. It is thus necessary to measure the entire impedance spectrum before computing the corrected impedance at a given frequency. The real component of the impedance devoid of the systematic error Vsine−ω0 , follows Eq. (7)

where E is the residual error after correction, which depends on the number of frequency points that were subtracted. There is, however, a practical limit to the number of points that can be subtracted considering that the impedance at that frequency must be measured (or extrapolated) beforehand, which might not be possible for high frequency samples. A similar process can be repeated for the quadrature component in Eq. (8). The corrected impedance magnitude and phase of Eqs. (9) and (10) can then be reconstructed

Printed circuit board. Considering that the microparticles that pass in the channel are microscopic, the sensor has to possess a high sensitivity. For the electronics, a thorough understanding of noise and best PCB design practices is required. The impedance-sensing system is made from a four-layer PCB, and has a size of 50 × 50 × 15 mm including the components. The substrate is FR-4 TG150, with minimum spacing of 0.1524 mm and a thickness of 1.5 mm. Finally, the surface finish is HASL with 1 oz copper. The final PCB with all components is shown in Fig. 2.

Microcontroller unit. The IFC system uses a MSP430F5529 as microcontroller unit (MCU). The MSP430F5529 is a mixed signal MCU used in low-power applications. It dissipates about 6 mW when active and 24 μW when in low-power mode. 6 channels of the 12-bit ADCs are used by the impedance-sensing system to measure the real and imaginary values of the outputs of the phase sensitive detectors ℜ(VPSD−ω0 ) and ℑ(VPSD−ω0 ) of the two electrode responses, as well as the 5V power-supply voltage VDD and the battery voltage. The I2C module is used to program the clock-system and modify the excitation signal frequency. The UART module is used with the external integrated chip FT232RL to transfer the data to a nearby computer using Bluetooth or USB 2.0. A couple of I/O pins are used to enable the power-supplies and status LEDs, modify the gains of the PGA on the fly, reset the phase of the measurement by enabling or disabling the flip-flops of the quadrature generator, and reset the MCU.

Microfluidics system. The microfluidics system created for this study encompasses the micro-electrodes designed on PCB and a PDMS microchannel squeezed hermetically between 3d-printed components.

Microchannel. The microfluidic system manufactured in this study has an inlet and an outlet, where the liquid respectively enters and exits the device, fitted to soft Tygon thermoplastic tubing57. The inlet tube is linked to a glass syringe connected to a precise motorized syringe pump by Cole-Parmer model CP-120 that compresses the syringe at a constant programmable rate. The SUT flows from the syringe to the tubes before entering the inlet. It then reaches the PDMS microchannel where it is sensed by the PCB microelectrodes. The liquid finally exits through the outlet tube, which is connected to a waste container. The microfluidics system is shown in Fig. 4.

The whole fabrication process is described in Bouzid38. A mold is initially drawn on a CAD software such as Solidworks. The drawn model is sent as a .STL file to the CADworks3D software to be meshed. This new meshed model is used by the stereolithography 3d-printer CADworks3d H50-405 to print a 3D-mold using Master Mold for PDMS Device Photopolymer Resin TM. The resin is then rinsed with IPA (90%) or methyl hydrate for 5 min and blow dried using an air gun. The mold is then cured using UV light around 400 nm in a LED light curing box for 50 min. PDMS ( SYLGARDTM 184 Silicone Elastomer Base) and a curing agent ( SYLGARDTM 184 Silicone Elastomer Curing Agent) are mixed at a ratio of 10:1 and degassed by letting the solution rest for 60 min. The mixture is put in the mold and cooked on a hot plate for 50 min at 70◦ , then dried overnight at 40 ◦C58. A scalpel is used to gently prick-off the channel from the mold. The surfaces of the PDMS microchannel are then exposed to plasma at 600 KPa for one min. Polyethylene glycol is immediately applied on the surface to keep its hydrophilicity for longer periods of times. After a 10 min wait time, the PDMS microchannel is cooked for 10 min at 130 ◦ C on a hot plate58. A dry gun is blown on the channel to take away any residues. The PCB electrodes are aligned on the microchannel and a pressure is applied to seal them. The PDMS channel and PCB electrodes are sealed tight by a system of 3d-printed squeezers that compress the channel and PCB electrodes. Those are tightened together by bolts, hermetically sealing the microchannel and PCB electrode due to the flexible nature of PDMS. Finally, tubes are inserted at the inlet and outlet of the PDMS microchannel, concluding on the whole process. The microfluidics system is thus hermetic and easy to handle.

The shape of the mold is shown in Fig. 4. The volume fraction (i.e. the ratio between the volume of the microparticle and volume of the fluid affected by the electrical field of the electrodes) should be maximized in IFC applications to obtain the highest sensitivity. Maximizing the volume fraction requires that the channel and electrodes size be identical to the microparticle of interest. In practice, the channel must be larger by a given safety factor to allow the fluid to circulate without clogging. Hemispherical bubble traps based on the work of Kang59 were added to the channel to reduce the quantity of air bubbles reaching the electrodes which could falsify the measured impedance. Using the stereolithography 3d-printer H50-405, a theoretical resolution of 30 μm is possible, although in practice, the minimum size of a printable channel without major defects is 90 μm. For the case of this study, a channel size of 180 μm was chosen. The PDMS cured in this mold solidifies into a structure with four openings where are placed the bolts. The electrode and squeezers share the same shape and openings for the bolts as the PDMS.

Electrodes. Coplanar electrodes are chosen in this design because they are 2D structures that can easily be made using a lithography mask or directly on a PCB28. The non-homogeneous electrical field distribution of coplanar electrodes does introduce errors in the measurement since the cell’s vertical position in the channel are subject to varying levels of electrical field. A higher particle in the channel typically experiences weaker electrical field than a low particle, which results in a lower perceived amplitude51,52, as was shown in Fig. 1. Since the amplitude of the spike is used to infer the cell properties, a significant error is thus observed. Three solutions can be used to counter this problem: (1) Using parallel facing electrodes placed diagonally opposed in the channel instead of coplanar ones52. (2) Using centering techniques such as dielectrophoresis, acoustophoresis, inertial focusing and sheath flows51. (3) Using coplanar electrodes with distinctive geometry to obtain additional information about the vertical position of the particle in the channel41,51. The 5-electrode configuration described in De Ninno51 is one of those distinctive geometry and has been chosen for this study, and is shown in Fig. 2. The relative prominence of the signal obtained from such a configuration can be used to correct the measured particle size. The downside of such a technique is that it reduces the sensibility of the sensor since the intricate electrode geometry increases the sensing volume60

The electrodes in this study are fabricated on a one-layer PCB. They have a size of 46 × 21 × 1.6 mm. The PCB uses the conventional substrate FR-4 TG130 in the exact shape of the microfluidics channel. The employed surface finish is immersion gold (ENIG) (1U”) with 1 oz copper. Inert metals such as gold or platinum are used for the electrodes because of their convenience in casting for small dimensions, for their unlimited lifetime, and since other types of electrodes such as Ag/AgCl are unsuitable for high excitation frequency28. The electrodes are 101.6 μm wide and are separated by 101.6 μm each. The PCB electrodes and their alignment with the microchannel are shown in Fig. 4.

Figure 5. Bode plot of the impedance magnitude and phase response of a 10-k discrete resistor in series with the parallel combination of a 4.47-k resistor and a 100-pF capacitor. 320 samples were taken from the SUT for each frequency, and the average and standard deviations are calculated and displayed on the error bar on the left. Four sets of data are displayed, the measured raw impedance, the raw impedance after calibration, the calibrated impedance after transformation using the square to sine spectroscopy algorithm, and the theoretical impedance of the SUT.

Results and discussion

To test and calibrate the performances of the device, EIS analysis of discrete resistors and capacitors forming complex circuits were measured. EIS analysis on saline water, as well as the detection and characterization results of microbeads and buccal cells using IFC also follow. Finally, the performance of the system is discussed.

Complex impedance circuit. The performances of the impedance-sensing system are measured from the EIS analysis of a 10 k discrete resistor in series with a parallel combination of a 4.7 k resistor and a 100 pF capacitor. The impedance-sensing system has a lowest excitation frequency of 20 kHz and a highest of 12 MHz. The square excitation signal is initialized at the lowest frequency, samples 64 data points, then the frequency is incremented logarithmically until the end frequency is reached. When that is the case, the frequency is reinitialized to the lowest frequency, and the process begins anew. The impedance magnitude and phase are shown in Fig. 5, and were recorded for about 34 s at a sampling rate of 655 Sps, which amounts to about 320 data points per frequency.

To solve some of these issues and linearize the sensor, a calibration using a look-up table is realized using resistors of known values. Since the parasitics are singular to the electronics, the same nonlinearity will be found for different values of resistance, which can be used as a frequency-dependent factor to linearize the magnitude and phase curves. The square-to-sine conversion adapted from Subhan54 is then performed on the calibrated dataset. The raw, calibrated and converted-to-sine results are shown in Fig. 5 for the median impedance at each frequency points. At high frequencies, a bias is observed both in the impedance and phase since higher frequency data points are not available to perform the square to sine conversion from Eqs. (7) and (8). A way to solve this issue would be to extrapolate the behavior of the system from the previous points and use that extrapolation in the square to sine conversion. For the sake of simplicity, no such correction was attempted in this study. Apart from that bias, errors of less than 3% are observed for the magnitude and phase, for the frequency range considered in the spectroscopy. This is comparable to the commercial devices presented in Table 1, at a fraction of the power consumption and cost.

Saline solution. Following the proof of functioning and the calibration, we measure the impedance spectrum of a complex system. A solution of saline water at 22 ◦ C is passed in the 180 μm wide PDMS microchannel of the microfluidic system, and the EIS is measured. The calibrated and converted EIS curves for the two pairs of electrodes follow the behavior of a series capacitor and resistor, as expected from the Randles model. This dataset will be used to calculate the corrected impedance after using the square-to-sine transformation algorithm after the addition of microbeads and buccal cells

The measured impedance varies slightly according to the pressure exerted by the liquid flow. This difference is caused by a slight contraction of the PDMS walls caused by the liquid pressure, which also increases the liquid volume measured by the electrodes. The same effect can be observed for variations in temperature of the liquid. Thus, the liquid pressure and temperature are controlled for the whole duration of the experimentation.

Microbeads. In order to replicate more accurately the expected behavior from cells and microparticles, polyethylene microbeads are added to the previous saline solution. This SUT is kept at the same conditions as before, at 22 ◦ C and passed in a 180 μm wide microchannel. Only one square excitation signal is used this time, at a frequency of 1 MHz. The corrected impedance can then be calculated with Eq. (7) using the measured impedance of the microbeads as fundamental and the EIS of saline water as the harmonics. Considering the small difference in impedance spectroscopy between both tests, errors less than 1% are expected.

To avoid overloading the MCU, the impedance is sampled at a fixed high-frequency rate of 5461 Sps, while data is saved to memory only when a significant difference is observed in the real parts of the measured impedance of either electrodes. When that condition is detected, a burst of 64 consecutive measurement points is saved. This method produces regularly fixed data point with dense bursts of data when an event is detected. This event detection can be caused by a microparticle passing in the channel, or by sudden changes in liquid property or microchannel geometry. Signal processing is performed offline to retrieve only the events associated with a particle detection. Firstly, the average of the magnitude difference is removed using wavelet decomposition, the signal is then de-noised, low-pass filtered, and smoothed so that the impedance spikes caused by the particles are easier to recognize. A particle detection algorithm is used on this dataset to recover the positions of the peaks. A peak detection algorithm is first used, followed by a weak supervision approach using Snorkel61 to discriminate between the peaks obtained from microbeads, bubbles, or any other outliers. Most of the oddly shaped, or weird behaving particles are thus removed from the dataset automatically. With the particles peak locations, it is possible to recover the amplitude and width of the patterns, which are used to estimate the microbeads size using Eq. (2).

As an example, the pattern in (a) of Fig. 6 is studied. The first electrode has an impedance magnitude difference of 230 , while the second electrode has an impedance magnitude difference of 250 . Its G constant is estimated from the dataset to be around 10. This leads us to a diameter value of 78 μm. It is also possible to measure the particle velocity by dividing the distance between the electrode pairs L with the time it took for the particle to go from one electrode to the other t (which is the time difference between the two impedance maximums). The time it took for the particle to pass the electrodes is found to be 2.9 ms, while the distance between the electrode pairs is of 406 μm. This leads to a velocity around 14 cm/s. This flow rate has been found empirically to provide good and reliable measurements, since lower flow-rate can cause particles to stick to the walls of the channels or electrodes, and higher flow-rate are associated with a decreased time resolution. The fastest useful flow-rate for this sampling rate is when a minimum of 5 points are detected for a full particle. Any less than that is considered an outlier by the classifier. This leads to a maximum theoretical flow-rate of 89 cm/s.

Buccal cells. The proposed system works to detect cells of a maximum size fixed by the width of the microchannel, and of a minimum size fixed by the sensibility and inherent noise of the sensor. For the case of this study, this leads to a minimum and maximum cell size of 45 μm and 180 μm respectively. An easy to test cell that fits those size requirements are those found in the mouths, the so-called buccal cells, with sizes typically ranging between 50 and 60 μm62. Those cells were scraped from the tongue and cheek of the corresponding author, and mixed with the same saline solution used in the previous tests. An example of a cell detection is shown in Fig. 6, where the 1st and 2nd pair of electrodes each detected successive impedance events, which were used to characterize the cell size around 51 μm.

Collected datasets. The true potential of IFC sensors lies in how automatizable the sampling and testing process can be for biological studies. It could be imagined that a team of biologists collects the impedance data of millions of cells and particles using the portable device described in this study, and efficiently extract the

Figure 6. Magnitude difference of both electrode pairs (a) when a 78 μm polyethylene microbead passes in the 180 μm wide microchannel. (c) when a 51 μm buccal cell passes in the 180 μm wide microchannel. Distribution of (b) the 63–83 μm microbeads population and (d) the buccal cells population

important information of these cells using machine learning and high-end signal processing. As a small-scale example, two datasets were collected from the proposed system and passed into the peak detection and classification algorithm. 447 beads were detected from 617 detection events, and 360 buccal cells were successfully detected from 2823 detection events. The debris in the saline solutions from the cheek and tongue scraping, and the low impedance difference obtained from the buccal cells explain why so much detection events were detected by the algorithm compared to the number of actual buccal cells. The diameters of the microbeads and buccal cells were estimated as described previously and compiled in the histograms of Fig. 6. A minimum impedance difference threshold is used to classify what counts as a detection event from the measurement noise. This lower threshold means that the microparticles of sizes below 45 μm that are present in the solution are not registered by the algorithms. It could be said that the effective sensitivity of the impedance-sensing system to detect small particles is of 45 μm when used with a 180 μm wide microchannel

Measured performance. The performance and characteristics of the presented impedance-sensing device and microfluidic system are summarized in Table 2. The impedance-sensing system created for this study is the first found in the scientific literature to achieve great sensitivity level over wide frequency and impedance range while boasting a small size, low-cost, and low power-consumption. The impedance-sensing device coupled with the microfluidics systems are effectively capable of measuring and estimating the properties of the microparticles of sizes going as low as 45 μm when used within a 180 μm wide microchannel. The dimensions of the microchannel are fixed by the limitations of the 3D-printer, which could be improved for this study by using a 3D-printer such as the one designed by Gong63. This homemade 3D-printer is specifically made for microfluidics and can attain truly microscopic scales of 18 × 20 μm by modifying the type of resin used and optimizing the stereolithographic process. This higher resolution would help increase the sensitivity of our device for smaller particle detection. The impedancesensing device takes 50 mm × 50 mm × 15 mm of space, while the microfluidics system is 46 mm × 25 mm × 50 mm, with a combined weight of 300 g, making them portable enough to be put in a backpack for applications in the field. The electrode pairs in the microchannel are separated by 424 μm and each have a width of 106 μm compared to the microchannel size of 180 μm × 180 μm. The impedance-sensing system only needs 1.2 W to

function adequately, and is powered by a low-voltage battery of 2.5–3 V. The power consumption of the system is sufficient for portable applications and could be powered for a couple of hours at a time. The power consumption could, however, be greatly reduced by creating a custom ASIC instead of using discrete components. Indeed, the vast majority of the power (about 80%) is dissipated in the op-amps, while they serve only to do basic functions such as inverting and amplifying signals that could be replaced by optimized high frequency transistors. The impedance-sensing system costs around $300, while the microfluidics system costs only $10 per microchannel excluding the initial cost of the 3D printer. The impedance measurement range between 200 and 120 k is similar to the portable impedance analyzer described in the literature, such as the ones from Al-Ali37, and Radil34. The frequency range is adequate for IFC applications, with the important frequency range between 100 kHz and 10 MHz covered by the device. The upper frequency limit of 12 MHz observed in this work is fixed by the limitations of the op-amps used in the TIAs. The limited bandwidth of the op-amps attenuates the harmonics of the square signal, which progressively modifies the square excitation signal into a sinusoidal shape. This introduces significant disparity for frequencies higher than 12 MHz which goes above the 3% precision reported for the device. The device can theoretically be used with excitation frequencies as high as 100 MHz, but the reported error would increase significantly. Finally, an excitation voltage of only 100 mVpp is used, which is low enough to not affect most microorganisms in that size range.

Conclusion

This study succeeded in creating an autonomous device for the characterization of microorganisms in the fields. Using an inexpensive 3D printing manufacturing technique and standard printed circuit board technology, the presented device can detect and characterize microorganisms larger than 45 μm. The device succeeded in characterizing and differentiating between buccal cells and polyethylene microbeads. Future work will focus on improving the sensibility of the sensor to characterize microparticles of smaller sizes, as well as increasing the number of parameters that can be monitored to achieve a better characterization. Following the recent advances in micro-optical systems, adding a low-power 3D-imaging system to the device will be investigated.


Data availability
The Python, MATLAB, and C source code for this project, and the dataset obtained from the device are available upon reasonable request to the corresponding author. Python was used to sample the dataset from the device. C code was used to interact with the sensors and MCU. Post processing was done in Matlab and Python.

Tissue-Engineered Cochlear Fibrosis Model Links Complex Impedance to Fibrosis Formation for Cochlear Implant Patients

Tissue-Engineered Cochlear Fibrosis Model Links Complex Impedance to Fibrosis Formation for Cochlear Implant Patients

Simone R. de Rijk, Alexander J. Boys, Iwan V. Roberts, Chen Jiang, Charlotte Garcia, Róisín M. Owens, Manohar Bance

Cochlear implants are a life-changing technology for those with severe sensorineural hearing loss, partially restoring hearing through direct electrical stimulation of the auditory nerve. However, they are known to elicit an immune response resulting in fibrotic tissue formation in the cochlea that is linked to residual hearing loss and suboptimal outcomes. Intracochlear fibrosis is difficult to track without postmortem histology, and no specific electrical marker for fibrosis exists. In this study, a tissue-engineered model of cochlear fibrosis is developed following implant placement to examine the electrical characteristics associated with fibrotic tissue formation around electrodes. The model is characterized using electrochemical impedance spectroscopy and an increase in the resistance and a decrease in capacitance of the tissue using a representative circuit are found. This result informs a new marker of fibrosis progression over time that is extractable from voltage waveform responses, which can be directly measured in cochlear implant patients. This marker is tested in a small sample size of recently implanted cochlear implant patients, showing a significant increase over two postoperative timepoints. Using this system, complex impedance is demonstrated as a marker of fibrosis progression that is directly measurable from cochlear implants to enable real-time tracking of fibrosis formation in patients, creating opportunities for earlier treatment intervention to improve cochlear implant efficacy.

We kindly thank the researchers at Laval University for this collaboration, and for sharing the results obtained with their system.

1 Introduction

Hearing loss affects 20% of the world's population with an ≈5% needing clinical intervention.[1] Cochlear implants (CIs) are life-changing technology that allows people with severe hearing loss to hear and achieve speech perception.[2] CIs, arguably the most successful neural prostheses to date, transform sounds into electrical pulses that directly stimulate the auditory nerve.[3] The intracochlear[4, 5] multi-electrode array of CIs takes advantage of tonotopic, frequency-dependent, organization of the cochlea by electrically stimulating different parts of the auditory nerve processes to convey different sounds.[3] However, these implants are known to cause fibrosis when placed, which can limit their efficacy long-term.

Implants are known to elicit an inflammatory response, associated with fibrotic encapsulation.[6-8] Fibrosis is of particular concern for neural implants, as the fibrotic capsule can limit electrical signal transduction to surrounding tissues.[8-11] For CIs, the inflammatory response is driven by mechanical trauma during insertion, which results in protein absorption, particularly fibrin,[9, 12-15] extracellular matrix (ECM) deposition, and subsequent cell-mediated contraction into a dense fibrotic capsule around an implant.[11, 16, 17] This process is initiated by various immune cells, such as macrophages, before infiltration by fibroblastic cells that lay down further ECM.[9, 12-15] The extent of new tissue formation in the cochlea can vary from a thin fibrous sheath surrounding the electrode array, to new bone formation.[18-22] This overall inflammatory response to CI insertion has been associated with the loss of intracochlear hair cells and auditory neurons[4, 9, 23-25] and restriction of basilar membrane vibration,[26] which subsequently results in residual acoustic hearing loss.[18, 23, 27-29] Studying cochlear fibrosis can inform new treatments, such as efficacy for drug-eluting electrode arrays, and may provide insight into current treatments to prevent fibrosis[4, 5, 27, 30-32] or emerging treatments, such as cell and gene therapies and optogenetic stimulation.[33, 34]

Despite the relevance of intracochlear fibrosis to residual hearing loss, we possess few methods for tracking fibrosis in CI patients.[18] One indirect method is the measurement of contact “impedances,” an increase of which has been associated with fibrotic tissue formation and residual hearing loss in patients.[5, 19, 35-37] While not actual electrical impedance measurements, contact impedances are voltage responses at a single timepoint to a biphasic charge-balanced current pulse, normalized to the amplitude of input current.[38] These have been investigated in preclinical models with posthumous evaluation[4, 5] but lack real-time measurements and vary significantly from human anatomy. Other studies have explored fibrosis using 2D in vitro models,[36, 39] but these studies lack the complexity of the 3D matrix deposition and contraction seen in vivo.[40] Tissue-engineered models provide a potential in-roads for examining the relationship of fibrosis to electrical measurements, given their capabilities for simulating cellular phenomena in 3D, such as tissue contraction.[41, 42] Further, these models could be coupled to clinical-grade implants, where the overall frequency and responses of the model can be studied. While replication of the immune system is challenging in vitro, resultant tissue-engineered ECMs can possess similar properties to in vivo fibrotic tissue by harnessing the capabilities of cells to remodel tissue-engineered matrices.[43, 44] Electrochemical impedance spectroscopy (EIS) has long been used to measure cell and tissue behavior such as proliferation, differentiation, cell adhesion, detect various forms of malignancies, monitor 3D cell cultures, and detect liver fibrosis.[45-47] By measuring frequency response of impedance, EIS provides higher-content information on tissues, lending promise for tracking fibrosis progression.

In this study, we investigate complex impedance as a biomarker for fibrosis progression by developing a 3D tissue-engineered model of cochlear fibrosis. We replicate the intracochlear fibrotic environment by encapsulating clinical-grade CI electrode arrays inside tissue-engineered fibroblast-seeded fibrin gel constructs. We show significant and consistent changes in complex impedance over time, with which we produce a realistic electrical circuit model for fibrosis development. We also utilize full voltage waveform measurements to propose an electrical marker of fibrosis development that could be implemented clinically, finding similar electrical behavior in our measurements of patient samples. The results presented in this study and the markers we propose will enable us to track cochlear fibrosis progression in real-time, allowing for earlier treatment intervention for combating residual hearing loss for CI patients.

2 Results

2.1 Development of a Tissue Engineered Model of Cochlear Implant Fibrosis

 We modeled cochlear fibrosis by producing a fibrous sheath around a clinical-grade cochlear electrode array. To generate this model, we injected molded fibrin gels containing fibroblasts into a 3D-printed mold, with the cochlear electrode array centered on the axis. These electrode arrays with cell-seeded gel constructs were suspended in culture media inside a conical bioreactor, to set the electrode array and ground electrode location for consistent electrical measurement (Figure 1). Fibrin was chosen as the biological scaffold as it is the provisional matrix laid down during wound healing,[48] both post-implantation of cochlear implants[9, 12-15] and other implanted biomaterials scenarios.[49] Given the composition of the constructs, cells were expected to interact with and contract fibrin gels into a denser conformation around the array.[42] To promote increased interaction between fibroblasts and gel, a contractile medium was formulated, along with media supplementation of TGF-β1 to promote fibroblast differentiation into a more fibrotic-like, contractile phenotype.[49-51] Images were captured to track contraction throughout the experiment. We measured EIS and voltage waveforms at six timepoints over the course of 14 days (days 2, 4, 7, 9, 11, and 14) and took concurrent images beginning on day 0 (Figure 1A; Figure S1A,B, Supporting Information). To examine the effects of electrical stimulation, from our measurement criteria, we also utilized an unstimulated control.

Figure 1 Open in figure viewer PowerPoint Schematic of 3D bioreactor setup. A) Schematic of the tissue-engineered cochlear fibrosis model construct including a cochlear implant electrode array encapsulated with a fibrin gel with 3D-seeded fibroblasts. e1 represents the first/most apical electrode. B) Image of the three apical electrode contacts including connecting wires.

Figure 2 Open in figure viewer PowerPoint Contraction and histology analysis of the constructs. A) Representative image set showing contraction of the construct over the course of the experiment. The length of the construct was calculated using the known mid-to-mid contact spacing of the electrode arrays to calculate the scale of the images for each electrode array and timepoint separately, which was then used to calculate the length of the construct. B) Relative contraction of the constructs, normalized to day 0 absolute length, over time. Single datapoints are shown in grey lines with open circles. Mean ± standard deviation is shown for the stimulated and unstimulated groups (ns = not significant, univariate n-way ANCOVA) in bold as well as all constructs. Relative contraction is significant over time (p < 0.001, univariate n-way ANCOVA) with the inflection point between days 7 and 9 (Tukey's post hoc test). C) Hematoxylin & eosin (H&E) and polarized picrosirius red (PSR) stained histology slices, transverse and longitudinal sections, of two constructs. “*” represents the location of the electrode array. H&E staining shows higher density of cells at the lateral and medial edges of the construct. PSR reveals birefringence and thus collagen formation. No differences between the stimulated and unstimulated constructs are visible.

Next, we performed histology to retrieve information on cellular orientation and extracellular matrix morphology (Figure 2C; Figure S2, Supporting Information). Hematoxylin and Eosin (H&E) staining revealed a higher lateral density of cells with denser medial ECM, indicating cellular repositioning with respect to available nutrients. We performed picrosirius red (PSR) staining for oriented fibrillar collagen[52, 53] to examine for collagen orientation. Some coloration is evident (Figure 2C), indicating that the fibroblasts are producing dense, fiber-like collagen bundles, most likely via mechanical boundary conditions,[54] which are inherently applied by the presence of the CI array. We also utilized a Ki-67 stain, a marker of cell proliferation,[55] to confirm that cells within the constructs where proliferating in all cases (Figure S2, Supporting Information).

Use of electrical stimulation as a method to prevent CI fibrosis has sparked recent interest.[35, 36, 56, 57] Therefore, we tested the effect of stimulation on contraction, while correcting for additional sources of variation. No significant effect of stimulation was found (F = 1.59, p = 0.23, df = 1, univariate n-way ANOVA, relative contraction on day 14-dependent variable, stimulation, electrode design-fixed factors, and experiment number-random factor). We also did not observe any differences from our histological analysis (H&E, PSR, Ki-67). To confirm these similarities, we performed a Hoechst fluorescence assay to quantify DNA at day 14 (Figure S3, Supporting Information). No significant effect of stimulation was found (F = 0.08, p = 0.78, df = 1, univariate n-way ANOVA, DNA per µg dry weight as dependent variable (n = 10), stimulation-fixed factor (n = 4 simulated, n = 6 unstimulated), experiment number-random factor (n = 2 Exp1, n = 4 Exp2, n = 4 Exp3)). These results agree with studies investigating the effects of early switch-on and more extensive stimulation post-operatively, which show little effect on long-term markers of fibrosis formation.[58, 59]

Given that constructs axially contract, we found in some cases, constructs would contract away from electrodes that were covered on day 0. To understand the 3D structure of the constructs relative to positioning along the arrays, samples (n = 2, 1 stimulated, 1 unstimulated) were stained for DNA and actin. The edge and center of the constructs are visible, showing dense cells attached to the arrays (Figure 3A). This allowed us to visualize areas that had become uncovered during contraction, where we observed no evidence of construct remnants. We also observed cellular spreading on an exposed electrode at the trailing edge of the construct (Figure 3B). This observation indicates that cells can adsorb directly onto electrodes, potentially effecting electrode–electrolyte (EE) interface during stimulation. However, as no residual construct remained in areas of arrays that had become uncovered during contraction, this interface is potentially recoverable.

Apparatus &Materials

Master Mold Resin

M Series

Figure 3 Open in figure viewer PowerPoint Confocal fluorescence imaging of the construct. 3D orientation of the construct, as fixed on day 14, related to the electrode array. The nuclei are stained blue via DNA staining with Hoechst 33258, while actin was stained with phalloidin-iFluor 594 showing in red. A) Edge and mid-construct images without stimulation. B) Total and close-up of a stimulated construct. Both constructs show attachment of the cells on the electrode surfaces. Actin fibers in the cells can be seen spreading out over the surface of the electrodes and numerous cells are attached to a singular electrode alone.

Within the statistical tests described in this section, an effect of experiment number was found on relative contraction and DNA quantification (Figures S1C and S3, Supporting Information), possibly indicating some variance within the fibroblast cell line used for this study.

2.2 Electrochemical Impedance Spectroscopy Shows Significant Changes in the Bulk of the Gel

 We hypothesized that complex impedance would change as measured via EIS with cellular contraction and construct remodeling. To test this hypothesis, we tracked complex impedance spectra over six timepoints to day 14. These spectra were fitted to a circuit model (Figure 4A), consisting of a constant phase element (CPE) representing EE interface, a resistor (R1) in parallel with a capacitor (C) representing the bulk of the construct, and an additional resistor (R2) representing the resistance of the media and ground. Since we did not expect a major contribution to overall impedance with changes in cell media and pathway to ground, R2 was fixed based on the earliest available timepoint for each electrode. The EE interface and bulk of the construct have been hypothesized to change during cochlear fibrosis.[35, 36, 39, 60, 61] So, these elements were fitted without constraints. The average weighted sum-of-squares, proportional to the average percentage error between original and fitted data, was <1% for most fittings and at least <5% for all fittings (Figure S4A, Supporting Information). An example of impedance magnitude and phase angle over time, for both measured and modeled data, for 1 electrode with the construct on throughout the experiment can be seen in Figure 4B and without construct in Figure S4B (Supporting Information). An increase in absolute impedance magnitude is seen at higher frequencies (>10 kHz), while phase angle decreased across most frequencies in this example.

Figure 4 Open in figure viewer PowerPoint Complex impedance measured with electrochemical impedance spectroscopy (EIS). EIS was measured on all electrodes, regardless of having an open circuit (e.g., air bubble or broken electrode). The exclusion criteria, as described in the materials & methods section, led to n = 231 recordings with construct and n = 153 without construct being included (of a combined total of n = 528 recordings). A) Proposed equivalent circuit of the 3D bioreactor model with a constant phase element (CPE) representing the electrode-electrolyte (EE) interface, a resistor (R1) in parallel with a capacitor (C) representing the bulk of the construct, and an additional resistor (R2) representing the media and ground (GND). B) Absolute impedance magnitude and phase angle of an example electrode over time, showing both measured and modeled values. Measured data are shown as mean ± standard deviation. C) Modeled circuit elements over time of all timepoints and electrodes with construct (and thus modeling fibrosis) on the electrode. Individual data are shown in grey. The arithmetic mean ± standard deviation is shown in bold black, except for C where the geometric mean and standard deviation is shown. CPE-P and CPE-T show no significant (ns) changes from day 2 to day 14 (univariate n-way ANOVA, Tukey's post hoc test). R1 shows a significant increase from day 2 to day 14 (****p < 0.001, univariate n-way ANOVA, Tukey's post hoc test), while C shows a significant decrease (****p < 0.001, univariate n-way ANOVA, Tukey's post hoc test).

Fitted circuit elements over time with construct on showed CPE phase (CPE-P) and magnitude (CPE-T) stay constant, while circuit element R1 increased and C decreased (Figure 4C). This change is not seen for electrodes without constructs on them (Figure S4C). R1 showed a large significant effect of time (F = 21.51, p < 0.001, df = 5), with the inflection point between days 7 and 9 as revealed by Tukey's post hoc test, and overall significant change between day 2 and day 14 (p<0.001). C also showed a large significant effect of time (F = 15.12, p < 0.001, df = 5), with the inflection point between days 4 and 7 as revealed by Tukey's post hoc test, and an overall significant decrease between days 2 and 14 (p < 0.001). CPE-P and CPE-T remained largely. When comparing to fitted circuit elements for electrodes without construct, no significant effect of time is found for CPE-P (F = 1.06, p = 0.38, df = 5), CPE-T (F = 0.79, p = 0.56, df = 5), R1 (F = 0.78, p = 0.56, df = 5), and C (F = 1.85, p = 0.11, df = 5). Overall, these data suggest changes in EIS can be explained by an increase in resistance and decrease in capacitance of the bulk of the construct with no significant changes in EE interface seen.

A commonly studied circuit to model contact impedances in relation to cochlear fibrosis was introduced by Tykocinski et al. and includes a resistor in parallel with a capacitor representing EE interface and a single resistor in series representing the bulk of tissue (Figure S5A, Supporting Information).[62] This circuit is extracted from a voltage waveform (contact impedance timepoints) and models access resistance, initial increase in voltage at the start of the waveform, and polarization impedance, the capacitive build-up after access resistance.[62] Changes in polarization impedance have since been linked to protein adsorption (increase) and resorption (decrease) on the electrode.[35, 39, 57] Changes in access resistance are more commonly associated with changes in bulk tissue surrounding the electrode, where an increase in access resistance is linked to an increase in tissue formation.[36, 39, 56, 60] However, changes are not specific to new tissue formation only, as an increase in access resistance has also been associated with electrode-modiolus distance, translocation of the electrode from one scala to another intracochlearly, extracochlear electrodes, and electrode failure.[63-67] We fitted this circuit to our example data (Figure 4B; Figure S5B, Supporting Information) mainly showing a large error in phase angle for complex impedance. Average weighted sum-of-squares was >10% in all six timepoints (Figure S5C, Supporting Information), suggesting this circuit is too simple to model complex impedance for our model of fibrosis.

2.3 Contact Impedances and Second Phase Peak Ration (SPPR) of Voltage Waveforms Increase Significantly Over Time

To translate the changes in complex impedance to an electrical measurable in patients, we measured voltage waveforms at all timepoints for electrodes with and without construct on (Figure 5A; Figure S6A, Supporting Information). Generally, an increase in voltage over time is observed with construct on the electrode, while no changes are seen without construct on the electrode. When the construct contracts off an electrode, the voltage waveform was seen to normalize back to the level of the waveform at day 2 (Figure S6B).

Figure 5 Open in figure viewer PowerPoint Measured voltage waveforms, contact “impedances” and SPPR over time. Voltage waveforms were only measured when EIS measurements were included and a single pulse did not elicit high voltage waveform responses, leading to n = 221 with construct and n = 115 without construct. A) Example mean voltage waveforms over time for the same electrode as in Figure 4B, with a cathodic-leading biphasic pulse and anodic-leading biphasic pulse as an input. B) Absolute and relative contact “impedances” over time. Individual traces are shown in grey, while mean ± standard deviation is shown in bold black. Absolute contact “impedances” significantly increased over time from day 2 to day 14 (****p < 0.001, univariate n-way ANOVA, Tukey's post hoc test). C) Absolute and relative SPPR shown over time. Individual traces are shown in grey, while the mean ± standard deviation is shown in bold black. A schematic of how the SPPR is calculated is shown (second phase peak as a percentage of the first phase peak). Absolute SPPR significantly increased over time from day 2 to day 14 (****p < 0.001, univariate n-way ANOVA, Tukey's post hoc test). (D) Example of linear function fitting to SPPR over time including the output slope are shown for a relatively good (blue) and bad (green) fit. The linear function is shown as a dashed black line. The slope of the linear fit is significantly positively correlated with R1 as fitted with EIS and significantly negatively correlated with C as fitted with EIS (Pearson's correlation coefficient).

Contact impedances, voltage at the end of the first phase of a cathodic-leading pulse normalized to the input current, were calculated with (Figure 5B) and without (Figure S6C, Supporting Information) constructs on the electrode. Contact impedances with construct on the electrode were normalized to day 2. Contact impedances significantly increased over time when construct was on the electrode (F = 23.91, p < 0.001, df = 5, univariate n-way ANOVA corrected for experiment number (random factor)), with inflection point between days 7 and 9 as revealed by Tukey's post hoc test, and an overall significant change between days 2 and 14 (p < 0.001). Without construct on the electrode, no significant changes were seen over time (F = 1.32, p = 0.26, df = 5).

We hypothesized that with an increase in R1 and a decrease in C over time for electrodes with construct on, ratio of the second peak as a percentage of the first peak would change over time, as the contribution of capacitive discharge to the second phase peak would decrease. Therefore, we calculated the second phase peak ratio (SPPR) as shown in Figure 5C, which describes second peak voltage as a percentage of the first peak voltage. The SPPR increased significantly over time when the construct was on the electrode (F = 38.05, p < 0.001, df = 5, univariate n-way ANOVA corrected for experiment number (random factor)), with the inflection point between days 4 and 7 (Tukey's post hoc test). Here, a significant difference was found between days 2 and 14 (p < 0.001). Without construct on the electrode (Figure S6D, Supporting Information), no significant changes were seen with time (F = 1.07, p = 0.38, df = 5).

To compare change in SPPR over time with a single measure to EIS-fitted circuit elements R1 and C, we calculated slope of SPPR over time with a linear function. Two examples of such slopes can be seen in Figure 5D, with both SPPR that shows a linear increase over time and one that does not. We only fitted data to a linear function when >3 datapoints and datapoints after day 7 (inflection point of R1) were available, leading to n = 33 slopes. These slopes were correlated with the final available timepoint used for the linear fit (Figure 5D) of EIS-fitted circuit elements. A significant positive correlation was found between the SPPR and R1 (Pearson's r = 0.37 (95% CI: 0.03 to 0.63), p < 0.05, n = 33), while a significant negative correlation was found between SPPR and C (Pearson's r = −0.51 (95% CI: −0.20 to −0.72), p < 0.005, n = 33).

2.4 Voltage Waveform-Fitted Circuit Elements Correlate Significantly with EIS-Fitted Circuit Elements

To expand information extraction from voltage waveforms, we reverse fitted (voltage waveform (VW) fitted) (Figure S7A, Supporting Information) our chosen circuit (Figure 4A) to the voltage waveforms (Figure 6A). R1 and C show similar trends, yet capacitance is higher for the VW-fitted example. Additionally, C is capped at its upper limit (102 nF) for days 2 through 7.

Figure 6 Open in figure viewer PowerPoint Reverse fitting of voltage waveforms (VW) to electrical circuit. A) EIS-fitted and VW-fitted voltage waveforms (top row), absolute impedance magnitude (middle row), and phase angle (bottom row) of electrical circuit in Figure 4A on example data shown in Figures 4B and 5A. B) Example of direct comparison between EIS-fitted (blue) and VW-fitted (pink) circuit element sizes for the example shown in (A). C) VW-fitted circuit elements over time of all timepoints and electrodes with construct on the electrode (n = 193). Individual data is shown in grey. The arithmetic mean ± standard deviation is shown in bold black, except for C where the geometric mean and standard deviation are shown. R1 shows a significant increase from day 2 to day 14 (****p < 0.001, univariate n-way ANOVA, Tukey's post hoc test), while C shows a significant decrease (****p < 0.001, univariate n-way ANOVA, Tukey's post hoc test). D) Correlation between VW-fitted and EIS-fitted R1 and C, excluding uncapped values (including n = 89 for R1 and n = 79 for C, compared to n = 193 for both) as part of the bimodal distribution seen in Figure S7C (Supporting Information). A significant but modest correlation was found for both circuit elements (Pearson's correlation coefficient). For R1, the correlation is stronger when VW-fitted elements >2 kΩ are excluded, suggesting outliers are more likely with R1 > 2 kΩ in VW-fitting.

CPE-P and CPE-T were fixed based on day 2 values in addition to fixed R2 values, and so only data with EIS fitting available on day 2 was included (Figure 6C). R1 increased significantly over time (F = 17.83, p < 0.001, df = 5, univariate n-way ANOVA corrected for experiment number (random factor)), whilst C decreased significantly over time (F = 23.98, p < 0.001, df = 5). The inflection point, as shown by Tukey's post hoc test, was in between days 7 and 9 for R1 and days 4 and 7 for C. Significant differences from days 2 to 14 were present for both R1 (p < 0.001) and C (p < 0.001).

A percentage of VW-fit output shows capped values where R1 caps its lowest bound of 50 Ω and C caps its upper bound of 10−7 F. Capping mainly happens when the voltage waveform peak is at its lowest, since 60.6% of the output values is capped in at least one element over all timepoints, whilst from days 9 to 14, only 21% of the VW-fittings is capped (Figure S7B, Supporting Information). This leads to a bimodal distribution for output values of VW-fitted R1 and C with the element bounds used (Figure S7C, Supporting Information). Widening the element bounds, however, leads to capping at both bounds for both circuit elements (Figure S7D, Supporting Information). To compare EIS-fitting with VW-fitting we correlated EIS-fitted values of circuit elements R1 and C to VW-fitted values of the same electrode and timepoint. A significant positive correlation was found between EIS-fitted R1 and VW-fitted R1 (Pearson's r = 0.48 (95% CI: 0.30 to 0.63), p < 0.001, n = 89). However, outliers were seen when VW-fitted R1 reached >2 kΩ. Excluding VW-fitted R1 > 2kΩ showed a stronger correlation between VW-fitted and EIS-fitted R1 (Pearson's r = 0.72 (95% CI: 0.59 to 0.82), p < 0.001, n = 73). A significant positive correlation was also found between EIS-fitted C and VW-fitted C (Pearson's r = 0.42 (95% CI: 0.22 to 0.59), p < 0.001, n = 79).

2.5 Changes in Contact Impedances and SPPR in CI Patients Postoperatively are in Line with Changes Found in the Tissue Engineered Model

Based on our findings of SPPR changes in our tissue-engineered model, we wanted to test this marker in recently implanted CI patients. We used the CI company's software function to measure mutliple timepoints along voltage waveform response, to measure an altered version of SPPR (6 µs into each phase) as well as compare this to the contact impedances (Figure 7A) over 2 and 3 timepoints, respectively, in four patients. We assumed no or little fibrosis was present before cochlear implantation, since these were new CI patients, and at least some fibrosis formation to occur within 5 months postoperatively. It should be noted that, given the inability of currently-used diagnostics to monitor fibrosis progression, we have no independent information about fibrosis status at the collected timepoints.

Figure 7 Open in figure viewer PowerPoint Contact “impedances” and altered SPPR of four recently implanted CI patients. A) Schematic of input current (dashed line), measured timepoints (circles), and analyzed timepoints (filled circles). An example response from a patient is shown in turquoise. B) Contact “impedances”, as measured at the end of the first phase (25 µs), intraoperatively, 3 months postoperatively, and 5 months postoperatively for four patients. Individual data are shown in grey, the mean of each patient is shown in bold. A significant increase in contact “impedances” is seen from intra-op to post-op on group level (****p < 0.001, univariate n-way ANOVA, Tukey's post hoc test), but not from 3 to 5 months postoperatively (ns = not significant, univariate n-way ANOVA, Tukey's post hoc test). C) Altered SPPR (6 µs into each phase) is shown for two post-operative timepoints. Individual data are shown in grey, the mean of each patient is shown in bold. A significant increase in SPPR is seen from 3 to 5 months post-op on group level (**p < 0.01, univariate n-way ANOVA, Tukey's post hoc test).

Contact impedances showed a significant increase from intraoperative to postoperative timepoints (F = 139.1, p < 0.001, df = 2, n = 264 datapoints across four patients, univariate n-way ANOVA, p < 0.001 Tukey's post hoc test), when correcting for patient as a random factor and electrode number as a fixed factor (Figure 7B). No significant effect of patient (F = 2.36, p = 0.07, df = 3–4 patients) nor electrode number (F = 0.35, p = 0.997, df = 21,22 electrodes) were found. Postoperative contact impedances at 3 and 5 months were not significantly different from each other (p = 0.995, Tukey's post hoc test). SPPR was only available for two postoperative timepoints (Figure 7C). The altered SPPR showed a significant increase over time (F = 6.83, p < 0.01, df = 1, n = 176 datapoints across four patients, univariate n-way ANOVA), when correcting for patient and electrode number. No significant effects for patient (F = 1.99, p = 0.118, df = 3–4 patients) or electrode number (F = 1.41, p = 0.120, df = 21–22 electrodes) were found. The largest increase in mean altered SPPR is seen for patient 4 (P4, Figure 7C), while a modest increase is seen for P1 and P3 and a modest decrease for P2. While an increase in altered SPPR is found for all 22 electrodes for P4, a variation of increase and decrease is found for the other three patients when looking at electrode level changes.

Mean comfortable loudness levels (MCLs), expressed in the log-scale unit of current levels (CLs), were available for the 3 and 5 months postoperative timepoints for a subset of electrodes. MCLs are complex and likely to vary due to patient-related factors such as, but not limited to, auditory nerve survival and central factors. Therefore, we looked at the relationship between change in SPPR and contact impedances with change in MCLs, as to correct for between-patient differences in absolute MCL. When looking at changes in contact impedances and SPPR (Figure S8, Supporting Information) from 3 to 5 months per electrode, no significant correlations were found with changes in MCLs (change in contact impedances: Pearson's r = 0.23 (95% CI: −0.15 to 0.54), p = 0.23, n = 30, change in altered SPPR: Pearson's r = −0.32 (95% CI: −0.61 to 0.05), p = 0.09, n = 30). The negative correlation between change in altered SPPR with change in MCL per electrode was mainly driven by three datapoints of P4 that showed a large positive change in altered SPPR and a decrease in MCL. An overview of all the patient data can be found in Table S1 (Supporting Information).

3 Discussion

In this study, we tissue engineered a 3D model of cochlear fibrosis that behaves similarly to data we collected from a postoperative population of patients with cochlear implants. This model was designed to improve our understanding of the fibrotic response that occurs during cochlear implantation and ideally will be used in conjunction with large-scale human data collection and animal models to improve outcomes for patients experiencing the effects of fibrosis from the placement of a cochlear implant. We used a tissue-engineered, cell-seeded gel to simulate the electrical environment of a fibrosing cochlear implant on a clinical cochlear electrode array. We analyzed these data both biologically and electrically to confirm the usefulness of this system as a model for cochlear fibrosis. Finding that we could recreate some of the conditions that we observed in a patient population, we developed a new marker based on our electrical data that was also found to increase in our postoperative patient-derived data at group level. Cochlear implants are known to cause fibrosis formation in the cochlea that can lead to residual hearing loss for cochlear implant patients.[4, 9, 23-26] An electrical marker of fibrosis progression could create an early window for treatment intervention to reduce the residual hearing loss for patients.

Our tissue-engineered model of cochlear fibrosis has the advantage of including the electrode array that is used in the clinical setting for CI patients, as well as incorporating the 3D aspect of fibrous tissue encapsulation that is known to behave differently from 2D tissue.[40] Using this model, we were able to examine some cellular behaviors for which the field has only been able to previously speculate.[39, 68] We found attachment of the fibroblasts to the electrode surfaces, where numerous cells were situated on an electrode. This is in line with what is thought to happen in vivo[39, 68] and is important to detect any changes in electrode–electrolyte interface that might be caused by this attachment. As these cells are seeded into a tissue-engineered gel, the cells can also remodel and change this construct. In line with previous studies,[42, 69] the cells cause significant contraction, ultimately resulting in contraction of the construct away from some electrodes that were originally embedded in construct at the beginning of the experiment. These electrodes show full recovery from an electrical perspective (data included in Figure S4, Supporting Information). This result is very promising for patients in that we also show electrodes can return to their original state, indicating that the development of treatments for the reduction or reversal of fibrosis has the potential to restore degradation in stimulation efficiency in clinical scenarios.

To design a new electrical marker of fibrosis progression, we first needed to understand the complex impedance changes over time in our model. We proposed a new electrical circuit to represent the changes in our model of cochlear fibrosis and showed significant changes in complex impedance over time. The elements representing the bulk of the construct (R1 and C) showed significant changes over time, while the CPE representing the electrode–electrolyte interface did not. This suggests that biological changes affecting electrical impedance can be explained by changes in the bulk of the construct, such as ECM formation and reorganization, rather than changes in the electrode-electrolyte interface. In line with the complex impedance results, full voltage waveform recordings showed significant changes in the clinically measurable contact impedances over time, as well as in a newly proposed electrical marker, SPPR. The SPPR is directly measurable in patients and could allow for earlier detection of fibrosis formation and progression allowing for earlier treatment intervention. This marker, in addition to the further information we show to be retrievable from fitting full voltage waveforms, could also be utilized as a measurement tool in drug developing and testing studies.

EIS revealed changes in both impedance magnitude and phase angle over time and when modeled with our proposed circuit, revealed significant changes for those circuit elements representing the bulk of the construct. The changes in R1, however, are of a larger magnitude than the changes in C, suggesting absolute impedance magnitude changes are due to an increase in the resistance of the construct. No significant changes from day 2 to endpoint for the CPE representing the EE interface, even with cells visibly attached on the electrode surface, were found. A recent study by Fuentes-Vélez et al. used the same electrical circuit as presented in the current study as a marker of liver fibrosis in mice.[70] Liver fibrosis follows a wound-healing response similar to what is thought to happen intracochlearly post-implantation and includes an increase in ECM deposition.[71] The authors saw an increase in bulk resistance, similar to that presented in the current study, when stimulating liver fibrosis and correlated this increase in resistance to the formation of ECM. This supports the use of our presented circuit model and suggests changes seen in this study could be due to cellularly mediated ECM alterations. Furthermore, our findings are in line with previous patient studies modeling fibrosis through voltage waveforms with the Tykocinski et al. circuit, where a change in access resistance is found over time.[36, 56, 60, 62] We also tested this circuit model on our EIS data and found a large error for fitting across multiple frequencies, indicating that this model oversimplifies complex impedance. This has been previously described by Mesnildrey et al., who found a large residual error when using the simple RC circuit for the EE interface and proposed the use of a CPE instead for both EIS and voltage waveform fitting.[72] Combining these observations, the circuit model presented in this study provides a more accurate picture of the electrical changes present during cochlear fibrosis formation. This model could potentially be used to study other types of input pulses, such as triphasic of pseudomonophasic pulses, for which SPPR could also be sensitive.

To allow for easily measurable data in CI patients using current clinical software and to provide a comparison with currently collected data from patients, we measured voltage waveform responses at each electrode. The clinically measurable contact impedances showed an increase over time, which is in line with studies examining contact impedances and fibrosis formation.[4, 19, 35, 56] As mentioned above, we present a new electrical marker that would require only one extra timepoint to be measured and so provides an opportunity to easily expand data collection in patients. Interestingly, the inflection point for our SPPR marker was at an earlier timepoint than for the measured contact impedances. We were able to test our SPPR in patients at 2 timepoints postoperatively, which revealed a significant change on a group level in SPPR from 3 to 5 months postoperative while no significant change was found for contact impedances between these timepoints. However, interpretation of the statistical tests on this data should be done with caution as the sample size is small. Additionally, no control measure for fibrosis is present. To further test SPPR as a marker for fibrosis formation in patients, as well as test its correlation with residual hearing, a large patient study with intra-operative and post-operative timepoints of SPPR and auditory thresholds should be done. This would allow for a clearer indication of no fibrosis present (intra-operatively) to fibrosis present (post-operatively) than at only post-operative timepoints as presented here.

We fitted our circuit model of fibrosis encapsulation to voltage waveforms measured and found significant correlations with the output of complex impedance fitting, showing an opportunity for additional information extraction from voltage waveforms in CI patients, which is possible with research software.[38, 61, 73] However, proposed fitting of full voltage waveforms needs to be optimized further and needs to include a circuit model fitting to CI patients rather than an in vitro model. Our VW fitting had a large percentage of values capped at the limits and needed fixed values for the EE interface.

The model presented in this paper could be used as a drug-testing platform, where changes in complex impedance, SPPR, and contraction could be used to test ways to inhibit or even reverse fibrosis. Patient biopsies could be used to build patient-specific models of cochlear fibrosis. In our study, we did not observe any effects on our cells from applied electrical stimulation, despite contrary observations in some studies.[74] Analysis of different stimulation regimens could yield different results, which would be easily achievable using our model. However, this criterion for examination was outside the range of our goals for this study.

As our system is meant to represent the immune response to CI implantation, we have notably not included immune cells within this model. Fibrosis in vivo is complex and involves other cell types beyond just fibroblasts.[15, 16, 75, 76] Immune cells play a major role in the development and progression of fibrotic tissue. Previous animal studies indicate that after CI implantation, fibrin is first adsorbed onto electrodes. This matrix is infiltrated with macrophages and leukocytes, whose presence is reduced upon the infiltration of fibroblasts, which has been shown to occur around 7 days post-implantation.[12, 14, 15] Our model focuses on this latter stage of development after fibroblast infiltration. Of note, the presence of these immune cells plays a major role in the development of fibrotic response in vivo and would, therefore, likely have an effect on our model if present. We would speculate that the addition of immune cells would produce a more accurate timeline for fibrosis with possible changes in tissue morphology and structure. However, given that our model mostly focuses on the electrical response from the CI electrodes, these changes are unlikely to result in a different outcome from that which we observed.

One limitation of this model is that clinical fibrous encapsulation is attached to the walls of the cochlea, making longitudinal contraction to the levels seen in our study less likely.[29, 77] This effect is also influenced by the positioning of the electrode array positioning in the tapered 3D structure of the cochlea, which could influence baseline complex impedance.[65] On the next iterations of this model, we envision incorporating different characteristics of the cochlear environment using a tapered conical model of the cochlea. In this cochlea-shaped bioreactor, we can incorporate testing of current spread towards the auditory nerve with fibrosis development to help understand CI performance changes due to bulk tissue formation. Nevertheless, this study shows significant and large changes from baseline in complex impedance and allowed us to present electrical changes in real-time on a clinical electrode, which we were able to translate to a directly measurable electrical marker for fibrosis in CI patients.

4 Conclusion

In conclusion, this study presents a tissue-engineered model of fibrosis progression on a clinical cochlear implant array. It demonstrates complex impedance as a marker of fibrosis progression and applies the changes found in complex impedance to directly measurable cochlear implant patient data. A new marker, the SPPR, provides a potential mechanism for gauging cochlear implant fibrosis formation progress in patients, with no additional software or equipment needed. These findings can be used to track fibrosis formation in patients in real-time, allowing for earlier treatment intervention, and can be used in drug-testing platforms to test and develop new treatments inhibiting fibrosis and therefore combating residual hearing loss. The findings in this study hold the potential for generalization to other neural implants with fibrosis formation, opening up new areas of exploration and treatment, for improving implant science.

Skin-interfaced microfluidic systems with spatially engineered 3D fluidics for sweat capture and analysis

A 3D printed epifluidic device called a "sweatainer" used for sweat capture and analysis

Skin-interfaced microfluidic systems with spatially engineered 3D fluidics for sweat capture and analysis

by Chung-Han Wu, Howin Jian Hing Ma, Paul Baessler, Roxanne Kate Balanay and Tyler Ray

Abstract: Skin-interfaced wearable systems with integrated microfluidic structures and sensing capabilities offer powerful platforms for monitoring the signals arising from natural physiological processes. This paper introduces a set of strategies, processing approaches, and microfluidic designs that harness recent advances in additive manufacturing [three-dimensional (3D) printing] to establish a unique class of epidermal microfluidic (“epifluidic”) devices. A 3D printed epifluidic platform, called a “sweatainer,” demonstrates the potential of a true 3D design space for microfluidics through the fabrication of fluidic components with previously inaccessible complex architectures. These concepts support integration of colorimetric assays to facilitate in situ biomarker analysis operating in a mode analogous to traditional epifluidic systems. The sweatainer system enables a new mode of sweat collection, termed multidraw, which facilitates the collection of multiple, independent sweat samples for either on-body or external analysis. Field studies of the sweatainer system demonstrate the practical potential of these concepts.

We kindly thank the researchers at University of Hawai'i at Mānoa for this collaboration, and for sharing the results obtained with their CADworks3D system.

A 3D printed epifluidic device called a "sweatainer" used for sweat capture and analysis

Introduction

Eccrine sweat is an attractive class of biofluid suitable for the noninvasive monitoring of body chemistry. Sweat contains a rich composition of biomarkers relevant to physiological health status including electrolytes (1), metabolites (24), hormones (56), proteins (7), and exogenous agents (8). Studies demonstrate the intermittent or continuous assessment of these, and other sweat biomarkers offer time dynamic insight into the metabolic processes of the body relevant to applications ranging from athletic performance (911) to medical diagnostics (21214).

Recent advances in soft microfluidics, sensing technologies, and electronics establish the foundations for a unique class of skin-like epidermal microfluidic (“epifluidic”) systems. Adapting concepts from traditional lab-on-chip technologies, these wearable microfluidic platforms comprise sophisticated networks of channels, valves, and reservoirs embedded in elastomeric substrates (1520). The thin, flexible device construct facilitates a conformal, fluid-tight skin interface by virtue of skin-compatible adhesives to collect sweat directly from sweat glands. The integration of colorimetric, fluorometric, and electrochemical measurement techniques enable such platforms to measure sweat constituents in situ across a wide array of applications and environments (21).

Traditional approaches for sweat collection use absorbent pads (22) or microbore tubes (23) pressed against the epidermis by virtue of bands or straps to capture sweat as it emerges from the skin. Requiring trained personnel, special handling, and costly laboratory equipment, such methods are incompatible with real-time sweat analysis and prone to sample contamination or loss (24). Epifluidic devices eliminate external sample contamination by virtue of the intrinsic encapsulation of the microfluidic network and conformal skin interface. Such systems are vulnerable to surface contamination from exogenous agents present on the epidermis, such as cosmetics or natural oils, without careful preparation of the skin surface before device attachment. Furthermore, the dependence on an adhesive interface for skin attachment limits these devices to single-use applications. Upon removal, the risk of contamination, potential sample loss, and active sweat response of previously covered glands pose substantial challenges to reapplication and continued sweat collection.—Traditional approaches for sweat collection use absorbent pads (22) or microbore tubes (23) pressed against the epidermis by virtue of bands or straps to capture sweat as it emerges from the skin. Requiring trained personnel, special handling, and costly laboratory equipment, such methods are incompatible with real-time sweat analysis and prone to sample contamination or loss (24). Epifluidic devices eliminate external sample contamination by virtue of the intrinsic encapsulation of the microfluidic network and conformal skin interface. Such systems are vulnerable to surface contamination from exogenous agents present on the epidermis, such as cosmetics or natural oils, without careful preparation of the skin surface before device attachment. Furthermore, the dependence on an adhesive interface for skin attachment limits these devices to single-use applications. Upon removal, the risk of contamination, potential sample loss, and active sweat response of previously covered glands pose substantial challenges to reapplication and continued sweat collection.

The typical epifluidic fabrication pathway uses soft lithography techniques (25) to produce devices with microfluidic components and complex geometries. A common, well-established process for fabricating lab-on-chip microfluidic devices (26), soft lithography, requires high-precision molds to form discrete, patterned layers of an elastomeric material [e.g., poly(dimethylsiloxane) (PDMS)] that when bonded together yield a sealed device. Traditionally, producing molds with sufficient feature resolution (>20 μm) requires expensive, time-consuming processing methods [micromachining (27) and micromilling (28)] and access to specialized environments (cleanroom). Such requirements result in elongated device design cycles, inequitable access to equipment necessary for innovation, and additional challenges for commercial deployment due to incompatibilities with large-scale manufacturing.

Additive manufacturing (AM), or three-dimensional (3D) printing, represents an attractive alternative to conventional planar (2D) fabrication methods. AM offers powerful capabilities for producing structurally complex objects with true 3D architectures through a rapidly expanding library of printing methods. In general, these methods create solid objects in a sequential, layer-by-layer manner directly from a digital computer-aided design (CAD) file. In the context of microfluidics, the use of 3D printing is well established (29) for the rapid, cost-effective fabrication of high-resolution templates for soft lithography. In particular, vat photopolymerization techniques [e.g., resin-based printing, stereolithography, digital light processing (DLP), and continuous liquid interface polymerization] (30) enable rapid production of microscale features (>100 μm) over large areas (>600 mm2) with high precision (31). Innovations in printer hardware, software processing, and materials chemistry further extend these 3D printing capabilities to enable the direct production of enclosed microfluidic channels for lab-on-chip applications. Although manufacturers advertise printers with high resolution (xy resolution: >50 μm and z-resolution: >5 μm), in practice, the obtainable channel dimensions and device complexity are typically limited to millifluidic features (>250 μm) (29). Printer specifications represent only one key constraint to printing devices with micron-scale internal fluidic features (<100 μm). Successful fabrication requires optimization of other critical factors including printing technology (e.g., vat photopolymerization versus extrusion), feature design and spatial location, and printer-dependent parameters. AM process optimization, particularly for vat photopolymerization, demands careful attention to the chemistry of printed materials (3032). Resin formulations must simultaneously satisfy application specific requirements, such as biocompatibility or optical clarity, while preserving printability. Recent reports (3233) leverage specialized DLP-based printers and customized resins to fabricate devices containing microfluidic components with <50-μm dimensions.

Apparatus Used

Clear Microfluidic Resin

Curezone

The CADworks3D Pr110 3D Printer with a 385nm wavelength projector

PR110
3D Printer

Legacy

In general, wearable system designs must address the inherent mismatch between the mechanical properties of skin and rigid, planar device components. The most advanced platforms fabricated by conventional (non-AM) methods exploit sophisticated strategies, combining complex device geometries and soft (low modulus) materials to establish a seamless, nonirritating epidermal interface. Recent advances in soft materials chemistry support 3D printing approaches to fabricating wearable devices for applications spanning biophysical (34), biochemical (3536), and environmental (37) monitoring. However, such capabilities remain limited for the 3D fabrication of epifluidic devices as a result of the high Young’s moduli of the primary material chemistries (i.e., methacrylate-based resins) (38) suitable for printing high-resolution microfluidics. Current efforts to fabricate skin-interfaced 3D printed microfluidics use alternative printing methods [e.g., fused deposition modeling (34) and direct ink writing (39)] that support fabrication with low modulus materials at the expense of printer resolution (>200 μm). In the context of epifluidics, the ideal fabrication scheme would use resin-based printing to fabricate devices with feature sizes comparable to conventional methods with biologically compliant form factors. Such an approach would transform the fluidic design space with truly 3D device architectures while enabling a rapid, iterative design process, facilitating individual-specific device customization, and reducing the cost for low-volume production.

This paper introduces a set of strategies, processing approaches, and microfluidic designs that support such fabrication capabilities using a commercial DLP 3D printer in a simple manner of operation. A modular 3D printed epifluidic platform, termed a “sweatainer,” demonstrates several unique aspects of an AM approach to fabricating epifluidic systems. This platform, to our knowledge, represents the first 3D printed epifluidic platform with true microfluidic dimensions. Specifically, the results highlight the potential of a true 3D design space for microfluidics through the fabrication of fluidic components (channels and valves) with previously inaccessible complex architectures. Printer optimization strategies and systematic experiments enable realization of micron-scale feature sizes (<100 μm) and enhancement of optical transparency of 3D printed channels. In combination, these concepts support integration of colorimetric assays to facilitate in situ biomarker analysis operating in a mode analogous to traditional epifluidic systems. Drawing inspiration from the vacutainer blood collection tube, the sweatainer system introduces a novel mode of sweat collection, termed “multidraw.” This method overcomes the inherent limitations of single-use devices by enabling the collection of multiple, independent pristine sweat samples during a single collection period. Field studies of the sweatainer system demonstrate the practical potential of these concepts.

Results

Sweatainer system design

Figure 1A shows a schematic illustration of the two primary modules of the sweatainer system: (i) the sweatainer device and (ii) an epidermal port interface. The sweatainer consists of a 3D printed microfluidic network of enclosed channels and unsealed reservoirs, a reservoir capping layer of PDMS (thickness: 200 μm), and a gasket formed from ultrathin biomedical adhesive (3M 1524; thickness: 60 μm). The bonded 3D printed photocurable resin structure and PDMS capping layer, as presented in Materials and Methods, define a closed microfluidic structure. Introduction of either dye or colorimetric assay before bonding enables sweat visualization or chloride concentration analysis, respectively. The cross-sectional width and thickness of the filleted serpentine channels presented here are 1200 and 1000 μm, respectively. The width and height of the rectangular-shaped internal microfluidic channels are 600 and 400 μm, respectively. The filamentary design of the rigid 3D printed structure (Young’s modulus: ~975 MPa) follows from the well-established principles of stretchable electronics (40) to impart sufficient stretchability to form a mechanically robust conformal interface. The gasket establishes a temporary, fluid-tight seal with the epidermal port interface permitting facile sweatainer application and removal via reversible adhesion to the PDMS surface.

Figure 1. Schematic illustrations and optical images of the 3D printed epidermal microfluidic devices for the collection and analysis of sweat. (A) An exploded render highlights key components of the sweatainer system and epidermal interface (port). PDMS, poly(dimethylsiloxane). (B) A photograph of the sweatainer mounted on the ventral forearm of an individual before the onset of sweat collection. (C) The construct of the sweatainer eliminates uncontrolled fluid transport under mechanical loading (e.g., finger pressure and device removal). (D) Illustration of the sweatainer highlighting key device aspects including the inlet, capillary burst valves (CBVs; blue and red dashed area), collection reservoir, and ventilation holes to eliminate backpressure. (E) Renders of three-dimensional (3D) CBV designs enabled by 3D printing with diverging angles of 90° (top) and 135° (bottom). (F) 3D printing enables fabrication of device geometries in a true 3D space as shown by the computer-aided design (CAD) render (top) and photograph of actual device (bottom). Location of sweat appears in blue. (G) Photographic sequence highlighting the complete filling of a sweat collection reservoir.
Figure 1. Schematic illustrations and optical images of the 3D printed epidermal microfluidic devices for the collection and analysis of sweat. (A) An exploded render highlights key components of the sweatainer system and epidermal interface (port). PDMS, poly(dimethylsiloxane). (B) A photograph of the sweatainer mounted on the ventral forearm of an individual before the onset of sweat collection. (C) The construct of the sweatainer eliminates uncontrolled fluid transport under mechanical loading (e.g., finger pressure and device removal). (D) Illustration of the sweatainer highlighting key device aspects including the inlet, capillary burst valves (CBVs; blue and red dashed area), collection reservoir, and ventilation holes to eliminate backpressure. (E) Renders of three-dimensional (3D) CBV designs enabled by 3D printing with diverging angles of 90° (top) and 135° (bottom). (F) 3D printing enables fabrication of device geometries in a true 3D space as shown by the computer-aided design (CAD) render (top) and photograph of actual device (bottom). Location of sweat appears in blue. (G) Photographic sequence highlighting the complete filling of a sweat collection reservoir.

The epidermal port interface comprises a thin film of pigmented PDMS (white, thickness: 400 μm) and a medical-grade adhesive layer (3M 1524) with laser-patterned openings. The adhesive layer facilitates a biocompatible, fluid-tight interface with the epidermis in which the patterned opening defines the sweat collection region (~180 mm2). An aligned access point on the backside of the sweatainer allows sweat to enter the system directly from the skin with flow driven by the natural pressures created by the sweat glands. The sweatainer design can support collection of 50.8 μl of sweat (10.8 μl per reservoir and 18.4 μl of channel network). A fully assembled representative system appears in Fig. 1B, where it is shown worn on the ventral forearm. Figure 1C demonstrates the insensitivity of the sweatainer to mechanical deformation through the absence of uncontrolled fluid flow during physical handling (finger pressure). The schematic illustration in Fig. 1D shows the microfluidic network within the 3D printed sweatainer. Sweat enters the device by the central inlet and flows through a microfluidic channel leading to a series of capillary burst valves (CBVs) and corresponding reservoirs. The CBV at the ingress of each reservoir permits fluid flow only after exceeding a set pressure, thereby enabling time-sequential sweat collection (20). Integrated ventilation holes (width: 100 μm and height: 200 μm) on the reservoir eliminate the backpressure that would evolve from trapped air and impede flow. The high-barrier properties of the photocurable resin support a low sweat evaporation rate with minimal mass loss over a 24-hour period (fig. S1 and table S1).

A key feature of this system is the use of AM to enable fully 3D, monolithic microfluidic designs comprising sophisticated nonplanar internal channel structures, spatially graded geometries, and 3D CBVs. Representative examples of 3D CBVs and the spatially graded, nonplanar geometries enabled by this fabrication method appear in Fig. 1 (E and F, respectively). By comparison, soft lithography fabrication methods restrict the design space of traditional lab-on-chip and epifluidic devices to planar (2D) channel configurations. Although lamination of multiple channel layers can yield elaborate 3D microfluidic networks, each component layer is inherently a planar geometry. As detailed in the sections that follow, the 3D fabrication expands the design space for CBVs with finer control over resultant burst pressure in comparison to planar CBVs. In a similar manner, spatially graded geometries improve sweat collection efficiency by permitting a continuous transition between the microfluidic channel and reservoir (Fig. 1F). This engineered interface, in combination with ventilation holes, ensures a uniform fluid front during reservoir filling (Fig. 1G, blue dye for visualization), thereby eliminating trapped air bubbles that result from a rapid expansion.

Design and DLP printing considerations for optimized fabrication of 3D printed epifluidic devices

Successful fabrication of a fully enclosed microfluidic channel with feature sizes at the xy plane resolution limit of current DLP printers (~30 to 50 μm) depends on several related factors including: design aspects (e.g., channel vertical position), print process parameters [e.g., layer height, layer cure time (LCT), and print speed], and printer hardware (e.g., projector light power and wavelength). Optimization of user-adjustable factors results in a robust print process suitable for producing microfluidic devices with sufficient optical clarity, dimensional fidelity, and mechanical performance for use in epifluidic applications.

As expected, epifluidic device performance is dependent on the dimensional accuracy of a fabrication process. If not quantified, then unintended deviation from designed dimensions can adversely affect component performance (i.e., CBV burst pressure) or measurement accuracy (i.e., sweat volume, sweat rate). Fabrication of test structures (Fig. 2A) comprising a sequence of square channels (width and height range: 100 to 900 μm, 100-μm increments; length: 5 mm) embedded in a square base (width and height: 1 mm) facilitate determination of the minimum printable channel dimensions and sidewall thickness (minimum of 50 μm). The asymmetric vertical position of the channels establishes a uniform capping layer (100 μm) across all dimensions tested. Because the DLP printer fabricates the structure in an inverse manner (Fig. 2A, base prints first), the channel position minimizes photopolymerizing resin trapped in the channel during the printing process.

Figure 2. Optimized design strategy for fabricating 3D printed epifluidic devices with prescribed channel geometries. (A) Photograph of 3D printed test channels [100 to 900 μm, square; 2-s layer cure time (LCT)]. (B) Plot of variation of printed channel height from designed dimensions as a function of LCT. (C) Plot of variation of printed channel width from designed dimensions as a function of LCT. (D) Plot highlighting the printable region of the digital light processing (DLP) printer used in this work for various channel dimensions relevant to epifluidic devices.
Figure 2. Optimized design strategy for fabricating 3D printed epifluidic devices with prescribed channel geometries. (A) Photograph of 3D printed test channels [100 to 900 μm, square; 2-s layer cure time (LCT)]. (B) Plot of variation of printed channel height from designed dimensions as a function of LCT. (C) Plot of variation of printed channel width from designed dimensions as a function of LCT. (D) Plot highlighting the printable region of the digital light processing (DLP) printer used in this work for various channel dimensions relevant to epifluidic devices.

Experimental studies reveal the similarly strong influence of LCT on print success and device quality. The LCT defines the energy dose used to cross-link the photopolymer given in time (seconds). The projector wavelength is hardware defined (385 nm for this work), and varying the power is not typically user accessible. Systematic studies of four LCT settings—selected starting from the minimum (0.54 s) to maximum values (2.0 s; 0.6-s interval) beyond which channels could not be fabricated successfully—establish a relationship among print performance (i.e., channel printed successfully), dimensional accuracy, and optical clarity. Measurement results from optical microscope images, shown in Fig. 2B for channel height and Fig. 2C for channel width, highlight the relationship between LCT and printed channel dimensions. The proportional relationship between increasing LCT and light propagation into the z dimension (thickness) of the masked regions (i.e., channels) results in smaller than designed channel heights. By comparison, the dimensional accuracy for a given channel width depends primarily on the size of the DLP printer pixels (xy plane resolution) rather than LCT. The observed positive channel width variation with decreasing LCT indicates incomplete photopolymerization. Subsequent postprocessing removal of uncured resin yields channels with dimensions greater than designed. In combination, these results establish the printable region for an epifluidic design as a function of LCT. As shown in Fig. 2 (B and C), successful fabrication of a 100-μm square channel requires a short LCT (i.e., 0.54 and 0.8 s), whereas a longer LCT results in photopolymerization of the otherwise unreacted resin. Conversely, for large dimensions (>700 μm square channels), a short LCT produces channels too fragile to survive printing and postprocessing due to incomplete photopolymerization. These results establish an LCT of 0.8 s as the optimal setting for balancing printability with dimensional accuracy for the printed epifluidic devices described in subsequent sections.

Additional systematic experiments establish the DLP-printable design space for epifluidic-relevant dimensions (100 to 600 μm). Evaluation of print success as a function of channel dimensions (width and height) for an enclosed microfluidic channel (length: 30 mm) identifies the printable region (Fig. 2D). An encapsulated microfluidic channel capable of supporting unrestricted fluid flow, in contrast to a sealed or partially restricted channel, defines a successful print. Intuitively, print failure rate increases as the enclosed channel dimensions approach the printer xy plane resolution limit (~32-μm x 32-µm square pixel). Results show a channel dimension of 100 μm (either width or height) corresponds to the lower limit for a successful printed device.

Print process optimization to support colorimetric analysis in 3D printed epifluidic systems

The optical transparency of a 3D printed microfluidic device depends on several factors including material selection, printer hardware (e.g., build plate and vat surface material), postprocessing, and surface roughness. In contrast to the typical surface roughness feature size necessary for optical transparency (<10 nm) (41), DLP printers produce parts with microscale surface roughness, resulting in a semi-translucent appearance (32).

As mentioned previously, the digital micromirror device (DMD) pixel size governs the xy plane resolution of a DLP printer. Minute gaps between individual DMD elements locally reduce reflected light intensity, yielding a surface roughness with features corresponding to DMD pixel size and layer height. While specialized printing methods (grayscale) (42) or printer hardware (oscillating lenses) (43) offer sophisticated strategies to reduce aliasing and improve surface roughness, the fundamental approach to eliminating this defect mode is enhancing the uniformity of projected light to ensure complete photopolymerization. Figure 3A illustrates that increasing the exposure dose by lengthening the LCT eliminates the observed grid pattern defects (from DMD element gaps) and improves optical transparency. Ultraviolet-visible (UV-Vis) spectroscopy experiments examine the transmission properties of 3D printed microcuvettes in comparison to a commercial plastic cuvette (Fig. 3B). While results show substantial modulation of light transmission with increasing LCT, ranging from ~20 (LCT: 0.54 s) to ~60% (LCT: 2 s), the reference commercial plastic cuvette offers higher light transmission (~80%). Intuitively, there is no observed wavelength dependence for light transmission within the Vis spectrum (400 to 1100 nm) beyond the anticipated strong absorbance within the UV region (<400 nm, necessary for photopolymerization) for the 3D printed samples. As a consequence of the presence of both the UV absorber and photoinitiator in the resin, green parts (i.e., before curing) have a light yellow hue. As presented in Materials and Methods, completion of the postprocessing sequence eliminates part coloring (fig. S2).

Figure 3. Optimized design strategy for enabling colorimetric analysis in 3D-printed epifluidic systems. (A) Optical micrographs of the surface of parts printed with different LCT settings. (B) Plot of light transmission of commercial and resin-printed cuvettes measured with ultraviolet-visible (UV-Vis) spectrometer. (C) Photographs of epifluidic reservoirs fabricated using static (0.54 s, 2-s LCT) and adaptive (AP1 and AP2) printing processes illustrating differences in optical transparency. (D) Calibration curves as a function of LCT highlighting improvement in optical transparency (and thus colorimetric performance) with increasing LCT. A.U., arbitrary units.
Figure 3. Optimized design strategy for enabling colorimetric analysis in 3D-printed epifluidic systems. (A) Optical micrographs of the surface of parts printed with different LCT settings. (B) Plot of light transmission of commercial and resin-printed cuvettes measured with ultraviolet-visible (UV-Vis) spectrometer. (C) Photographs of epifluidic reservoirs fabricated using static (0.54 s, 2-s LCT) and adaptive (AP1 and AP2) printing processes illustrating differences in optical transparency. (D) Calibration curves as a function of LCT highlighting improvement in optical transparency (and thus colorimetric performance) with increasing LCT. A.U., arbitrary units.

In addition to LCT, layer height affects both overall device quality (e.g., vertical resolution, optical clarity, and channel roughness) and print time, which corresponds to device yield. Conventional approaches to vat photopolymerization use constant values for a given print run (i.e., fixed layer height and LCT). At present, only one manufacturer [Formlabs (44)] supports an adaptive layer height process to increase print speed by adjusting layer height as a function of model detail (i.e., small layers for fine features and thick layers for coarse features). Adaptive printing is an attractive process for obtaining expanded design flexibility for 3D printed epifluidic systems. Although not supported by default, a combination of custom software and manual geometric code programming in this work enables definition of both layer height and LCT as a function of model dimensions. The representative example shown in fig. S3 illustrates the capabilities of this adaptive printing process to fabricate a cube (all dimensions: 2 mm) using four layer heights (5, 10, 30, and 50 μm) in an arbitrary order. By comparison to a constant LCT and layer height setting printing process, this approach enables successful, time-efficient fabrication of epifluidic systems with complex geometries and superior device quality.

Colorimetric assays facilitate passive, battery-free in situ quantitative measurement of sweat biomarkers. A chemical reagent reacts with a target species to generate an optical signal proportional to analyte concentration (45). Accurate colorimetric analysis requires channels with uniform height (i.e., path length), a high degree of optical transparency, and integrated color reference markers to support reliable image processing under variable ambient lighting conditions (46). The layer-by-layer control over LCT and layer height parameters enabled by an adaptive printing process is critical for fabricating microfluidic devices with the requisite surface finish and optical transparency to support colorimetric analysis. Figure 3C illustrates the influence of an adaptive LCT print process on the optical transparency of microfluidic channels. The optical clarity for two representative sweatainer reservoirs manufactured using a layer-constant LCT (0.54 and 2 s) increases with longer LCT (Fig 3C). While beneficial for reducing nonuniform illumination, the increased UV dose results in undesirable curing of resin in enclosed features (channels and CBVs). By comparison, an adaptive printing process (AP 1) using an LCT of 0.54 s for the reservoir surface and an LCT of 2 s for subsequent layers facilitates fabrication of a sweatainer with a translucent imaging plane, a transparent device, and preservation of internal channel features. An inverse adaptive printing process (AP 2; base LCT: 2 s and subsequent layer LCT: 0.54 s) results in an optically transparent imaging plane and a translucent device.

Systematic benchtop experiments evaluate the suitability of devices fabricated by adaptive printing for colorimetric analysis. The colorimetric assay silver chloranilate produces a dark violet color response proportional to chloride concentration. Imaging the device with a smartphone camera enables color extraction and subsequent quantification of color response. The inclusion of a color balance chart facilitates color calibration for each image. As in previous reports (4748), converting images from native red, green, blue (RGB) color space to CIELAB color space—which expresses color as lightness (L), amount of green to red (a*), and amount of yellow to blue (b*)—ensures device-independent color sampling. Conversion of the a* and b* components to chroma (C*) by the relation

Expression in Plain Text: C* = ((a*)^2 + (b*)^2)^(1/2)

yields a calibration curve with chloride concentration by a power-law relation (fig. S4). Figure 3D shows calibration charts created from 3D printed sweatainers with different LCT parameters and reference colorimetric assay solutions. This plot reveals that the improvement in optical clarity with increasing LCT provides a corresponding enhancement in the range of detectable color measurements. As these findings indicate, an adaptive printing process is essential for fabricating epifluidic devices with an optical transparency sufficient to support colorimetric analysis.

3D CBV designs for sequential sweat analysis

CBVs are a key component for the sequential analysis of sweat biomarkers in many epifluidic platforms. The time dynamic variations in sweat rate arising from physical (e.g., sweat gland density), physiological (e.g., exertion and emotion), and external factors (e.g., temperature and pH) result in corresponding changes in analyte concentration. As previously described, CBVs prevent flow for fluid pressure conditions below a designed threshold [bursting pressure (BP)]; when the fluid pressure exceeds the BP, the CBV immediately bursts. Operating without use of actuation or moving components, CBV BP is governed by valve geometry.

The Young-Laplace equation describes the BP for a CBV (rectangular channel) as (49)

Expression in Plain Text: BP = -2σ (cos(θ*_{I})/b) + (cos(θ_{A})/h)

where σ is the fluid surface tension, θA is the critical advancing contact angle for the channel (material dependent, θA = 120° for PDMS) (50), θ*I is the minimum of either θA + β or 180°, β is the channel diverging angle, and b and h are the diverging channel width and height, respectively. As the second term of Eq. 2 is constant for a planar (2D) CBV, channel width and diverging angle govern the BP for a given CBV. In practice, epifluidic device designs use geometric restrictions (i.e., modifications to channel width) to control valve BP.

The 3D printing concept for epifluidic devices presented here expands CBV capabilities by enabling a full 3D CBV design. As a consequence, Eq. 2 can be written as

Expression in Plain Text: BP = -2σ (cos(θ*_{I})/b) + (cos(θ_{J})/h)

for a 3D CBV, where θ*J is the minimum of either θA + γ or 180° and γ is the channel diverging angle (z axis). It follows that for a microfluidic channel with fixed dimensions, the CBV BP becomes a function of the channel diverging angles (β and γ). Computational predictions of four representative CBV designs, presented as a schematic in Fig. 4A with parameters specified in Table 1, illustrate this relationship. Figure 4B shows the theoretical BP versus channel size (square channel) for the four CBV designs with σ = 0.072 N/m (surface tension of water) and θA = 120° (PDMS) for the 2D CBV (type 1) and θA = 60° for the 3D CBVs (resin, types 2 to 4). It is shown that BP is inversely proportional to channel size. As expected, the analytical model reveals that for a given channel size BP increases for 3D CBV designs (resin) in comparison to a 2D CBV (PDMS). Within the subset of 3D CBV designs, the channel diverging angles (β and γ) dictate the valve BP (BPType4 > BPType3 > BPType2).

Figure 4. 3D CBV designs for sequential sweat analysis. (A) Schematic renders highlighting four design types of CBVs used in this work. Areas highlighted in blue indicate differences between CBV designs. (B) Plot of the theoretical maximum bursting pressures (BPs) calculated from the Young-Laplace equation as a function of channel size for a square geometry. (C) Sequence of photographs illustrating the performance of different CBV designs (labels 1 to 8). Use of backside illumination for the overview photograph facilitates visualization of valves and channels. (D) A sequence of photographs shows a 3D printed H channel with one central inlet and four CBVs (color indicates CBV design and fixed channel geometry) filling sequentially, highlighting the fluid control enabled by a true 3D CBV. (E) Plot of the theoretical BP as a function of diverging angle β for a channel with a fixed geometry (width: 600 μm and height: 400 μm). The CBV designs are identical to (B). (F) A sequence of photographs highlighting performance of the 3D printed sweatainer design used in human participant testing.
Figure 4. 3D CBV designs for sequential sweat analysis. (A) Schematic renders highlighting four design types of CBVs used in this work. Areas highlighted in blue indicate differences between CBV designs. (B) Plot of the theoretical maximum bursting pressures (BPs) calculated from the Young-Laplace equation as a function of channel size for a square geometry. (C) Sequence of photographs illustrating the performance of different CBV designs (labels 1 to 8). Use of backside illumination for the overview photograph facilitates visualization of valves and channels. (D) A sequence of photographs shows a 3D printed H channel with one central inlet and four CBVs (color indicates CBV design and fixed channel geometry) filling sequentially, highlighting the fluid control enabled by a true 3D CBV. (E) Plot of the theoretical BP as a function of diverging angle β for a channel with a fixed geometry (width: 600 μm and height: 400 μm). The CBV designs are identical to (B). (F) A sequence of photographs highlighting performance of the 3D printed sweatainer design used in human participant testing.
Table 1. Diverging angle parameters for CBV type. CBV, capillary burst valve; 2D, two-dimensional; N/A, not applicable
Table 1. Diverging angle parameters for CBV type. CBV, capillary burst valve; 2D, two-dimensional; N/A, not applicable

Benchtop experiments yield measurements of CBV BPs by means of a positive pressure displacement pump apparatus that perfuses water (dyed blue for visualization) into the microfluidic network at defined pressures. Figure 4C shows a representative test of the sequential filling performance of a network of 2D and 3D CBV-gated reservoirs, labeled chronologically in order of increasing BP. Table 2 and fig. S5 detail the CBV design parameters, theoretical CBV BPs, and effective theoretical BPs, which consider the theoretical CBV BP and fluidic resistance of the microfluidic channel network. Imperfections resulting from the 3D printing process result in experimental BP values below theoretical limits.

Table 2. Design parameters for CBVs. BP, bursting pressure.
Table 2. Design parameters for CBVs. BP, bursting pressure.

The 3D design space provides attractive capabilities for fine-scale control over CBV performance to enable compact fluid control features within epifluidic devices. Varying the diverging angle design parameters (β and γ) for a 3D CBV results in substantial differences in BP for valves with similar dimensions and form factors. Systematic experiments performed in similar manner as described previously verify the correlation between diverging angle and BP for the 3D CBV architectures illustrated in Fig. 4A with identical channel dimensions. Figure 4D shows a representative test of 3D CBV performance via a 3D printed microfluidic device with channels arrayed in an H configuration (channel dimensions: 600-μm width and 400-μm height). As Fig. 4E highlights, BP increases with β for a resin-based 3D CBV in contrast to a PDMS-based 2D CBV baseline reference. Material properties limit the valve design space on account of the BP dependence on contact angle. For hydrophobic materials such as PDMS (i.e., contact angle >120°), β values greater than 60° reduce to 180°, resulting in BP value dependent only on channel width (b). By comparison, the expanded design range, in which β governs valve BP, results from the smaller contact angle of hydrophilic materials (i.e., resin). The experimental results support these trends predicted by the analytical model with the variation between measured and predicted values attributed to geometric imperfections inherent to the fabrication process (i.e., slight rounding of corners) (51). A similar trend occurs for valve designs in which γ varies with respect to a fixed β.

Additional studies demonstrate 3D CBV performance in a device architecture relevant to practical use. Robust operation requires CBV designs with BP within the physiologically relevant range of sweat secretory pressure (0.5 to 2 kPa) (51). Tests of the sweatainer design shown in Fig. 4F proceed in the same manner whereby water enters the device through a central inlet. Reservoirs fill sequentially in the order indicated as the CBVs at entrance of reservoirs no. 2 and no. 3 prevent fluid flow until reservoir no. 1 fills completely. Variation of CBV diverging angle defines the BP for CBV no. 1 (blue, 0.66 kPa) and CBV no. 2 (red, 0.86 kPa). These results validate the design of the sweatainer for use in on-body testing.

Field studies of the sweatainer

A pilot study comprising healthy adult volunteers (N = 8) exercising on a stationary bike explores the on-body performance of the sweatainer system. Following the protocol detailed in Materials and Methods, the sweatainer intimately couples to the ventral forearm of a participant by means of the epidermal port (PDMS/skin-safe adhesive). Participants cycled at moderate intensity for a period of 50 min under controlled environmental conditions [22°C, 59% relative humidity (RH)]. Upon entering the device from the skin, sweat proceeds to sequentially fill the microfluidic reservoirs. The addition of chloride-free dye at the sweatainer inlet aids in visualization. Periodic imaging with a smartphone camera during exercise facilitates monitoring fill performance. The sweatainer typically fills within 40 min from the initiation of exercise; after filling, the device is exchanged mid-exercise with a new sweatainer in a seamless manner. Figure 5A highlights this event sequence with sweatainers distinguished by distinct visualization dyes (device no. 1: blue and device no. 2: orange). The simplicity of the exchange facilitates a rapid replacement time (<30 s), thereby minimizing potential interruption to the sweat collection process. In all tests, the adhesive gasket maintains a robust, water-tight interface between the sweatainer and epidermal port evidenced by the absence of observed leaks. The 3D printed sweatainer resists mechanical deformation during detachment, thereby eliminating unconstrained fluid flow. In combination, these features support the multidraw collection of pristine sweat samples and reduce the risk of sample contamination during collection process.

Figure 5. Sweatainer field studies. (A) Sequence of photographs highlighting operation of the sweatainer system. A sweatainer (device no. 1) collects sweat during an active exercise period, which, upon filling, is rapidly exchanged (
Figure 5. Sweatainer field studies. (A) Sequence of photographs highlighting operation of the sweatainer system. A sweatainer (device no. 1) collects sweat during an active exercise period, which, upon filling, is rapidly exchanged (<30 s) for a second sweatainer (device no. 2) facilitating multidraw sweat collection. (B) Photograph of sweatainer position during exercise trials (blue box). (C) A magnified view of the same sweatainer devices shown in (B) before the onset of sweating. The sweatainer shown on left is for collection (control) with the device on the right for colorimetric analysis. (D and E) respectively show the collection and colorimetric sweatainers at the conclusion of the exercise period. (F) Plot showing the concentration of sweat chloride from the collection (chloridometer) and colorimetric sweatainers for three independent exercise trials for a single participant (stationary cycling, 50 min, constant power). (G) Plot of showing sweat chloride concentration from two different colorimetric sweatainers worn sequentially (i.e., replaced during trial) during a predefined exercise period (stationary cycling, 50 min, constant power). The total sweat volume lost by a given individual during this exercise period corresponds to the total number of filled sweatainer chambers. Scale bars, 5 mm.

A second set of exercise tests focuses on the in situ measurement of the concentration of sweat chloride by colorimetric analysis. A sweatainer configured with an integrated colorimetric assay (replacing visualization dye) enables measurement of chloride concentration in collected sweat during exercise. Figure 5B shows the sweatainer mounted on the ventral forearm of a volunteer participant. Simultaneous deployment of a collection sweatainer (orange dye alone) in close spatial proximity of a colorimetric sweatainer (Fig. 5C) facilitates comparison of colorimetric chloride measurements with gold standard clinical methods for chloride analysis (chloridometer). The collection sweatainer operates in a similar mode to microbore tubes (i.e., Macroduct) traditionally used in clinical settings for collecting sweat for chemical analysis. Representative photographs of the colorimetric and collection sweatainers at the conclusion of an exercise period appear in Fig. 5 (D and E, respectively). As shown in Fig. 5F, for a representative participant, chloride concentrations measured using colorimetric sweatainers correlate well for given individual (Fig. 5F reports three independent exercise trials), within experimental uncertainties, to values determined using coulometry and are within the normal physiological range (48). The chronological sampling capabilities of the sweatainer enable monitoring of the time dynamic variation of sweat biomarkers. Figure 5G demonstrates the multidraw sweatainer operation for three participants (field study data for remaining five participants shown in table S2) during a fixed exercise period (stationary cycling, 50 min, constant power). In both sets of trials, the observed increase in sweat chloride concentration during exercise is consistent with results from previous studies (52). Here, an inverse relationship exists between the sweat duct efficiency in reabsorbing chloride and rate of sweat loss, resulting in a corresponding increase in sweat chloride. Factors such as fitness level, training status, and heat acclimation affect this relationship for a given individual. These findings demonstrate the sweatainer system as a viable platform for colorimetric-based biomarker analysis with reported values comparable to established clinical methods.

Discussion

The sweatainer system reported here introduces an AM approach to fabricating epidermal microfluidic devices to collect and analyze sweat. AM enables true 3D design of microfluidic channels and fluid control components, such as valves, with architectures typically inaccessible to planar (2D) fabrication methods. The detailed characterization and optimization of print parameters provides a pathway to fabricate microfluidic devices with enhanced optical transparency and feature sizes below 100 μm. Field studies using stationary cycling provide a practical demonstration of key concepts of the sweatainer platform including multidraw sample collection scheme and in situ colorimetric analysis of chloride concentration. Future studies will seek to investigate the generalizability of the sweatainer platform beyond clinical applications to sweat collection during more vigorous and dynamic physical activities through the development of optimized designs capable of supporting a broader spectrum of physical exertion.

The sweatainer platform represents a pivotal advancement in the collection and analysis of sweat samples. Inspired by the versatility of the vacutainer for blood collection, the sweatainer allows for the acquisition of multiple, independent aliquots of sweat from a single collection period. This collection mode enables an array of possibilities for sweat-based studies, including remote and at-home diagnostics, biobanking for future clinical research, and the integration of sweat analysis into existing clinical chemistry methods. Moreover, the utilization of AM for fabricating the sweatainer allows for customized geometries and streamlined integration into clinical workflows, further enhancing the potential of the platform for facilitating the quantification of ultralow concentration sweat biomarkers. The realization of multidraw sweat collection, enabled by the sophisticated sample collection strategies and customizable designs reported here, represents a major step forward in the field of sweat-based analysis.

Materials and Methods

Fabrication of 3D printed epifluidic devices

Each epifluidic device design (3D) was created using CAD software (AutoCAD 2019, Autodesk, CA, USA). Subsequent export to a stereolithography file format (.stl) yielded a file suitable for direct use by the DLP resin printer (Prime 110, 385 nm, MiiCraft, Taiwan and Creative CADworks, ON, Canada). The included printer control software (Utility, version 6.3.0.t3) provided direct control over print parameters for each file including layer height (5 to 50 μm), dose, and lamp power. High-fidelity printing was achieved by application of a removable Kapton polyimide tape over the surface of the polished aluminum build plate. The applied tape was free of bubbles/wrinkles to ensure a smooth build surface free of defects.

Devices were printed using transparent resin (MiiCraft BV-007A, Creative CADworks, ON, Canada) and a 10-μm layer height (six devices per build plate, ~20 min total print time). Gentle removal of printed parts from the build plate, soaking in 1% detergent solution (Alconox-1232-1, Alconox, NY, USA) under sonication (CPX2800, Fisher Scientific, PA, USA) for 10 min, drying of device using clean dry air (CDA), postprint UV cure for 10 min (CureZone, MiiCraft, Taiwan), and postcure bake at 70°C for 30 min (Model 40E Lab Oven, Quincy Lab Inc., IL, USA) yielded a 3D printed epifluidic device suitable for direct use or integration with PDMS.

A three-step process (fig. S6) facilitated printing fully enclosed 3D printed devices. Printing epifluidic devices with open reservoirs (step 1) and postprint removal of uncured liquid resin by CDA (step 2) enabled enclosure of the devices with a thin capping layer (30 μm) by means of a second print process (step 3). The printed device remains fixed to the build plate during the printing process to ensure feature alignment. Following the previously described postprocessing steps yielded a fully enclosed epifluidic device.

Apparatus Used

Clear Microfluidic Resin

Curezone

The CADworks3D Pr110 3D Printer with a 385nm wavelength projector

PR110
3D Printer

Legacy

Fabrication of ultrathin capping layer for microfluidic channels in hybrid devices

Pouring liquid PDMS (10:1 base:curing agent; Sylgard 184, Dow Inc., MI, USA) with white pigment (3% w/w; Ignite White, Smooth-On Inc., PA, USA) onto a sacrificial mylar film (2 mil thickness), spin coating for 30 s [400 revolutions per minute (rpm) for reservoir capping layer and 200 rpm for epidermal interface layer], and curing in an oven (70°C, 2 hours) formed films with thicknesses of 200 and 400 μm, respectively. A CO2 laser cutter (30 W Epilog Mini 24, Epilog Laser, CO, USA) patterned the PDMS films into the final geometries used in the epifluidic devices. A medical-grade adhesive (1524, 3M Inc., MN, USA) is patterned in the same manner and bonded to the PDMS interfacial layer, established the epidermal interface for the device.

Hybrid 3D printed epifluidic devices use bonded PDMS capping layers to enclose 3D printed microfluidic reservoirs. Modification of a previously reported method (53) facilitated a strong bond between PDMS and the printed device. Specifically, rinsing with isopropyl alcohol (2-propanol, A416, Fisher Scientific, MA, USA), soaking in deionized (DI) water (Direct-Q 3 UV Water Purification System, MilliporeSigma, MO, USA) for 30 min, corona treating with air plasma (BD-20, Electro-Technic, IL, USA) for 30 s followed by immediate immersion in a 12% v/v solution of (3-aminopropyl)triethoxysilane (APTES; 440140, MilliporeSigma, MO, USA) for at least 20 min, rinsing in DI water, and drying with CDA prepared the oven-baked 3D printed device for bonding to PDMS. Pipetting colorimetric reagents or flow visualization dye (Soft Gel Paste, AmeriColor Corp., CA, USA) into predetermined regions occurred before sealing of the 3D printed device. After a 30-s corona treatment, laminating the PDMS capping layer to the APTES-modified printed surface sealed the epifluidic device. Heat treating the assembled device on a hotplate (70°C) under applied weight (3 kg) for 30 min formed a permanent bond. Removal of the sacrificial mylar layer, release from excess PDMS via laser cutting, and opening the central sweat ingress points using a 1.5-mm diameter circular punch (reusable biopsy punch, World Precision Instruments) yielded a final hybrid epifluidic device.

Measurement of evaporation rate for 3D printed microfluidic networks

3D printed microfluidic devices (N = 7) with theoretical capacity (~101 ml) facilitated the measurement of the rate of evaporation. Sealing of the inlet and outlet of a device with parafilm after filling with DI water (dyed blue for visualization) formed the complete device for testing. Measurement of the initial sealed device mass (inclusive of water, film, and printed microfluidic system) using a microbalance (Sartorius Quintix 224-1S, Germany) enabled recording of mass loss at 2 and 24 hours. Devices were maintained at room temperature in a controlled laboratory environment reflective of anticipated use environment (22°C, 55% RH). An optical camera (Canon 90D, Canon EF 100 mm f/2.8L USM lens) facilitated observation of visual changes to fluid levels at each measurement interval.

Characterization of 3D CBVs

A digital microscope (VHX-7100, Keyence Corp., Japan) produced micrographs of the devices. An optical camera (Canon 90D, Canon EF 100 mm f/2.8L USM lens) provided video capture capabilities (30 frames per second) for device analysis. Measurement of the CBV burst pressure consisted of a “fill test” in which water (dyed blue for visualization) entered a device until flow stopped the CBV. A modular, calibrated pressure displacement flow system (Flow EZ, Fluigent, France) controlled the fluid pressure and permitted near-instantaneous stepwise increase in pressure (0.1-mbar interval, 10-s dwell time). Video observation identified the pressure threshold for fluid to burst the valve.

Measurements of transmission properties of 3D printed devices

A UV-Vis spectrophotometer (UV-1900i, Shimadzu, Japan) enabled quantification of the optical transmission properties of the printed devices (300 to 1000 nm, 0.5-nm interval). A commercial plastic cuvette (path length: 10 mm; Shimadzu) served as a reference (control). Four sets of 3D printed cuvettes (N = 3 per set) using a different LCT setting (0.54, 0.8, 1.4, and 2 s) enable quantification of the relationship between LCT and optical transmission (dimensions: height, 50 mm; width, 8 mm; path length, 1 mm; and volume, ~21 μl).

Colorimetric assay for chloride

The chloride colorimetric assay solution resulted from thoroughly vortexing 50 mg of silver chloranilate (MP Biomedicals, CA, USA) in 200 μl of a solution of 2% (w/v) polyhydroxyethylmethacrylate (529265, MilliporeSigma, MO, USA) in methanol (A412, Fisher Scientific, MA, USA) to yield a homogenous suspension. Spotting 2 μl of this solution via laboratory pipette onto the 3D printed device near the central sweat ingress point, followed by drying in an oven for 30 min before encapsulation, prepared the epifluidic device for colorimetric chloride measurements.

Standard color development and color reference marker preparation

Mixing sodium chloride (S271, Fisher Scientific, MA, USA) in DI water produced standard test solutions (0, 10, 20, 30, 50, 75, 90, 110, 130, and 150 mM). Clinical-grade chloridometer measurements (ChloroChek, ELITech Group Inc.) yielded validated test solution concentrations. Digital imaging and analysis of sample reservoirs (N = 7) containing one standard solution reacted with the silver chloranilate assay under uniform illumination formed a set of reference images. The sample reservoirs were of the same depth as the epifluidic channels to ensure accurate color representation.

Digital image analysis for the evaluation of sweat chloride concentrations

A smartphone camera (iPhone 11 Pro Max, Apple, CA, USA) captured images during on-body field tests. A color calibration card (ColorChecker Classic, X-Rite, MI, USA) in the frame of each image facilitated accurate color extraction under various illumination conditions. An open-source photography software package (Darktable 3.0.0, Darktable.org) served as the platform for calibrating images using the color reference card. Analysis of calibrated images using MATLAB (R2019b, MathWorks Inc., MA, USA) enabled cropping multiple regions of interest (N = 3) from images and extraction of CIELAB color values (La*, and b*) for chroma analysis. Mapping of chroma values from colorimetric samples of known reference chloride solutions yielded colorimetric calibration charts with a power-law relationship. This calibration chart supported quantification of the sweat chloride concentration in on-body field testing.

Human participant sweat analysis

The purpose of this pilot study was to evaluate the performance of the 3D printed epifluidic device and use in collecting and analyzing sweat. Testing involved healthy young adults (N = 8, six male and two female) as volunteers during normal physical activity (stationary cycling) with no additional human participant risk. The study was International Review Board (IRB) approved through the University of Hawaiʻi (IRB no. 2018-1440). Informed consent was obtained after explanation of the nature and possible consequences of study participation.

Cleaning of the forearm of each individual with an alcohol wipe prepared the skin for robust adhesion to the device. The exercise regime comprised stationary cycling under approximately constant working load for 50 min in a controlled laboratory environment (22°C, 55% RH).

Evaluation of the colorimetric sweatainer performance required individual participants (N = 3) to wear two separate sweatainers, one colorimetric and one collection (as a control), located in close proximity on the same arm. Before device removal, a photograph of the colorimetric sweatainer was recorded at the conclusion of the collection period for image processing and chloride analysis. Extraction of sweat from the individual reservoirs of the collection sweatainer at the conclusion of the exercise period facilitated chloride measurements using a ChloroChek Chloridometer.

Evaluation of sequential generation of aliquots of sweat required periodic monitoring the filling of the epifluidic device (N = 8). Once all reservoirs filled, as determined by visual observation, the initial device (attached at the start of the exercise period) was removed from the interfacial layer and replaced with a new device while simultaneously continuing to exercise.

Unidirectional imaging using deep learning–designed materials

Unidirectional imaging using deep learning–designed materials

JINGXI LI, TIANYI GAN, YIFAN ZHAO, BIJIE BAI, , CHE-YUNG SHEN, SONGYU SUN, MONA JARRAHI AND AYDOGAN OZCAN

A unidirectional imager would only permit image formation along one direction, from an input field-of-view (FOV) A to an output FOV B, and in the reverse path, B → A, the image formation would be blocked. We report the first demonstration of unidirectional imagers, presenting polarization-insensitive and broadband unidirectional imaging based on successive diffractive layers that are linear and isotropic. After their deep learning–based training, the resulting diffractive layers are fabricated to form a unidirectional imager. Although trained using monochromatic illumination, the diffractive unidirectional imager maintains its functionality over a large spectral band and works under broadband illumination. We experimentally validated this unidirectional imager using terahertz radiation, well matching our numerical results. We also created a wavelength-selective unidirectional imager, where two unidirectional imaging operations, in reverse directions, are multiplexed through different illumination wavelengths. Diffractive unidirectional imaging using structured materials will have numerous applications in, e.g., security, defense, telecommunications, and privacy protection.

We kindly thank the researchers at University of California for this collaboration, and for sharing the results obtained with their system.

Introduction

Optical imaging applications have permeated every corner of modern industry and daily life. A myriad of optical imaging methods have flourished along with the progress of physics and information technologies, resulting in imaging systems such as super-resolution microscopes (1, 2), space telescopes (3–5), and ultrafast cameras (6, 7) that cover various spatial and temporal scales at different bands of the electromagnetic spectrum. With the recent rise of machine learning technologies, researchers have also started using deep learning algorithms to design optical imaging devices based on massive image data and graphics processing units, achieving optical imaging designs that, in some cases, surpass what can be obtained through physical intuition and engineering experience (8–14).

Standard optical imaging systems composed of linear and time-invariant components are reciprocal, and the image formation process is maintained after swapping the positions of the input and output fields of view (FOVs). If one could introduce a unidirectional imager, then the imaging black box would project an image of an input object FOV (A) onto an output FOV (B) through the forward path (A → B), whereas the backward path (B → A) would inhibit the image formation process by scattering the optical fields outside the output FOV (see Fig. 1A).

Fig. 1. Schematic of a diffractive unidirectional imager. (A) Concept of unidirectional imaging, where the imaging operation can be performed as the light passes along a certain specified direction (A → B), while the image formation is blocked along the opposite direction (B → A). (B and C) Illustration of our diffractive unidirectional imager, which performs imaging of the input FOV with high fidelity in its forward (B) direction and blocks the image formation in its backward (C) direction. This diffractive unidirectional imager is a reciprocal device that is linear and time invariant and provides asymmetric optical mode processing in the forward and backward directions. Its design is insensitive to light polarization and leaves the input polarization state unchanged at its output. Furthermore, it maintains its unidirectional imaging functionality over a large spectral band and works under broadband illumination.

To design a unidirectional imager, one general approach would be to break electromagnetic reciprocity: One can use, e.g., magneto-optic effect (the Faraday effect) (15–17), temporal modulation of the electromagnetic medium (18, 19), or other nonlinear optical effects (20–27). However, realizing such nonreciprocal systems for unidirectional imaging over a sample FOV with many pixels poses challenges due to high fabrication costs, bulky and complicated setups/materials, and/or high-power illumination light sources. Alternative approaches have also been used to achieve unidirectional optical transmission from one point to another without using optical isolators. One of the most common practices is using a quarter-wave plate and a polarization beam splitter; this approach for point-to-point transmission is polarization sensitive and results in an output with only circular polarization. Other approaches include using asymmetric isotropic dielectric gratings (28–31) and double-layered metamaterials (32) to create different spatial mode transmission properties along the two directions. However, these methods are designed for relatively simple input modes and face challenges in off-axis directions, thus making them difficult to form imaging systems even with relatively low numerical apertures.

Despite all the advances in materials science and engineering and optical system design, there is no unidirectional imaging system reported to date, where the forward imaging process (A → B) is permitted and the reverse imaging path (B → A) is all optically blocked.

Here, we report the first demonstration of unidirectional imagers and design polarization-insensitive and broadband unidirectional imaging systems based on isotropic structured linear materials (see Fig. 1, B and C). Without using any lenses commonly used in imaging, here, we optimize a set of successive dielectric diffractive layers consisting of hundreds of thousands of diffractive features with learnable thickness (phase) values that collectively modulate the incoming optical fields from an input FOV. After being trained using deep learning (33–46), the resulting diffractive layers are physically fabricated to form a unidirectional imager, which performs polarization-insensitive imaging of the input FOV with high structural fidelity and power efficiency in the forward direction (A → B), while blocking the image transmission in the backward direction, not only penalizing the diffraction efficiency from B → A but also losing the structural similarity or resemblance to the input images. Despite being trained using only Modified National Institute of Standards and Technology (MNIST) handwritten digits, these diffractive unidirectional imagers are able to generalize to more complicated input images from other datasets, demonstrating their external generalization capability and serving as a general-purpose unidirectional imager from A → B. Although these diffractive unidirectional imagers were trained using monochromatic illumination at a wavelength of λ, they maintain unidirectional imaging functionality under broadband illumination, over a large spectral band that uniformly covers, e.g., 0.85 × λ to 1.15 × λ.

We experimentally confirmed the success of this unidirectional imaging concept using terahertz waves and a three-dimensional (3D) printed diffractive imager and revealed a very good agreement with our numerical results by providing clear and intense images of the input objects in the forward direction and blocking the image formation process in the backward direction. Using the same deep learning–based training strategy, we also designed a wavelength-selective unidirectional imager that performs unidirectional imaging along one direction (A → B) at a predetermined wavelength and along the opposite direction (B → A) at another predetermined wavelength. With this wavelength-multiplexed unidirectional imaging design, the operation direction of the diffractive unidirectional imager can be switched (back and forth) based on the illumination wavelength, improving the versatility and flexibility of the imaging system.

The optical designs of these diffractive unidirectional imagers have a compact size, axially spanning ~80 to 100λ. Such a thin footprint would allow these unidirectional imagers to be integrated into existing optical systems that operate at various scales and wavelengths. While we considered here spatially coherent illumination, the same design framework and diffractive feature optimization method can also be applied to spatially incoherent scenes. Polarization-insensitive and broadband unidirectional imaging using linear and isotropic structured materials will find various applications in security, defense, privacy protection, and telecommunications among others.

Results

Diffractive unidirectional imager using reciprocal structured materials

Figure 1A depicts the general concept of unidirectional imaging. To create a unidirectional imager using reciprocal structured materials that are linear and isotropic, we optimized the structure of phase-only diffractive layers (i.e., L1, L2, …, L5), as illustrated in Fig. 1 (B and C). In our design, all the diffractive layers share the same number of diffractive phase features (200 by 200), where each dielectric feature has a lateral size of ~λ/2 and a trainable/learnable thickness providing a phase modulation range of 0 to 2π. The diffractive layers are connected to each other and the input/output FOVs through free space (air), resulting in a compact system with a total length of 80λ (see Fig. 2A). The thickness profiles of these diffractive layers were iteratively updated in a data-driven fashion using 55,000 distinct images of the MNIST handwritten digits (see Materials and Methods). A custom loss function is used to simultaneously achieve the following three objectives: (i) minimize the structural differences between the forward output images (A → B) and the ground truth images based on the normalized mean square error (MSE), (ii) maximize the output diffraction efficiency (overall transmission) in the forward path, A → B, and (iii) minimize the output diffraction efficiency in the backward path, B → A. More information about the architecture of the diffractive unidirectional imager, loss functions, and other training-related implementation details can be found in Materials and Methods. After the completion of the training, the phase modulation coefficients of the resulting diffractive layers are shown in Fig. 2C. Upon closer inspection, it can be found that the phase patterns of these diffractive layers have stronger modulation in their central regions, while the edge regions appear relatively smooth, with weaker phase modulation. This behavior can be attributed to the size difference between the smaller input/output FOVs and the relatively larger diffractive layers, which causes the edge regions of the diffractive layers to receive weaker waves from the input, as a result of which their optimization remains suboptimal.

Fig. 2. Design schematic and blind testing results of the diffractive unidirectional imager. (A and B) Layout of the diffractive unidirectional imager when it operates in the forward (A) and backward (B) directions. (C) The resulting diffractive layers of a diffractive unidirectional imager. (D) Exemplary blind testing input images taken from Modified National Institute of Standards and Technology (MNIST) handwritten digits that were never seen by the diffractive imager model during its training, along with their corresponding diffractive output images in the forward and backward directions. a.u., arbitrary units. (E) Same as (D), except that the testing images are taken from the Extended MNIST (EMNIST) and Fashion-MNIST datasets, demonstrating external generalization to more complicated image datasets.

This diffractive unidirectional imager design was numerically tested using the MNIST test dataset, which consists of 10,000 handwritten digit images that were never seen by the diffractive model during the training process. We report some of these blind testing results in Fig. 2D for both the forward and backward directions, clearly illustrating the internal generalization of the resulting diffractive imager to previously unseen input images from the same dataset. We also quantified the performance of this diffractive unidirectional imager for both the forward and backward directions based on the following metrics: (i) the normalized MSE and (ii) the Pearson correlation coefficients (PCCs) between the input and output images (denoted as “output MSE” and “output PCC”) and (iii) the output diffraction efficiencies; these metrics were calculated using the same set of MNIST test images, never seen before. As shown in Fig. 3 (A and B), the forward (A → B) and backward (B → A) paths of the diffractive unidirectional imager shown in Fig. 2C provide output MSE values of (5.68 ± 1.56) × 10−5 and (0.919 ± 0.048) × 10−3, respectively, and their output PCC values are calculated as 0.9740 ± 0.0065 and 0.3839 ± 0.0685, respectively. A similar asymmetric behavior between the forward and backward imaging directions is also observed for the output diffraction efficiency metric as shown in Fig. 3C: The output diffraction efficiency of A → B is found as 93.50 ± 1.56%, whereas it is reduced to 1.57 ± 0.44% for B → A, which constitutes an average image power suppression ratio of ~60-fold in the reverse direction compared to the forward imaging direction. Equally important as this poor diffraction efficiency for B → A is the fact that the weak optical field in the reverse direction does not have spatial resemblance to the input objects as revealed by a poor average PCC value of ~0.38 for B → A. These results demonstrate and quantify the internal generalization success of our diffractive unidirectional imager: The input images can be successfully imaged with high structural fidelity and power efficiency along the forward direction of the diffractive imager, while the backward imaging operation B → A is inhibited by substantially reducing the output diffraction efficiency and distorting the structural resemblance between the input and output images.

Fig. 3. Performance analysis of the diffractive unidirectional imager shown in Fig. 2. (Aand B) Normalized mean square error (MSE) (A) and Pearson correlation coefficient (PCC) (B) values calculated between the input images and their corresponding diffractive outputs in the forward and backward directions. (C) The output diffraction efficiencies of the diffractive unidirectional imager calculated in the forward and backward directions. In (A) to (C), the metrics are benchmarked across the entire MNIST test dataset and reported here with their mean values and SDs added as error bars. (D) Left: The power of the different spatial modes propagating in the diffractive volume during the forward and backward operations, shown as percentages of the total input power. Right: Schematic of the different spatial modes propagating in the diffractive volume. FOV, field of view.

To better understand the working principles of this diffractive unidirectional imager, next, we consider the 3D space formed by all the diffractive layers and the input/output planes as a diffractive volume and categorize/group the optical fields propagating within this volume as part of different spatial modes: (i) the optical modes that lastly arrive at the target output FOV, i.e., at FOV B for A → B and at FOV A for B → A; (ii) the optical modes arriving at the output plane but outside the target output FOV; and (iii) the unbounded optical modes that do not reach the output planes; since the diffractive layers are axially separated by >10λ, there are no evanescent waves being considered here. We calculated the power distribution percentages of each one of these types of optical modes for both A → B and B → A for each test image and reported their average values across the 10,000 test images in Fig. 3D (see Materials and Methods for details). The results summarized in Fig. 3D clearly reveal that, in the forward path (A → B) of the diffractive unidirectional imager, the majority of the input power (>93.5%) is coupled to the imaging modes that arrive at the output FOV B, forming high-quality images of the input objects with a mean PCC of 0.974, while the optical modes that fall outside the FOV B and the unbound modes are minimal, accounting for only ~2.95 and ~3.54% of the input total power, respectively. In contrast, the backward imaging path (B → A) of the same diffractive unidirectional imager steers most of the input power into the nonimaging modes that fall outside the FOV A or escape out of the diffractive volume through the unbounded modes, which correspond to power percentages of ~34.8 and ~63.6%, respectively. For B → A, the optical modes that arrive at the FOV A only constitute, on average, ~1.57% of the input total power; however, these optical modes are not only weak but also substantially aberrated by the diffractive unidirectional imager, resulting in very poor output images, with a mean PCC value of ~0.38.

 The underlying reason for these contrasting power distributions in the two imaging directions stems from the different order of the diffractive layers as the light passes through them. This can be further confirmed through the analyses reported in fig. S1, where we provided a visualization of the variation of the optical fields propagating within the same diffractive design. As shown in fig. S1C, in the forward operation, A → B, the diffractive layers arranged in the order of L1 to L5 manage to maintain most of the light waves at the central regions throughout the wave propagation such that the input image is efficiently focused within output FOV B to form high-quality output images. In contrast, in the backward operation, B → A, as shown in fig. S1D, the same set of diffractive layers arranged in the reversed order (i.e., L5 to L1) scatter the transmitted input optical fields and couple them into nonimaging modes (i.e., unbound modes that leave the diffractive imager volume and modes that end up outside the output image FOV); both of these set of modes never arrive at the output image FOV A. In addition to this, for the backward operation, B → A, the diffractive layers ordered in the reverse direction (L5 to L1) scramble the distributions of the optical fields that arrive at FOV A, suppressing their structural resemblance to the input images.

Note that, since the presented diffractive unidirectional imager is composed of linear, time-invariant and isotropic materials, it forms a reciprocal system that is polarization insensitive. In experimental implementations (reported below) due to absorption-related losses, a diffractive unidirectional imager also exhibits time-reversal asymmetry.

To further highlight the capabilities of our diffractive unidirectional imager (which was trained using handwritten digits), we also tested its external generalization using other datasets: The Extended MNIST (EMNIST) dataset that contains images of handwritten English letters and the Fashion-MNIST dataset that contains images of various fashion products. The blind testing results on these two additional datasets using the diffractive unidirectional imager of Fig. 2C are exemplified in Fig. 2E, which once again confirm its success. As another demonstration of the external generalization of our diffractive unidirectional imager, we reversed the contrast of the images in these test datasets, where the light transmitting and blocking regions of the input images were swapped, further deviating from our training image set. The results of this analysis are presented in fig. S2, demonstrating successful unidirectional imaging using our diffractive design, irrespective of the test image dataset and the contrast of the input image features.

In addition to these, we quantified the imaging resolution performance of this diffractive unidirectional imager using gratings as resolution test targets, which were also never used in the training phase (see Fig. 4). Our results reveal that the diffractive unidirectional imager can resolve a minimum linewidth of ~4λ in the forward path, A → B, while successfully inhibiting the image formation in the reverse path, B → A, as expected. These results once again prove that the training of the diffractive unidirectional imager is successful in approximating a general-purpose imaging operation in the forward path, although we only used handwritten digits during its training.

Fig. 4. Spatial resolution analysis for the diffractive unidirectional imager shown in Fig. 2. Resolution test target images composed of grating patterns with different periods and orientations and their corresponding diffractive output images are shown for both the forward and backward imaging directions. The red lines indicate the one-dimensional (1D) cross-sectional profiles calculated by integrating the intensity of the grating patterns in the diffractive output images along the direction perpendicular to the grating.

Spectral response of the diffractive unidirectional imager

Next, we explored the spectral response of the diffractive unidirectional imager reported in Fig. 2 under different illumination wavelengths that deviate from the training illumination wavelength (λtrain = λ). The results of this analysis are reported in Fig. 5 (A and B), where the output image PCC and diffraction efficiency values of the diffractive unidirectional imager of Fig. 2 were tested as a function of the illumination wavelength. Although this diffractive unidirectional imager was only trained at a single illumination wavelength (λ), it also works well over a large spectral range as shown in Fig. 5 (A and B). Our results reveal that the imaging performance in the forward path (A → B) remains very good with an output image PCC value of ≥0.85 and an output diffraction efficiency of ≥85.5% within the entire spectral range [λL : λR], where λL = 0.92 × λ and λR = 1.11 × λ (see Fig. 5, A and B). Within the same spectral range defined by [λL : λR], the power suppression ratio between the forward and backward imaging paths always remains ≥17.4×, and the output diffraction efficiency of the reverse path (B → A) remains ≤5.49% (see Fig. 5B), indicating the success of the diffractive unidirectional imager over a large spectral band, despite the fact that it was only trained with monochromatic illumination at λ. Figure 5D further reports examples of test objects (never seen during the training) that are simultaneously illuminated by a continuum of wavelengths, covering two different broadband illumination cases: (i) [0.92 × λ : 1.11 × λ] and (ii) [0.85 × λ : 1.15 × λ]. The forward and backward imaging results for these two broadband illumination cases shown in Fig. 5D clearly illustrate the success of the diffractive unidirectional imager under broadband illumination.

Fig. 5. Spectral response of the diffractive unidirectional imager design shown in Fig. 2. (A and B) Output image PCC (A) and diffraction efficiency (B) of the diffractive unidirectional imager in the forward and backward directions as a function of the illumination wavelength used during the blind testing. The values of the power suppression ratio are also reported in (B), which refers to the ratio between the output diffraction efficiency of the forward operation and the backward operation. The shaded areas indicate the SD values calculated based on all the 10,000 images in the testing dataset. (C) Examples of the output images in the forward and backward directions when using different illumination wavelengths during the testing, along with the corresponding input test images (never used during the training). (D) Broadband illumination results for several test objects are shown for the forward and backward imaging directions. Two different broadband illumination cases are shown, uniformly covering (i) 0.92 × λ to 1.11 × λ and (ii) 0.85 × λ to 1.15 × λ, where λ is the training illumination wavelength, λtrain = λ.

We should emphasize that these broadband unidirectional imaging results can be further enhanced by training the diffractive layers using a set of wavelengths sampled from a desired spectral band, as an alternative to using a single training wavelength. The validity of this approach is confirmed by an additional analysis reported in fig. S7 (A and B), which compares the spectral response of the model shown in Fig. 2 to that of another diffractive model trained using the same configuration and hyperparameters, but with the operational wavelength selected randomly within the spectral range of [λL : λR] during the training process, where λL = 0.92 × λ and λR = 1.11 × λ. As shown in fig. S7C, this training approach with a wide continuum of wavelengths substantially improves the output PCC values in the forward direction at illumination wavelengths far away from the center wavelength λ. These advantages of the broadband design also come at the expense of relatively reduced peak PCC values of the forward output images at λ and a small reduction in the forward diffraction efficiency.

Experimental validation of the diffractive unidirectional imager design
We experimentally validated our diffractive unidirectional imager using a monochromatic continuous-wave terahertz illumination at λ = 0.75 mm, as shown in Fig. 6A. A schematic diagram of the terahertz setup is shown in Fig. 6B, and its implementation details are reported in Materials and Methods. For this experimental validation, we designed a diffractive unidirectional imager composed of three diffractive layers, where each layer contains 100 by 100 learnable diffractive features, each with a lateral size of 0.64λ (dictated by the resolution of our 3D printer). The axial spacing between any two adjacent layers (including the diffractive layers and the input/output planes) is chosen as ~26.7λ. Different from earlier designs, here, we also took into account the material absorption using the complex-valued refractive index of the diffractive material in our optical model, such that the optical fields absorbed by the diffractive layers are also considered in our design (which will be referred to as the “absorbed modes” in the following discussion). Moreover, to overcome the undesired performance degradation that may be caused by the misalignment errors in an imperfect physical assembly of the diffractive layers, we also adopted a “vaccination” strategy in our design by introducing random displacements applied to the diffractive layers during the training process, which enabled the final converged diffractive unidirectional imager to become more resilient to potential misalignment errors (see Materials and Methods).

After the training was complete, we conducted numerical performance analysis for this converged diffractive design using blind testing objects, with the results shown in fig. S3. Upon comparison to the earlier model presented in Fig. 2, which had a high output diffraction efficiency of >90% in its forward direction, we found that this experimental design exhibits a relatively lower diffraction efficiency of ~21.33% in the forward imaging direction, A → B. This power efficiency reduction can be attributed to two main factors: (i) The existence of absorption by the diffractive layers caused ~27% of the input power to be lost through the absorbed modes; and (ii) our experimental design choice of using fewer diffractive layers (i.e., three layers) resulted in a reduced number of trainable diffractive features, leading to a larger portion of the input power (~46%) converted to the unbound modes. Nevertheless, this experimental design still maintains a substantially higher forward diffraction efficiency when compared to the backward direction, where ~1.8% of the input energy enters the output FOV (FOV A) in the reverse direction, B → A. Moreover, the forward and backward PCC values for this experimental design stand at 0.9618 ± 0.0100 and 0.4859 ± 0.0710, respectively, indicating the success of the unidirectional imager design.

After the training, the resulting diffractive layers were fabricated using a 3D printer (Fig. 6, C and D). In our experiments, we tested the performance of this 3D fabricated diffractive unidirectional imager along the forward and backward directions, as illustrated in Fig. 7 (A and B). Ten different handwritten digit samples from the blind testing set (never used in the training) were used as the input test objects, also 3D printed. These experimental imaging results for A → B and B → A are shown in Fig. 7C, which present a good agreement with their numerical simulated counterparts, very well matching the input images. As expected, 3D printed diffractive unidirectional imager faithfully imaged the input objects in its forward direction and successfully blocked the image formation in the backward direction; these results constitute the first demonstration of unidirectional imaging.

Fig. 7. Experimental results. (A and B) Layout of the diffractive unidirectional imager that was fabricated for experimental validation when it operates in the forward (A) and backward (B) directions. (C) Experimental results of the unidirectional imager using the fabricated diffractive layers.

Wavelength-multiplexed unidirectional diffractive imagers
Next, we consider a more challenging task: combining two diffractive unidirectional imagers that operate in opposite directions, where the direction of imaging is controlled by the illumination wavelength. The resulting diffractive system forms a wavelength-multiplexed unidirectional imager, where the image formation from A → B and B → A is maintained at λ1 and λ2 illumination wavelengths, respectively, whereas the image formation from B → A and A → B is blocked at λ1 and λ2, respectively (see Figs. 8 and 9). To implement this wavelength-multiplexed unidirectional imaging concept, we designed another diffractive imager that operates at λ1 and λ2 = 1.13 × λ1 wavelengths and used an additional penalty term in the training loss function to improve the performance of the image blocking operations in each direction, A → B and B → A. More details about the numerical modeling and the training loss function for this wavelength-multiplexed diffractive design can be found in Materials and Methods.

Fig. 8. Illustration of the wavelength-multiplexed unidirectional diffractive imager. In this diffractive design, the image formation operation is performed along the forward direction at wavelength λ1 and the backward direction at λ2, while the image blocking operation is performed along the backward direction at λ1 and the forward direction at λ2. This diffractive imager works as a unidirectional imaging system at two different wavelengths, each with a reverse imaging direction with respect to the other. λ2 = 1.13 × λ1.

Fig. 9. Design schematic and blind testing results of the wavelength-multiplexed unidirectional diffractive imager. (A and B) Layout of the wavelength-multiplexed unidirectional diffractive imager when it operates in the forward (A) and backward (B) directions. (C) Exemplary blind testing input images taken from MNIST handwritten digits that were never seen by the diffractive imager model during its training, along with their corresponding diffractive output images at different wavelengths in the forward and backward directions. (D) Same as (C) except that the testing images are taken from the EMNIST and Fashion-MNIST datasets, demonstrating external generalization.

We trained this wavelength-multiplexed unidirectional diffractive imager using handwritten digit images as before; the resulting, optimized diffractive layers are reported in fig. S4. Following its training, the diffractive imager was blindly tested using 10,000 MNIST test images that were never used during the training process, with some representative testing results presented in Fig. 9C. These results indicate that the wavelength-multiplexed diffractive unidirectional imager successfully performs two separate unidirectional imaging operations, in reverse directions, the behavior of which is controlled by the illumination wavelength; at λ1, A → B image formation is permitted and B → A is blocked, whereas at λ2, B → A image formation is permitted and A → B is blocked.

We also analyzed the imaging performance of this wavelength-multiplexed unidirectional diffractive imager as shown in Fig. 10 (A to C). At the first wavelength channel λ1, the output PCC values for the forward (A → B) and backward (B → A) directions are calculated as 0.9428 ± 0.0154 and 0.1228 ± 0.0985, respectively, revealing an excellent image quality contrast between the two directions (see Fig. 10B). Similarly, the output diffraction efficiencies for the forward and backward directions at λ1 are quantified as 65.82 ± 3.57 and 3.62 ± 0.72%, respectively (Fig. 10C). In contrast, the second wavelength channel λ2 of this diffractive model performs unidirectional imaging along the direction opposite to that of the first wavelength, providing output PCC values of 0.9378 ± 0.0187 (B → A) and 0.0840 ± 0.0739 (A → B) (see Fig. 10B). Similarly, the output diffraction efficiencies at λ2 were quantified as 51.81 ± 3.77 (B → A) and 2.57 ± 0.36% (A → B). These findings can be further understood by investigating the power distribution within this wavelength-multiplexed unidirectional diffractive imager, which is reported in Fig. 10D. This power distribution analysis within the diffractive volume clearly shows how two different wavelengths (λ1 and λ2) along the same spatial direction (e.g., A → B) can result in very different distributions of spatial modes, performing unidirectional imaging in opposite directions, following the same physical behavior reported in Fig. 3D, except that this time it is wavelength-multiplexed, controlling the direction of imaging. Such an exotic wavelength-multiplexed unidirectional imaging system cannot be achieved using simple spectral filters such as absorption or thin-film filters, since the use of a spectral filter at one wavelength channel (for example, to block A → B at λ2) would immediately also block the reverse direction (B → A at λ2), violating the desired goal.

Fig. 10. Performance analysis of the wavelength-multiplexed unidirectional diffractive imager shown in Fig. 9 and fig. S2. (A and B) Normalized MSE (A) and PCC (B) values calculated between the input images and their corresponding diffractive outputs at different wavelengths in the forward and backward operations. (C) The output diffraction efficiencies of the diffractive imager calculated in the forward and backward operations. In (A) to (C), the metrics are benchmarked across the entire MNIST test dataset and shown with their mean values and SDs added as error bars. (D) Left: The power of the different spatial modes at the two wavelengths propagating in the diffractive volume during the forward and backward operations, shown as percentages of the total input power. Right: Schematic of the different spatial modes propagating in the diffractive volume.

We should also note that, since this wavelength-multiplexed unidirectional imager was trained at two distinct wavelengths that control the opposite directions of imaging, the spectral response of the resulting diffractive imager, after its optimization, is vastly different from the broadband response of the earlier designs, reported in, e.g., Fig. 5. Figure S5 reveals that the wavelength-multiplexed unidirectional imager (as desired and expected) switches its spectral behavior in the range between λ1 and λ2, since its training aimed unidirectional imaging at opposite directions at these two predetermined wavelengths. Therefore, this spectral response that is summarized in fig. S5 is in line with the training goals of this wavelength-multiplexed unidirectional imager. However, it still maintains its unidirectional imaging capability over a range of wavelengths in both directions. For example, fig. S5 reveals that the output image PCC values for A → B remain ≥0.85 within the entire spectral range covered by 0.975 × λ1 to 1.022 × λ1 without any considerable increase in the diffraction efficiency for the reverse path, B → A. Similarly, the output image PCC values for B → A remain ≥0.85 within the entire spectral range covered by 0.968 × λ2 to 1.029 × λ2 without any noticeable increase in the diffraction efficiency for the reverse path, A → B, within the same spectral band. These results highlighted in fig. S5 indicate that the wavelength-multiplexed unidirectional imager can also operate over a continuum of wavelengths around λ1 (A → B) and λ2 (B → A), although the width of these bands are narrower compared to the broadband imaging results reported in Fig. 5.

Last, we also tested the external generalization capability of this wavelength-multiplexed unidirectional imager on different datasets: handwritten letter images and fashion products as well as the contrast-reversed versions of these datasets. The corresponding imaging results are shown in Fig. 9D and fig. S6, once again confirming that our diffractive model successfully converged to a data-independent, generic imager where unidirectional imaging of various input objects can be achieved along either the forward or backward directions that can be switched/controlled by the illumination wavelength.

DISCUSSION
Our results constitute the first demonstration of unidirectional imaging. This framework uses structured materials formed by phase-only diffractive layers optimized through deep learning and does not rely on nonreciprocal components, nonlinear materials, or an external magnetic field bias. Because of the use of isotropic diffractive materials, the operation of our unidirectional imager is insensitive to the polarization of the input light, also preserving the input polarization state at the output. As we reported earlier in Results (Fig. 5), the presented diffractive unidirectional imagers maintain unidirectional imaging functionality under broadband illumination, over a large spectral band that covers, e.g., 0.85 × λ to 1.15 × λ, despite the fact that they were only trained using monochromatic illumination at λ. This broadband imaging performance was further enhanced, covering even larger input bandwidths, by training the diffractive layers of the unidirectional imager using a set of illumination wavelengths randomly sampled from the desired spectral band of operation as illustrated in fig. S7.

By examining the diffractive unidirectional imager design and the analyses shown in Fig. 2 and fig. S1, one can gain more insights into its operation principles from the perspective of the spatial distribution of the propagating optical fields within the diffractive imager volume. The diffractive layers L1 to L3 shown in Fig. 2C exhibit densely packed phase islands, similar to microlens arrays that communicate between successive layers. Conversely, the diffractive layers L4 and L5 have rapid phase modulation patterns, resulting in high spatial frequency modulation and scattering of light. Consequently, the propagation of light through these diffractive layers in different sequences leads to the modulation of light in an asymmetric manner (A → B versus B → A). To gain more insights into this, we calculated the spatial distributions of the optical fields within the diffractive imager volume in fig. S1 (C and D) for a sample object. We observe that, in the forward direction (A → B), the diffractive layers arranged with the order of L1 to L5 ensured that these optical fields propagated forward through the focusing by the microlens-like phase islands located in the diffractive layers L1 to L3, and as a result, the majority of the input power was maintained within the diffractive volume, creating a power efficient image of the input object at the output FOV. However, for the backward operation (B → A) where the diffractive layers are arranged in the reversed order (L5 to L1), the optical fields in the diffractive volume are initially modulated by the high spatial frequency phase patterns of the diffractive layers (i.e., L5 and L4), and during the early stages of the propagation within the diffractive volume, this leads to a large amount of radiation being channeled to the outer space aside the diffractive volume, in the form of unbound modes (see the green shaded areas in fig. S1, A and B). For the remaining spatial modes that managed to stay within the diffractive volume (propagating from B to A), they were guided by the subsequent diffractive layers (i.e., L3 to L1) to remain outside the output FOV (i.e., ending up within the orange shaded areas in fig. S1B).

One should note that the intensity distributions formed by these modes that lie outside the output FOV can be potentially measured by using, for example, side cameras that capture some of these scrambled modes. Such side cameras, however, cannot directly lead to meaningful, interpretable images of the input objects, as also illustrated in fig. S1. With the precise knowledge of the diffractive layers and their phase profiles and positions, one could potentially train a reconstruction digital neural network to make use of such side-scattered fields to recover the images of the input objects in the reverse direction of the unidirectional imaging system. This “attack” to digitally recover the lost image of the input object through side cameras and learning-based digital image reconstruction methods would not only require precise knowledge of the fabricated diffractive imager but can also be mitigated by surrounding the diffractive layers and the regions that lie outside the image FOV (orange regions in fig. 1, A and B) with absorbing layers/coatings that would protect the unidirectional imager against “hackers,” blocking the measurement of the scattered fields, except the output image aperture. Such absorbing layers also break the time-reversal symmetry of the imaging system, which help mitigate the risk of deciphering and decoding the original input in the backward direction.

Throughout this manuscript, we presented diffractive unidirectional imagers with input and output FOVs that have 28 by 28 pixels, and these designs were based on transmissive diffractive layers, each containing ≤200 by 200 trainable phase-only features. To further enhance the unidirectional imaging performance of these diffractive designs, one strategy would be to create deeper architectures with more diffractive layers, also increasing the total number (N) of trainable features. In general, deeper diffractive architectures present advantages in terms of their learning speed, output power efficiency, transformation accuracy, and spectral multiplexing capability (39, 44, 47, 48). Suppose an increase in the space-bandwidth product (SBP) of the input FOV A (SBPA) and the output FOV B (SBPB) of the unidirectional imager is desired, for example, due to a larger input FOV and/or an improved resolution demand; in that case, this will necessitate an increase in N proportional to SBPA × SBPB, demanding larger degrees of freedom in the diffractive unidirectional imager to maintain the asymmetric optical mode processing over a larger number of input and output pixels. Similarly, the inclusion of additional diffractive layers and features to be jointly optimized would also be beneficial for processing more complex input spectra through diffractive unidirectional imagers. In addition to the wavelength-multiplexed unidirectional imager reported in Figs. 8 to 10, an enhanced spectral processing capability through a deeper diffractive architecture may permit unidirectional imaging with, e.g., a continuum of wavelengths or a set of discrete wavelength across a desired spectral band. Furthermore, by properly adjusting the diffractive layers and the learnable phase features on each layer, our designs can be adapted to input and output FOVs that have different numbers and/or sizes of pixels, enabling the design of unidirectional imagers with a desired magnification or demagnification factor.

Although the presented diffractive unidirectional imagers are based on spatially coherent illumination, they can also be extended to spatially incoherent input fields by following the same design principles and deep learning–based optimization methods presented in this work. Spatially incoherent input radiation can be processed using phase-only diffractive layers optimized through the same loss functions that we used to design unidirectional imagers reported in our Results. For example, each point of the wavefront of an incoherent field can be decomposed, point by point, into a spherical secondary wave, which coherently propagates through the diffractive phase-only layers; the output intensity pattern will be the superposition of the individual intensity patterns generated by all the secondary waves originating from the input plane, forming the incoherent output image. However, the simulation of the propagation of each incoherent field through the diffractive layers requires a considerably increased number of wave propagation steps compared to the spatially coherent input fields, and as a result, the training of spatially incoherent diffractive imagers would take longer.

MATERIALS AND METHODS

Numerical forward model of a diffractive unidirectional imager

In the forward model of our diffractive unidirectional imager design, the input plane, diffractive layers, and output plane are positioned sequentially along the optical axis, where the axial spacing between any two of these layers (including the input and output planes) is set as d. For the numerical and the experimental models used here, the value of d is empirically chosen as 10 and 20 mm, respectively, corresponding to 13.33λ and 26.67λ, where λ = 0.75 mm. In our numerical simulations, the diffractive layers are assumed to be thin optical modulation elements, where the mth neuron on the kth layer at a spatial location (xm, ym, zm) represents a wavelength-dependent complex-valued transmission coefficient, tk, given by

where n(λ) and κ(λ) are the refractive index and the extinction coefficient of the diffractive layer material, respectively; these correspond to the real and imaginary parts of the complex-valued refractive index n~(λ) , i.e., n~(λ)=n(λ)+jκ(λ) (34). For the diffractive unidirectional imager validated experimentally at λ = 0.75 mm, the values of n~(λ) are measured using a terahertz spectroscopy system to reveal n(λ) = 1.700 and κ(λ) = 0.017 for the 3D printing material that we used. The same refractive index value n(λ) = 1.700 is also used in all the diffractive imager models used in our numerical analyses with κ = 0. hkm denotes the thickness value of each diffractive feature on a layer, which can be written as

where hlearnable refers to the learnable thickness value of each diffractive feature and is confined between 0 and hmax. The additional base thickness, hbase, is a constant that serves as the substrate (mechanical) support for the diffractive layers. To constrain the range of hlearnable, an associated latent trainable variable hv was defined using the following analytical form

where Sigmoid(hv) is defined as

Note that before the training starts, hv values of all the diffractive features were initialized as 0. In our implementation, hmax is chosen as 1.07 mm for the diffractive models that use λ = 0.75 mm so that the phase modulation of the diffractive features covers 0 to 2π. For the diffractive imager model that performs wavelength-multiplexed unidirectional imaging, hmax was empirically selected as 1.6 mm, still covering 0 to 2π phase range for both wavelengths (λ1 = 0.75 mm and λ2 = 0.85 mm). The substrate thickness, hbase, was assumed to be 0 in the numerical diffractive models and was chosen as 0.5 mm in the diffractive model used for the experimental validation. The diffractive layers of a unidirectional imager are connected to each other by free space propagation, which is modeled through the Rayleigh-Sommerfeld diffraction equation (33, 49)

where fkm(x,y,z,λ) is the complex-valued field on the mth pixel of the kth layer at (x, y, z), which can be viewed as a secondary wave generated from the source at (xm, ym, zm), r=(x−xm)2+(y−ym)2+(z−zm)2−−−−−−−−−−−−−−−−−−−−−−−−−−−√ , and j=−1−−−√ . For the kth layer (k ≥ 1, assuming that the input plane is the 0th layer), the modulated optical field Ek at location (xm, ym, zm) is given by

where S denotes all the diffractive features located on the previous diffractive layer. In our implementation, we used the angular spectrum approach (33) to compute Eq. 6, which can be written as

where F and F −1 denote the 2D Fourier transform and the inverse Fourier transform operations, respectively, both implemented using a fast Fourier transform. H(xn, yn, zm − zn, λ) is the transfer function of free space

where fx and fy represent the spatial frequencies along the x and y directions, respectively.

Training loss functions and image quantification metrics

 We first consider a generic form of a diffractive unidirectional imager, where the image formation is permitted in one direction (e.g., A → B), and it is inhibited in the opposite direction (e.g., B → A) at a single training wavelength, λ. The training loss function for such a diffractive unidirectional imager was defined as

where I(λ) stands for the input image illuminated at a wavelength of λ and OImg(λ) and OBlk(λ) denote the output images in the forward and backward directions, respectively. All the input and output images have the perspective of the illumination beam direction, flipping them left to right as one switches the illumination direction, A → B or B → A. L ImgMSE penalizes the normalized MSE between the OImg(λ) and its ground truth, which can be written as

where *(x, y, λ) indexes the individual pixels at spatial coordinates (x, y) and wavelength λ and V denotes the defined FOV that has Nx × Ny pixels at the input or output plane. σ is a normalization constant used to normalize the energy of the diffractive output, thereby ensuring that the computed MSE value is not influenced by the errors arising from the output diffraction efficiency (50), and it is given by the following expression

L EffBst is used to improve the output diffraction efficiency along the imaging direction (e.g., A → B), which is defined as

where η(·) is the output diffraction efficiency of the diffractive unidirectional imager and βEffBst is an empirical weight coefficient, which was set as 1.0 during the training of all the diffractive models. η was defined as

L ImgBlk is defined to penalize the structural resemblance between the input image and the diffractive imager output along the image blocking direction (e.g., B → A)

where PCC stands for the Pearson correlation coefficient, defined as

L EffSqz in Eq. 9 is used to penalize the output diffraction efficiency in the backward direction

αImgBlk, αEffBst, and αEffSqz in Eq. 9 are the empirical weight coefficients associated with L ImgBlk, L EffBst, and L EffSqz, respectively. We denote the diffractive unidirectional imager output images for A → B and B → A as OA→B(λ) and OB→A(λ), respectively. For the diffractive unidirectional imaging models that were trained using a single illumination wavelength (e.g., in Figs. 2 and 7), the image formation is set to be maintained in the forward direction (A → B) and inhibited in the backward direction (B → A), i.e., OImg(λ) = OA→B(λ) and OBlk(λ) = OB→A(λ). Therefore, the loss function for training these models can be formulated as

where L (·) refers to the same loss function defined in Eq. 9. During the training of the unidirectional imager models with five diffractive layers and a single training wavelength channel, the empirical weight coefficients αImgBlk, αEffBst, and αEffSqz were set as 0, 0.001, and 0.001, respectively; during the training of the other model with three diffractive layers used for the experimental validation, the same weight coefficients were set as 0, 0.01, and 0.003, respectively.

 For the wavelength-multiplexed unidirectional diffractive imager model shown in Fig. 9, at λ1, the image formation is permitted in the direction A → B and inhibited in the direction B → A, whereas at λ2, the image formation is permitted in the direction B → A and inhibited in the direction A → B, respectively; i.e., OImg(λ1) = OA→B(λ1), OBlk(λ1) = OB→A(λ1), OImg(λ2) = OB→A(λ2), and OBlk(λ2) = OA→B(λ2). Accordingly, we formulated the loss function used for training this model as

where L (·) refers to the loss function defined in Eq. 9. During the training of this model, the weight coefficients αImgBlk, αEffBst, and αEffSqz were empirically set as 0.0001, 0.001, and 0.001, respectively.

For quantifying the imaging performance of the presented diffractive imager designs, the reported values of the output MSE, output PCC, and output diffraction efficiency were directly taken from the calculated results of L ImgMSE, PCC, and η, respectively, revealing the averaged values across the blind testing image dataset. When calculating the power distributions of different optical modes within the diffractive volume, the power percentage of the output FOV modes takes the same value as η, and the power percentage outside the output FOV is computed by subtracting the total power integrated within the output image FOV from the total power integrated across the entire output plane. The power in the absorbed modes is calculated by summing up the power loss before and after the optical field modulation by each diffractive layer. After excluding the power of the above modes from the total input power, the remaining part is calculated as the power of the unbound modes.

Training details of the diffractive unidirectional imagers

For the numerical models used here, the smallest sampling period for simulating the complex optical fields is set to be identical to the lateral size of the diffractive features, i.e., ~0.53λ for λ = 0.75 mm. The input/output FOVs of these models (i.e., FOV A and B) share the same size of 44.8 by 44.8 mm2 (i.e., ~59.7λ × 59.7λ) and are discretized into 28 by 28 pixels, where an individual pixel corresponds to a size of 1.6 mm (i.e., ~2.13λ), indicating a four-by-four binning performed on the simulated optical fields.

For the diffractive model used for the experimental validation of unidirectional imaging, the sampling period of the optical fields and the lateral size of the diffractive features are chosen as 0.24 and 0.48 mm, respectively (i.e., 0.32λ and 0.64λ). This also results in a two-by-two binning in the sampling space where an individual feature on the diffractive layers corresponds to four sampling space pixels that share the same dielectric material thickness value. The input and output FOVs of this model (i.e., FOV A and B) share the same size of 36 by 36 mm2 (i.e., 48λ × 48λ) and are sampled into arrays of 15 by 15 pixels, where an individual pixel has a size of 2.4 mm (i.e., 3.2λ), indicating that a 10-by-10 binning is performed at the input/output fields in the numerical simulation.

During the training process of our diffractive models, an image augmentation strategy was also adopted to enhance their generalization capabilities. We implemented random translation, random up-to-down, and random left-to-right flipping of the input images using the transforms.RandomAffine function built-in PyTorch. The translation amount was uniformly sampled within a range of [−10, 10] and [−5, 5] pixels in the diffractive unidirectional imager models used for numerical analysis and the model used for the experimental validation, respectively. The flipping operation is set to be performed at a probability of 0.5.

All the diffractive imager models used in this work were trained using PyTorch (v1.11.0, Meta Platforms Inc.). We selected AdamW optimizer (51, 52), and its parameters were taken as the default values and kept identical in each model. The batch size was set as 32. The learning rate, starting from an initial value of 0.03, was set to decay at a rate of 0.5 every 10 epochs, respectively. The training of the diffractive models was performed with 50 epochs. For the training of our diffractive models, we used a workstation with a GeForce GTX 1080Ti graphical processing unit (Nvidia Inc.) and Core i7-8700 central processing unit (Intel Inc.) and 64 GB of RAM, running Windows 10 operating system (Microsoft Inc.). The typical time required for training a diffractive unidirectional imager is ~3 hours.

Vaccination of the diffractive unidirectional imager against experimental misalignments

During the training of the diffractive unidirectional imager design for experimental validation, possible inaccuracies imposed by the fabrication and/or mechanical assembly processes were taken into account in our numerical model by treating them as random 3D displacements (D) applied to the diffractive layers (53). D can be written as

where Dx and Dy represent the random lateral displacement of a diffractive layer along the x and y directions, respectively, and Dz represents the random perturbation added to the axial spacing between any two adjacent layers (including diffractive layers, input FOV A, and output FOV B). Dx, Dy, and Dz of each diffractive layer were independently sampled based on the following uniform (U) random distributions

where Δ*,tr denotes the maximum amount of shift allowed along the corresponding axis, which was set as Δx,tr = Δy,tr = 0.48 mm (i.e., 0.64λ) and Δz,tr = 1.5 mm (i.e., 2λ) during the training process. Following the training under this vaccination strategy, the resulting diffractive unidirectional imager shows resilience against possible misalignments in the fabrication and assembly of the diffractive layers.

Note that, in addition to the 3D displacements of the diffractive layers, there may also exist other types of alignment errors in our experimental setup, such as 3D rotational misalignments of the diffractive layers. However, since the holders used to fix the diffractive layers are, in general, manufactured with high structural precision and surface flatness, we did not incorporate these types of misalignments into our forward model considering their negligible impact in our case. In the event that such rotational misalignments of the diffractive layer become an important factor in the experimental results, the undesired in-plane rotations of the diffractive layers can be readily modeled through applying a 2D coordinate transformation based on unitary rotation matrices, while the out-of-plane rotation of the diffractive layers can be addressed by modifications to the formulation of the wave propagation between tilted diffractive planes (53–55).

Experimental terahertz imaging setup

We fabricated the diffractive layers using a 3D printer (PR110, CADworks3D). The test objects were also 3D printed (Objet30 Pro, Stratasys) and coated with aluminum foil to define the light-blocking areas, with the remaining openings defining the transmission areas. We used a holder that was also 3D printed (Objet30 Pro, Stratasys) to assemble the printed diffractive layers along with input objects, following the relative positions of these components in our numerical design.

A terahertz continuous-wave scanning system was used for testing our diffractive unidirectional imager design. According to the experimental setup illustrated in Fig. 6B, we used a terahertz source in form of a WR2.2 modular amplifier/multiplier chain (AMC), followed by a compatible diagonal horn antenna (Virginia Diodes Inc.). A 10-dBm radiofrequency (RF) input signal at 11.1111 GHz (fRF1) at the input of AMC is multiplied 36 times to generate the output radiation at 400 GHz, corresponding to a wavelength of λ = 0.75 mm. The AMC output was also modulated with a 1-kHz square wave for lock-in detection. The assembled diffractive unidirectional imager is placed ∼600 mm away from the exit aperture of the horn antenna, which results in an approximately uniform plane wave impinging on its input FOV (A) with a size of 36 by 36 mm2 (i.e., 48λ × 48λ). The intensity distribution within the output FOV (B) of the diffractive unidirectional imager was scanned at a step size of 1 mm by a single-pixel mixer/AMC (Virginia Diodes Inc.) detector on an xy positioning stage that was built by combining two linear motorized stages (Thorlabs NRT100). The detector also receives a 10-dBm sinusoidal signal at 11.083 GHz (fRF2) as a local oscillator for mixing to down-convert the output signal to 1 GHz. The signal is then fed into a low-noise amplifier (Mini-Circuits ZRL-1150-LN+) with a gain of 80 dBm, followed by a band-pass filter at 1 GHz (± 10 MHz) (KL Electronics 3C40-1000/T10-O/O), so that the noise components coming from unwanted frequency bands can be mitigated. Then, after passing through a tunable attenuator (HP 8495B) used for linear calibration, the final signal is sent to a low-noise power detector (Mini-Circuits ZX47-60). The detector output voltage is measured by a lock-in amplifier (Stanford Research SR830) with the 1-kHz square wave used as the reference signal. Last, the lock-in amplifier readings were calibrated into a linear scale. In our postprocessing, linear interpolation was applied to each measurement of the intensity field to match the pixel size of the output FOV (B) used in the design phase, resulting in the output measurement images shown in Fig. 7C.

Materials

3D-Printed Microinjection Needle Arrays via a Hybrid DLP-Direct Laser Writing Strategy

3D-Printed Microinjection Needle Arrays via a Hybrid DLP-Direct Laser Writing Strategy

Sunandita Sarker, Adira Colton, Ziteng Wen, Xin Xu, Metecan Erdi, Anthony Jones, Peter Kofinas, Eleonora Tubaldi, Piotr Walczak, Miroslaw Janowski, Yajie Liang, Ryan D. Sochol

Microinjection protocols are ubiquitous throughout biomedical fields, with hollow microneedle arrays (MNAs) offering distinctive benefits in both research and clinical settings. Unfortunately, manufacturing-associated barriers remain a critical impediment to emerging applications that demand high-density arrays of hollow, high-aspect-ratio microneedles. To address such challenges, here, a hybrid additive manufacturing approach that combines digital light processing (DLP) 3D printing with “ex situ direct laser writing (esDLW)” is presented to enable new classes of MNAs for fluidic microinjections. Experimental results for esDLW-based 3D printing of arrays of high-aspect-ratio microneedles—with 30 µm inner diameters, 50 µm outer diameters, and 550 µm heights, and arrayed with 100 µm needle-to-needle spacing—directly onto DLP-printed capillaries reveal uncompromised fluidic integrity at the MNA-capillary interface during microfluidic cyclic burst-pressure testing for input pressures in excess of 250 kPa (n = 100 cycles). Ex vivo experiments perform using excised mouse brains reveal that the MNAs not only physically withstand penetration into and retraction from brain tissue but also yield effective and distributed microinjection of surrogate fluids and nanoparticle suspensions directly into the brains. In combination, the results suggest that the presented strategy for fabricating high-aspect-ratio, high-density, hollow MNAs could hold unique promise for biomedical microinjection applications.

We kindly thank the researchers at University of Maryland for this collaboration, and for sharing the results obtained with their system.

Introduction

Microinjection technologies underlie a diversity of biomedical applications, such as in vitro fertilization, intraocular injection, therapeutic drug and vaccine delivery, developmental biology, and transgenics.[1-4] Historically, microinjection protocols have relied on using a single hollow microneedle to deliver target substances (e.g., cells, DNA, RNA, micro/nanoparticles) to a singular location of interest.[5-7] Recently, however, alternatives in the form of microneedle arrays (MNAs) have garnered increasing interest due to a wide range of benefits over their single-needle counterparts, including the ability to rapidly deliver target material over a large, distributed area, which has proven to be particularly beneficial for transdermal and intradermal drug delivery.[8-11] Despite the significant potential of MNAs for microinjection applications, the majority of current MNA developments are founded on solid (e.g., coated and/or dissolvable) microneedles that are inherently incompatible with active fluidic microinjection protocols.[12-14] This focus on solid MNAs is, in part, due to the considerable challenges associated with manufacturing arrays comprising hollow microneedles at small scales. Specifically, although researchers have demonstrated that conventional clean room-based micromachining approaches can be adapted to fabricate arrays of hollow microneedles,[15-17] such protocols can be exceedingly time-, cost-, and labor-intensive, while restricting the architectures of the microneedles to low-aspect-ratio “2.5D” geometries.[18-20] The geometric limitations, in particular, represent a significant barrier to extending the benefits of MNAs to emerging microinjection applications, such as for treatments of neurological conditions.

One example of such a treatment in which MNAs could potentially offer benefits over single-needle injection strategies is stem cell therapy (SCT). A crucial obstacle to the clinical efficacy of SCT is the poor viability of stem cells following delivery into the brain.[21-23] One challenge associated with conventional needles is cell crowding at the injection site due to the high concentrations of donor cells (e.g., up to 100 000 cells µL−1),[2425] which can lead to large cell spheroids with undesirable conditions (e.g., decreased access to O2 and nutrients for interior cells) that contribute to the low survival rates of implanted stem cells.[26-29] It is possible that simultaneous, distributed cell delivery via MNAs could provide novel means to improve cell survival rates by reducing cell crowding; however, no MNA yet exists to enable such studies. For instance, even in the case of mice—a widely used disease model[30] with a relatively shallow (≈1 mm) cerebral cortex compared to other animal models[31]—the ability to penetrate into the cerebral cortex for therapeutics delivery would necessitate hollow microneedles that not only comprise outer diameters (ODs) on the order of tens of micrometers but also include heights in excess of 500 µm. Consequently, new strategies for manufacturing MNAs composed of such high-aspect-ratio, hollow microneedles are in critical demand.

Additive manufacturing (or colloquially, “3D printing”) technologies offer distinctive benefits for applications that require a high degree of geometric control in component fabrication.[32-34] Previously, researchers have demonstrated a wide range of 3D printing techniques for the fabrication of needle arrays at various scales. For example, at larger scales, Derakhshandeh et al. used extrusion-based 3D printing (e.g., “direct ink writing”) to manufacture arrays of hollow, millimeter-scale needles for drug delivery,[35] which facilitated enhanced wound healing.[36] For mesoscale needles, however, the print speed and geometric limitations of extrusion-based methods at smaller scales[37-39] have motivated investigators to instead focus on fabricating MNAs via vat photopolymerization approaches, such as stereolithography and digital light processing (DLP) 3D printing.[40-42] Unfortunately, these printing techniques are poorly suited for printing hollow MNAs that comprise needles with sub-100 µm ODs, which has led to increasing interest in the use of “direct laser writing (DLW)” for such cases.

DLW entails scanning a femtosecond pulsed IR laser in a point-by-point, layer-by-layer manner to selectively crosslink a photocurable material in target locations via two-photon (or multiphoton) polymerization to ultimately produce 3D objects comprising cured photomaterial with feature resolutions down to the 100 nm range.[43-46] Previously, researchers have demonstrated the utility of using DLW to print MNA master molds, which can then be used to replicate solid MNAs with drug coatings[47-50] or solid MNAs that are fully dissolvable.[5152] Additionally, Rad et al. reported the use of DLW to print molds and MNAs directly that include open (i.e., unenclosed) side channels.[53-55] For realizing hollow microneedles that are a requisite for microinjection applications, one key challenge inherent to the submicrometer-scale resolution of the DLW-printing volume element (i.e., “voxel”) is that it is ill suited for constructing the larger macro-to-microinterfaces (e.g., input ports) required for delivering fluids to the needles.[565792] To avoid the undesirable costs and time associated with fabricating macro-to-microinterfaces in their entirety via DLW,[58] researchers have instead DLW-printed hollow singular microneedles (aspect ratios ≈4–5)[59] and MNAs (aspect ratios ≈2–5)[60] as isolated entities, and then used adhesives (e.g., glue) to manually connect the printed components to macroscale fluidic interfaces. Trautmann et al. bypassed such protocols by employing a fabrication methodology that combines femtosecond laser irradiation, annealing, grinding, and polishing to produce microchips with external openings, and then DLW-printing truncated cone-shaped MNAs (aspect ratios ≈1.3–3) directly onto the chips.[61] In contrast to the aforementioned approaches, printing MNAs directly onto fluidic connectors (e.g., at the end of capillaries) would overcome many of the current interface-associated barriers to MNA utility. Furthermore, to our knowledge, no report yet exists (for conventional or additive manufacturing-based approaches) in which MNAs are fabricated with hollow, high-aspect-ratio (e.g., ≥10) microneedles with microscale ODs (e.g., <100 µm) and high array densities (e.g., ≤100 µm needle-to-needle spacing) relevant to emerging microinjection applications, such as the delivery of therapeutic fluidic payloads directly into brain tissue.

In this work, we introduce a novel hybrid additive manufacturing strategy that entails using DLP 3D printing to fabricate batches of capillaries in set positions (Figure 1a,b), and then employing an “ex situ DLW (esDLW)” approach to DLW-print hollow, high-aspect-ratio, high-density MNAs directly onto—and notably, fluidically sealed to—the DLP-printed capillaries (Figure 1c,d). Thereafter, individual MNA-capillary assemblies can be selectively released by disrupting the connections to the batch (Figure 1e, arrows) and then interfaced with injector systems for microinjection applications. As an exemplar, we investigate the utility of the MNAs for performing microinjections into brain tissue (Figure 1f) by using excised mouse brains to not only evaluate MNA penetration into and retraction from the tissue with respect to microneedle integrity but also explore the efficacy of MNA-mediated delivery of microfluidic cargo (e.g., aqueous fluids and nanoparticle suspensions) into brain tissue ex vivo.

Conceptual illustrations of the hybrid additive manufacturing strategy for 3D microprinting hollow, high-aspect-ratio microneedle arrays (MNAs) for microinjection applications. a,b) Digital light processing (DLP)-based 3D printing of batch capillaries. a) A liquid-phase photocurable material is UV-crosslinked in designated locations in a layer-by-layer manner to produce a batch of arrayed capillaries comprising cured photomaterial. b) The DLP-printed batch of prealigned capillaries following the development process. c–e) “Ex situ direct laser writing (esDLW)” of MNAs directly atop—and fluidically sealed to—each DLP-printed capillary. c) A femtosecond pulsed IR laser is scanned to selectively initiate two-photon polymerization of a liquid-phase photocurable material in a point-by-point, layer-by-layer manner to produce MNAs comprising cured photomaterial. d) A batch array of MNA-capillary assemblies following the DLW-associated development process. e) Individual MNA-capillary assemblies within the array can be released on demand by manually severing the supporting structures (arrows). f) Example application of integrating MNA-capillary assemblies with nanoinjector systems to facilitate MNA-mediated simultaneous, distributed microinjections of target fluidic substances/suspensions into brain tissue.

Materials

Clear Microfluidics Resin V7.0a

2 Results and Discussion

Hybrid Additive Manufacturing of Hollow MNAs

The presented hybrid additive manufacturing strategy consists of two fundamental stages: i) DLP 3D printing of batch arrays of capillaries and ii) esDLW-based printing of the MNAs directly atop each capillary. DLP 3D printing is a vat photopolymerization approach in which a DLP projector is used to UV-crosslink a liquid-phase photocurable material in designated locations in a layer-by-layer manner to ultimately produce 3D objects composed of cured photomaterial.[62] Here, we leveraged DLP 3D printing to fabricate batches of arrayed capillaries in a single print run to overcome several drawbacks of recent esDLW approaches for printing 3D micro/nanostructured objects onto mesoscale fluidic components, such as micropiston-based microgrippers[63] and liquid biopsy systems[64] onto fluidic capillaries. First, the geometric control afforded by DLP 3D printing allows for each capillary to be designed with a variable OD to match the dimensions of the capillary base to those of the desired injector system. This capillary-specific geometric customization capability obviates the need for additional fluidic adapters and/or sealants (e.g., glues) often required to couple the mesoscale capillaries to macroscale fluidic equipment (e.g., injector systems).[63-65] Second, the outer dimensions of the batch array can be designed to support facile loading into the DLW 3D printer, which eliminates the time, labor, and costs associated with manufacturing and employing custom-built capillary holders typically needed for esDLW approaches.[63-65] Lastly, the ability to print all of the capillaries in predefined array locations—with uniform surface positions and rotational orientations—addresses critical deficits associated with the use of custom-built capillary holders that rely on undesired manual (e.g., by hand and/or eye) alignment protocols for each individual capillary.[65]

For DLP 3D printing of the batch capillary arrays, we used a Miicraft M50 microfluidics DLP 3D printer (CADworks3D, Toronto, ON, Canada) to fabricate two batches (i.e., 18 capillaries in total) per print run, which corresponded to a total print time of less than 45 min (Movie S1, Supporting Information). To enable direct integration with the nanoinjector system (MO-10, Narishige International USA, Inc., Amityville, NY), we designed each capillary with a consistent inner diameter (ID) of 650 µm, but with a variable OD that was set at 1.2 mm for the top 1.5 mm and then gradually increased to 2.4 mm for the remainder of the 10 mm length of the capillary (Figure S1, Supporting Information). Fabrication results revealed effective construction of the arrayed capillaries—each attached to the batch via five connecting structures (400 µm in width and depth; 1.5 mm in length) (Figure 2a,b). In addition, the outer dimensions of the overall batch resolved such that the print could be readily loaded into the multi-DiLL holder of the DLW system (Photonic Professional GT2, Nanoscribe GmbH, Germany) (Figure S2, Supporting Information) to facilitate esDLW-based 3D printing.

Fabrication results for DLP 3D printing of batch arrays of capillaries and esDLW-based printing of MNAs. a,b) DLP prints of batch arrays of capillaries. a) Photograph of a complete batch with nine arrayed capillaries. Scale bar = 5 mm. Inset shows two batches attached to the build plate directly after DLP 3D printing (see Movie S1 in the Supporting Information). b) Low-vacuum scanning electron microscopy (SEM) images of a representative DLP-printed capillary attached to the batch via five connecting structures. Scale bars = 500 µm. c,d) The esDLW approach for printing MNAs directly onto DLP-printed capillaries in a single print run. c) Computer-aided manufacturing (CAM) simulations and d) corresponding images of the esDLW printing process. Scale bar = 250 µm (see Movie 2 in the Supporting Information). e–g) Low-vacuum SEM images of representative fabrication results showing: e) an esDLW-printed MNA atop a DLP-printed capillary following release from the batch array (see Movie S3 in the Supporting Information); f) a magnified view of the MNA; and g) a magnified view of a single microneedle tip in the array. Scale bars = e) 250 µm, f) 100 µm, and g) 25 µm.

We designed the MNAs to include identical hollow microneedles—each with an ID of 30 µm, an OD of 50 µm, and a height of 550 µm—with needle-to-needle spacing of 100 µm (Figure S3, Supporting Information). For the esDLW printing process, we initiated the print with 50 µm of overlap with the top surface of the capillary to ensure bonding at the interface. Computer-aided manufacturing (CAM) simulations and brightfield images of a corresponding esDLW process for printing the MNA directly onto a DLP-printed capillary are presented in Figure 2c,d, respectively (see also Movie S2, Supporting Information). The total esDLW printing process was completed in ≈10 min. Following development, we retrieved target MNA-capillary assemblies from the batch by manually severing the five connecting structures (Movie S3, Supporting Information). Images of the released MNA-capillary assemblies captured using low-vacuum scanning electron microscopy (SEM) revealed effective alignment and integration of the esDLW-printed MNAs with the DLP-printed capillaries, without any visible signs of physical defects along the MNA-capillary interface (Figure 2e). In addition, images of the esDLW-printed MNA and needle tips suggest that the manual release process did not appear to affect MNA integrity (Figure 2f,g).

In Silico and In Vitro Investigations of MNA Mechanical Performance

The critical first steps of MNA-based microinjection protocols involve the effective puncture and penetration into a target medium (e.g., biological tissue), which can impart significant mechanical forces on the microneedles.[66] Thus, the potential utility of MNAs is predicated on their ability to successfully withstand such mechanical loading conditions. To evaluate this capability for the esDLW-printed high-aspect-ratio MNAs, we employed both numerical and experimental approaches to elucidate the mechanical performance of the MNAs. We performed finite element analyses (FEA) to provide insight into the mechanical failure behavior of the MNAs when subjected to a compressive load applied longitudinally with respect to the needles. The simulation results revealed that each arrayed microneedle exhibited a buckling-like deformation with the largest displacements observed around the midpoint of the heights; however, needles positioned in the outer region (i.e., the needles radially arrayed farthest from the center of the MNA) displayed larger deformations compared to those located in the more central array positions (Figure 3a). This behavior arises from the load distribution caused by the disc-like base of the MNA, which deforms more in its central region than its peripherical region, thereby allowing the centrally located microneedles to rigidly displace more in the axial direction than their outer-region counterparts. According to the stress–strain curve generated from the FEA compressive loading simulations (Figure 3b), the overall MNA exhibited an effective Young's Modulus (E) of 4.31 MPa and yield strength (σy) of 135 kPa. We also numerically modeled MNA mechanics associated with puncture into the brain tissue. By characterizing the nonlinear response at the interface between the tips of the microneedles and the brain substrate, we found that the forces associated with the needles located in the outer region were larger than those in the central regions (Figure S4, Supporting Information), which is in agreement with the compressive loading analyses (Figure 3a).

The critical first steps of MNA-based microinjection protocols involve the effective puncture and penetration into a target medium (e.g., biological tissue), which can impart significant mechanical forces on the microneedles.[66] Thus, the potential utility of MNAs is predicated on their ability to successfully withstand such mechanical loading conditions. To evaluate this capability for the esDLW-printed high-aspect-ratio MNAs, we employed both numerical and experimental approaches to elucidate the mechanical performance of the MNAs. We performed finite element analyses (FEA) to provide insight into the mechanical failure behavior of the MNAs when subjected to a compressive load applied longitudinally with respect to the needles. The simulation results revealed that each arrayed microneedle exhibited a buckling-like deformation with the largest displacements observed around the midpoint of the heights; however, needles positioned in the outer region (i.e., the needles radially arrayed farthest from the center of the MNA) displayed larger deformations compared to those located in the more central array positions (Figure 3a). This behavior arises from the load distribution caused by the disc-like base of the MNA, which deforms more in its central region than its peripherical region, thereby allowing the centrally located microneedles to rigidly displace more in the axial direction than their outer-region counterparts. According to the stress–strain curve generated from the FEA compressive loading simulations (Figure 3b), the overall MNA exhibited an effective Young's Modulus (E) of 4.31 MPa and yield strength (σy) of 135 kPa. We also numerically modeled MNA mechanics associated with puncture into the brain tissue. By characterizing the nonlinear response at the interface between the tips of the microneedles and the brain substrate, we found that the forces associated with the needles located in the outer region were larger than those in the central regions (Figure S4, Supporting Information), which is in agreement with the compressive loading analyses (Figure 3a).

Numerical and experimental results for MNA mechanical characterizations. a,b) Finite element analysis (FEA) results for a) microneedle deformations and b) stress–strain curve corresponding to MNA mechanics under compressive loading conditions. c,d) Experimental results for MNA compression testing. c) Sequential images of the MNA during axial compression test. Inset shows an SEM image of an MNA following compressive failure. Scale bars = 250 µm (see Movie S4 in the Supporting Information). d) Stress–strain curve generated from compressive loading experiments (n = 3 MNAs). e–g) Sequential images of representative MNA penetration and retraction operations corresponding to hydrogels with agarose concentrations of: e) 2.4%, f) 5%, and g) 10%. Scale bars = 500 µm (see Movie S5 in the Supporting Information).

To experimentally examine the mechanical performance of the esDLW-printed MNA, we conducted two sets of puncture and penetration-associated studies. First, we performed axial compression tests with esDLW-printed MNAs (n = 3), which revealed buckling-type deformations of the microneedles with increasing loading until complete mechanical failure (Figure 3c and Movie S4, Supporting Information). From SEM images of MNAs following compressive testing, we observed several cases of complete fracture, but the majority of the arrayed microneedles remained intact with the caveat that the tips and the overall shapes of the needles exhibited plastic deformation (Figure 3c, inset). Quantified results for the stress–strain relationships for the esDLW-printed MNAs revealed an average E of 2.12 ± 0.35 MPa and σy of 155 ± 30 kPa (Figure 3d). Although these results provide insight into the upper boundaries of mechanical loading, compression testing using an impenetrable plate is limited in its direct relevance to microinjection applications that rely on microneedle penetration into a target medium. Thus, we also investigated the capacity for the esDLW-printed MNAs to puncture and penetrate into surrogate hydrogels with increasing concentrations of agarose that correspond to varying degrees of biologically relevant stiffness. In particular, we performed experiments with agarose concentrations of: i) 1.2% (E = 12.8 ± 1.1 kPa), which would support penetration into liver and breast tissue; ii) 2.4% (E = 27.5 ± 1.0 kPa), which is relevant to brain, heart, kidney, arterial, and prostate tissue; and iii) both 5% (E = 223 ± 14 kPa) and 10% (E = 268 ± 31 kPa), which are relevant to cartilage tissues (Figure S5, Supporting Information).[67-70] Experimental results revealed that the MNA successfully penetrated into the 1.2%, 2.4%, and 5% agarose gels; however, we observed buckling of the microneedles and failure to penetrate into the 10% agarose gel (Figure 3e–g and Movie S5, Supporting Information). These results suggest that the esDLW-printed MNA is sufficient for penetration into brain tissue as well as a variety of other tissues (e.g., liver, breast, heart, kidney, arterial, and prostate tissues), but alternative photomaterials (with stronger mechanical properties) and/or microneedles with geometrically enhanced strength (e.g., by increasing the OD) would be needed for microinjection applications involving target mediums with E in excess of 250 kPa.

In Vitro Microfluidic Interrogations of MNA-Capillary Interface Integrity

One of the most catastrophic failure modes for esDLW-based prints—whether for optical,[71] photonic,[72] mechanical,[73] or fluidic[63-65] structures—is the potential for the DLW-printed objects to detach from the meso/macroscale components on which they are additively manufactured. For biomedical MNA applications, the consequences of this type of failure could be particularly serious, such as an MNA detaching from the capillary while embedded in brain tissue following microinjection. To investigate the potential for this failure mode and, in turn, provide insight into the mechanofluidic integrity of the interface between the esDLW-printed MNAs and the DLP-printed capillaries, we performed microfluidic cyclic burst-pressure tests with the MNA-capillary assemblies. Initially, using an applied pressure set at 5 kPa, we gradually infused blue-dyed deionized (DI) water into the MNA-capillary assembly via the opposing end of the capillary (i.e., the side without the printed MNA) until the fluid began exiting the tips of the arrayed microneedles (Figure 4a and Movie S6, Supporting Information). Thereafter, we performed separate sets of cyclic burst-pressure experiments (n = 100 cycles per experiment) corresponding to applied pressures set at 100, 200, and 300 kPa (Figure 4b–d). Throughout the burst-pressure testing, we monitored the MNA-capillary interface under brightfield microscopy for visible signs of undesired leakage phenomena (e.g., fluid exiting at any point along the interface rather than out of the tops of the microneedle tips); however, we did not observe any instances of such flow behavior. Similarly, quantified results of fluid flow through the MNA-capillary assembly recorded during the burst-pressure tests did not exhibit any indications of burst events—i.e., large increases in flow rates after a certain point, despite the applied pressure remaining constant—nor signs of gradual leakage phenomena associated with the flow rates increasing from pressure cycle to pressure cycle over the course of the experiment. Rather, the flow rate magnitudes corresponding to the applied input pressures remained consistent throughout the burst-pressure experiments (Figure 4b–d), suggesting uncompromised fluidic integrity of the MNA-capillary interface for all cases examined.

Experimental results for MNA microfluidic investigations. a) Sequential images during fluidic infusion. Scale bar = 500 µm (see Movie S6 in the Supporting Information). b–d) Quantified results for representative cyclic burst-pressure experiments (n = 100 cycles) corresponding to input pressures targeting: b) 100 kPa, c) 200 kPa, and d) 300 kPa.

Ex Vivo Mouse Brain Studies of MNA Penetration, Microinjection, and Retraction Functionalities

As an exemplar with which to interrogate the penetration, microinjection, and retraction capabilities of the esDLW-printed MNAs, we excised brains with intact dura mater from euthanized 6-month-old male mice (Wildtype C57BL/6 J, Jackson Laboratory) for experimentation ex vivo (Figure 5a). We performed three sets of experiments to elucidate these fundamental MNA functionalities. First, we investigated the ability to execute penetration and retraction operations (but not fluidic microinjections) with the MNAs as critical measures of performance with respect to three potential failure modes that would critically limit the efficacy of the esDLW-printed MNAs: i) the sharpness of the tips of the microneedles—governed by the resolution of the DLW 3D printer—is insufficient to puncture the brain tissue without inducing significant deformation of the brain; ii) the mechanical properties of the high-aspect-ratio microneedles lead to buckling and/or fracture of the microneedles prior to effective penetration into the brain tissue; and/or iii) the forces during the penetration or retraction processes fracture the microneedles, causing microneedles (or fragments of microneedles) to remain embedded in the brain tissue after retraction completion. To facilitate the penetration and retraction studies, we interfaced each MNA-capillary assembly examined with a nanoinjector system fixed to a stereotactic frame as a means to enable precise position control while optically monitoring the MNA-brain tissue interactions. Experiments performed with three distinct MNA-capillary assemblies (n = 3 penetration and retraction operations for each distinct MNA-capillary assembly) revealed that the MNAs could successfully puncture the brain tissue within 1 mm of total displacement from initial contact and, importantly, without any visible signs of mechanical failure during any of the penetration or retraction operations (Figure 5b and Movie S7, Supporting Information). Images of the MNAs (captured after completion of the retraction process) corroborated these results, without any indications of microneedle-associated failure modes (e.g., buckling or fracture) or MNA detachment from the capillary (Figure 5c).

Experimental results for ex vivo MNA penetration, microinjection, and retraction operations using an excised mouse brain. a) Experimental setup including the MNA-capillary assembly interfaced with a custom-built nanoinjector and an excised mouse brain on ice. b,c) Brain tissue puncture and retraction results. b) Sequential images of MNA insertion into (≤20 s) and retraction from (≥20 s) the brain tissue. Scale bar = 1 mm (see Movie S7 in the Supporting Information). c) SEM image of the MNA after retraction from the brain tissue. Scale bar = 250 µm. d–f) MNA-mediated microinjection results. d) Sequential images of a representative MNA penetration, microinjection, and retraction process for a surrogate fluid (blue-dyed DI water) injected into brain tissue. Scale bar = 1 mm (see Movie S8, Supporting Information). e) Magnified view of the postinjection site. Scale bar = 250 µm. f) SEM image of the MNA following microinjection into the brain tissue. Scale bar = 250 µm.

After validating the penetration and retraction capabilities, we then initially investigated the microinjection functionality of the MNAs based on the ability to deliver a surrogate microfluidic payload into the brain tissue. In this case, we preloaded the MNA-capillary assembly with blue-dyed (1.5% Evan's Blue) DI water, and then interfaced the assembly with the nanoinjector (Figure 5a, expanded view) for control of both the MNA position and fluidic microinjection dynamics. Although the results for the cyclic microfluidic burst-pressure experiments performed in vitro (Figure 4b–d) suggested that the MNA-capillary interface should withstand the forces associated with microinjections into the brain tissue, we optically monitored the overall MNA-capillary assembly during the microinjection process for potential signs of undesired leakage via the interface. Akin to the tissue penetration and retraction studies, we used the stereotaxic frame to guide the descent of the MNA into the brain tissue (Figure 5d, top and Movie S8, Supporting Information). Following completion of the penetration process, we then used the pneumatically controlled nanoinjector to dispense the surrogate dyed fluid through the MNA-capillary assembly and, in turn, deliver the fluid into the brain tissue. Thereafter, we retracted the MNA from the brain (Figure 5d, bottom and Movie S8, Supporting Information), and then washed the surface of the injection site with phosphate buffered saline (PBS) to eliminate any residual surrogate fluid from the surface, such that the only remaining fluid was located beneath the tissue surface (Figure 5e). Throughout the microinjection process, we did not observe any undesired leakage phenomena (Movie S8, Supporting Information), with optical characterizations of the postinjection site indicating effective, distributed MNA-mediated delivery of the surrogate fluid well below the surface of the excised brain (Figure 5e). Furthermore, SEM images of the MNA-capillary assembly following tissue penetration, fluidic microinjection, and retraction revealed uncompromised structural integrity (Figure 5f).

Lastly, we evaluated the microinjection performance of the esDLW-printed MNA compared to a conventional needle (Hamilton 33G) widely used for delivering therapeutics into brain tissue.[74] In this case, we used a suspension of fluorescently labeled nanoparticles (100 nm in diameter) as the surrogate microfluidic payload. As an initial positive experimental control for the esDLW-printed MNA, we performed microinjections (n = 3 MNAs) of the nanoparticle suspension into 0.6% agarose gel in vitro (Figure 6a and Movie S9, Supporting Information) and visualized the particle distributions using two-photon (Figure 6b S10) and widefield fluorescence microscopy (Figure 6c). We observed injected nanoparticles corresponding to each microneedle in the array—which included one microneedle in the center of the array, six needles arrayed radially in a middle region (150 µm from the center), and six needles arrayed radially in an outer region (260 µm from the center)—but to determine if microneedle array position influenced injection behavior, we analyzed the fluorescence intensities associated with each arrayed needle. Quantified results revealed that the fluorescence intensities were statistically indistinguishable, with no discernable difference for the microneedle injection sites between the center and either the middle (p = 0.66) or outer regions (p = 0.61), nor between the middle and outer regions (p = 0.72) (Figure 6d). Thereafter, we performed microinjections of the nanoparticle suspension into excised mouse brains using both the conventional needle and the esDLW-printed MNA (Figure 6e and Movie S10, Supporting Information). Two-photon fluorescence images of the injection sites revealed stark differences in the nanoparticle distributions associated with each needle system. In the conventional needle case, the nanoparticles accumulated tightly within the single needle track (Figure 6f,g). For example, quantified fluorescence intensity results revealed that the majority of the fluorescence signal was detected within an ≈150 µm region (Figure 6h). In contrast, MNA-associated microinjection sites exhibited a more homogeneous distribution of injected nanoparticles over a larger area (Figure 6i,j)—with particles detected at sites corresponding to each arrayed microneedle—resulting in a more consistent fluorescence signal along the length of the injection site (Figure 6k). These results suggest that MNAs offer an effective means to distribute fluidic payloads more uniformly over a larger area compared to conventional single-needle systems. In combination, these experimental results for MNA penetration, surrogate fluid/suspension delivery, and retraction functionalities using an ex vivo mouse brain provide an important foundation for the utility of the presented hybrid DLP-DLW-enabled MNAs for microinjection applications.

Experimental results for microinjections of fluorescent nanoparticles a–d) in vitro in 0.6% agarose gels and e–k) ex vivo using excised mouse brains. a) Sequential images of nanoparticle microinjection and retraction. Scale bar = 250 µm (see Movie S9 in the Supporting Information). b,c) Fluorescence images of the postinjection site captured using b) two-photon and c) widefield fluorescence microscopy. Scale bars = 250 µm. d) Mean fluorescence intensities of injection sites corresponding to microneedles in distinct array regions (n = 3 MNAs). Error bars = S.D. e) Sequential images of a representative MNA penetration, microinjection, and retraction process for a suspension of fluorescent nanoparticles injected into brain tissue. Scale bar = 1 mm (see Movie S10 in the Supporting Information). f–k) Postinjection results for fluorescent nanoparticles delivered via f–h) a conventional Hamilton 33G needle, and i–k) an esDLW-printed MNA. f,g,i,j) Fluorescence images of the postinjection site captured using two-photon fluorescence microscopy visualized in f,i) side and g,j) cross-sectional views. Scale bars = 250 µm. h,k) Quantified fluorescence intensities along the length of the corresponding cross-sectional views of the postinjection sites.

3.Conclusion

Microneedle-based microinjection protocols are essential to wide-ranging fundamental research and clinical applications across biological and biomedical fields, with MNAs providing numerous benefits over their single-needle counterparts in many scenarios.[75-77] Unfortunately, manufacturing-associated limitations have heretofore impeded researchers from leveraging the potential benefits of high-density MNAs comprising hollow, high-aspect-ratio microneedles at small length scales.[78-80] In this work, we introduced the concept of using esDLW to 3D print MNAs directly atop DLP-printed capillaries in batch arrays and demonstrated this approach by fabricating arrays of 50 µm OD, 30 µm ID, 550 µm tall hollow microneedles with 100 µm needle-to-needle spacing. Because the presented strategy is founded on two additive manufacturing technologies, the inherent geometric versatility can be harnessed to tailor both the DLP-printed capillaries and the esDLW-based MNAs to target experimental setups and applications. For the DLP-printed capillary, the shape and size need not be uniform along the length of the capillary as is the predominant case for conventional and/or commercially available fluidic capillaries. Here, for instance, we designed the OD of the base of the capillary to yield facile, direct integration with the nanoinjector, thereby circumventing the need for additional fluidic adapters or sealants. Similarly, although the presented design for the esDLW-printed MNAs included identical microneedles with dimensions based on a specific exemplar—i.e., fluidic microinjection into the cerebral cortex of a mouse brain—the high architectural control and submicrometer-scale resolution of DLW can be leveraged to customize the size, shape, and position of each individual microneedle in an array as desired (Figure S4, Supporting Information). For example, future efforts could increase the microneedle heights substantially to target different regions of the brain and/or additional animal models. Conversely, while this work centered on printing hollow microneedles (with 30 µm IDs) to support fluidic delivery operations, given the recent developments for the utility of solid MNAs in other cases, the presented strategy could also be extended to print MNAs composed of solid microneedles, such as those fabricated using DLW-compatible biodegradable materials,[81, 82] or potentially hybrid MNAs that comprise both hollow and solid microneedles.

The presented strategy also provides an important foundation for future academic and industrial translation through four pathways. First, in contrast to prior esDLW efforts, DLP-printing of the batch arrays of fluidic capillaries allows for facile loading into the DLW 3D printer, obviating the need for custom-built capillary holders as well as the time- and labor-intensive protocols required to manually load each individual capillary into such holders. Furthermore, because each capillary is printed in a designated array position with specified orientations, the setup for initiation of the esDLW-printing process is minimized, which could provide a promising avenue to scalable and automated production. Second, although we employed a layer-by-layer DLP printer to manufacture the batch arrays of fluidic capillaries, numerous vat photopolymerization approaches could be used instead to increase production speed, including continuous liquid interface production to print parts in minutes[83] and various volumetric 3D printing strategies to fabricate parts in tens of seconds.[84-86] Third, for esDLW-based printing of the MNAs, while the voxel size remained constant throughout the printing process with a scan speed of ≈120 mm s−1, future efforts can harness recent advancements for state-of-the-art DLW printers that can not only dynamically tailor the size of the voxel to target features but also allow for scan speeds up to 1,250 mm s−1 (e.g., with 5× objective lens configurations) in order to dramatically enhance print efficiency and speed. Lastly, recent improvements in the available build area for commercial DLW printers could be extended to print multiple MNAs simultaneously in a single pass (in contrast to the serial MNA printing strategy reported here), which would further increase the attainable production volume.

The numerical and experimental mechanical characterizations of the esDLW-printed MNA suggest that, in addition to brain tissue, the MNA described in this work could be used to facilitate microinjections for a wide range of additional biological tissues, including those associated with the liver, breast, heart, kidney, veins, arteries, and prostate.[67-70] For future efforts based on different injection targets with higher stiffness (e.g., E > 250 kPa), however, the inability of the presented MNA to successfully penetrate into the 10% agarose gel indicates that, for the current design, alternative photomaterials should be used for esDLW-based printing. In particular, researchers have reported DLW-compatible fused silica glass-based photomaterials,[87] which are now available commercially and would provide an order of magnitude increase in E of the fabricated MNAs. Alternatively, while we designed each microneedle with 10-µm-thick walls and 50 µm ODs, both dimensions could be readily increased to improve the mechanical strength. For excised mouse brains specifically, the ex vivo investigations in the current study revealed effective MNA-mediated penetration, microinjection, and retraction operations without any instances of microneedle-associated mechanical failures (e.g., buckling or fracture). In addition, throughout both in vitro microfluidic cyclic burst-pressure characterizations (with applied pressures in excess of 250 kPa) and ex vivo brain tissue experiments, the MNA-capillary interface exhibited consistent fluidic integrity, without any signs of undesired leakage phenomena or MNA detachment from the capillary.

We envision that future efforts could extend the methodology reported here to achieve novel MNA designs that remediate the deficits of single-needle injection strategies by expanding the delivery range via simultaneous, distributed microinjection. For example, as both in vitro and ex vivo experiments for MNA-mediated microinjections of nanoparticle suspensions revealed homogeneous distributions of implanted particles, such capabilities could offer new means to address the cell crowding challenges of SCT associated with single-needle delivery systems that contribute to low cell viability and, thus, limited therapeutic efficacy.[88-91] Such a pathway to improved SCT could hold distinctive promise for treating a diversity of medical conditions and neurodegenerative diseases, but further studies are needed to explore the potential for MNAs at this scale to enhance therapies that rely on fluidic microinjections—not only for stem cells, but also additional therapeutic payloads (e.g., growth factors and viruses for gene therapy)—into the brain. Nonetheless, given the vast diversity of scientific and clinical applications that are founded on microinjections and/or microneedles, the presented hybrid additive manufacturing strategy offers unique potential as an enabling technology for realizing entirely new classes of MNAs to advance scientific discovery and promote human health and well-being.

4.Experimental Section

Batch Capillary Array Fabrication via DLP 3D Printing

The computer-aided design (CAD) software, SolidWorks (Dassault Systèmes, France), was used to generate models of batch arrays of capillaries (Figure S1, Supporting Information). Models were exported as STL files and then imported into the CAM (slicer) software for the Miicraft M50 DLP 3D printer (CADworks3D, Canada) to define the print parameter settings (Table S1, Supporting Information). The batch capillary arrays were DLP-printed using Clear Microfluidics Resin V7.0a (CAdworks3D) with the layer height set to 50 µm. Following the DLP printing process, the build plate was removed and the prints were manually detached from the build plate using a razor blade. The prints were developed in methanol for ≈10 s and then methanol was perfused through each capillary to eliminate any residual resin from the interiors. After one additional rinse with methanol, the prints were washed with 90% isopropyl alcohol (IPA). The prints were then dried with pressurized air and postcured under UV light for 20 s (flipping the device after 10 s to cure both sides equally).

MNA Fabrication Atop the Capillaries via esDLW

The microneedle arrays—modeled using SolidWorks (Dassault Systèmes)—were designed with identical needles (ID = 30 µm; OD = 50 µm; height = 550 µm) and arrayed with 100 µm needle-to-needle spacing (Figure S3, Supporting Information). MNA models were exported as STL files and then imported into the CAM software, DeScribe (Nanoscribe), to define the print parameter settings (Table S2, Supporting Information), which included a hatching distance of 800 nm and a layer height of 2.5 µm. Initially, IP-Q photoresist (Nanoscribe) was dispensed directly atop the DLP-printed capillaries and the batch was then loaded into the Nanoscribe Photonic Professional GT2 DLW 3D printer (Figure S2, Supporting Information). For esDLW printing, the dip-in laser lithography (DiLL) mode was used with a 10× objective lens, a laser power of 27.5 mW, and a laser scanning speed of 120 000 µm s−1. The printing process was initiated with 50 µm of overlap with the top capillary surfaces. Following the esDLW process, the batch assembly (with MNAs printed atop the capillaries) was removed from the DLW printer for development. The prints were developed using propylene glycol monomethyl ether acetate (PGMEA) for 30 min and IPA for 5 min, and then dried using a gentle stream of N2 gas. Individual MNA-capillary assemblies were removed from the batch by manually severing the five connecting structures arrayed radially around each capillary (Movie S3, Supporting Information).

Finite Element Analysis (FEA)

Numerical simulations of the MNA compression test were performed using the commercially available software, ABAQUS/Standard (Abaqus Inc., Palo Alto, CA). Initially, the complete 3D CAD model of the MNA (i.e., including both the base and needles) was imported into the FEA software, and then the distinct material properties were set. Specifically, the MNA was modeled as a linear elastic homogeneous material (E = 250 MPa; ν = 0.49). The mesh was constructed using four-node, linear, 3D-stress-tetrahedra elements (ABAQUS element type C3D4H), and the accuracy was verified by mesh convergence. During all studies, the circular bottom surface orthogonal to the loading direction was modeled to be perfectly fixed. A static analysis (*STATIC step with NLGEOM = ON in ABAQUS) was conducted to characterize the nonlinear response and loaded the structure by linearly increasing the applied tip force. To characterize the nonlinear response at the interface between the needle tips and the brain substrate, the bottom surface of the cylinder mimicking the brain sample was modeled to be fully clamped while a displacement was applied to the MNA's cylindrical base. A surface-to-surface contact was defined between the brain substrate and the MNA needle tips. Both tangential and normal contact behaviors were defined. The MNA was modeled as a linear elastic homogeneous material, while the brain substrate was modeled as a hyperelastic Neo-Hookean material. To characterize the nonlinear response at the interface between the needle tips and the brain substrate, a dynamic implicit analysis (*DYNAMIC step with NLGEOM = ON in ABAQUS) was conducted.

MNA Mechanical Characterization

Mechanical testing on the MNAs was conducted using a Q800 Dynamic Mechanical Analysis (DMA) system (TA Instruments, New Castle, DE) equipped with a compression clamp. Samples were compressed at a rate of 0.1 N min−1 until the failure was confirmed via optical microscopy. Values for E and σy of MNAs were calculated from the linear region of the resulting stress-strain curve. To evaluate the puncture ability of the MNAs, hydrogels with different stiffness were prepared by mixing agarose gel powder in 1% PBS (Sigma-Aldrich, Saint Louis, MO) at four different concentration levels: 1.2%, 2.4%, 5%, and 10%. The solutions were heated to a boiling temperature and then cooled down until the hydrogels were set at room temperature. Before each MNA puncture, the top surface of the hydrogel was rinsed with PBS. The MNA was mounted on a stereotaxic manipulator, slowly inserted into the hydrogel samples, and optically monitored for any signs of failure.

Ex Vivo Mouse Brain Extraction and Experimentation

Brain tissues excised from 6-month-old male mice (Wild-type C57BL/6 J, Jackson Laboratory) were used for all ex vivo experiments. Each brain with an intact dura mater was excised within 10 min of euthanasia and stored in cold PBS on ice prior to testing. To maintain tissue integrity, the tissue samples were handled gently before and during the experiment. Each MNA-capillary assembly was interfaced with a custom-built nanoinjector (Narishige) and mounted on a stereotax with a digital display (#68807, RWD, China) to control the displacement and perform microinjections. In separate experiments, blue-dyed water and green fluorescent nanoparticles (505/515, 100 nm diameter, #F8803, Thermofisher) diluted with PBS were injected into the freshly dissected mouse cerebral cortex (or agarose gel) using either an MNA-capillary assembly connected to a micromanipulator (#MO10, Narishige) or a Hamilton syringe with a 33G needle connected to a motorized pump (#78–8130, KD Scientific, Holliston, MA). The injection depth was 500 µm with an extra 200 µm overshoot. The injection duration was ≈2 min for both MNA and Hamilton syringe-mediated injections. After injection with fluorescent nanoparticles, the fresh mouse brains were fixed with 4% paraformaldehyde for 2 d, rinsed, and mounted on glass slides for imaging under a two-photon microscope. These studies were performed in accordance with the National Institutes of Health (NIH) Guide for Care and Use of Laboratory Animals and the University of Maryland, School of Medicine, Animal Care and Use Committee.

Optical Characterizations

SEM images were captured using a TM4000 Tabletop SEM (Hitachi, Tokyo, Japan) under low vacuum, which allowed for imaging of uncoated samples. The mechanical tests were recorded using a Monocular Max 300× microscope objective and a 41MP USB C-Mount Industry Microscope Camera Set (Hayear Electronics Co. Ltd., Shenzhen, China). Brightfield microscopy during microfluidic testing was performed using an inverted microscope (Motic AE31, Motic, Canada) connected to a CCD camera (Moticam Pro 285B, Motic). For ex vivo microinjection experiments, the injection process was recorded using the Monocular microscope while the fluorescent images of the top view of the gel injection site were captured using a DMi8 automated fluorescence microscope (Leica Microsystems, Wetzlar, Germany). The 3D stack images of the injection sites were acquired using the Modular In Vivo Multiphoton Microscopy System designed by Janelia Research Campus, Howard Hughes Medical Institute. A 900 nm laser (≈5 mW) was used for excitation of the green fluorescent nanoparticles. The 3D stacks from the top of the brain to the bottom of the needle track were acquired at a step size of 2 µm under a water-immersion 25× objective (numerical aperture of 1.05, Olympus). Fluorescence emission was collected by two GaAsP photomultiplier tubes after being split by a dichroic mirror (560 nm, T560pxrxt, Chroma) with an emission filter green (510/84 nm, 84–097, Edmund) fluorescence. A similar acquisition setting was used for imaging the needle tracks in hydrogels injected with the fluorescent nanoparticles. Fluorescence images were processed and visualized with ImageJ (NIH, Bethesda, MD). BigDataViewer was used to adjust the tilting angle of the 3D stack for optimized visualization. For comparisons of needle-to-needle injection sites within the MNAs as well as injection distributions between the MNA and Hamilton injections, ImageJ was used to quantify the fluorescence intensities.

Statistical Analysis

Statistical significance was quantified via unpaired Student's t-tests, with two-tailed p values greater than 0.05 considered statistically indistinguishable. A minimum of three samples were used to quantify any means reported, with data presented in the text as mean ± standard deviation (S.D.).

Acknowledgements

The authors greatly appreciate the contributions of Olivia Young, Michael Restaino, and Chen-Yu Chen, as well as additional members of the Bioinspired Advanced Manufacturing (BAM) Laboratory and the William Bentley Laboratory. The authors appreciate the help and support of staff members at the University of Maryland Terrapin Works and the Micro/Nanofabrication Center at the Princeton Institute of Materials. This work was supported in part by the Maryland Robotics Center, the Center for Engineering Concepts Development (CECD), U.S. NIH Award Numbers 1R01EB033354-01, 1R03NS123733-01, 1R21AG077631-01, 1R03NS128459-01, 1R01EB019963, and F31DK129021, the Maryland Stem Cell Research Fund 2022-MSCRFL-5893, and U.S. National Science Foundation (NSF) Award Number 1943356. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH. Correction added on 10th March 2023 after initial publication: The affiliation for P. Walczak was corrected.]

Conflict of Interest

The authors declare no conflict of interest.

3D-printed capillaric ELISA-on-a-chip with aliquoting

3D-printed capillaric ELISA-on-a-chip with aliquoting

Azim Parandakh, Oriol Ymbern,  William Jogia, Johan Renault, Andy Ng and  David Juncker

Sandwich immunoassays such as the enzyme-linked immunosorbent assay (ELISA) have been miniaturized and performed in a lab-on-a-chip format, but the execution of the multiple assay steps typically requires a computer or complex peripherals. Recently, an ELISA for detecting antibodies was encoded structurally in a chip thanks to the microfluidic chain reaction (Yafia et al. Nature, 2022, 605, 464–469), but the need for precise pipetting and intolerance to commonly used surfactant concentrations limit the potential for broader adoption. Here, we introduce the ELISA-on-a-chip with aliquoting functionality that simplifies chip loading and pipetting, accommodates higher surfactant concentrations, includes barrier channels that delay the contact between solutions and prevent undesired mixing, and that executed a quantitative, high-sensitivity assay for the SARS-CoV-2 nucleocapsid protein in 4×-diluted saliva. Upon loading the chip using disposable pipettes, capillary flow draws each reagent and the sample into a separate volumetric measuring reservoir for detection antibody (70 μL), enzyme conjugate (50 μL), substrate (80 μL), and sample (210 μL), and splits washing buffer into 4 different reservoirs of 40, 40, 60, and 20 μL. The excess volume is autonomously drained via a structurally encoded capillaric aliquoting circuit, creating aliquots with an accuracy of >93%. Next, the user click-connects the assay module, comprising a nitrocellulose membrane with immobilized capture antibodies and a capillary pump, to the chip which triggers the step-by-step, timed flow of all aliquoted solutions to complete the assay in 1.5 h. A colored precipitate forming a line on a nitrocellulose strip serves as an assay readout, and upon digitization, yielded a binding curve with a limit of detection of 54 and 91 pg mL−1 for buffer and diluted saliva respectively, vastly outperforming rapid tests. The ELISA chip is 3D-printed, modular, adaptable to other targets and assays, and could be used to automate ELISA in the lab; or as a diagnostic test at the point of care with the convenience and form factor of rapid tests while preserving the protocol and performance of central laboratory ELISA.

We kindly thank the researchers at McGill University for this collaboration, and for sharing the results obtained with their system.

Introduction

The enzyme-linked immunosorbent assay (ELISA) is utilized for the detection and quantification of proteins, antibodies, or antigens. The sandwich format with a capture antibody immobilized on the surface and a detection antibody applied in solution is used for assays requiring high sensitivity and specificity. Laboratory microplate ELISA still serves as a gold standard for assays and benefits from high sensitivity thanks to enzymatic signal amplification (down to sub-picomolar concentration for the best antibody pairs), quantitative results, standardized operations, off-the-shelf consumables, and a comparably high throughput thanks to the use of 96-well plates. Long incubation times and copious washing between different steps to reduce non-specific binding and assay background are critical to achieving high assay sensitivity. However, the ELISA suffers from several downsides, such as being laborious, lengthy (∼2–12 h depending on the protocol), requiring precise timing for each step, dependence on technical skills notably for adding and removing reagents (and thus susceptible to inter-operator variation) and necessitating a plate reader for signal readout.1,2

The miniaturization of ELISA has proceeded thanks to microfluidic lab-on-a-chip systems that can also automate the protocol.3,4 Microfluidics successfully reduced the consumption of reagents and the total assay time while preserving assay performance. However, whereas the chips are small, they rely on bulky peripherals such as syringe pumps3 or control motors,4 and a computer or an instrument for operation.5,6 Capillary phenomena and gravity have been harnessed to automate simple liquid manipulation, reducing or obviating the need for an external/active power supply.7–9 For instance, a disk-like microfluidic platform (powered by a combination of centrifugal and capillary forces)8 and a microfluidic siphon platform (powered by gravitational forces)9 have been developed to carry out the common steps of a conventional ELISA with reduced reagents consumption and assay time while preserving assay sensitivity. Yet, both examples require multiple precise pipetting steps and timed user interventions for operation.

Sandwich assays can also be performed at point-of-care using so-called lateral flow assays (LFAs), also called rapid diagnostics, and are used globally for pregnancy tests and COVID-19 (coronavirus disease 2019) diagnosis. LFAs replace the enzyme amplification with conjugated colorimetric particles (either gold nanoparticles or polystyrene beads) that become visible to the naked eye upon accumulation. LFAs are simple to use as they only require the application of the sample, which flows thanks to capillarity without the need for peripherals, and produce a test result within a few minutes. However, LFAs offer only qualitative yes-no results, their sensitivity is typically lower compared to that of laboratory ELISA and are not suitable for archival as the readout must be completed within a few minutes of the test, because otherwise, the result can be compromised.10–12 Enzymatic amplification has been implemented in the LFAs,13,14 for instance by using a microfluidic interface,14 to improve sensitivity. Yet, they cannot implement various fluidic handling tasks of common ELISA such as timed incubation of reagents and multiple rinsing steps between each incubation interval.

Paper-based microfluidics has been developed to introduce more advanced fluidic functions such as sequential delivery, additional rinsing step, and enzymatic amplification that collectively help improve assay sensitivity compared to LFAs.15–17 Sponge actuators that upon swelling connect or disconnect different parts of a paper-based microfluidic circuit, along with flow paths with different lengths and resistance, have been used to time the delivery of multiple reagents for completing a bona fide ELISA.17 However, these systems lacked the intermediate washing steps characteristic of classical ELISA, and undesired mixing of consecutive reagents occur at their mutual interface. Enzyme–substrate mixing may limit the potential for higher sensitivity as it could contribute to non-specific signal amplification.

Capillaric circuits (CCs) are capillary microfluidics in microchannels designed and built using capillaric elements which can automate liquid handling operations by pre-programming them structurally using capillary phenomena and powering them by capillary flow, without the need for peripheral equipment.18,19 Multiple CCs have been designed to perform and automate ELISA with new functionality including flow reversal,18 timing, reagent lyophilization,20 and portable readers,21,22 but they skip intermediate washing steps. Aliquoting a single solution into multiple reservoirs has been shown for a nucleic acid test.23 For an ELISA in a CC, multiple solutions including sample, buffer, and reagents must be serially delivered, and fluidically connected to effect fluid flow by hydraulic transmission of pressure differentials, which would subject them to unwanted mixing due to diffusion and/or convection, negatively affecting assay sensitivity and reliability.

The microfluidic chain reaction (MCR) introduces conditional initiation of capillary flow events, whereby event n is triggered after the preceding event n − 1 has been completed, and completion of n, in turn, initiates event n + 1; the condition is encoded using the so-called capillary domino valves.24 MCR can drive hundreds of sequential flow operations robustly, thus opening new opportunities for CCs. CCs and MCRs are susceptible to failure in the presence of surfactants that reduce surface tension and contact angles. Yet surfactants are essential ingredients to assays, and often 0.05% Tween 20 is used to prevent non-specific binding.25 In the initial MCR demonstration,24 only 0.0125% was used because higher concentrations led to corner flow and trapping of air bubbles19 and failure of stop valves. The initial MCR was used for a SARS-CoV-2 (severe acute respiratory syndrome coronavirus 2) antibody assay that could help determine the immune or vaccination status or detect prior infection, but which does not typically require high sensitivity needed for antigen assays where insufficient assay sensitivity often limits clinical usefulness.

User-friendliness is a critical feature for a device to be used at the point of need,26–28 and accurate aliquoting and volumetric consistency are essential to reliable immunoassays.29,30 Whereas in the lab they can be met using precision pipettes operated by technicians, they are difficult to achieve in a point-of-care setting. Both ELISA and the MCR CC introduced previously are dependent on precision pipetting. In particular, for the MCR chips,24 reagent loading occurred via miniature inlets that require precise positioning of the pipette, precise volumetric control and hence precision laboratory pipettes, and controlled pipetting rates to match the capillary flow rate of the reservoir being filled. LFAs for COVID-19 often include a dropper and instructions for delivering a precise number of droplets.31 Droppers are prone to occasional miscounting,31 while for more complex assays multiple solutions with different volumes are required that could not be serviced using droppers to be supplied and aliquoted with different volumes.

Here, we introduce the ELISA chip that automates ELISA protocol on a chip using an MCR CC while preserving the washing steps used in classical ELISA. The ELISA chip eases the pipetting procedure thanks to the automated aliquoting of solutions and can accommodate higher surfactant concentrations commonly used in immunoassays. Akin to measuring spoons that are used to size ingredients in cooking, measuring reservoirs with different volumetric capacities are used to aliquot reagents, buffer, and sample. Upon loading, solutions spontaneously fill their respective measuring reservoir, while an integrated capillaric aliquoting circuit (CAC) autonomously drains excess liquid from all reservoirs simultaneously, forming precise aliquots. We describe the capillaric circuitry and its components, and the step-by-step automated capillary flow operations of both aliquoting of all solutions and the sequential MCR-controlled ELISA protocol. We characterize the volumetric accuracy of aliquoting, the timing precision of delivery of the reagents for the ELISA, and the performance and limit of detection (LOD) of an assay for the detection of the SARS-CoV-2 nucleocapsid (N) protein spiked in buffer and 4×-diluted human saliva.

Materials

Results and discussion

ELISA chip

We designed a capillaric ELISA chip without a moving part that automates aliquoting of sample, reagents, and buffer, and autonomously executes an ELISA protocol by flowing eight solutions in sequence according to predetermined flow rates using an MCR (Fig. 1A and S1†). The solutions are delivered for the ELISA in the order of (i) sample, (ii) buffer, (iii) biotinylated detection antibody, (iv) buffer, (v) streptavidin poly-HRP (horseradish peroxidase), (vi) buffer, (vii) colorimetric substrate DAB (3,3′-diaminobenzidine) and (viii) buffer. The ELISA was developed for measuring the SARS-CoV-2 N protein in natural saliva and includes a strip of nitrocellulose membrane with a positive control line and a test line that will produce a permanent colorimetric signal proportional to the concentration of the N protein.

Fig. 1 3D-printed ELISA chip with aliquoting for SARS-CoV-2 N protein assay in saliva. (A) Picture of the chip with superposed process flow graphics. (i1) Reagents (R1–3) and (i2) washing buffer (W) are added to the inlets using disposable, low-precision pipettes shown in (B) and fill the measuring reservoirs by capillary flow; the washing buffer is split into four reservoirs automatically (W1–4). (i3) Addition of the sample brings the drainage capillary pump to a fluidic connection, initiating drainage of excess solutions from all inlets via the capillaric aliquoting circuit (CAC, see C), and producing (ii) final aliquots schematized by graduated tubes. (iii) Connection of the nitrocellulose strip triggers the ELISA via the sequential flow of sample, reagents, and washing buffer through the chip according to the propagation of the MCR. (iv) Following the enzymatic conversion of the substrate into a permanent line of a brown precipitate, the result can be read out by eye, and quantified following digitization. (B) Picture of a disposable pipette used to load the chip. (C) 3D design view of the ELISA chip with the structurally encoded capillaric elements without moving parts. The CAC is highlighted in yellow, and insets showcase (i) inlet barrier channel and supply/drainage conduit and air vent, (ii) reservoir outlet, outlet barrier channel, main assay conduit, and air vent (iii) sample inlet and reservoir, main drainage conduit and connection to supply/drainage conduit, and drainage capillary pump.

The need for the use of laboratory micropipettes is circumvented by integrating CAC into the chip which accepts the delivery of larger-than-needed volumes into the adequately sized inlets, followed by the spontaneous flow of the solutions via capillarity into serpentine measuring reservoirs for each solution. To complete the aliquoting process, the CAC removes excess volumes into a drainage capillary pump. The volume of each reservoir was set following assay optimization.

Simple squeeze pipettes, shown in Fig. 1B, can be used to load the chip while visually monitoring filling progression, and the measuring reservoirs form the aliquots (symbolized by graduated tubes) after draining excess liquid. The ELISA chip has five inlets servicing the eight reservoirs, three for the reagents, one for the buffer, and one for the sample, which need to be added in this order. The buffer is automatically split into four separate measuring reservoirs on the chip. The addition of the sample simultaneously fills the sample reservoir and initiates the drainage of excess volumes via the CAC. Next, upon click-connection of the main capillary pump, the step-by-step execution of the ELISA protocol is triggered, and the eight reservoirs chained by capillary domino valves are drained one by one as the MCR progresses. Finally, the signal is read out via the naked eye or using a scanner for quantification. A fluidic flow chart to explain the step-by-step manual user interventions and the ELISA chip's autonomous functions is provided in Fig. S2.

The ELISA chip is made by additive manufacturing using an off-the-shelf 3D printer and light-based photopolymerization, followed by a series of post-processing steps for a total time of ∼1 h (Fig. 2; see Experimental section for additional details).

Fig. 2 Additive manufacturing of ELISA chip and post-processing steps. The manufacturing includes (A) ELISA chip design in computer-aided design software, (B) digital light processing-based 3D printing, (C) rinsing with isopropanol (IPA) to remove uncured resin, (D) drying under a stream of pressurized nitrogen gas (N2), (E) 1 min UV curing, (F) 10 s air plasma treatment and (G) sealing with a microfluidic tape with cut-out openings for servicing sample, buffer, and reagent inlets.

Step-by-step loading and aliquoting

The ELISA chip loading and aliquoting are shown in Video S1.† Using a low-precision pipette (Fig. 1B), the user first deposits each reagent in the respective inlet in an arbitrary order with an excess of solution. The three serpentine reservoirs for the detection antibody, streptavidin poly-HRP, and DAB solutions have nominal measuring volumes of 70, 50, and 80 μL, respectively. Fig. 3A and B show the loading of the detection antibody and are representative of the other reagents. Capillary flow fills the reservoir up to the trigger valve at the empty outlet barrier conduit, while excess solution remains in the inlet and fills the trigger valve connecting to the conduit at the inlet (called inlet barrier channel) that is part of the CAC (see Fig. 1C). Next, the washing buffer is delivered to the common inlet to the right of the ELISA chip and flows by capillarity through the supply/drainage conduit into the four washing buffer measuring reservoirs with a nominal volume of 40, 40, 60, and 20 μL (Fig. 3C). The buffer simultaneously fills the main assay conduit, which is supplied by the vertical channel adjacent to the buffer inlet. The buffer distribution and splitting simplify chip loading as only one buffer solution needs to be added. However, the chip design imposes contact between buffer and reagents both at the inlet and outlet of the reagent reservoirs which could lead to unwanted mixing by diffusion and convection between various solutions.

Fig. 3 ELISA chip loading, filling into measuring reservoirs, and drainage of excess liquid by the CAC. Most figures show the part of the chip outlined by the dashed rectangle in Fig. 1C. (A and B) Loading of the detection antibody reagent into the inlet followed by capillary flow into the measuring reservoir and stop of the liquid at the capillary stop valves located next to the inlet and at the outlet. Loading of the other two reagents, i.e., streptavidin poly-HRP and DAB, follows the same procedure. (C and D) Loading of buffer in buffer inlet and splitting into the buffer reservoirs supplied by the supply/drainage conduit. (E–H) Sample loading with the filling of the sample reservoir and simultaneous triggering of the drainage of all excess solutions on the chip via the CAC. See Video S1† and text for further details.

Suppressing unwanted mixing and bubble trapping

We isolated each of the reagents measuring reservoirs to prevent unwanted mixing between them and/or the sample prior to assay completion. We equipped them with two diffusion barrier channels located upstream (at the inlet) and downstream (at the outlet) of each reagent reservoir. By isolation, we suppress undesired preliminary mixing of reagents that could affect assay performance and reproducibility during both aliquoting (i.e., filling and excess volume drainage), and subsequent ELISA sequential delivery (e.g. premature reaction between enzyme and colorimetric substrate DAB could lead to unwanted background signal). Because diffusion will occur for a long time at the outlet of each reservoir, and for almost the entire assay duration particularly for the DAB solution, the last reagent to flow in the ELISA protocol, a long diffusion barrier is needed. By implementing diffusion barrier channels, we increase the distance between ELISA reagents to prevent premixing that could occur by both diffusion and/or convection (e.g. small convective flows due to evaporation). The outlet barrier channel was designed with a U-turn and a separation length between the outlet of the reservoir and the main assay conduit with a length of 13–17 mm. The characteristic one-dimensional diffusion time is given as t = L2/D with L = length, D = diffusion constant, and t = time.32 For an IgG antibody with D ≅ 36 μm2 s−1 and DAB with D ≅ 570 μm2 s−1 (calculated by the Stokes–Einstein law for diffusion),32 the time for diffusion across 10 mm is ∼16 days and ∼1 day, respectively. In our case, the functional connection connecting the reservoir and the barrier forms a constriction,24 hence the diffusion time would be further increased relative to the calculation. The diffusion barriers thus ensure that no unwanted mixing occurs among reagents during the ELISA.

The inlet and outlet barrier channels form a separation between each reagent reservoir and the supply/drainage conduit (upstream), and main assay conduit (downstream) respectively. Both barrier channels remain empty during the autonomous filling of the reagents due to the action of trigger valves (see Fig. 3A), and autonomously fill by buffer after buffer injection (Fig. 3D). The inlet of the inlet (upstream) barrier conduit is located to the left of the reagent reservoir and branches out from the serpentine buffer reservoir, close to the outlet of the buffer reservoir. Upon reaching the branching point, capillary flow filling the buffer reservoir immediately branches into the barrier channel and fills it up to the extremity, which forms a dead-end. To prevent bubble trapping, an air vent is included that is connected to the channel via four stop valves, including three positioned upstream of the dead-end. At the reservoir outlet, the outlet barrier channel, which forms an extension to the buffer reservoir, is similarly filled by the buffer. However, the buffer reservoir to the right of the reagent channel supplies the corresponding outlet barrier channel and fills only after the buffer reservoir is filled completely (Fig. 3D bottom close-up, Video S1†). Akin to the inlet barrier channel, the extremity of the outlet barrier channels also forms a dead-end and is connected by three stop valves to a venting opening. Note that the design ensures that only pristine buffer fills each of the buffer measuring reservoirs. The inlet and outlet barrier channels are empty as the buffer starts flowing, and hence there is no contact or mixing between the reagents and the buffer flowing into the reservoirs.

The provision of multiple venting connections/stop valves is needed to accommodate liquids with low surface tension such as buffer containing a surfactant that is used in ELISA (e.g., Tween 20) and other assays. Low surface tension liquids have a lower contact angle than water (which has a high surface tension), which induces corner flow preceding the main filling front. The corner flow reaches the extremity of the conduit before the main filling front and can thus clog small air vents located at the extremity of dead-end channels before all the air can escape (see Fig. S3†). Upstream connections to the vent allow air to escape even after corner flow reaches the dead-end and here prevent air bubbles from being trapped. For the ELISA chip, we evaluated the chip fluidic functionality under Tween 20 concentrations of 0.0125%, 0.025%, 0.05%, and 0.1%, and because of occasional failure at 0.1%, limited it to 0.05% for subsequent experiments.

Excess volume draining

Drainage of the excess volumes is initiated by supplying the sample to the sample inlet (Fig. 3F). The sample inlet is connected to both the sample measuring reservoir and the main drainage conduit, both filling simultaneously by capillary flow. The outlet of the sample reservoir connects to a now pre-filled outlet barrier channel supplemented with an air vent to permit complete filing. The sample flowing into the main drainage conduit also initiates drainage of the excess volumes by triggering a trigger valve connecting the filled supply/drainage conduit to the main drainage conduit, and subsequently flowing to the drainage capillary pump, all part of the CAC. Initially, most of the drainage flow is supplied by the sample owing to excess solution in the sample inlet, the absence of a retention capillary pressure, and lower flow resistance. Once the sample inlet is emptied to the point where only corners remain filled, the drainage flow rates from the inlets containing excess buffer and reagents increase. The inlet barrier channel, in addition to minimizing the diffusion between the main drainage conduit and each reservoir, also forms the fluidic connection through which the excess reagent is drained from each reagent inlet.

Flow path routing during drainage

Whereas conceptually simple, in CCs, drainage must both be triggered and stopped without external intervention. In addition, multiple possible drainage paths exist when the circuit is operating in the absence of active valves to close off sections of the circuit, yet to avoid unwanted mixing and drainage of solutions in the reservoirs, the liquid flow must flow through a pre-designed flow path, Fig. 4A. In the desired path, the excess reagents flow from the common inlet to the drainage capillary pump as follows: reagent inlet → inlet barrier channel → supply/drainage conduit → main drainage conduit → drainage capillary pump. To facilitate flow and favor this fluidic path while preserving other capillaric functionality, we included 4 parallel connection conduits (0.1 × 0.2 × 0.75 mm3) between the inlet barrier channel and the supply/drainage conduit (Fig. 1C close-up i), which yielded an overall flow resistance of 135 Pa s mm−3 for path i.

Fig. 4 Drainage of excess volumes via the CAC. (A) The flow path for draining excess volumes in sample, reagent, and buffer inlets is shown with the black arrows. (B) In addition to the desired flow path i, three alternative, parasitic paths ii, iii, and iv are schematized using colored arrows. Drainage of reagent via a parasitic flow path could lead to mixing with other solutions and jeopardize the function of the ELISA chip. The design minimizes parasitic flow by ensuring that their flow resistance is much higher because they are longer and if necessary, sections with a small cross-section (and therefore a high flow resistance) are included. Note that in panel B, all flow paths are shown for only one reagent, and the same exist for other reagents as well.

For each reagent inlet, three unwanted, leaky flow paths exist, as illustrated for the detection antibody in Fig. 4B. Flow through any of the parasitic paths should be disabled as they would result in unwanted reagents mixing and/or deviation from the pre-programmed volumes. Path ii proceeds from reagent inlet → inlet barrier channel → washing buffer reservoir → supply/drainage conduit, and path iii from reagent inlet → reagent reservoir → reagent outlet barrier channel → washing buffer reservoir → supply/drainage conduit to the main drainage conduit and drainage capillary pump. Both paths would result in reagents flowing into, and mixing with, the washing buffer in the adjacent reservoirs. To impede drainage through path ii, the resistance at the inlet of the inlet barrier channel as well as that of the corresponding washing buffer reservoir was increased, resulting in overall flow resistance of 250 Pa s mm−3 for path ii. Likewise, to impede drainage through path iii, the resistance at the inlet of the outlet barrier channel as well as that of the corresponding washing buffer reservoir was increased, thereby the overall resistance of path iii was 550 Pa s mm−3.

Path iv follows the reagent inlet → reagent reservoir → reagent outlet barrier channel → main assay conduit → sample outlet barrier channel → sample reservoir → sample inlet → main drainage conduit → capillary drainage pump (Fig. 4B, path iv), with an overall resistance of 700 Pa s mm−3. This drainage path would lead to mixing with the sample, yet the last part of it being in charge of the excess drainage of the sample. This path only exists until the point where the excess sample is drained to the drainage capillary pump through the main drainage conduit (with an overall resistance of 50 Pa s mm−3) and is disconnected from the sample reservoir as the inlet is emptied.

Here we discussed the overall flow resistance of each of the four paths, but the parasitic flow is governed by the difference between the point where a given parasitic path splits from the drainage path i to the point where it merges again, which is a shorter distance. The ratio of flow resistance for the split section between path i, and each of paths ii, iii, and iv was calculated to be 7.8, 8.4, and 6.9 times higher respectively (Table S1†). Path iv is only active for a short time, and thus only contributes marginally. This design hence ensures that most of the excess volume will flow through path i.

The automated ELISA protocol

The ELISA is initiated by the user click-connecting the main capillary pump to the chip (Fig. 5A), which wicks liquid from the CC and the reservoirs, and initiates the MCR that propagates from one reservoir to the next via the capillary domino valves that chain the reservoirs24 (Video S2†). Briefly, the first reservoir is the sample reservoir that is capped by a retention burst valve (RBV) with a nominal capillary/burst pressure of −200 Pa (i.e., it will burst when the negative sucking pressure in absolute value is >200 Pa), which quickly bursts and results in draining of the reservoir. Air drawn in from the outside replaces liquid, and once all liquid is displaced, the capillary domino valve that forms an air link to the next reservoir is now connected to the outside via the just-emptied reservoir. At the top of the buffer reservoir, there is another RBV (here with the nominal capillary pressure of −215 Pa), which in turn bursts, leading to the draining of the first buffer reservoir, and in turn, opening the air link to the next (reagent) reservoir, and so on. The sequence of flow is thus sample, buffer, detection antibody, buffer, streptavidin poly-HRP, buffer, DAB, and buffer (Fig. 5B–F). Drainage of the main assay conduit is not controlled via the MCR because the chip lacked space for an additional capillary domino valve. Instead, we implemented an additional RBV with a nominal burst pressure of −550 Pa. That is the RBV with the highest threshold within the CC, and hence the last one to burst. The ELISA chip illustrates the possibility of combining an MCR with an RBV to control the sequence of liquids flowing through an assay conduit. Upon draining the main assay conduit at the last step, the main capillary pump and therefore the nitrocellulose membrane, is fluidically disconnected from the microfluidic chip to complete the ELISA.

Fig. 5 MCR progression of the ELISA chip to complete ELISA steps. (A) Click-connecting the main capillary pump to the chip (the region indicated by the black circle and arrow) triggers (B–G) sequential delivery of the sample, reagents, and corresponding washes to the sensor. Black arrows indicate the direction of flow. The black rectangle in panel A shows the small serpentines implemented in the ELISA chip to avoid mixing the sample with other reagents due to evaporation-induced backflow during sample delivery.

ELISA chip over-loading capacity and overflow

We evaluated the volumetric operational window of the ELISA chip that preserves aliquoting and overall ELISA chip functionality. We analyzed the video of four ELISA chips loaded using precision pipettes with known volumes of sample, reagents, and buffer, determined the minimum volume required to fill the measuring reservoirs, tested a range of excess volumes, and verified flow functionality while also monitoring unwanted mixing.

The nominal aliquot volume corresponds to the capacity of the measuring reservoir. However, in practice, some additional volume is needed to account for dead volumes in the chip. Conversely, if the inlets are overfilled, leakage into other parts of the chip could occur. We verified the nominal aliquoting accuracy by mapping the levels of the measuring reservoirs filled with food dyes after aliquoting was completed, and calculating the volumetric error by image analysis (see Experimental and Fig. S4† for detailed explanation). For the sample, the nominal aliquot volume is 210 μL but at least 300 μL is required because the sample triggers the CAC (as explained above). As this ELISA chip is designed for saliva, which was collected in an amount of 1 mL per individual, the additional volume requirement could be accommodated easily. We verified that the chips preserved their functionality and nominal aliquoting accuracy for a volume of 400 μL (maximum tested). Within the 300–400 μL range, the nominal accuracy of sample aliquoting was 99.5% with the coefficient of variation (CV) of 1.1%.

In the case of detection antibody, streptavidin poly-HRP, and DAB, the nominal aliquot volume is 70, 50, and 80 μL respectively, while loading the chip with a micropipette showed that at least an extra of ∼1 μL (for a total of 71, 51, and 81 μL, respectively) is needed to ensure complete filling of each reservoir with desired flow rate and without bubble entrapment. The maximum volumes that reagent inlets could accommodate while avoiding pre-mixing of reagents with buffer were 110, 90, and 120 μL for the detection antibody, streptavidin poly-HRP, and DAB, respectively. Under these conditions, the nominal accuracy of aliquoting was found to be 99.7 (CV = 2.1%), 93.4 (CV = 1.5%), and 99.9% (CV = 3.5%) respectively. In a case that overloading exceeds the maximum volumes, the chip will continue to function, but some part of the excess may spill into other reservoirs, which was observed when loading 140 μl of detection antibody led to spilling into the adjacent buffer reservoir (Fig. S5†).

For the washing buffer, ∼300 μL is needed to fill all washing buffer reservoirs as well as the supply/drainage conduit, inlet/outlet barrier channels, and main assay conduit. We tested its operation for a volume of up to 400 μl, successfully. Within this operating range, the buffer was reliably aliquoted into the four reservoirs of 40 μL, 40 μL, 60 μL, and 20 μL with a nominal accuracy of 98.7, 97.4, 98.3, and 94.7%, respectively, all with the CV of <1%. The results are summarized in Table 1. Considering these values, the ELISA chip outperforms the bulky pipetting robots utilized in laboratory ELISA, and indicates superior performance over the previously developed microfluidic devices while requiring significantly less user intervention.29,30 The computed tomography data provided in our previous study also confirm the high dimensional/volumetric precision of 3D-printed CCs with CV values of ≤3.9% in width and ≤1.2% in depth for the range of 400–1000 μm.24 It should be noted that due to the dimensional precision, the error in accuracy can be compensated by fine-tuning the length of a corresponding reservoir to adjust the volume.

Flow timing

We characterized the reliability of the timing of the ELISA protocol by measuring the duration of each step pre-programmed into the ELISA chip. The timing of the assay steps is an important parameter when considering the accuracy and reproducibility of ELISA. We analyzed the video of four ELISA chips loaded with assay reagents spiked with food dye colors for visualization purposes. Table 1 summarizes the draining/delivery duration for the sample, reagents, and buffer. Remarkably, we found the timing to be reproducible within a CV of 3.5% which confirms print-to-print precision on the smallest conduits which have a remarkably high flow resistance relative to large conduits, and where imprecision could thus disproportionally affect flow timing. We consider these results, both for the volumes and incubation times, excellent for a capillary-driven microfluidic system and suitable for conducting ELISA. Sequential delivery of all solutions takes ∼1.15 h. To reduce the effect of evaporation-induced backflow and the resultant mixing of the sample with other reagents, particularly during the ∼25 min of sample draining, the size and length of the main assay conduit were expanded (see serpentines in the main assay conduit in Fig. 5A).

SARS-CoV-2 N protein ELISA

To evaluate the performance of the ELISA chip, we used it to run a saliva-based SARS-CoV-2 N protein assay. We performed an in-depth optimization process including testing various concentrations of capturing antibody, detection antibody, streptavidin poly-HRP, and Tween 20 as well as the extent of saliva dilution on the minimum detectable signal and assay background. In particular, we observed that increasing Tween 20 concentration improved the assay background, with 0.05% and 0.1% concentrations having comparable performance (Fig. S6†). Saliva dilutions of 1×, 2×, 4×, and 10×, were tested, and 4× was selected as it eased flow through the nitrocellulose membrane, preserved a low assay background, and yielded good assay-to-assay reproducibility without significantly compromising sensitivity (see Table S2† for a summary of the optimization process).

Akin to classical microplate ELISA, the ELISA chip allowed for the implementation of enzymatic amplification that necessitates a two-step process (i.e., the addition of enzyme, followed by the substrate) which is not possible for the common LFAs that are typically carried out in one-step. DAB is the substrate used here which is chromogenic and oxidized in the presence of HRP, forming a brown precipitate at HRP locations. Whereas in laboratory ELISA the assay produces a colored solution, here it forms a precipitate on the paper strip which can be read out by the naked eye or digitized with a scanner.24 While the read-out time for both ELISA and LFAs is limited to a window of a few minutes, the precipitate in the ELISA chip is stable and can be read out later, thus potentially also serving as an archival record.

We generated binding curves by spiking-in N protein of SARS-CoV-2 across 6 orders of magnitude of dilution from 1 to 106 pg mL−1 in 4×-diluted pooled human saliva and ELISA buffer. Fitting the experimental data using a 4-parameter logistic regression, we obtained an LOD of 54 pg mL−1 and 91 pg mL−1 for the N protein in buffer and 4×-diluted pooled saliva, respectively (Fig. 6). The difference in LOD between buffer and saliva is due to the use of saliva as the diluent in the saliva binding curve, which leads to a higher background signal and greater variation than buffer alone. The small dilution of human saliva together with the high sensitivity of the assay is a practical advantage of the developed ELISA-on-chip given the higher performance needed for reliable COVID-19 diagnosis based on saliva testing.33

Fig. 6 Calibration curves of the SARS-CoV-2 N protein assay enabled by the ELISA chip. Different concentrations of SARS-CoV-2 N protein were spiked in ELISA buffer or 4×-diluted pooled saliva. The resultant time-insensitive colorimetric signals were then captured by a regular scanner and yielded a binding curve that was fitted with a 4-parameter logistic regression. The number of replicates for each concentration is 3 for each calibration curve.

We benchmarked the ELISA chip against two commercially available microplate-based ELISA for SARS-CoV-2 N protein detection (SinoBiological, Inc. and RayBiotech Life, Inc.). These ELISA kits use blood serum as the sample and have a time-to-result of ∼5 h with an LOD of 35 pg mL−1 and 70 pg mL−1, respectively, as reported by the manufacturer's protocol (Table 2). Note that the LODs of these serum ELISA were established based on a binding curve using buffer, and are thus comparable to the ELISA chip LOD for buffer; besides, RayBiotech calculated the LOD based on 2 × SD (standard deviation) above the blank instead of the widely used 3 × SD. The LOD of the ELISA chip rivals one of the classical ELISA kits while being four times faster with less hands-on time and no time-sensitive manipulations of liquids, and in a format that is compatible with the point-of-care setting.

A study compared the performance of seven LFA rapid antigen tests for N protein spiked in phosphate buffer saline.34 The tests notably included the widely used Abbott Panbio COVID-19 Ag Rapid Test and the Roche-SD Biosensor SARS-CoV Rapid Antigen Test. The most sensitive one was reported to be the R-Biopharm RIDA QUICK SARS-CoV-2 Antigen Test which yielded a line visible to the naked eye for a concentration as low as 2.5 ng mL−1.34 The LOD of the ELISA chip is ∼50 and ∼25 times higher than this LFA test in buffer and 4×-diluted pooled saliva, respectively.

Conclusions and future work

We presented an integrated ELISA chip that miniaturized and automated an ELISA-on-a-chip using capillarics and an MCR with an encoded aliquoting function, enabling the ELISA chip to be serviced with disposable pipettes. The ELISA chip generated aliquots with various volumes and timed the assay steps both within the CV of ≤3.5%, rivaling the common pipetting robots utilized in laboratory ELISA and other automated microfluidic ELISA.29,30 The ELISA chip could operate with 0.05% Tween 20 commonly used in assays. The LOD of the ELISA chip for the SARS-CoV-2 N protein in 4×-diluted saliva was 91 pg mL−1, in line with classical microplate ELISA and outperforming conventional lateral flow assays by ∼25×.

The ELISA chip could be 3D-printed and assembled in less than 1 h, and ∼1200 chips were manufactured as part of this work. ELISA chips were designed with superficial channels only and may thus be adaptable to mass production by injection molding with much lower mass manufacturing costs than 3D printing.

In the future, the chip may be validated with patient samples in retrospective and possibly prospective studies. The ELISA-on-chip introduced here will be particularly attractive for point-of-care applications where higher sensitivity than an LFA is needed while a longer assay time can be tolerated. Shortening assay time and further simplifying operations by pre-drying reagents and rehydrating them with a buffer solution20–22 would further increase the usefulness of the ELISA chip. A more systematic optimization to accommodate higher surfactant concentrations might expand the range of assays that could be automated on the chip. Finally, the effect of temperature, which is known to modulate the enzymatic turnover, will need to be studied and accounted for prior to clinical use. Following these improvements, ELISA Chips could be deployed at the point-of-need and used by non-experts, and using a cell phone for imaging and quantifying the assay results,21,22,24 quantitative, point-of-care tests with the performance of a central laboratory ELISA become available for everyone.

Experimental

ELISA chip fabrication and preparation

The chips were designed in AutoCAD (Autodesk), exported as “STL” files, and printed via an automated stereolithography 3D printer with the LED wavelength of 405 nm (Pr 110, Creative CADworks, Concord, Canada) using a monocure 3D rapid clear resin (Monocure 3D, NSW, Australia) with the following printing parameters: exposure time per layer: 2.5 s (10 s for the base layer); transition buffer layers: 2; layer thickness: 20 μm; printing delay: 1 min; and gap adjustment: 0.1 mm. Once printed, the chips were rinsed with isopropanol (Fisher Scientific, Saint-Laurent, Quebec, Canada) to wash away uncured resin, dried under a stream of pressurized nitrogen gas, cured for 1 min in a UV lamp (CureZone; Creative CADWorks; Concord; Canada), plasma treated for 10 s at 100% power (PE50 plasma chamber, Plasma Etch, Carson City, USA), and sealed with a microfluidic diagnostic tape (catalog number: 9795R; 3M Science. Applied to Life.™, Ontario, Canada).

A strip of Whatman CF4 paper (Cytiva, Marlborough, Massachusetts, United States) was clamped between 2 absorbent pads (Electrophoresis and Blotting Paper, Grade 320, Ahlstrom-Munksjo Chromatography, Helsinki, Finland) from the back end to collectively serve as the capillary pump. For the main capillary pump, a strip of Vivid™ 120 lateral flow nitrocellulose membrane (catalog number: VIV1202503R; Pall Corporation, Port Washington, USA) was clamped between the same absorbent pads from the back end and to a G041 glass fiber conjugate pad (Millipore Sigma, Oakville, Ontario, Canada) from the front end to facilitate connection to the chip.

Nitrocellulose membrane

The strips of Vivid™ 120 lateral flow nitrocellulose membranes were designed in AutoCAD with the dimensions of 5.2 mm wide and 12 mm long and cut using the Silhouette Portrait paper cutter (Silhouette, Lindon, USA). Membranes were stripped with a 5 mm-wide test line of SARS-CoV-2 N protein mouse monoclonal antibody (catalog number: 40143-MM08; Sino Biological, Inc., Beijing, China) at the concentration of 1 mg mL−1 and a 5 mm-wide control line of bovine serum albumin (BSA)–biotin solution at the concentration of 50 μg mL−1, both delivered using a programmable inkjet spotter (sciFLEXARRAYER SX, Scienion, Berlin, Germany). The membranes were dried for 1 h at 37 °C and blocked by dipping into the blocking buffer solution (1% BSA and 0.1% Tween 20 in PBS) until completely wet, followed by shaking on a rocker for 60 min at 75 rpm. The membranes were then retrieved, dried in an oven for 1 h at 37 °C, and stored with a desiccant at 4 °C until use on the next day.

SARS-CoV-2 N protein ELISA

Fresh saliva specimens were collected from three individuals using oral cotton swabs (Salivette, Sarstedt, Numbrecht, Germany) with their informed consent. The collected saliva samples were then pooled at a 1 : 1 : 1 ratio, filtered through a 0.22-micron filter, and diluted by a factor of 4 in the ELISA buffer solution (0.1% BSA and 0.05% Tween 20 in 1× PBS). The sample solutions were prepared by spiking SARS-CoV-2 N protein (catalog number: 40588-V08B; Sino Biological, Inc., Beijing, China) at the concentrations of 0, 1, 5, 10, 50, 102, 103 104, 105, and 106 pg mL−1 in either the ELISA buffer solution or 4×-diluted pooled saliva solution. The biotinylated SARS-CoV-2 N protein rabbit monoclonal antibody (catalog number: 40143-R004-B; Sino Biological, Inc.; Beijing, China) and streptavidin poly-HRP (Pierce; catalog number: 21140; ThermoFisher; Ottawa, Canada) solutions were prepared in the ELISA buffer solution both with the concentration of 7.5 μg mL−1. The substrate solution was prepared by dissolving SIGMAFAST™ DAB tablets (catalog number: D4293-50SET; Sigma-Aldrich; Oakville, Canada) in Milli-Q water. The washing buffer solution was the same as the ELISA buffer solution.

For benchmarking, the developed SARS-CoV-2 N protein assay in buffer was compared with the SARS-CoV-2 (2019-nCoV) Nucleocapsid Detection ELISA Kit (catalog number: KIT40588; Sino Biological, Inc.; Beijing, China) and the RayBio® COVID-19/SARS-COV-2 Nucleocapsid Protein ELISA Kit (catalog number: ELV-COVID19N; RayBiotech Life, Inc.; Peachtree Corners, United States).

Nitrocellulose membranes image analysis and LOD calculation

After completion of the ELISA, the nitrocellulose strips were removed from the ELISA chip, left to dry at room temperature, and scanned at 1200 dpi in TIFF format (Epson Perfection V600) (see Fig. S7†). The images were imported in Photoshop (version: CS5 ME) and superposed with guide structures to locate the region of interest (2.5 × 0.4 mm) for the test line as well as the bottom background and top background, each located 1.5 mm below and above the test line respectively. The superposed images were then imported into Fiji to measure the gray value of the three regions of interest for each nitrocellulose membrane (see Fig. S8†). For each concentration and the negative control, the local signal intensity of the test line was calculated by subtracting the gray value of the test line from the average gray value of the top and bottom local backgrounds. The relative signal intensity was then calculated by subtracting the local signal intensity of the test line from the average of the local signal intensity of the negative controls.

The experimental data were fitted using a 4-parameter logistic regression with the following equation:35

where a and d are theoretical responses at zero and infinity, respectively, b denotes the slope factor (i.e., Hill slope), and c represents the mid-point concentration (inflection point).35

The LOD was then determined by adding 3 standard deviations to the mean relative signal intensity of the bank samples (i.e., zero antigen concentration) and calculating the corresponding concentration from the established calibration curve.36

ELISA chip video recording and still image capture

For video recording (Panasonic Lumix DMC-GH3K), the ELISA chips were loaded with the ELISA buffer solution in the reagents and washing buffer inlets and with the 4×-diluted pooled saliva solution in the sample inlet. Both solutions were colored with food dye for visualization purposes unless stated otherwise.

Videos were edited in Adobe Premiere Pro (version: 22.1.2) to adjust brightness, contrast, sharpness, and speed. Still images were captured using the Sony α7R III camera and edited in Adobe Photoshop for brightness, contrast, and sharpness.

Characterization of ELISA chip aliquoting and maximum over-loading capacity

To characterize aliquoting, screenshots of the chips following the completion of aliquoting were analyzed in Fiji. The regions corresponding to the extra or lost solution shown in Fig. S4† were mapped, and the equivalent volume was calculated based on the 3D design file. The visual aliquoted volume and the nominal aliquoting accuracy were then calculated as follows:

where the nominal aliquot volume is equal to the capacity of the reservoir in the 3D design.

To characterize the maximum overloading capacity, the chips were loaded with exact volumes of sample, reagents and washing buffer using laboratory micropipettes with an increment of 5 μL for each solution. The recorded videos were analyzed visually to investigate any unwanted mixing or spilling of reagents to the adjacent reservoirs.

Characterization of ELISA chip timing

The videos of the ELISA chips were recorded, and the timing of each step in the MCR read from the video data was tabulated and the duration of different steps was calculated.

Capillary pressure and resistance calculation

Capillary pressure was calculated using the Young–Laplace equation as follows:19


where P denotes the capillary pressure, γ represents the liquid surface tension, h and w are the microchannel height and widths, and θt, θb, θl, and θr respectively are the contact angle of the liquid with the top, bottom, left, and right wall of the microchannel.19 For the ELISA buffer and 4×-diluted saliva, the liquid surface tension γ was measured to be 37.8 and 38.0 mN m−1 respectively using a dynamic tensiometer (DCAT11; Filderstadt; Germany). A custom-built apparatus was utilized for the contact angle measurement. Each θb, θl, and θr was replaced by the contact angle of ELISA buffer (10° ± 1.5°) (mean ± SD) or 4×-diluted saliva (12.5° ± 2°) on the plasma-treated, flat 3D printed samples. θt was replaced by the contact angle of the ELISA buffer (86.2° ± 3.4°) or 4×-diluted saliva (88.7° ± 1.9°) on the microfluidic diagnostic tape. As reported in our previous study, flat 3D printed monocure has a native contact angle of 77° ± 3° before, and 33° ± 5° after plasma treatment for MilliQ water.24 To calculate the resistance of a fluidic path, a lumped-element model was created where each section of the circuit was assigned a resistance calculated using the following equation:19

where R denotes the resistance, η represents the liquid viscosity, and h is the height of the microchannel. Then, for circuit elements in series or parallel, the equivalent resistance (Req) was calculated as follows:

where N denotes the number of capillaric elements included in the corresponding fluidic path. The liquid viscosity, η, was measured to be 1.11 mPa s (temperature: 16.9 °C) for the ELISA buffer and 1.07 mPa s (temperature: 17.7 °C) for the 4×-diluted saliva using a viscometer (SV-10; A&D Company Ltd; Tokyo; Japan).

Author contributions

A. P. and O. Y. designed the ELISA chip. A. P. characterized the ELISA chip. A. P., J. R. W. J., and O. Y. optimized the SARS-CoV-2 N protein assay. A. P. and D. J. wrote the initial draft of the manuscript. A. P. and O. Y. prepared the figs. A. P., O. Y., A. N., and D. J. reviewed and edited the manuscript. A. P. and D. J. analyzed the data. D. J. conceptualized, administered, and supervised the work.

Conflicts of interest

Authors declare that they have no conflicts of interest.

Acknowledgements

We thank Geunyong Kim and Molly Shen for their assistance in assay optimization. We thank Mohamed Yafia for his constructive feedback regarding chip design. We thank Galyna Shul from NanoQAM, UQAM for her assistance in operating the viscometer and tensiometer. This work was supported by the NSERC Alliance COVID-19 Grant (grant number: ALLRP 551058-20), NSERC Discovery Grant (grant number: RGPIN-2016-06723), and the McGill MI4 Emergency COVID-19 Research Funding Grant. D. J. acknowledges support from a Canada Research Chair in Bioengineering (grant number: CRC-232159).

Footages

Electronic supplementary information (ESI) available. See DOI: https://doi.org/10.1039/d2lc00878e 

These authors contributed equally to this work.

Thermomechanical Soft Actuator for Targeted Delivery of Anchoring Drug Deposits to the GI Tract

Thermomechanical Soft Actuator for Targeted Delivery of Anchoring Drug Deposits to the GI Tract

Joshua A. Levy, Michael A. Straker, Justin M. Stine, Luke A. Beardslee, Vivian Borbash, and Reza Ghodssi

Current systemic therapies for inflammatory gastrointestinal (GI) disorders are unable to locally target lesions and have substantial systemic side effects. Here, a compact mesoscale spring actuator capable of delivering an anchoring drug deposit to point locations in the GI tract is demonstrated. The mechanism demonstrated here is intended to complement existing ingestible capsule-based sensing and communication technologies, enabling treatment based on criteria such as detected GI biomarkers or external commands. The 3D-printed actuator has shown on command deployment in 14.1 ± 3.0 s, and a spring constant of 25.4 ± 1.4 mN mm−1 , sufficient to insert a spiny microneedle anchoring drug deposit (SMAD) into GI tissue. The complementary SMAD showed a 22-fold increase in anchoring force over traditional molded microneedles, enabling reliable removal from the actuator and robust prolonged tissue attachment. The SMAD also showed comparable drug release characteristics (R2 = 0.9773) to penetrating molded microneedles in agarose phantom tissue with a drug spread radius of 25 mm in 168 h. The demonstrated system has the potential to enable on command delivery and anchoring of drug-loaded deposits to the GI mucosa for sustained treatment of GI inflammation while mitigating side effects and enabling new options for treatment.

We kindly thank the researchers at University of Maryland for this collaboration, and for sharing the results obtained with their system.

1. Introduction

Inflammatory bowel disease (IBD), that is, Crohn’s disease and ulcerative colitis (UC) is a class of inflammatory gastrointestinal (GI) disorder that impacts 3.1 million adults in the United States and involves chronic inflammation and eventual damage of the gastrointestinal tract.[1–3] Conventional treatment for inflammatory bowel disease (IBD) is primarily achieved through oral or intravenous delivery of therapeutic agents.[4,5] A wide variety of drugs can be employed, including aminosalicylic acids, corticosteroids, immunosuppressants, and various biological macromolecules.[6–8] These medications have a myriad of adverse side effects, limiting the course of treatment for patients.[9–11] For example, corticosteroid treatment duration is limited to approximately 3 months to mitigate the potential for conditions like osteoporosis,[12,13] and immunosuppressants increase susceptibility to opportunistic infections.[14] The presence of substantial side effects can be partly attributed to the large systemic doses needed to achieve effective therapeutic concentrations within the GI tract. Localizing treatment to inflammatory lesions using topically active agents, like corticosteroids, is one method to reduce the necessary drug dose and combat the adverse systemic side effects associated with intravenous and oral non-site-specific therapeutic delivery.[13,15–17] Highly localized topical treatment may also offer a path to mitigate drug costs by reduced dosing, making room for costs associated with innovative delivery modalities. Commercial technologies exist that can improve the localization of drug release within the GI tract. One such technology is pH-sensitive enteric coatings, like Evonik Eudragit L100, that swell and release drugs in pH-specific regions in the GI tract (stomach [pH 1.5–3], small intestine [pH 6–7.4], caecum [pH 5.7], etc.).[18,19] Such coatings help to localize delivery; either focusing the release of systemic drugs to the most absorbent regions of the GI tract or releasing locally active drugs in the most afflicted GI regions enabling site-specific treatment. Another technology that can be applied in conjunction with pH-sensitive polymers is mucoadhesive coatings.[5,20] These coatings bind to the intestinal mucus layer via valence forces or interlocking action slowing tablet transit; thus, localizing the release and delivery of the contained therapeutics.[21] Both technologies help to localize to regions in the GI tract, but do not allow the targeting of highly specific locations of disease affliction. Furthermore, the physiological attributes that determine localization (pH and mucus) can vary from patient to patient. With the advancement of microsystem fabrication technology enabling the miniaturization of remote electronics, sensors and actuators, the treatment of diseases within isolated locations in the human body has become more attainable. Recently, a focus of many researchers on ingestible capsule devices has enabled drug delivery to several regions within the GI tract to treat systemic diseases.[22] For example, the spring-loaded SOMA capsule can deliver millimeter-scale dissolving needles to the stomach triggered by hydration-dependent polymers,[23] and the Rani Pill can do the same using pH-responsive polymers.[24] The LUMI capsule also uses a pH-responsive unfolding mechanism to inject microneedle patches into small intestine tissue.[25] While these passive mechanisms support locational delivery, they do not permit fully closed-loop deployment in response to sensors or explicit commands, and hence only offer regional control over delivery.

Sensing technologies such as optical sensing,[26] gas sensing,[27–29] pH sensing,[30] temperature sensing,[31] and electrochemical impedance spectroscopy (EIS)[32] have been readily integrated into ingestible capsules in recent years. These sensing modalities can give insights into the current state of inflammation in the GI tract and be used to inform active targeted drug delivery to afflicted tissue locations. However, due to the motion in the GI tract, rapid on command actuation is a key requirement to feedback-driven localized therapeutic treatment. Previously, MEMS-based actuators have been developed to achieve on command payload release in ingestible fluid drug delivery and endoscopic location tagging capsules.[33] Various investigators have employed heating elements[34,35] and combustion-based microthrusters[36,37] to achieve a rapid release of fluid drugs from a reservoir. Goffredo et  al. even integrated a ring electrode electrochemical impedance spectroscopy (EIS) sensor with a fluid drug release reservoir.[38] Yet, localization using this fluid payload release method is limited due to fluid dispersion after release from the capsule reservoir, which varies by case and allows little opportunity for control over the release profile. An alternative method capable of further localizing delivery of therapeutics in the GI tract is active delivery of drug-loaded dissolving microneedles to the GI tissue. Lee et al. demonstrated this type of site-specific microneedle delivery using a magnetic locomotion and actuation system.[39] Though they were able to achieve highly localized delivery ex vivo, utilizing such a system in vivo requires external electromagnetic devices that may be clinically impractical. Furthermore, Lee and co-authors observed challenges with tissue adherence of the therapeutic-loaded molded microneedles. A variety of passive and active microsystems have been used previously to enable tissue attachment by latching on to tissue with different forms of thermo and hydration responsive microgrippers,[40] theragrippers,[41,42] microinjectors,[43] and anchoring microneedles[44–48] on benchtop and in vivo. The tissue attachment achieved using these technologies enables long-term residency for extended-release therapeutic delivery, however locational control is still limited. Various forms of thermoresponsive soft meso-scale actuators are also prevalent in literature,[49–51] however the application of these systems for ingestible capsulebased localized drug delivery has yet to be demonstrated. Furthermore, a true combination of thermally triggered and highly localized actuation with active or passive anchoring techniques remains elusive and is a powerful coupling for achieving superlative control over drug localization and release.

In this paper, we look to address the challenges of sitespecific and on command drug delivery in the GI tract (Figure 1a) using a thermomechanical spring actuator paired with a biomimetic drug anchoring structure, termed the spiny microneedle anchoring drug deposit (SMAD). The compact system shown in Figure  1 and Figure S1, Supporting Information, is compatible with sensing and communication ingestible capsule technology for closed-loop detection and therapeutic delivery to lesions in the GI tract. Release of a dissolving drug-loaded deposit anchored in the GI tissue allows prolonged delivery of therapeutics in the target region. A novel 3D-printable spring was designed to imitate a wave spring, which is used in applications requiring large travel distance with less consumed space—a necessity for ingestible capsule devices. In addition, this type of spring provides greater lateral stability when compared to a standard conical coil spring. The spring is fixed in compression using polycaprolactone (Tm = 60 °C), and releases by melting the polycaprolactone using a resistive heating element fabricated on a Kapton substrate. The drug-loaded SMAD in Figure 1b attaches to the top of the spring actuator with a water-soluble polymer to enforce reliable release after actuation. The SMAD exploits recently developed biomimetic barbed microneedle technology that has been demonstrated by our group[44,48] and others[45–47] for improved tissue anchoring. The hybrid system developed here can be combined with sensors to enable on command delivery of a drug-loaded anchoring deposit for early, focused, and prolonged treatment of GI lesions and has the potential to be applied for various other localized treatment applications in the gastrointestinal tract.

2. Experimental Section

2.1. Design and Fabrication of Multi-Coil Spring

Springs were designed using Autodesk Fusion 360 (Autodesk, San Rafael, CA, USA) with a base diameter of 3 mm on center, an outer tip diameter of 2.1 mm, and a height of 8 mm. The conical design had four overlapping coils (two clockwise and two counterclockwise) with a pitch of 2.66 mm, a width of 400 µm, and a thickness of 150 µm. COMSOL Multiphysics 5.4 (Stockholm, Sweden) was used to compare the overlapping conical springs to a traditional conical coil spring. The single-coil spring was designed with the same 400 µm width as the superimposed spring, however with a 600 µm thickness, 4× that of the superimposed spring, yielding a comparable stiffness. A 50 mN axial force was applied to each spring, and lateral deflection was measured to assess the stability under axial compression/decompression. A 4 mm post (∅ = 1 mm) extends down from the underside of the spring tip to affix compression of the spring with a low melting point PCL polymer (Figure 2a).

2.2. Design and Microfabrication of Resistive Heaters

A resistive heater was designed using Autodesk AutoCAD (San Rafael, CA, USA) for an intended resistance of 50 Ω with a 100 nm thickness of deposited thin-film Au on a 1 mil Kapton polyimide film (McMaster-Carr, Elmhurst, IL, USA). The circular heater coil pattern had an outer diameter of 2.18 mm with trace width of 100 µm and a trace length of 20.6 mm, yielding the theoretical 50 Ω resistance with a Au resistivity of 2.44 × 10−8 Ω⋅m.[53] The Kapton film, used for its low thermal and electrical conductivities, was mounted to a carrier wafer with 3M double sided tape (St. Paul, MN, USA), then cleaned using acetone, methanol, isopropanol and DI water (18 MΩ). Cr/Au (20 nm/155 nm) was deposited on the surface at ≈2 Å s−1 using an Angstrom NexDep Electron Beam Evaporator (Kitchener, ON, Canada) and revealed using a liftoff process.

2.3. Molding of PVA Drug Disk and Microneedle Array

Drug disks were cast from a 20% w/v aqueous solution of polyvinyl alcohol (PVA) (Mw 31–50 kDa, Sigma Aldrich, St. Louis, MO, USA) containing FD&C blue #1 dye. Molding of the drug disks was done by solvent casting of the dye-PVA onto a polystyrene (PS) tray. The casting solution was poured into the container to a depth of 2.5 mm, such that the final film thickness would be 500 µm after the ≈1:5 volume reduction. The solvent was then allowed to evaporate for 24 h in the ambient environment yielding a 500 µm PVA film. The film was peeled from the PS casting tray, and a ∅ = 2 mm punch was used to punch the drug disks from the film.

To form 3 × 3 molded microneedle (MMN) arrays for comparison with the SMAD (Figure S5, Supporting Information), an 11 × 11 microneedle array mold was acquired from Blueacre Technology Ltd. (Dundalk, Co Louth, Ireland). Microneedles molded from the 11 × 11 mold had a height of 600 µm, a base diameter of 300 µm and an interspacing of 600 µm on center. 500 µL of the dye loaded PVA solution was deposited on the microneedle mold, then the mold was placed in vacuum for 15 min to evacuate air from the needle mold, prompting filling of the mold. Molds were then removed from vacuum, and the solvent was allowed to evaporate in the ambient environment for 24 h. Arrays were then segmented into 3 × 3 needle sections.

2.4. Fabrication of Biomimetic Barbed Microneedles

Barbed microneedles previously demonstrated in the authors’ group by Liu et  al.[44,48] were fabricated by direct laser writing (DLW) (Figure  2b). The microneedles were 650 µm in height with a 74 µm tip diameter and were adapted to include a 300 µm flared base for greater surface contact and enhanced adhesion to the drug disk (Figure S6, Supporting Information). Each needle contains a total of 72 backward-facing barbs with high sharpness (≈1 µm) that promote robust tissue anchoring. DLW was performed using the Dip-in Laser Lithography (DiLL) mode on a fused silica substrate with the Nanoscribe Photonic Professional GT (Karlsruhe, Germany). IP-S photoresist was used with a 25× objective and a slicing distance of 1 µm to fabricate a 3-needle array. Needles were printed upside down in a triangular pattern, each needle being 600 µm from the array center and 120° separated from the adjacent needle. This predetermined spacing supports reliable attachment to the drug disk and control over the spatial arrangement of the needles on the fabricated structure. After printing, needle arrays were cleaned in propylene glycol monomethyl ether acetate (PGMEA) for 15 min, followed by 5 min in isopropyl alcohol and 2 min on a hot plate at 60 °C.

2.5. Package Assembly

Polycaprolactone (PCL) flakes (Mw ≈ 14 000, Sigma Aldrich, St. Louis, MO, USA) were melted and compressed between two Kapton sheets to a thickness of 150 µm. A 4 mm biopsy punch was used to punch a disk from this film. This disk was melted on top of the resistive heater, then the spring was placed atop the melt, and compressed as shown in Figure 2a. The package was then removed from heat and held compressed until the PCL solidified. The drug disk was then attached to the top of the actuator using ≈1.5 µg of melted polyethylene glycol (PEG) that solidifies after cooling. The assembly was then lowered onto the barbed 3-needle array and adhered using a film of Loctite M-21HP biocompatible (ISO-10993) epoxy adhesive. The adhesive was allowed to cure for 2 h then the assembly was raised to mechanically detach the needles from the fused silica substrate (Figure 2c).

2.6. Mechanical Characterization of Spring Actuator

The spring actuator underwent mechanical compression testing using an Instron 5942 universal testing system (Norwood, MA, USA) equipped with a 10 N load cell. The force profile was obtained during loading and unloading at a crosshead speed of 1 mm min−1 . Springs were attached to the platen, then the crosshead was lowered until contacting the spring. Springs were compressed to a displacement of 3 mm, then decompressed until reaching the test origin. Two test groups were evaluated using this protocol: 1) springs compressed for ca. 18 h before testing (n = 3); and 2) springs untouched after printing (n = 8).

2.7. Characterization of SMAD and MMN Anchoring

Mechanical tests were performed to compare SMAD and MMN tissue anchoring and removal forces (Figure S7, Supporting Information). This was done using the same Instron 5942 universal testing apparatus with a 10 N load cell. All tests were performed using a crosshead speed of 1 mm min−1 . Spring actuators fitted with MMN or SMAD tip structures were lowered onto tissue samples until reaching 75 mN of compressive force. Tissue was ordered frozen from Animal Biotech Industries (Boylestown, PA, USA) with mucus attached and without mesentery. Tissue samples were pre-coated with a ≈2 mm layer of 1× phosphate buffered saline solution (PBS) (Sigma Aldrich, St. Louis, MO, USA) to simulate the presence of mucus and aqueous intestinal media on the tissue surface. Upon reaching the 75 mN force, the tissue was moved 2 mm laterally to reproduce the longitudinal motion experienced in the GI tract. The sample was then retracted from the tissue. For samples that removed from the tissue before detaching from the actuator, the tissue removal force was measured. For samples that detached from the actuator and remained in the tissue, the detachment force was measured. An additional group of SMAD samples were permanently adhered to the actuator to determine the SMAD tissue retention force because the SMAD remained anchored in tissue for all other cases.

2.8. Model Drug Delivery

MMN and SMAD samples were compared using a quantitative method to evaluate the diffusion of dye from each structure into agarose phantom tissue. A ≈1 mm agarose film was created in petri dishes, and dye-loaded MMNs and SMADs were applied to the film to evaluate the 2D diffusion profile from each sample type. Images were captured with controlled lighting at set time points from 0 to 168 h after insertion into the agarose phantom. The images were processed in MATLAB R2021b (MathWorks Corporation, Natick, MA, USA) to quantitatively determine the diffusion radius for each time point (Figure S8, Supporting Information). The red channel image was obtained for each image, and the image was binarized using a 40% intensity threshold. Pixels were scaled and counted, and the radius of diffusion was calculated using the number of 0 pixels representing the area of dye spread. Radial diffusion was then plotted, and correction for initial dye mass in each sample was performed in the context of the diffusion equation.

2.9. Deployment Characterization in a Simulated Environment

The heating element, spring actuator, and attached drug disk were packaged in a polyethylene terephthalate glycol (PETG) 3D printed capsule shell to evaluate properties of actuator deployment. Capsules (n = 7) were mounted on a custom-made testing apparatus designed to control translation speed of the capsules. Contact was made between the capsule body and an agar hydrogel bed, which was intended to capture the location of contact with the dye-loaded drug disk based on dye remnants in the hydrogel. The capsules were set to translate at 1.4 cm min−1 across the hydrogel surface, the mean speed of motion in the small intestine.[54] The actuator was fired at a set displacement along the bed and the deployment translation distance was then measured as the distance between firing the spring actuator and the appearance of the dye marker on the agar bed.

Assessment of the fully assembled SMAD and actuator was performed in a similar manner to the deployment characterizations, however ex vivo porcine intestinal tissue with mucus still attached (Animal Biotech Industries, Boylestown, PA) was used instead of the agar bed. The ex vivo tissue was placed in a 3D-printed PLA semicylindrical cutout (∅ = 25 mm) to replicate the curvature of the small intestine. Test capsules were translated at 1.4 cm min−1 and the actuator was deployed via current flow through the resistive heater, and the detachment process of the SMAD from the actuator was monitored. SMAD removal was also evaluated during lateral translation across ex vivo intestinal tissue outside of a capsule to verify removal of the SMAD from the actuator upon capsule translation in the small intestine. The actuator and attached SMAD were lowered onto intestinal tissue (Figure S9, Supporting Information), then the actuator was moved laterally at ≈1.4 cm min−1 while monitoring the detachment process via stereomicroscope.

3. Results and Discussion
3.1. Fabrication and Assembly
Figure  2 shows the fabrication and assembly process for the thermomechanical spring actuator and SMAD system. Polycaprolactone (PCL) is melted on the microfabricated resistive heater, then the spring is compressed and held in place when

the PCL solidifies (Figure 2a). The SMAD is then built on top of the spring. First, the biomimetic anchoring microneedles are printed upside down on a fused silica substrate using DLW (Figure  2b). The drug-loaded polyvinyl alcohol (PVA) disk is then attached to the spring with polyethylene glycol (PEG) and the microneedles are transferred to the SMAD via biocompatible epoxy resin (Figure  2c), yielding the fabricated structure (Figure 2d).

3.2. Simulated Characteristics of the 3D-Printed Soft Spring Actuator

Wave springs offer high lateral stability and space conservation in the axial direction, but their fabrication using 3D-printing is challenging due to the low level of connectivity between the stacked coils. To address this challenge, we developed a wave-like spring design that utilizes similar principles to improve lateral stability and axial compression characteristics but is compatible with DLP 3D-printing because of the reinforced joints between coils. Figure 3 shows a comparison in COMSOL Multiphysics between an 8 mm conical coil spring and the wave-like spring under axial compression with a force of 50 mN. Lateral deflection experienced by the superimposed spring was reduced to approximately 1/8th that of the standard conical coil spring (1341 to 172 µm). This result validates this design, indicating higher stability under axial loading—a necessary attribute for reliable injection of microneedles into the mucosal tissue using a freestanding soft spring actuator. Furthermore, the lateral stability achieved by this design is critical for the repeatable 3D printing of the springs in a vat polymerization process like DLP. Springs with less lateral stability tended to sway with movements of the build plate and resin, causing misalignments between the crosslinked layers, and print failure in some cases. DLP 3D fabrication is advantageous because it permits the fabrication of soft polymeric springs with blunt surfaces and tunable mechanical characteristics by varying the volume ratio of the Tuff:Flex100 resins that were used. The balance of compliance and stiffness achieved here is critical to attaining sufficient spring stiffness while safely interfacing with the delicate GI mucosal tissue.

Apparatus &Materials

M Series

3.3. Mechanical Characterization of the 3D-Printed Spring

Mechanical properties of the spring were characterized via mechanical compression tests to evaluate the spring stiffness and force application profile (Figure 4). Additionally, a comparison was performed between freshly printed springs and springs compressed overnight (≈18 h) to examine the impact of prolonged compression on deployment reliability. Further experimentation on the effects of compression and ageing is shown in Figure S2, Supporting Information. Samples were loaded between chucks and the springs were compressed to a displacement of 3 mm at a rate of 1 mm min−1 , then decompressed at 1 mm min−1 until reaching the origin. Force exerted on the 10 N load cell was plotted versus displacement and averaged for each group to yield the mean spring compression curves. The mean spring stiffness for each group was calculated using the maximum force, minimum force, and displacement through force loading for each sample.

The mean spring stiffness within the control group was found to be 25.4 ± 1.4 mN mm−1 (n = 8), while stiffness among the overnight compression group was 27.0 ± 2.5 mN mm−1 (n = 3). Comparing these figures to the 0.6 mN insertion force for the biomimetic barbed microneedles previously shown by Liu et al,[44,48] this force is sufficient to insert many microneedles into the GI mucosa at only small deflection values. Furthermore, the pressure exerted on the tissue at full compression, ≈24 kPa, is expected to be non-destructive to the GI tissue as previous reports of pressure application on porcine intestinal tissue showed no significant tissue trauma below 100 kPa.[55] The use of soft polymeric materials has advantages for tissue compatibility, because the soft material is less destructive than metal springs to the delicate GI tissue. However, soft polymeric materials can exhibit viscoelastic characteristics and plastic deformation that make them challenging to use for a spring mechanism. The mechanical tests performed here give insight into the effects of viscoelastic properties on the spring actuator. The first notable consequence is that at the compression–decompression rate of 1 mm min−1 , the spring exhibited ≈600 µm relaxation throughout the compression. This degree of relaxation is still not prohibitive to the project objectives, as only ≈1 mm of the full 3 mm actuation distance needs to be realized for microneedle penetration into tissue. The comparison here also gives insight into the effects of long-term compression on the spring characteristics. It was observed that the overnight compression springs experience ≈390 µm of relaxation due to the prolonged compression and showed comparable relaxation to the as-printed springs during the compression cycle. Though this still enables more than the ≈1 mm actuation distance needed for SMAD anchoring, the challenge of viscoelasticity in such a soft actuator deserves more attention. In the future, selection of low-creep polymers and process refinement may significantly improve these characteristics.

3.4. Heater Characterization

Power supply for ingestible electronics is a significant limitation, and coin cells like the 2L76 are commonly used, which can produce a sustained current of up to 60 mA. Heaters were designed to maximize power dissipation under these constraints. Accordingly, the design resistance of 50 Ω limits peak current draws to ≈60 mA by ohms law with a 3.3 V source after consideration of additional internal resistances of the system. The linear dependence of conductance on cross sectional area begins to fail as material properties change below ≈100 nm trace thickness,[56] therefore the heater was designed to have the desired 50 Ω resistance with a 100 nm film deposition thickness to minimize material cost while still enabling predictable 
changes in resistance with changes in trace deposition thickness. Given the constraints on resistance and trace thickness, the critical parameter of trace Length Width-1 must have the value of 208.3. Au heaters were deposited to 100 nm design trace thickness (Figure 5), but resistance testing revealed a systematic process error, resulting in a mean resistance of 77.5 Ω, a 55% deviation from the design value. The origin of this error is unclear; however, it could be a result of a systematic error in deposition instrumentation, lithography process, substrate topography or deposition uniformity that may have impacted the cross-sectional trace area in some regions, yielding higher than expected resistance values.

To address the deviation, a linear correction was made using the design and experimental resistance values, indicating that a 155 nm deposition thickness would return the desired 50 Ω trace resistance. The revised fabrication process was performed, yielding an experimental resistance of 49.8 ± 1.8 Ω, a 0.4% deviation from the intended resistance. The melt time of PCL when in contact with the resistive heater was evaluated by supplying current from a 2L76 coin cell battery regulated to 3.3 V. A mean melt time of 3.3 ± 0.2 s was achieved, indicating the ability of the mechanism to fire rapidly to achieve on command delivery of the drug-loaded SMAD to the GI tissue. Moreover, this melt duration at 60 mA corresponds to only 0.03% of the 2L76 160 mAh capacity.[57]

3.5. Spring Deployment Testing

Following the validation of the independent spring and heater systems under capsule-relevant constraints, the combined system was evaluated to understand characteristics of spring deployment (Figure  5). The spring actuator was deployed outside of a capsule (Figure S3, Supporting Information) and packaged in a capsule (Figure 5a) to image deployment using a 2L76 coin cell battery. Then, to determine the deployment time and distance, a capsule was translated on an intestinal simulator and the spring was fired into an agar bed. The spring was capped with a dye-loaded PVA disk, and fired at a predetermined location, eventually marking the point of contact between the dye-loaded disk and agar bed (Figure 5c). The distance between firing and dye marking was measured and the time to contact was calculated using the translation speed. The mean measured transit distance before contacting the phantom agar medium was 3.3 ± 0.7 mm (n = 7). Using this data and the translation speed, a deployment time of 14.1 ± 3.0 s (n = 7) was determined. Based on the deployment time demonstrated here and the polymer melt times, it is evident that the major contribution to deployment time is the decompression of the spring. This deployment time contribution is the experimental manifestation of the previously discussed viscoelastic spring properties. Nevertheless, spring deployment was found to be reliable and predictable, as indicated by the 3.0 s standard error in deployment time, which is acceptable relative to the mean intestinal translation speed of 1.4 cm min−1 . [54] Furthermore, instances of failed deployment did not occur during the controlled experiment. In early testing and development, instances of failed deployment were a result of fabrication error, for example, spring-heater misalignment, excess of PCL or poor electrical attachment to the thin-film heater—errors that would largely be resolved by mass-fabrication approaches.

The actuator deployment distance demonstrated here is an important metric to determine both the accuracy and precision of location targeting. The precision of delivery location should be high in comparison with the radius of drug spread after delivery to ensure that the delivered drug reaches the target location. This point will be addressed below in the discussion of model drug delivery experiments. Another significant consideration is the locational accuracy, or mean deployment distance, compared to the size of the ingestible capsule. For sensor-informed delivery to locations of interest, the accuracy of drug delivery location can be augmented by the positioning of sensors in front of the drug delivery actuator to correct for the deployment distance. The Food and Drug Administration (FDA) recommended 22 mm maximum capsule size[58] is significantly larger than the 3.3 mm deployment translation distance shown here, therefore sensor placement can be used to account for the deployment time, yielding much higher accuracy in delivery. These calculations rely on the mean translation speed in the GI tract, however motion in the GI tract is not continuous, thus a more comprehensive evaluation of delivery timing could improve the precision of delivery. With sensing modalities like optical sensing, one can even estimate the current translation speed and fire the actuator at a suitable time to improve locational accuracy.

3.6. Characterization of SMAD Anchoring in Ex Vivo Tissue

To quantitatively evaluate the axial tensile removal properties of the SMAD when compared to the widely demonstrated MMN technology,[59–67] mechanical removal experiments were performed on the SMAD and MMNs attached to the spring actuator. Figure 6a shows a representative sample at each stage of testing, and Figure  6b illustrates the force loading and unloading during experimentation, including a red arrow indicating the point of detachment or removal. Figure  6c,d shows the mechanical removal and tip detachment data of the SMAD compared to MMNs and the corresponding renderings of each structure. ‘Tip detachment force’ refers to the removal force of the SMAD or MMN structure from atop the actuator, while ‘anchoring force’ refers to the force required to remove the SMAD or MMN structure from the tissue sample. The conical MMNs showed an anchoring force of 0.8 ± 0.1 mN (n = 4) compared to the 17.2 ± 2.6 mN (n = 4), a 22-fold improvement over the conical MMNs. Furthermore, the 17.2 mN anchoring force of the SMAD is significantly higher than the 3.3 ± 1.1 mN (n = 4) force required to detach the tip structure from the actuator. Conversely, the MMNs demonstrated an anchoring force that was insufficient to remove the array from the actuator. In all cases, we found that tissue removal or SMAD/MMN removal occurred before rupture of the spring. The firm tissue anchoring achieved here is critical for removal of the SMAD from the actuator, but it also enables robust adherence to the target region and, consequently, reliable prolonged therapeutic delivery. The exceptional anchoring ability of the structure demonstrated here compared to the MMNs generates more reliable tissue anchoring and system operation. Furthermore, the biocompatibility of the IP-S microneedles, Epoxy resin, PVA, and PEG materials enables prolonged attachment without significant harm to tissue and surrounding organs.

3.7. Model Drug Delivery

To demonstrate the efficacy of drug delivery in agarose phantom tissue, the diffusion of a model drug (FD&C Blue #1 Dye) from SMAD and MMN samples was compared. Figure 7a shows the release and subsequent diffusion of dye from a SMAD sample at 0, 48, and 168 h as representative time points for the dye diffusion profile. At 48 h, the apparent perimeter of dye diffusion is at a radial distance of ≈1.8 cm, while this expands to ≈2.5 cm after 168 h. Five samples of each SMAD and MMN were characterized using this diffusion approach and represented quantitatively as the mean diffusion radius at each measured time point in Figure  7b. Upon initial observation, there exists a discrepancy between the extent of diffusion from the MMN and SMAD. However, a difference in outcomes is to be expected due to variations in initial dye mass between MMN and SMAD structures. The thin agarose diffusion medium constrains diffusion to two dimensions; thus, this case most closely resembles 2D Fickian diffusion. The time-dependent concentration profile pertaining to this diffusion case is described by Equation (1), where C is concentration, C0 is initial concentration, D is the diffusion constant in the given medium, t is time and r is radial distance:

The extent of diffusion from each sample can be measured by the radial diffusion distance at which the dye concentration exceeds a threshold value. With consistent lighting, the threshold concentration value corresponds to a constant light intensity value in the dye spreading image. Thus, the red channel light intensity from a blue dye sample can be used as a direct indicator of the perimeter of constant dye concentration—enabling a quantitative treatment of the diffusion to account for differences in initial dye content across samples. For a given intensity threshold (T), the squared radial diffusion distance (r2 ) is predicted by Equation (2):

At one specific measurement time, D, t, and T are constant and r2 carries a logarithmic dependence on the initial concentration; therefore, one time point can be used to compare the relationship between r2 and C0 for the SMAD and MMN cases. Figure  7c uses t = 168 h to compare the radial diffusion distance with initial dye mass in the context of Equation (2). The data across both groups obeys the equation with R2 = 0.9773. To validate the logarithmic character, a diffusion coefficient of D = 2.6 × 10−10 m2 s−1 was calculated from the logarithmic coefficient using Equation  (2). This value is in strong agreement with previously reported values for the diffusion of dye in agar gel (D = [2.5 ± 0.2] × 10−10 m2 s−1 ),[68] corroborating the logarithmic data trend. These results indicate that the SMAD is capable of effusion of drug into tissue in a comparable manner to penetrating MMNs that have been widely validated in literature.

Revisiting the spread radius of dye in the agarose phantom, the radius of dye spread in tissue is a critical parameter because containment enables increased local concentration with reduced dosing. However, radial diffusion distance must be sufficient to target an inflammatory location with high repeatability using the actuation mechanism demonstrated here. The intestinal translation distance between firing and tissue contact varied with a standard error of 0.7 mm and a 99.9% confidence level of repeatability within 2.24 mm. Compared to the dye spread radius demonstrated in Figure  7 of up to 25 mm, the repeatability is more than sufficient to ensure therapeutic coverage to the inflammatory target site. Moreover, there is room to increase the localization, which can be done using material selection. The expulsion of model drug from the drug disk relies primarily on the diffusion of a dye from the PVA following the polymer hydration and swelling. Once hydrated, the diffusion from the SMAD to the phantom tissue is rapid compared to diffusion through the phantom and the concentration in the SMAD is approximately equal to the directly adjacent phantom. In this sense, the disk acts as a directly connected reservoir of therapeutic agent, thus the drug spread relies largely on diffusion from the SMAD. Alternatively, low-solubility materials could be used that would offer the ability to regulate and significantly slow the release of drug from the SMAD. Consequently, the system developed here provides a platform for modulating the release profile utilizing a variety of materials with different dissolution or swelling rates in intestinal fluid to fit the needs of the desired application, potentially spanning to broader applications than only gastrointestinal inflammation.

3.8. Spring Deployment and SMAD Release in Tissue

To demonstrate the combined system operation in a controlled ex vivo environment, the entire packaged system was placed in a test capsule and fired into porcine intestinal tissue (Figure 8). Power was supplied to the heater and limited to 3.3 V and 60 mA, emulating capsule conditions. Translation on the ex vivo tissue occurred at a speed of 1.4 cm min−1 , and the spring was fired during translation. No leakage of dye into the tissue and surrounding fluid was observed before the SMAD was released into the tissue. Following the deployment of the actuator, blue dye could be seen behind the capsule, then the SMAD appeared from behind the capsule. This result validates the reliable location-specific release that the actuator and SMAD can achieve. Notably, the presence of simulated intestinal fluid media increases the rate of drug spread and impacts the level of achievable localization. Further investigation will be required to understand the degree of drug spread with varying drug disk composition and intestinal fluidic conditions. Nevertheless, the prolonged direct tissue contact made by the SMAD enables the greatly increased localization of effused drug when compared to an instantaneous release of fluid.

Throughout deployment a significant component of displacement and force on the system will be applied perpendicular to the actuation direction. To model and document SMAD release in this case, the SMAD was attached to an actuator outside of a capsule, applied to tissue and translated laterally (Figure 8c) until the SMAD structure was removed. Removal occurred at ≈3 mm deflection, corresponding to 3 mm of capsule transit within the intestine. This experiment further validates the structures removal in tissue under lateral force application and gives insight into the removal characteristics.

Overall, the significantly localized model drug delivery achieved using the SMAD has the potential to considerably increase the level of focused treatment for early intervention of GI disorders. Directed therapeutic delivery to the 2.5 cm delivery radius demonstrated here compared to the ≈32 m2 gastrointestinal surface area[69] would result in a ≈16  000-fold higher areal drug concentration. This drastic enhancement in local concentration could serve to mitigate or effectively eliminate drug side effects with lower dosing and achieve enhanced targeted treatment of affected inflammatory regions, enabling early intervention of GI disorders, and limiting the spread and consequences of the disease while minimizing drug side effects.

It is critical to note that the localization of drug release could vary significantly when performed in vivo. The introduction of peristaltic and segmentation movements,[70] as well as varying fluid and mucus content will result in lower predictability in delivery localization. Still, location specific delivery is significantly enhanced using this mechanism. There are also tissue compatibility challenges within the dynamic GIT environment. The SMAD has the potential to become permanently lodged within tissue via envelopment in intestinal folds, which could lead to complications including blockage and infection. One mitigating circumstance is the solubility of the drug disk, resulting in complete dissolution on the scale of days, leaving only the microscale biocompatible needles to be cleared by cellular mucosal cell turnover.[71] Further experimentation is also needed to evaluate the impacts of prolonged retention on tissue integrity and inflammation, as well as blockage for passing luminal contents including the capsule itself. Microneedle devices in general have been established as exhibiting low tissue damage,[59,60,65] however more analysis is needed to evaluate the impacts of the biomimetic needles on tissue. Nevertheless, the system demonstrated here has the potential to meaningfully enrich the collection of available targeted drug delivery technologies.

The hybrid fabrication process demonstrated here utilizes several technologies in concert to achieve precise actuation and robust anchoring. Scaling of these processes also presents a challenge, as 3D printing methods in particular are of notoriously low throughput. However, micromolding of 3D printed components from this proof-of-concept system presents a possible mass scale fabrication strategy that would further enable versatility in material selection. Fabrication reliability and throughput would also be augmented by robotic assembly machines for film casting, stamping, microfabrication, and component placement. Evaluating the aptitude of such largescale processes will be a critical investigation for future application of this technology.

4. Conclusion

In this paper we introduce a compact thermomechanical 3D-printable actuator combined with the first application of biomimetic barbed microneedles toward drug delivery in the GI tract. This is accomplished using a 50 Ω microfabricated thin film Au resistive heating element that melts a PCL adhesive layer, firing the spring actuator. Gastrointestinal anchoring is achieved by a SMAD structure, composed of spiny microneedles adhered to a dissolving drug loaded PVA deposit. The high resolution (≈1 µm) of the DLW process enables high sharpness and a firm tissue anchoring force of 17.2 ± 2.6 mN (n = 4). The actuator demonstrated reliable and repeatable deployment with a mean deployment time of 14.1 ± 3.0 s (n = 7), and SMADs with the barbed microneedles anchored with a 22-fold higher tissue retention force than conical molded microneedles—reliably overcoming the force required for removal of the SMAD from the spring actuator. SMADs also demonstrated comparable drug delivery characteristics to standard MMNs through model drug delivery in an agarose phantom tissue. Diffusion of dye from SMADs reached a radial distance of 25 mm. After correction for the initial dye mass in each sample, the SMAD and MMN data showed high correlation (R2 = 0.9773) indicating predictable and comparable performance with a diffusion constant of D = 2.6 × 10−10 m2 s−1 . Furthermore, the vast material selection available for a simple disk structure introduces a means for direct modulation of drug release profiles and spread characteristics. Overall, the reliable actuation and robust anchoring provided by this system enables location-specific long-term delivery and anchoring of therapeutics to facilitate prolonged treatment of target locations in the GI tract, opening new possibilities for early therapeutic treatment of GI diseases and other local gastrointestinal conditions.

Microfluidics system. The microfluidics system created for this study encompasses the micro-electrodes designed on PCB and a PDMS microchannel squeezed hermetically between 3d-printed components.