This work is supported by a fund from Bourns College of Engineering of University of California, Riverside. BK received a scholarship from the Lung-Wen Tsai Memorial Award in Mechanical Design.

1. Planar 4-layer Device (Stacked Layers) Construction
<ol>
	<li>Print arrays of each layer of the device9 onto each piece of filter paper using a solid ink printer.11,12 Place each filter paper on a hotplate at 170 &#176;C for 2 min. This will melt the wax-based ink and allow it to fully penetrate the thickness of the paper, forming hydrophobic barriers. 
	NOTE: The exact designs used are available as supplemental files.</li>
	<li>Remove filter paper from hotplate and allow it to cool to RT.</li>
	<li>Deposit 4 &#181;l of 5 mM dye (red: Allura Red; yellow: tartrazine; blue: erioglaucine disodium salt; green: 10:1 mix of tartrazine:erioglaucine disodium salt) in each branch (one color per branch) of layer 3 (third layer from the top of the completed device) using a micropipette.</li>
	<li>Begin with the bottom-most layer. Clamp the filter paper between the stencil and a stiff backing, such as a piece of plate glass, using binder clips, or another similar temporary method. Ensure that the stencil is flat against the paper. This will minimize any spray shadows cast by the stencil onto the paper.</li>
	<li>Apply adhesive (see list of materials and equipment) with an approximately 1.33 sec (a four-count at 180 bpm) spray from about 24 cm.9,10 During this time, move the can of adhesive across the stencil at a medium pace. Too slow of travel across the stencil will cause adhesive to accumulate on the stencil itself, clogging it. Too rapid of travel will fail to deposit sufficient adhesive on the paper. Four passes during this time (up-down-up-down) are sufficient in preventing spray shadows.</li>
	<li>Remove stencil and place the next layer of the device (numbered layers are available as supplemental files) atop the freshly sprayed layer, aligning the edges of the paper. Firmly press the two layers together.</li>
	<li>Replace stencil and repeat the spraying process for each layer of the device. Remove the stack of devices and place packing tape across the bottom layer. This prevents any fluid leakage from the device. Cut individual devices from the sheet using scissors, following the edge of the printed region.</li>
</ol>
2. Planar 4-layer Device (Origami Folded Layers) Construction
<ol>
	<li>Print sheets containing all layers of the device onto filter paper using a solid ink printer. Place filter paper on a hotplate at 170 &#176;C for 2 min. Remove filter paper from hotplate and allow it to cool to RT. 
	NOTE: The exact designs used are available as supplemental files.</li>
	<li>Deposit 4 &#181;l of 5 mM dye (red: Allura Red; yellow: tartrazine; blue: erioglaucine disodium salt; green: 10:1 mix of tartrazine:erioglaucine disodium salt) in each branch (one color per branch) of layer 3 (third layer from the top of the completed device) via micropipette.</li>
	<li>Clamp the sheet of devices between the stencil and a stiff backing, such as a piece of plate glass, using binderclips, or another similar temporary method. Ensure that the stencil is flat against the paper.</li>
	<li>Apply adhesive (see list of materials and equipment) with an approximately 1.33 sec (a four-count at 180 bpm) spray from about 24 cm. Four passes during this time (up-down-up-down) are sufficient in preventing spray shadows.</li>
	<li>Remove stencil and turn the sheet over. Replace stencil and spray back side of the paper. Remove the sheet of devices and begin folding in an accordion pleat, as depicted in Figure 1. Cut each device out from sheet using scissors, following the edge of the printed region. Place packing tape across the bottom layer.</li>
</ol>
3. Nonplanar (Origami) Device Construction
<ol>
	<li>Print device (Figure 2A) onto filter paper using a solid ink printer and place the filter paper on a hot plate at 170 &#176;C for 2 min. Remove device from hotplate and allow it to cool to RT. 
	NOTE: The exact designs used are available as supplemental files.</li>
	<li>Print crease pattern (Figure 2C) onto printer paper using a solid ink printer and cut to the size of the filter paper. Place crease pattern on a hotplate at 170 &#176;C for 2 min, to melt the wax, causing the pattern to be visible from both sides of the paper. Remove crease pattern from hotplate and allow it to cool to RT.</li>
	<li>Align the edges of the crease pattern to the edges of the paper containing the channel patterns and attach the two pieces of paper using binder clips, or another similar temporary method.</li>
	<li>Trace the crease pattern with a blunt stylus, applying enough force that marks appear on the device sheet, but not so hard that the crease pattern paper rips. If that occurs, the device risks being damaged. Precreasing causes the paper to fold much more easily and allows for greater accuracy and precision in folding.</li>
	<li>Begin folding the device with mountain and valley folds according to the crease pattern. Once the adhesive has been applied, the entire device must be assembled very quickly, so folding the device as much as possible before adhesive application is very helpful.</li>
	<li>Once the device is folded, unfold the device to expose the portions of the device that require adhesive. Cut out masks (Figure 2D) that limit where on the device adhesive may be applied, using a razor blade.</li>
	<li>Clamp the device between the stencil and mask and a stiff backing, such as a piece of plate glass. Ensure that the stencil is flat against the device. Apply adhesive (see list materials and equipment) with an approximately 1.33 sec (a four-count at 180 bpm) spray from about 24 cm. Four passes during this time (up-down-up-down) are sufficient in preventing spray shadows. Remove stencil and turn the sheet over. Replace stencil and mask and spray back side of the paper.</li>
	<li>Immediately remove device from stencil and begin folding the device. Once the device is completely folded, apply pressure to the adhesive containing portion until the adhesive has dried. 
	NOTE: The drying time of the adhesive is very sensitive to ambient humidity, so in locations with low humidity, folding in a humidity controlled chamber allows more time to fold the device.</li>
</ol>
4. Wicking Test for 4-layer Devices
<ol>
	<li>Randomly select 20 devices, previously assembled according to the above protocols. Place devices in a location shielded from any wind or breezes to minimize evaporation. Deposit 40 &#181;l water at the inlet of each device. Record the time it takes for each device to have all of its outlets completely filled with dye.</li>
</ol>
5. Origami Wicking Comparison
<ol>
	<li>Construct two origami peacocks &#8212; one according to the above protocol (Section 3), and the other without the use of a stencil during adhesive application.</li>
	<li>Insert one end of a small paper lead (approximately 5 mm wide by 5 cm long) into the body of each peacock.</li>
	<li>Place both peacocks in a chamber kept at a high relative humidity (&#62;90%) to minimize evaporation. Place each leg and lead of each peacock into a container filled with 5 mM dye (red: Allura Red, yellow: tartrazine, blue: erioglaucine disodium salt). Record wicking process with a digital camera.</li>
</ol>

<ol><li>Li, X., Ballerini, D. R., Shen, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+perspective+on+paper-based+microfluidics%3A+Current+status+and+future+trends%5BTitle%5D)+AND+%22Biomicrofluidics%22%5BJournal%5D)">A perspective on paper-based microfluidics: Current status and future trends.</a> Biomicrofluidics. 6, 11301-11313 (2012).</li>
<li>Yetisen, A. K., Akram, M. S., Lowe, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Paper-based+microfluidic+point-of-care+diagnostic+devices%5BTitle%5D)+AND+%22Lab+Chip%22%5BJournal%5D)">Paper-based microfluidic point-of-care diagnostic devices.</a> Lab Chip. 13, 2210-2251 (2013).</li>
<li>Cate, D. M., Adkins, J. A., Mettakoonpitak, J., Henry, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Recent+developments+in+paper-based+microfluidic+devices%5BTitle%5D)+AND+%22Anal+Chem%22%5BJournal%5D)">Recent developments in paper-based microfluidic devices.</a> Anal Chem. 87, 19-41 (2015).</li>
<li>Martinez, A. W., Phillips, S. T., Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Three-dimensional+microfluidic+devices+fabricated+in+layered+paper+and+tape%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">Three-dimensional microfluidic devices fabricated in layered paper and tape.</a> Proc Natl Acad Sci U S A. 105, 19606-19611 (2008).</li>
<li>Fu, E., Ramsey, S. A., Kauffman, P., Lutz, B., Yager, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Transport+in+two-dimensional+paper+networks%5BTitle%5D)+AND+%22Microfluid+Nanofluidics%22%5BJournal%5D)">Transport in two-dimensional paper networks.</a> Microfluid Nanofluidics. 10, 29-35 (2011).</li>
<li>Govindarajan, A. V., Ramachandran, S., Vigil, G. D., Yager, P., Bohringer, K. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+low+cost+point-of-care+viscous+sample+preparation+device+for+molecular+diagnosis+in+the+developing+world%3B+an+example+of+microfluidic+origami%5BTitle%5D)+AND+%22Lab+Chip%22%5BJournal%5D)">A low cost point-of-care viscous sample preparation device for molecular diagnosis in the developing world; an example of microfluidic origami.</a> Lab Chip. 12, 174-181 (2012).</li>
<li>Schilling, K. M., Jauregui, D., Martinez, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Paper+and+toner+three-dimensional+fluidic+devices%3A+programming+fluid+flow+to+improve+point-of-care+diagnostics%5BTitle%5D)+AND+%22Lab+Chip%22%5BJournal%5D)">Paper and toner three-dimensional fluidic devices: programming fluid flow to improve point-of-care diagnostics.</a> Lab Chip. 13, 628-631 (2013).</li>
<li>Liu, H., Crooks, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Three-dimensional+paper+microfluidic+devices+assembled+using+the+principles+of+origami%5BTitle%5D)+AND+%22J+Am+Chem+Soc%22%5BJournal%5D)">Three-dimensional paper microfluidic devices assembled using the principles of origami.</a> J Am Chem Soc. 133, 17564-17566 (2011).</li>
<li>Lewis, G. G., DiTucci, M. J., Baker, M. S., Phillips, S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((High+throughput+method+for+prototyping+three-dimensional%2C+paper-based+microfluidic+devices%5BTitle%5D)+AND+%22Lab+Chip%22%5BJournal%5D)">High throughput method for prototyping three-dimensional, paper-based microfluidic devices.</a> Lab Chip. 12, 2630-2633 (2012).</li>
<li>Kalish, B., Tsutsui, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Patterned+adhesive+enables+construction+of+nonplanar+three-dimensional+paper+microfluidic+circuits%5BTitle%5D)+AND+%22Lab+Chip%22%5BJournal%5D)">Patterned adhesive enables construction of nonplanar three-dimensional paper microfluidic circuits.</a> Lab Chip. 14, 4354-4361 (2014).</li>
<li>Carrilho, E., Martinez, A. W., Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Understanding+wax+printing%3A+a+simple+micropatterning+process+for+paper-based+microfluidics%5BTitle%5D)+AND+%22Anal+Chem%22%5BJournal%5D)">Understanding wax printing: a simple micropatterning process for paper-based microfluidics.</a> Anal Chem. 81, 7091-7095 (2009).</li>
<li>Lu, Y., Shi, W., Jiang, L., Qin, J., Lin, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rapid+prototyping+of+paper-based+microfluidics+with+wax+for+low-cost%2C+portable+bioassay%5BTitle%5D)+AND+%22Electrophoresis%22%5BJournal%5D)">Rapid prototyping of paper-based microfluidics with wax for low-cost, portable bioassay.</a> Electrophoresis. 30, 1497-1500 (2009).</li>
<li>Maekawa, J. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aMaekawa%2C+J+%22Genuine+Japanese+origami%22">Genuine Japanese origami</a>. , Dover Publications, Inc.. Dover edition (2012).</li>
<li>Schonhorn, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+device+architecture+for+three-dimensional%2C+patterned+paper+immunoassays%5BTitle%5D)+AND+%22Lab+Chip%22%5BJournal%5D)">A device architecture for three-dimensional, patterned paper immunoassays.</a> Lab Chip. 14, 4653-4658 (2014).</li>
<li>Guan, J. J., He, H. Y., Hansford, D. J., Lee, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Self-folding+of+three-dimensional+hydrogel+microstructures%5BTitle%5D)+AND+%22J+Phys+Chem+B%22%5BJournal%5D)">Self-folding of three-dimensional hydrogel microstructures.</a> J Phys Chem B. 109, 23134-23137 (2005).</li>
</ol>

The authors have nothing to disclose.

The above protocols use perforated metal sheets as stencils for applying aerosol adhesives to construct planar and nonplanar 3D paper microfluidic devices. In planar devices, this has the advantage of allowing devices to be completely unfolded after the adhesive has dried without destroying the device. In other adhesive based construction techniques, this is almost impossible, although, some designs allow for partial destructive disassembly by unpeeling two halves held together with a removable adhesive.14 Adhesiveless construction does allow devices to be unfolded after use, but requires custom clamps or housings for each device.8
In devices with primarily lateral wicking, adhesives can significantly slow wicking. By patterning the adhesive, the amount of adhesive applied to the wicking regions can be significantly reduced, limiting any potential wicking interference. Devices with predominantly vertical wicking also exhibit similarly slow wicking caused by adhesive, although to a much lesser extent. Design of a stencil that completely blocks out all wicking regions, limiting adhesive application to hydrophobic regions only, may eliminate any potential wicking interferences, but may also add considerable alignment time to the construction process.
In nonplanar devices, the patterned adhesive dramatically eases folding, as the quantity of adhesive that is applied to the paper is decreased, making folding significantly easier than with a uniformly applied adhesive layer. Paper completely covered in adhesive is far more difficult to fold when any incidental contact between different areas of the paper causes adhesion that must be undone before continuing.
For planar 3D multilayer devices that have a large wicking area relative to the hydrophobic area, origami folding paired with an aerosol adhesive is likely not the optimal construction technique due to the adhesive&#39;s inability to hold wetted paper layers together while overcoming the tendency of the creases to unfold. Devices with designs that include sufficient hydrophobic borders will increase the success rate of origami-folded devices. Using a stronger bond-strength adhesive may also help to solve this issue, preventing water from weakening the paper-adhesive bond.
Stacked layer devices overall performed better, as they lack creases, which tend to unfold the device. Further, the use of a stencil during adhesive application reduces the total amount of adhesive applied, dramatically reducing the time required for fluid to wick between layers.
In designing nonplanar 3D paper microfluidic devices, there are a number of issues to consider. It is important to compare the crease pattern of the folded device to the layout of the channels, as placing channels along a crease will force the crease open upon water imbibition, due to swelling cellulose fibers. Depending on the design of the specific device, though, this may or may not be desired behavior. Device storage at ambient conditions is not favorable to device viability,10 thus long-term storage under dry air is recommended to prevent the adhesive bond between layers from weakening.
As previously noted by Lewis et al.,9 the use of aerosol adhesives provide an efficient means to rapidly produce large quantities of 3D paper microfluidic devices. By patterning such adhesives, new devices can be more rapidly developed that take advantage of being able to be unfolded after use.
Further, patterning enables the construction and development of nonplanar 3D paper microfluidic devices. Such devices are expected to be able to provide functionality not previously found in planar paper microfluidics, such as integrated actuation and sensing. For example, actuation can be achieved by creating a bilayer from a water reactive polymer film15 and a patterned paper substrate. In a device constructed from such a bilayer, actuation would be generated when water wicks along the device&#39;s channels and interacts with the film. Once the film dries, the device would return to its initial configuration, leaving it ready to be used again.

In recent years, paper microfluidics has garnered considerable popularity for its potential to provide low-cost point of care (POC) diagnostic devices.1-3 POC devices offer functionality similar to those of lab-based tests in a format that allows results to be obtained relatively quickly. POC devices made from paper are low-cost, lightweight, and easy-to-use alternatives to expensive microfluidic chips and miniaturized laboratories, making them ideal for use in resource-limited settings. The most common paper microfluidic devices are one-dimensional lateral flow devices, but planar three-dimensional (3D) paper microfluidic devices hold promise to provide multiplexed diagnostic devices4 that take up a much smaller footprint than would be required by a 2D device5 and correspondingly use a smaller sample volume.
Initially, planar 3D paper microfluidic devices were assembled individually, layer-by-layer with patterned paper layers alternating with laser-cut double-sided tape. Carefully aligned holes cut in the tape layer were filled with cellulose powder to ensure inter-layer fluid transport.4 A number of alternate methods were subsequently developed,6-9 each improving different aspects of the devices. In particular, by eschewing adhesives, devices could be folded via origami techniques with layers held together by an external clamp.8 This eliminates any potential adhesive interference in a diagnostic test and allows the device to be unfolded post-use, potentially allowing even smaller sample volumes by displaying results internally. Alternatively, by using an aerosol adhesive applied between each paper layer, sheets of devices could be assembled simultaneously, without time-consuming patterning and alignment of tape.9
However, by applying an aerosol adhesive through a stencil, it is possible to gain the benefit of both of these techniques. By spraying the adhesive through a stencil, only a fraction of the adhesive is applied to the device, minimizing any potential interference with interlayer fluid transfer. Additionally, with careful stencil selection, a pattern of adhesive can be applied that results in semi-permanent adhesive bonding, allowing devices to be unfolded after use, while still providing sufficient interlayer contact to allow fluid to wick between layers.
Finally, applying aerosol adhesives through a stencil eases the construction of nonplanar 3D paper microfluidic devices, by minimizing the amount of adhesive applied to adjacent faces that may require frequent folding and unfolding during construction.10 Additionally, the use of patterned adhesive enables device to be unfolded after use for more convenient storage. Nonplanar 3D paper microfluidic devices are expected to be used for tasks that would otherwise be impossible in a planar 3D device. Figure 1 depicts the general process flow used to construct both planar and nonplanar 3D devices.

<table><tbody><tr><td>Camera</td><td>Nikon</td><td>D5100</td><td></td></tr><tr><td>Solid-ink printer</td><td>Xerox</td><td>ColorQube 8880</td><td></td></tr><tr><td>Hotplate</td><td>Torrey Pines</td><td>HS60</td><td></td></tr><tr><td>Humidity chamber</td><td>Electro-Tech Systems</td><td>5503-E</td><td></td></tr><tr><td>Spray adhesive</td><td>3M</td><td>62497749309</td><td>Super 77 (16.75 oz can)</td></tr><tr><td>Filter paper</td><td>Whatman</td><td>Grade 4</td><td></td></tr><tr><td>Perforated steel sheet</td><td>MetalsDepot</td><td>PS16116</td><td></td></tr><tr><td>Tartrazine</td><td>Sigma-Aldritch</td><td>T0388</td><td></td></tr><tr><td>Allura Red</td><td>Sigma-Aldritch</td><td>458848</td><td></td></tr><tr><td>Erioglaucine disodium salt</td><td>Sigma-Aldritch</td><td>861146</td><td></td></tr></tbody></table>

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.

The 4-layer device tests were performed in a sealed chamber, shielding them from any wind or breezes that might cause excessive evaporation of the limited deposited fluid volume. The majority of the wicking in the 4-layer devices is in the middle layers of the device, so differences in wicking speeds due to evaporation were expected to be minimal. Additionally, there is minimal lateral wicking, with only 13 mm between the inlet and any individual outlet, suggesting that variations in wicking times are likely due to vertical, interlayer fluid transfer. Average wicking times and success rates for 4-layer devices constructed with different amounts of applied adhesive are shown in Table 1.
In stacked devices, uniform adhesive coverage resulted in relatively high success rates that decreased as we increased the quantity of adhesive. Patterned adhesive coverage resulted in very low success rates when adhesive was only applied to one side, but had much higher success rates and faster wicking times when the patterned adhesive was applied to both sides. Typical successes are depicted in Figure 3A. There are several potential explanations for this observed behavior, any combination of which may be applicable. The applied adhesive may be physically blocking, either partially or completely, the pores at the surface of the paper, resulting in a smaller effective contact area between paper layers. Also, the adhesive itself may act as another porous substrate, so heavier coatings of adhesive result in a thicker adhesive layer that fluid must wick through, leading to longer wicking times. Patterning the adhesive, on the other hand, creates adhesive &#39;dots&#39; that only partially occlude the contact areas, allowing more fluid to wick from paper layer to paper layer directly, which decreases wicking times. However, this very reduction in adhesive coverage also decreases the strength of the adhesive bond between paper layers, resulting in decreased success rates when swelling fibers and unfolding creases cause layers to separate enough that they are no longer in contact. By doubling the size of the border around the channels (increasing the overall device area by ~30%), success rates for both single- and dual-sided adhesive applications increased. A comparison between the two sizes is shown in Figure 4. Typical stacked device failure was characterized by outlets that failed to completely fill with dye, or took longer than 5 min&#160;to fill. This is depicted in Figure 3B.
In origami folded devices, uniform adhesive coverage resulted in low success rates with complete failure resulting when applying the equivalent amount of adhesive present in the stacked, uniform, single-sided adhesive devices. Patterned adhesive coverage resulted in much lower success rates; however, this decrease was offset by using slightly larger devices that had 3 mm borders. Typical origami device failure was characterized by outlets that failed to fill with any amount of dye. These outlets were exclusively located along the two sides of the device that contained the creases. This is depicted in Figure 3C.
The masses of adhesive applied under different spray methods are shown in Table 2. The above-described spray duration of 1.33 sec (a four count at 180 bpm) deposits 0.26 mg/cm2 (dry mass) of adhesive when sprayed uniformly across the sheet of devices, while only depositing 0.02 mg/cm2 (dry mass) when sprayed through a stencil that was 23% open.
In nonplanar 3D structures, uniform adhesive coverage resulted in more difficult folding, as adjacent faces prematurely stuck together. The layers inside the structure could not be unfolded once the adhesive dried, and attempts to do so resulted in shredded paper. Patterned adhesive coverage made folding much easier, as any accidental adhesion was easily undone. Once the adhesive dried, the layers could be pulled apart without any ripping or tearing of the paper. Both methods of adhesive application resulted in devices that successfully routed liquid the length of their channels and without mixing; however, the device with uniformly applied adhesive was noticeably slower. A time-lapse of this wicking is shown in Figure 5. Wicking was performed in a humidity controlled chamber kept at &#62;90% relative humidity to minimize evaporation, as evaporation increases with decreasing relative humidity. Due to the long channels present in this design, up to 165 mm long, evaporation can significantly increase the wicking time, even with an infinite fluid reservoir.
<img alt="Figure 1" src="/files/ftp_upload/53805/53805fig1.jpg" /> 
Figure 1. Device Fabrication Process Flow. (A) Stacked device fabrication. (B) Origami device fabrication. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53805/53805fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53805/53805fig2.jpg" /> 
Figure 2. Peacock Patterns. (A) Channel pattern, where black indicates hydrophobic regions. (B) Arrows indicate the path taken by each dye. Circles indicate the point of contact between layers and the dotted lines indicate the vertical wicking paths. The length of each channel from its respective inlet to edge of the tail is indicated in millimeters. Channel widths averaged between 2 and 3 mm in the tail region. (C) Crease Pattern (modified from13). Red lines correspond to mountain folds in the final structure; black lines correspond to valley folds; blue lines correspond to creases that are not folded in the final structure, but aid in preliminary folding steps. (D) Masks placed between the origami device and the metal stencil during adhesive application, where the white portions are removed. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53805/53805fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53805/53805fig3.jpg" /> 
Figure 3. Typical Successes and Failures. (A) Typical Success &#8212; all outlets completely filled with dye. (B) Typical stacked failure &#8212; outlets that failed had no apparent pattern in their distribution. (C) Typical origami failure &#8212; all outlets that failed to fill were located along the left-most or right-most column, closest to the creases. All scale bars are 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53805/53805fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53805/53805fig4.jpg" /> 
Figure 4. Device Size Comparison. (A) Smaller device (1.6 mm border). (B) Larger device (3 mm border). All scale bars are 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53805/53805fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/53805/53805fig5.jpg" /> 
Figure 5. Time Lapse of Origami Peacock. Left: uniform adhesive coverage. Right: patterned adhesive coverage. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53805/53805fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Device Style</td>
			<td>Adhesive Type (Duration/Border/Sides)</td>
			<td>Average &#177; SD (sec)</td>
			<td>Success Rate</td>
		</tr>
		<tr>
			<td>Origami</td>
			<td>Uniform (1.33 sec / 1.6 mm / Double)</td>
			<td>44 &#177; 14</td>
			<td>45%</td>
		</tr>
		<tr>
			<td></td>
			<td>Uniform (0.67 sec / 1.6 mm / Double)</td>
			<td>0 &#177; 0</td>
			<td>0%</td>
		</tr>
		<tr>
			<td></td>
			<td>Patterned (1.33 sec / 1.6 mm / Double)</td>
			<td>41 &#177; 13</td>
			<td>15%</td>
		</tr>
		<tr>
			<td></td>
			<td>Patterned (1.33 sec / 3 mm / Double)</td>
			<td>64 &#177; 50</td>
			<td>40%</td>
		</tr>
		<tr>
			<td>Stacked</td>
			<td>Uniform (1.33 sec / 1.6 mm / Single)</td>
			<td>152 &#177; 66</td>
			<td>80%</td>
		</tr>
		<tr>
			<td></td>
			<td>Uniform (1.33 sec / 1.6 mm / Double)</td>
			<td>119 &#177; 68</td>
			<td>60%</td>
		</tr>
		<tr>
			<td></td>
			<td>Patterned (1.33 sec / 1.6 mm / Single)</td>
			<td>164 &#177; 75</td>
			<td>25%</td>
		</tr>
		<tr>
			<td></td>
			<td>Patterned (1.33 sec / 1.6 mm / Double)</td>
			<td>81 &#177; 25</td>
			<td>80%</td>
		</tr>
		<tr>
			<td></td>
			<td>Patterned (1.33 sec / 3 mm / Single)</td>
			<td>116 &#177; 63</td>
			<td>85%</td>
		</tr>
		<tr>
			<td></td>
			<td>Patterned (1.33 sec / 3 mm / Double)</td>
			<td>80 &#177; 55</td>
			<td>100%</td>
		</tr>
	</tbody>
</table>
Table 1. Four Layer Device Performance. Average wicking time and success rates for different adhesive application conditions. N=20.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Adhesive Coverage</td>
			<td>Spray Duration (sec)</td>
			<td>Average Mass &#177; SD (mg/cm&#178;)</td>
		</tr>
		<tr>
			<td>Uniform</td>
			<td>1.33</td>
			<td>0.26 &#177; 0.05</td>
		</tr>
		<tr>
			<td>Uniform</td>
			<td>0.67</td>
			<td>0.14 &#177; 0.03</td>
		</tr>
		<tr>
			<td>Patterned</td>
			<td>1.33</td>
			<td>0.02 &#177; 0.01</td>
		</tr>
		<tr>
			<td>None</td>
			<td>0</td>
			<td>-0.01 &#177; 0</td>
		</tr>
	</tbody>
</table>
Table 2. Applied Adhesive Amounts. Average adhesive thickness (dry mass) applied over a 9x9 cm square under different spray conditions. N=10.

Watch this Scientific Journal Video about Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices at JoVE.com

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

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 ...

using-adhesive-patterning-to-construct-3d-paper-microfluidic-devices

Nanyang Technological University

Research

JoVE Journal

Bioengineering

7.6K Views.  University of California, Riverside.  We demonstreren het gebruik van patroon aerosol kleefstoffen 3D papier microfluïdische inrichtingen te construeren. Deze methode van lijm aanvraagformulieren semi-permanente banden tussen de lagen, waardoor eenmalig gebruik apparaten op non-destructief gedemonteerd na gebruik en te vouwen complexe nonplanar structuren te verlichten zijn. 

Video: Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

Origami Inspired Self-assembly of Patterned and Reconfigurable Particles

There are numerous techniques such as photolithography, electron-beam lithography and soft-lithography that can be used to precisely pattern two dimensional (2D) structures. These technologies are mature, offer high precision and many of them can be implemented in a high-throughput manner. We leverage the advantages of planar lithography and combine them with self-folding methods1-20 wherein physical forces derived from surface tension or residual stress, are used to curve or fold planar structures into three dimensional (3D) structures. In doing so, we make it possible to mass produce precisely patterned static and reconfigurable particles that are challenging to synthesize.
In this paper, we detail visualized experimental protocols to create patterned particles, notably, (a) permanently bonded, hollow, polyhedra that self-assemble and self-seal due to the minimization of surface energy of liquefied hinges21-23 and (b) grippers that self-fold due to residual stress powered hinges24,25. The specific protocol described can be used to create particles with overall sizes ranging from the micrometer to the centimeter length scales. Further, arbitrary patterns can be defined on the surfaces of the particles of importance in colloidal science, electronics, optics and medicine. More generally, the concept of self-assembling mechanically rigid particles with self-sealing hinges is applicable, with some process modifications, to the creation of particles at even smaller, 100 nm length scales22, 26 and with a range of materials including metals21, semiconductors9 and polymers27. With respect to residual stress powered actuation of reconfigurable grasping devices, our specific protocol utilizes chromium hinges of relevance to devices with sizes ranging from 100 &mu;m to 2.5 mm. However, more generally, the concept of such tether-free residual stress powered actuation can be used with alternate high-stress materials such as heteroepitaxially deposited semiconductor films5,7 to possibly create even smaller nanoscale grasping devices.

We describe experimental details of the synthesis of patterned and reconfigurable particles from two dimensional (2D) precursors. This methodology can be used to create particles in a variety of shapes including polyhedra and grasping devices at length scales ranging from the micro to centimeter scale.

There are numerous techniques such as photolithography, electron-beam lithography and soft-lithography that can be used to precisely pattern two ...

Chemistry

Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as growth factors. Here, we outline a method to create opposing gradients of adhesive and soluble cues in a microfluidic chamber, which is compatible with live cell imaging. A copolymer of poly-L-lysine and polyethylene glycol (PLL-PEG) is employed to passivate glass coverslips and prevent non-specific adsorption of biomolecules and cells. Next, microcontact printing or dip pen lithography are used to create tracks of streptavidin on the passivated surfaces to serve as anchoring points for the biotinylated peptide arginine-glycine-aspartic acid (RGD) as the adhesive cue. A microfluidic device is placed onto the modified surface and used to create the gradient of adhesive cues (100% RGD to 0% RGD) on the streptavidin tracks. Finally, the same microfluidic device is used to create a gradient of a chemoattractant such as fetal bovine serum (FBS), as the soluble cue in the opposite direction of the gradient of adhesive cues.

A method for the assembly of adhesive and soluble gradients in a microscopy chamber for live cell migration studies is described. The engineered environment combines antifouling surfaces and adhesive tracks with solution gradients and therefore allows one to determine the relative importance of guidance cues.

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as ...

Cell Patterning on Photolithographically Defined 
Parylene-C: SiO2 Substrates

Cell patterning platforms support broad research goals, such as construction of predefined in vitro neuronal networks and the exploration of certain central aspects of cellular physiology. To easily combine cell patterning with Multi-Electrode Arrays (MEAs) and silicon-based &#8216;lab on a chip&#8217; technologies, a microfabrication-compatible protocol is required. We describe a method that utilizes deposition of the polymer parylene-C on SiO2&#160;wafers. Photolithography enables accurate and reliable patterning of parylene-C at micron-level resolution. Subsequent activation by immersion in fetal bovine serum (or another specific activation solution) results in a substrate in which cultured cells adhere to, or are repulsed by, parylene or SiO2 regions respectively. This technique has allowed patterning of a broad range of cell types (including primary murine hippocampal cells, HEK 293 cell line, human neuron-like teratocarcinoma cell line, primary murine cerebellar granule cells, and primary human glioma-derived stem-like cells). Interestingly, however, the platform is not universal; reflecting the importance of cell-specific adhesion molecules. This cell patterning process is cost effective, reliable, and importantly can be incorporated into standard microfabrication (chip manufacturing) protocols, paving the way for integration of microelectronic technology.

Cell Patterning on Photolithographically Defined 
Parylene-C: SiO2 Substrates

This protocol describes a microfabrication-compatible method for cell patterning on SiO2. A predefined parylene-C design is photolithographically printed on SiO2 wafers. Following incubation with serum (or other activation solution) cells adhere specifically to (and grow according to the conformity of) underlying parylene-C, whilst being repulsed by SiO2 regions.

Cell patterning platforms support broad research goals, such as construction of predefined in vitro neuronal networks and the exploration of certain ...

Microfluidic Genipin Deposition Technique for Extended Culture of Micropatterned Vascular Muscular Thin Films

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular behaviors on disease-relevant time scales. Previously, microcontact printing has been used to fabricate two-dimensional functional arterial mimics through patterning of extracellular matrix protein as guidance cues for tissue organization. Vascular muscular thin films utilized these mimics to assess functional contractility. However, the microcontact printing fabrication technique used typically incorporates hydrophobic PDMS substrates. As the tissue turns over the underlying extracellular matrix, new proteins must undergo a conformational change or denaturing in order to expose hydrophobic amino acid residues to the hydrophobic PDMS surfaces for attachment, resulting in altered matrix protein bioactivity, delamination, and death of the tissues.
Here, we present a microfluidic deposition technique for patterning of the crosslinker compound genipin. Genipin serves as an intermediary between patterned tissues and PDMS substrates, allowing cells to deposit newly-synthesized extracellular matrix protein onto a more hydrophilic surface and remain attached to the PDMS substrates. We also show that extracellular matrix proteins can be patterned directly onto deposited genipin, allowing dictation of engineered tissue structure. Tissues fabricated with this technique show high fidelity in both structural alignment and contractile function of vascular smooth muscle tissue in a vascular muscular thin film model. This technique can be extended using other cell types and provides the framework for future study of chronic tissue- and organ-level functionality.

We present a method for microfluidic deposition of patterned genipin and fibronectin on PDMS substrates, allowing extended viability of vascular smooth muscle cell-dense tissues. This tissue fabrication method is combined with previous vascular muscular thin film technology to measure vascular contractility over disease-relevant time courses.

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular ...

Construction of Modular Hydrogel Sheets for Micropatterned Macro-scaled 3D Cellular Architecture

Hydrogels can be patterned at the micro-scale using microfluidic or micropatterning technologies to provide an in vivo-like three-dimensional (3D) tissue geometry. The resulting 3D hydrogel-based cellular constructs have been introduced as an alternative to animal experiments for advanced biological studies, pharmacological assays and organ transplant applications. Although hydrogel-based particles and fibers can be easily fabricated, it is difficult to manipulate them for tissue reconstruction. In this video, we describe a fabrication method for micropatterned alginate hydrogel sheets, together with their assembly to form a macro-scale 3D cell culture system with a controlled cellular microenvironment. Using a mist form of the calcium gelling agent, thin hydrogel sheets are easily generated with a thickness in the range of 100 - 200 &#181;m, and with precise micropatterns. Cells can then be cultured with the geometric guidance of the hydrogel sheets in freestanding conditions. Furthermore, the hydrogel sheets can be readily manipulated using a micropipette with an end-cut tip, and can be assembled into multi-layered structures by stacking them using a patterned polydimethylsiloxane (PDMS) frame. These modular hydrogel sheets, which can be fabricated using a facile process, have potential applications of in vitro drug assays and biological studies, including functional studies of micro- and macrostructure and tissue reconstruction.

We describe the fabrication of micropatterned hydrogel sheets using a simple process, which can be assembled and manipulated in a freestanding form. Using these modular hydrogel sheets, a simple macro-scaled 3D cell culture system can be generated with a controlled cellular microenvironment.

Hydrogels can be patterned at the micro-scale using microfluidic or micropatterning technologies to provide an in vivo-like three-dimensional (3D) tissue ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

Patterning of Microorganisms and Microparticles through Sequential Capillarity-assisted Assembly

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, including studies of microbial physiology and interactions. At the simplest level, accurate spatial patterning of microorganisms would enable reliable, long-term imaging of large numbers of individual cells and transform the ability to quantitatively study distance-dependent microbe-microbe interactions. More uniquely, coupling accurate spatial patterning and full control over environmental conditions, as offered by microfluidic technology, would provide a powerful and versatile platform for single-cell studies in microbial ecology. 
This paper presents a microfluidic platform to produce versatile and user-defined patterns of microorganisms within a microfluidic channel, allowing complete optical access for long-term, high-throughput monitoring. This new microfluidic technology is based on capillarity-assisted particle assembly and exploits the capillary forces arising from the controlled motion of an evaporating suspension inside a microfluidic channel to deposit individual microsized objects in an array of traps microfabricated onto a polydimethylsiloxane (PDMS) substrate. Sequential depositions generate the desired spatial layout of single or multiple types of micro-sized objects, dictated solely by the geometry of the traps and the filling sequence. 
The platform has been calibrated using colloidal particles of different dimensions and materials: it has proven to be a powerful tool to generate diverse colloidal patterns and perform surface functionalization of trapped particles. Furthermore, the platform was tested on microbial cells, using Escherichia coli cells as a model bacterium. Thousands of individual cells were patterned on the surface, and their growth was monitored over time. In this platform, the coupling of single-cell deposition and microfluidic technology allows both geometric patterning of microorganisms and precise control of environmental conditions. It thus opens a window into the physiology of single microbes and the ecology of microbe-microbe interactions, as shown by preliminary experiments.

We present a technology that uses capillarity-assisted assembly in a microfluidic platform to pattern micro-sized objects suspended in a liquid, such as bacteria and colloids, into prescribed arrays on a polydimethylsiloxane substrate.

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, ...

Engineering

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 ...

A Versatile Method of Patterning Proteins and Cells

Substrate and cell patterning techniques are widely used in cell biology to study cell-to-cell and cell-to-substrate interactions. Conventional patterning techniques work well only with simple shapes, small areas and selected bio-materials. This article describes a method to distribute cell suspensions as well as substrate solutions into complex, long, closed (dead-end) polydimethylsiloxane (PDMS) microchannels using negative pressure. This method enables researchers to pattern multiple substrates including fibronectin, collagen, antibodies (Sal-1), poly-D-lysine (PDL), and laminin. Patterning of substrates allows one to indirectly pattern a variety of cells. We have tested C2C12 myoblasts, the PC12 neuronal cell line, embryonic rat cortical neurons, and amphibian retinal neurons. In addition, we demonstrate that this technique can directly pattern fibroblasts in microfluidic channels via brief application of a low vacuum on cell suspensions. The low vacuum does not significantly decrease cell viability as shown by cell viability assays. Modifications are discussed for application of the method to different cell and substrate types. This technique allows researchers to pattern cells and proteins in specific patterns without the need for exotic materials or equipment and can be done in any laboratory with a vacuum.

This report describes a simple, easy to perform technique, using low pressure vacuum, to fill microfluidic channels with cells and substrates for biological research.

Substrate and cell patterning techniques are widely used in cell biology to study cell-to-cell and cell-to-substrate interactions. Conventional patterning ...

Micro-Embossed Fabrication of Nanocellulose Paper-Based Analytical Devices  

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.

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.

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

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

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