The authors acknowledge the financial support from the programs of the Natural Science Foundation of the Jiangsu Higher Education (22KJB460033), and Jiangsu Science and Technology Programme - Young Scholar (BK20200251). This work is also partially supported by the XJTLU AI University Research Centre, Jiangsu Province Engineering Research Centre of Data Science and Cognitive Computation at XJTLU and SIP AI innovation platform (YZCXPT2022103). The support from State Key Laboratory for Manufacturing Systems Engineering via the open project (SKLMS2023019) and Key Laboratory of Bionic Engineering, Ministry of Education, are also acknowledged.

1. Microembossing process for microchannel patterning on nanopaper
<ol>
	<li>Mold preparation 
	NOTE: Refer to Yuan et al.12 for details on mold preparation.
	<ol>
		<li>Prepare a PTFE film as indicated in the Table of Materials.</li>
		<li>Laser-cut the prepared PTFE film to make a convex microchannel mold (Figure 1A-I). 
		NOTE: The dimensions of the PTFE mold determine the microchannel dimensions (Figure 2E,F) in a linear first-order function relationship.</li>
	</ol>
	</li>
	<li>Nanopaper preparation
	<ol>
		<li>Disperse 4.0 g of (2,2,6,6-tetramethylpiperidin-1-yl)oxyl (TEMPO)-oxidized NFC gel (see Table of Materials) in distilled water (final concentration of 0.1 wt%).</li>
		<li>Stir the suspension heavily at 120.8 x g for 30 min at room temperature until no cellulose floc is visible.</li>
		<li>Vacuum-filter the clear suspension to obtain a nanopaper gel (Figure 1A-II). 
		NOTE: In this example, the diameter of the obtained nanopaper gel is 4 cm. NanoPADs can be tailored for various applications by selecting suction filtration devices with different radii, enabling the design of NanoPADs at different scales.</li>
	</ol>
	</li>
	<li>Embossing of nanopaper gel
	<ol>
		<li>Place the PTFE mold on the surface of the nanopaper gel.</li>
		<li>Emboss the nanopaper gel (Figure 1A-III) using the PTFE mold by the hot press for 10 min each time under optimized pressure and temperature (Figure 2A-D). 
		NOTE: Higher embossing pressure (250 kPa to 1000 kPa) improves fabrication accuracy but should not exceed 1000 kPa to prevent damage to the cellulose structure. Higher embossing temperatures (25-100 &#176;C) enhance microchannel accuracy by promoting dehydration and decarburization, but temperatures should not exceed 75 &#176;C to avoid gel wrinkling and reduced light transmittance7. In this example, optimized embossing parameters were 750 kPa and 75 &#176;C.</li>
	</ol>
	</li>
	<li>Mold releasing
	<ol>
		<li>Peel off an additional layer of filter nanopaper gel from the filter membrane (Figure 1A-IV).</li>
	</ol>
	</li>
	<li>Bonding
	<ol>
		<li>Attach the peeled layer on top of the embossed layer of nanopaper gel, stacking the two layers to create a hollow microchannel structure (Figure 1A-V). 
		NOTE: The stronger hydrogen bonding in &#39;gel-like&#39; nanopaper compared to fiber suspension and dried nanopaper enhances the entanglement and adhesion of nanocellulose fibers. Consequently, two layers of &#39;gel-like&#39; nanopaper can bond tightly through self-diffusion without external force.</li>
	</ol>
	</li>
	<li>Drying
	<ol>
		<li>Place the two layers of nanopaper gel in a drying oven at 75 &#176;C for approximately 30 min (Figure 1A-VI).</li>
	</ol>
	</li>
</ol>
2. Construction of fundamental microfluidic devices
<ol>
	<li>Construction of laminar-mixer
	<ol>
		<li>Prepare the NanoPADs with straight and curved channels (Figure 3A) following step 1. 
		NOTE: In this example, the dimensions of channels are 1 mm width and 50 &#181;m depth.</li>
		<li>Add red and blue droplets in the inlet zones simultaneously, allowing flow through the hollow channel automatically. 
		NOTE: The successful independent flow of the red and blue solutions in a straight channel and their mixing at the end of the curved channel can be attributed to the low Reynolds number of the layers in microfluidic devices and the radial flow induced by shear stress35.</li>
	</ol>
	</li>
	<li>Construction of droplet generator
	<ol>
		<li>Prepare the two-inlet NanoPADs with a T-junction channel (Figure 3D) according to step 1.</li>
		<li>Introduce water and hexadecane (oil), two immiscible liquids, into the two-inlet zones of the T-junction channel to generate droplets (Figure 3E). 
		NOTE: In this example, the dimensions of the T-junction channel are 1 mm width, 25 mm length, and 50 &#181;m depth.</li>
		<li>Fix the speed of Q1 at 6 &#181;L/min, and the speed of Q2 at n &#215; Q1 (n = 1-6). Use two syringe pumps and set them at the above speed to inject water and oil. This behavior is governed by the one simple scaling equation (provided below). 
		NOTE: In this example, oil and colored water were poured into the channel36. 
		<img alt="Equation 1" src="/files/ftp_upload/65965/65965eq01.jpg" style="margin: auto;" /> 
		Where &#945; = 1, &#946; = 1, L is the length, W is the width of the droplet, and Q1 and Q2 are the flow rates of water and hexadecane, respectively37,38.</li>
	</ol>
	</li>
</ol>
3. In-situ AgNP growth
<ol>
	<li>NanoPADs preparation
	<ol>
		<li>Prepare two-inlet NanoPADs with a converging detection zone (Figure 4A) according to step 1.</li>
	</ol>
	</li>
	<li>Successive ionic layer adsorption and reaction process
	<ol>
		<li>Prepare a 20 mM AgNO3 solution and a 20 mM NaBH4 solution (see Table of Materials).</li>
		<li>Drop 5 &#181;L of the 20 mM AgNO3 solution into the left inlet zone of the flow channel.</li>
		<li>Allow the AgNO3 solution to remain in the reaction zone for 30 s. 
		NOTE: Repeat steps 3.2.2. and 3.2.3. five times to ensure the uniform distribution of AgNPs without agglomeration, which could explain the higher band intensity.</li>
		<li>Drop 5 &#181;L of distilled water into the left inlet zone of the flow channel for rinsing. 
		NOTE: Repeat step 3.2.4. three times to ensure the removal of excessive, unadsorbed Ag ions through washing.</li>
		<li>Add 5 &#181;L of the 20 mM NaBH4 solution to the right inlet zone of the flow channel. 
		NOTE: Repeat step 3.2.5. until the AgNPs are generated evenly in the reaction zone. The chemical reactions involved in step 3 are represented by the following formula39: 
		<img alt="Equation 2" src="/files/ftp_upload/65965/65965eq02.jpg" style="margin: auto;" /> 
		 
		In this example, dense, uniform, well-structured AgNP arrays were formed on the NanoPADs (Figure 4B). The average diameter of the AgNPs was 55 nm (Figure 4C).</li>
	</ol>
	</li>
</ol>
4. SERS measurement
<ol>
	<li>Raman spectroscopy system preparation
	<ol>
		<li>Turn on the laser and launch the accompanying software for the Raman spectrometer (see Table of Materials).</li>
		<li>Employ a 50x objective for focusing and collecting Raman signals and a 532 nm laser for excitation.</li>
		<li>Set the spectral resolution to 2 cm-1 for accurate measurement. Set Raman spectrum measurement range from 400 cm-1 to 600 cm&#8722;1.</li>
		<li>Calibrate the Raman spectrometer using a silicon wafer12. 
		NOTE: Perform step 4.1. for step 4.2.</li>
	</ol>
	</li>
	<li>Rhodamine B (RhB) measurement
	<ol>
		<li>Dissolve 4.7 mg of RhB (see Table of Materials) in 10 mL of ethanol to prepare a 1 mM RhB solution.</li>
		<li>Prepare a series of RhB solutions with concentrations ranging from 10 &#181;M to 0.1 pM by diluting the 1 mM RhB solution in the ethanol.</li>
		<li>Add 5 &#181;L of the RhB solution onto the inlet zone of the NanoPADs channel and let it dry. 
		NOTE: Repeat step 4.2.3. for RhB solutions of different concentrations indicated in step 4.2.2.</li>
		<li>Set the excitation time to 10 s, the grating to 2 cm&#8722;1, and the number of cycles to 1. Set Raman spectrum measurement range from 500 cm-1 to 1800 cm&#8722;1.</li>
		<li>Adjust the coarse focus screw and fine focus screw individually to achieve proper focus, then click on stop to save the position.</li>
		<li>Click on start to begin the measurement.</li>
		<li>Repeat the measurements seven times and save the collected data.</li>
		<li>Turn off the laser.</li>
	</ol>
	</li>
	<li>Data analysis
	<ol>
		<li>Import the saved data into the data analysis software (see Table of Materials).</li>
		<li>Calculate the average spectrum from the saved data.</li>
		<li>Select the line draft option to plot Raman spectra.</li>
		<li>Utilize the Peak analyzer tool to set the baseline of the spectra.</li>
		<li>Apply the Signal process - Smooth function to smooth the spectra for final results.</li>
	</ol>
	</li>
</ol>

Micro-Embossed Fabrication of Nanocellulose Paper-Based Analytical Devices  

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T. H., Dien, N. D., Vu, X. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Facile+synthesis+of+silver/gold+alloy+nanoparticles+for+ultra-sensitive+rhodamine+B+detection.">Facile synthesis of silver/gold alloy nanoparticles for ultra-sensitive rhodamine B detection.</a> RSC Advances. 11 (35), 21475-21488 (2021).</li><li>Li, D., Li, D. W., Li, Y., Fossey, J. S., Long, Y. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclic+electroplating+and+stripping+of+silver+on+Au@SiO2+core/shell+nanoparticles+for+sensitive+and+recyclable+substrate+of+surface-enhanced+Raman+scattering.">Cyclic electroplating and stripping of silver on Au@SiO2 core/shell nanoparticles for sensitive and recyclable substrate of surface-enhanced Raman scattering.</a> Journal of Materials Chemistry. 20 (18), 3688-3693 (2010).</li><li>Sun, C. H., Wang, M. L., Feng, Q., Liu, W., Xu, C. X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surface-enhanced+Raman+scattering+(SERS)+study+on+Rhodamine+B+adsorbed+on+different+substrates.">Surface-enhanced Raman scattering (SERS) study on Rhodamine B adsorbed on different substrates.</a> Russian Journal of Physical Chemistry A. 89 (2), 291-296 (2015).</li></ol>

The authors have nothing to disclose.

The primary focus of this study is to develop a simple method for fabricating microchannels on nanopaper. An efficient embossing technique was devised using PTFE as the mold to address this challenge12. By optimizing the temperature and embossing pressure, a series of experiments were conducted to establish a reliable fabrication process for NanoPADs. Additionally, the use of a quick-reference table to adjust the applications of NanoPADs in different fields was demonstrated. Although this method i...

Recently, nanofibrillated cellulose (NFC) paper (nanopaper) has emerged as a highly promising substrate material for various applications such as flexible electronics, energy devices, and biomedicals1,2,3,4. Derived from natural plants, nanopaper is cost-effective, biocompatible, and biodegradable, making it an appealing alternative to traditional cellulose paper5,6. Its exceptional properties include an ultra-smooth surface with a surface roughness of less than 25 nm and a dense cellulose matrix structure, allowing for the creation of highly structured nanostructures7. Abundant hydroxyl groups of nanopaper contribute to its compact and tightly packed nanocellulose structure8. Nanopaper exhibits excellent optical transparency and minimal optical haze, making it well-suited for optical sensors. Additionally, its inherent hydrophilicity enables pump-free flow, even with its thick structure, providing autonomous fluid motion9,10. Nanocellulose has diverse applications in biological sensors, conductive electronic devices, cell culture platforms, supercapacitors, batteries, and more, showcasing its versatility and potential11,12. Particularly, nanocellulose is promising for paper-based analytical microfluidic devices (&#181;PADs), offering unique advantages over conventional chromatography paper.
In the past decade, &#181;PADs have achieved significant attention due to their affordability, biocompatibility, pump-free operation, and ease of production13,14. These devices have emerged as effective point-of-care diagnostic tools, particularly in resource-limited settings15,16,17. A significant advancement in this field was the development of wax printing, pioneered by George Whitesides18 and the Bingcheng Lin group19, enabling the creation of functional &#181;PADs by incorporating microchannels on chromatography paper. Subsequently, &#181;PADs rapidly evolved, and various biosensing techniques, including electrochemical methods20, chemiluminescence21, and enzyme-linked immunosorbent assay (ELISA)22,23,24, were successfully implemented for the detection of diverse biomarkers such as proteins25,26, DNAs27,28, RNAs29,30, and exosomes31. Despite these achievements, &#181;PADs still face challenges, including slow flow speeds and solvent evaporation.
Several methods have been proposed for creating microchannels on nanopaper32,33,34. One approach involves 3D printing sacrificial ingredients into the material, but it requires a hydrophobic coating that limits pump-free operation33. Another technique involves manually stacking channel layers between nanopaper sheets using glue, which is labor-intensive32. Alternatively, spray-coating nanocellulose fibers onto pre-patterned molds can create microchannels, but it involves time-consuming and expensive mold preparation34. Notably, these methods are limited to millimeter-scale microchannels, compromising the advantages of microfluidic devices regarding reagent volume consumption and integration. Developing a simple nanopaper microchannel patterning process with micrometer-scale resolution remains a challenge.
This study presents a unique nanopaper microchannel patterning method based on practical microembossing. The approach offers several advantages over existing methods, as it requires no expensive or specialized equipment, is simple, cost-effective, and highly accurate. A convex microchannel mold is fabricated by laser cutting a polytetrafluoroethylene (PTFE) film, known for its chemical inertness and nonstick properties. This mold is then used to emboss microchannels onto a nanopaper gel membrane. A second layer of nanopaper gel is applied on top to create closed hollow channels. Using this patterning technique, fundamental microfluidic devices on nanopaper are developed, including a laminar mixer and droplet generator. Additionally, the fabrication of surface-enhanced Raman microscopy (SERS) NanoPADs is demonstrated. In-situ creation of a silver nanoparticle-based SERS substrate is achieved by introducing two chemical reagents (AgNO3 and NaBH4) into the channels, resulting in a remarkable performance with low limits of detection (LODs).

<table><tbody><tr><td>AgNO&lt;sub&gt;3&lt;/sub&gt;&amp;nbsp;</td><td>Hushi (Shanghai, China)</td><td>7761-88-8</td><td>&gt;99%</td></tr><tr><td>Ethanol</td><td>Hushi (Shanghai, China)</td><td>64-17-5</td><td>&gt;99%</td></tr><tr><td>Hexadecane</td><td>Macklin (Shanghai, China)</td><td>544-76-3</td><td>&gt;99%</td></tr><tr><td>LabSpec software</td><td>Horiba (Japan)</td><td>LabSpec5</td><td></td></tr><tr><td>Melamine</td><td>Macklin (Shanghai, China)</td><td>108-78-1</td><td>&gt;99%</td></tr><tr><td>NaBH&lt;sub&gt;4&lt;/sub&gt;</td><td>Aladdin (Shanghai, China)</td><td>16940-66-2</td><td>&gt;99%</td></tr><tr><td>Origin lab software</td><td>OriginLab (USA)</td><td></td><td></td></tr><tr><td>Polyethylene terephthalate (PET)&amp;nbsp;</td><td>Myers Industries (Akron, USA)</td><td></td><td></td></tr><tr><td>Polytetrafluoroethylene films</td><td>Shenzhen Huashenglong plastic material Co., Ltd. (Shenzhen, China)</td><td></td><td>Teflon film</td></tr><tr><td>PVDF filter membrane</td><td>EMD Millipore Corporation (USA)</td><td>VVLP04700</td><td>pore size: 0.1 &amp;mu;m</td></tr><tr><td>Raman spectrometer</td><td>Horiba (Japan)</td><td>Xplo RA</td><td></td></tr><tr><td>Rhodamine B</td><td>Macklin (Shanghai, China)</td><td>81-88-9</td><td>&gt;95%</td></tr><tr><td>Scanning electron microscopy (SEM)</td><td>FEI(USA)</td><td>Scios 2 HiVac</td><td></td></tr><tr><td>Silicon wafer</td><td>Horiba (Japan)</td><td></td><td>diameter: 5 mm</td></tr><tr><td>TEMPO-oxidized NFC slurry</td><td>Tianjin University of Science and Technology</td><td></td><td>1.0 wt% solid, carboxylate level 2.0 mmol/g solid, average nanofiber diameter: 10 nm</td></tr></tbody></table>

microembossing: a convenient process for fabricating microchannels on nanocellulose paper-based microfluidics

Nanopaper, derived from nanofibrillated cellulose, has generated considerable interest as a promising material for microfluidic applications. Its appeal lies in a range of excellent&#160;qualities, including an exceptionally smooth surface, outstanding optical transparency, a uniform nanofiber matrix with nanoscale porosity, and customizable chemical properties. Despite the rapid growth of nanopaper-based microfluidics, the current techniques used to create microchannels on nanopaper, such as 3D printing, spray coating, or manual cutting and assembly, which are crucial for practical applications, still possess certain limitations, notably susceptibility to contamination. Furthermore, these methods are restricted to the production of millimeter-sized channels. This study introduces a straightforward process that utilizes convenient plastic micro-molds for simple microembossing operations to fabricate microchannels on nanopaper, achieving a minimum width of 200 &#181;m. The developed microchannel outperforms existing approaches, achieving a fourfold improvement, and can be fabricated within 45 min. Furthermore, fabrication parameters have been optimized, and a convenient quick-reference table is provided for application developers. The proof-of-concept for a laminar mixer, droplet generator, and functional nanopaper-based analytical devices (NanoPADs) designed for Rhodamine B sensing using surface-enhanced Raman spectroscopy was demonstrated. Notably, the NanoPADs exhibited exceptional performance with improved limits of detection. These outstanding results can be attributed to the superior optical properties of nanopaper and the recently developed accurate microembossing method, enabling the integration and fine-tuning of the NanoPADs.

A unique method for creating microchannel patterns on nanopaper has been devised utilizing the practical plastic micro-molds through the convenient microembossing technique. Notably, this method accomplishes microchannel patterning at a scale as small as 200 &#181;m, which represents a fourfold improvement compared to existing methods32,33,34. After fine-tuning the patterning parameters, the provided guidelines exhibit excellent...

Watch this Scientific Journal Video about Revolutionizing Microfluidics Through Microchannel Fabrication on Nanopaper at JoVE.com

Author Spotlight: Revolutionizing Microfluidics Through Microchannel Fabrication on Nanopaper 

This protocol describes a straightforward process that utilizes convenient plastic micro-molds for simple microembossing operations to fabricate microchannels on nanofibrillated cellulose paper, achieving a minimum width of 200 &#181;m.

Microembossing: A Convenient Process for Fabricating Microchannels on Nanocellulose Paper-Based Microfluidics

Nanopaper, derived from nanofibrillated cellulose, has generated considerable interest as a promising material for microfluidic applications. Its appeal ...

microembossing-convenient-process-for-fabricating-microchannels-on

Research

JoVE Journal

Engineering

1.5K Views.  Xi’an Jiaotong - Liverpool University. This protocol describes a straightforward process that utilizes convenient plastic micro-molds for simple microembossing operations to fabricate microchannels on nanofibrillated cellulose paper, achieving a minimum width of 200 µm.

Video: Author Spotlight: Revolutionizing Microfluidics Through Microchannel Fabrication on Nanopaper 

Manufacturing of Three-dimensionally Microstructured Nanocomposites through Microfluidic Infiltration

Microstructured composite beams reinforced with complex three-dimensionally (3D) patterned nanocomposite microfilaments are fabricated via nanocomposite in&#64257;ltration of 3D interconnected microfluidic networks. The manufacturing of the reinforced beams begins with the fabrication of microfluidic networks, which involves layer-by-layer deposition of fugitive ink filaments using a dispensing robot, filling the empty space between filaments using a low viscosity resin, curing the resin and finally removing the ink.&#160;Self-supported 3D structures with other geometries and many layers (e.g.&#160;a few hundreds layers) could be built using this method. The resulting tubular micro&#64258;uidic networks are then infiltrated with thermosetting nanocomposite suspensions containing nanofillers (e.g.&#160;single-walled carbon nanotubes), and subsequently cured. The infiltration is done by applying a pressure gradient between two ends of the empty network (either by applying a vacuum or vacuum-assisted microinjection). Prior to the infiltration, the nanocomposite suspensions are prepared by dispersing nanofillers into polymer matrices using ultrasonication and three-roll mixing methods. The nanocomposites (i.e.&#160;materials infiltrated) are then solidified under UV exposure/heat cure, resulting in a 3D-reinforced composite structure. The technique presented here enables the design of functional nanocomposite macroscopic products for microengineering applications such as actuators and sensors.

Three-dimensional (3D) microstructured composite beams are fabricated through the directed and localized infiltration of nanocomposites into 3D porous microfluidic networks. The flexibility of this manufacturing method enables the utilization of different thermosetting materials and nanofillers in order to achieve a variety of functional 3D reinforced nanocomposite macroscopic products.

Microstructured composite beams reinforced with complex three-dimensionally (3D) patterned nanocomposite microfilaments are fabricated via nanocomposite ...

Chemistry

Fabrication of the Thermoplastic Microfluidic Channels

In our lab, we have successfully isolated nucleic acids directly from microliter and submicroliter volumes of human blood, urine and stool using polymer/nanoparticle composite microscale lysis and solid phase extraction columns. The recovered samples are concentrated, small volume samples that are PCRable, without any additional cleanup. Here, we demonstrate how to fabricate thermoplastic microfluidic chips using hot embossing and heat sealing. Then, we demonstrate how to use in situ light directed surface grafting and polymerization through the sealed chip to form the composite solid phase columns. We demonstrate grafting and polymerization of a carbon nanotube/polymer composite column for bacterial cell lysis. We then show the lysis process followed by solid phase extraction of nucleic acids from the sample on chip using a silica/polymer composite column. The attached protocols contain detailed instructions on how to make both lysis and solid phase extraction columns.

Here we demonstrate how to fabricate thermoplastic microfluidic chips using hot embossing and heat sealing. Then we demonstrate how to use in situ light directed surface grafting and polymerization through the sealed chip to form the composite solid phase columns.

In our lab, we have successfully isolated nucleic acids directly from microliter and submicroliter volumes of human blood, urine and stool using ...

Biology

Microfabrication of Chip-sized Scaffolds for Three-dimensional Cell cultivation

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell cultivation. These technologies offer a wide spectrum of advantages not only for manufacturing but also for different applications. The size and shape of formed cell clusters can be influenced by the exact and reproducible architecture of the microfabricated scaffold and, therefore, the diffusion path length of nutrients and gases can be controlled.1 This is unquestionably a useful tool to prevent apoptosis and necrosis of cells due to an insufficient nutrient and gas supply or removal of cellular metabolites.
Our polymer chip, called CellChip, has the outer dimensions of 2 x 2 cm with a central microstructured area. This area is subdivided into an array of up to 1156 microcontainers with a typical dimension of 300 m edge length for the cubic design (cp- or cf-chip) or of 300 m diameter and depth for the round design (r-chip).2
So far, hot embossing or micro injection moulding (in combination with subsequent laborious machining of the parts) was used for the fabrication of the microstructured chips. Basically, micro injection moulding is one of the only polymer based replication techniques that, up to now, is capable for mass production of polymer microstructures.3 However, both techniques have certain unwanted limitations due to the processing of a viscous polymer melt with the generation of very thin walls or integrated through holes. In case of the CellChip, thin bottom layers are necessary to perforate the polymer and provide small pores of defined size to supply cells with culture medium e.g. by microfluidic perfusion of the containers.
In order to overcome these limitations and to reduce the manufacturing costs we have developed a new microtechnical approach on the basis of a down-scaled thermoforming process. For the manufacturing of highly porous and thin walled polymer chips, we use a combination of heavy ion irradiation, microthermoforming and track etching. In this so called "SMART" process (Substrate Modification And Replication by Thermoforming) thin polymer films are irradiated with energetic heavy projectiles of several hundred MeV introducing so-called "latent tracks" Subsequently, the film in a rubber elastic state is formed into three dimensional parts without modifying or annealing the tracks. After the forming process, selective chemical etching finally converts the tracks into cylindrical pores of adjustable diameter.

We present two processes for the microfabrication of porous polymer chips for three-dimensional cell cultivation. The first one is hot embossing combined with a solvent vapour welding process. The second one uses a recently developed microthermoforming process combined with ion track technology leading to a significant simplification of manufacture.

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell ...

Microfluidic Fabrication of Polymeric and Biohybrid Fibers with Predesigned Size and Shape

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and used to dictate the shape as well as the diameter of a core stream.&#160;Grooves in the top and bottom of a microfluidic channel were designed to direct the sheath fluid and shape the core fluid.&#160;By matching the viscosity and hydrophilicity of the sheath and core fluids, the interfacial effects are minimized and complex fluid shapes can be formed.&#160;Controlling the relative flow rates of the sheath and core fluids determines the cross-sectional area of the core fluid.&#160;Fibers have been produced with sizes ranging from 300 nm to ~1 mm, and fiber cross-sections can be round, flat, square, or complex as in the case with double anchor fibers. Polymerization of the core fluid downstream from the shaping region solidifies the fibers.&#160;Photoinitiated click chemistries are well suited for rapid polymerization of the core fluid by irradiation with ultraviolet light.&#160;Fibers with a wide variety of shapes have been produced from a list of polymers including liquid crystals, poly(methylmethacrylate), thiol-ene and thiol-yne resins, polyethylene glycol, and hydrogel derivatives. Minimal shear during the shaping process and mild polymerization conditions also makes the fabrication process well suited for encapsulation of cells and other biological components.

Two adjacent fluids passing through a grooved microfluidic channel can be directed to form a sheath around a prepolymer core; thereby&#160;determining both shape and cross-section. Photoinitiated polymerization, such as thiol click chemistry, is well suited for rapidly solidifying the core fluid into a microfiber with predetermined size and shape.

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and ...

Bioengineering

One-Step Approach to Fabricating Polydimethylsiloxane Microfluidic Channels of Different Geometric Sections by Sequential Wet Etching Processes

Polydimethylsiloxane (PDMS) materials are substantially exploited to fabricate microfluidic devices by using soft lithography replica molding techniques. Customized channel layout designs are necessary for specific functions and integrated performance of microfluidic devices in numerous biomedical and chemical applications (e.g., cell culture, biosensing, chemical synthesis, and liquid handling). Owing to the nature of molding approaches using silicon wafers with photoresist layers patterned by photolithography as master molds, the microfluidic channels commonly have regular cross sections of rectangular shapes with identical heights. Typically, channels with multiple heights or different geometric sections are designed to possess particular functions and to perform in various microfluidic applications (e.g., hydrophoresis is used for sorting particles and in continuous flows for separating blood cells6,7,8,9). Therefore, a great deal of effort has been made in constructing channels with various sections through multiple-step approaches like photolithography using several photoresist layers and assembly of different PDMS thin sheets. Nevertheless, such multiple-step approaches usually involve tedious procedures and extensive instrumentation. Furthermore, the fabricated devices may not perform consistently and the resulted experimental data may be unpredictable. Here, a one-step approach is developed for the straightforward fabrication of microfluidic channels with different geometric cross sections through PDMS sequential wet etching processes, that introduces etchant into channels of planned single-layer layouts embedded in PDMS materials. Compared to the existing methods for manufacturing PDMS microfluidic channels with different geometries, the developed one-step approach can significantly simplify the process to fabricate channels with non-rectangular sections or various heights. Consequently, the technique is a way of constructing complex microfluidic channels, which provides a fabrication solution for the advancement of innovative microfluidic systems.

Several methods are available for the fabrication of channels of non-rectangular sections embedded in polydimethylsiloxane microfluidic devices. Most of them involve multistep manufacturing and extensive alignment. In this paper, a one-step approach is reported for fabricating microfluidic channels of different geometric cross sections by polydimethylsiloxane sequential wet etching.

Polydimethylsiloxane (PDMS) materials are substantially exploited to fabricate microfluidic devices by using soft lithography replica molding techniques. ...

A Microfluidic Platform to Study Bioclogging in Porous Media

Bacterial biofilms are found in several environmental and industrial porous media, including soils and filtration membranes. Biofilms grow under certain flow conditions and can clog pores, thereby redirecting the local fluid flow. The ability of biofilms to clog pores, the so-called bioclogging, can have a tremendous effect on the local permeability of the porous medium, creating a pressure buildup in the system, and impacting the mass flow through it. To understand the interplay between biofilm growth and fluid flow under different physical conditions (e.g., at different flow velocities and pore sizes), in the present study, a microfluidic platform is developed to visualize biofilm development using a microscope under externally-imposed, controlled physical conditions. The biofilm-induced pressure buildup in the porous medium can be measured simultaneously using pressure sensors and, later, correlated with the surface coverage of the biofilm. The presented platform provides a baseline for a systematic approach to investigate bioclogging caused by biofilms in porous media under flow conditions and can be adapted to studying environmental isolates or multispecies biofilms.

The present protocol describes a microfluidic platform to study biofilm development in quasi-2D porous media by combining high-resolution microscopy imaging with simultaneous pressure difference measurements. The platform quantifies the influence of pore size and fluid flow rates in porous media on bioclogging.

Bacterial biofilms are found in several environmental and industrial porous media, including soils and filtration membranes. Biofilms grow under certain ...

Environment

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol adhesive through a metal stencil, the overall amount of adhesive used in assembling paper microfluidic devices can be significantly reduced. We show on a simple 4-layer planar paper microfluidic device that the optimal adhesive application technique and device construction style depends heavily on desired performance characteristics. By moderately increasing the overall area of a device, it is possible to dramatically decrease the wicking time and increase device success rates while also reducing the amount of adhesive required to keep the device together. Such adhesive application also causes the adhesive to form semi-permanent bonds instead of permanent bonds between paper layers, enabling single-use devices to be non-destructively disassembled after use. Nonplanar 3D origami devices also benefit from the semi-permanent bonds during folding, as it reduces the likelihood that unrelated faces may accidently stick together. Like planar devices, nonplanar structures see reduced wicking times with patterned adhesive application vs uniformly applied adhesive.

We demonstrate the use of patterned aerosol adhesives to construct 3D paper microfluidic devices. This method of adhesive application forms semi-permanent bonds between layers, enabling single-use devices to be non-destructively disassembled after use and to ease folding complex nonplanar structures.

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol ...

Fabrication of Three-dimensional Paper-based Microfluidic Devices for Immunoassays

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and directed within a layer of paper. Moreover, stacking multiple layers of patterned paper creates sophisticated three-dimensional microfluidic networks that can support the development of analytical and bioanalytical assays. Paper-based microfluidic devices are inexpensive, portable, easy to use, and require no external equipment to operate. As a result, they hold great promise as a platform for point-of-care diagnostics. In order to properly evaluate the utility and analytical performance of paper-based devices, suitable methods must be developed to ensure their manufacture is reproducible and at a scale that is appropriate for laboratory settings. In this manuscript, a method to fabricate a general device architecture that can be used for paper-based immunoassays is described. We use a form of additive manufacturing (multi-layer lamination) to prepare devices that comprise multiple layers of patterned paper and patterned adhesive. In addition to demonstrating the proper use of these three-dimensional paper-based microfluidic devices with an immunoassay for human chorionic gonadotropin (hCG), errors in the manufacturing process that may result in device failures are discussed. We expect this approach to manufacturing paper-based devices will find broad utility in the development of analytical applications designed specifically for limited-resource settings.

We detail a method to fabricate three-dimensional paper-based microfluidic devices for use in the development of immunoassays. Our approach to device assembly is a type of multilayer, additive manufacturing. We demonstrate a sandwich immunoassay to provide representative results for these types of paper-based devices.

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and ...

Fabrication of Micro-Patterned Chip with Controlled Thickness for High-Throughput Cryogenic Electron Microscopy

A major limitation for the efficient and high-throughput structure analysis of biomolecules using cryogenic electron microscopy (cryo-EM) is the difficulty of preparing cryo-EM samples with controlled ice thickness at the nanoscale. The silicon (Si)-based chip, which has a regular array of micro-holes with graphene oxide (GO) window patterned on a thickness-controlled silicon nitride (SixNy) film, has been developed by applying microelectromechanical system (MEMS) techniques. UV photolithography, chemical vapor deposition, wet and dry etching of the thin film, and drop-casting of 2D nanosheet materials were used for mass-production of the micro-patterned chips with GO windows. The depth of the micro-holes is regulated to control the ice thickness on-demand, depending on the size of the specimen for cryo-EM analysis. The favorable affinity of GO toward biomolecules concentrates the biomolecules of interest within the micro-hole during cryo-EM sample preparation. The micro-patterned chip with GO windows enables high-throughput cryo-EM imaging of various biological molecules, as well as inorganic nanomaterials.

A newly developed micro-patterned chip with graphene oxide windows is fabricated by applying microelectromechanical system techniques, enabling efficient and high-throughput cryogenic electron microscopy imaging of various biomolecules and nanomaterials.

A major limitation for the efficient and high-throughput structure analysis of biomolecules using cryogenic electron microscopy (cryo-EM) is the ...

Computer Numerical Control Micromilling of a Microfluidic Acrylic Device with a Staggered Restriction for Magnetic Nanoparticle-Based Immunoassays

Microfluidic systems have greatly improved immunoassay techniques. However, many microfabrication techniques require specialized, expensive, or complicated equipment, making fabrication costly and incompatible with mass production, which is one of the most important preconditions for point-of-care tests (POCT) to be adopted in low-resource settings. This work describes the fabrication process of an acrylic (polymethylmethacrylate, PMMA) device for nanoparticle-conjugated enzymatic immunoassay testing using the computer numerical control (CNC) micromilling technique. The functioning of the microfluidic device is shown by performing an immunoassay to detect a commercial antibody using lysozyme as a model antigen conjugated to 100 nm magnetic nanoparticles. This device integrates a physical staggered restriction of only 5 &#181;m in height, used to capture magnetic microparticles that make up a magnetic trap by placing an external magnet. In this way, the magnetic force on the immunosupport of conjugated nanoparticles is enough to capture them and resist flow drag. This microfluidic device is particularly suitable for low-cost mass production without the loss of precision for immunoassay performance.

Microfluidics is a powerful tool for the development of diagnostic tests. However, expensive equipment and materials, as well as laborious fabrication and handling techniques, are often required. Here, we detail the fabrication protocol of an acrylic microfluidic device for magnetic micro- and nanoparticle-based immunoassays in a low-cost and simple-to-use setting.