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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+perspective+on+paper-based+microfluidics:+Current+status+and+future+trends.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Paper-based+microfluidic+point-of-care+diagnostic+devices.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+developments+in+paper-based+microfluidic+devices.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+microfluidic+devices+fabricated+in+layered+paper+and+tape.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transport+in+two-dimensional+paper+networks.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+low+cost+point-of-care+viscous+sample+preparation+device+for+molecular+diagnosis+in+the+developing+world;+an+example+of+microfluidic+origami.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Paper+and+toner+three-dimensional+fluidic+devices:+programming+fluid+flow+to+improve+point-of-care+diagnostics.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+paper+microfluidic+devices+assembled+using+the+principles+of+origami.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+throughput+method+for+prototyping+three-dimensional,+paper-based+microfluidic+devices.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterned+adhesive+enables+construction+of+nonplanar+three-dimensional+paper+microfluidic+circuits.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Understanding+wax+printing:+a+simple+micropatterning+process+for+paper-based+microfluidics.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+prototyping+of+paper-based+microfluidics+with+wax+for+low-cost,+portable+bioassay.">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://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Genuine Japanese origami. , (2012).</li><li>Schonhorn, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+device+architecture+for+three-dimensional,+patterned+paper+immunoassays.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Self-folding+of+three-dimensional+hydrogel+microstructures.">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 border="0" cellpadding="0" cellspacing="0" width="585">
	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Camera</td>
			<td style="width:143px;">Nikon</td>
			<td style="width:119px;">D5100</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Solid-ink printer</td>
			<td style="width:143px;">Xerox</td>
			<td style="width:119px;">ColorQube 8880</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Hotplate</td>
			<td style="width:143px;">Torrey Pines</td>
			<td style="width:119px;">HS60</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Humidity chamber</td>
			<td style="width:143px;">Electro-Tech Systems</td>
			<td style="width:119px;">5503-E</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Spray adhesive</td>
			<td style="width:143px;">3M</td>
			<td style="width:119px;">62497749309</td>
			<td style="width:160px;">Super 77 (16.75 oz can)</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Filter paper</td>
			<td style="width:143px;">Whatman</td>
			<td style="width:119px;">Grade 4</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Perforated steel sheet</td>
			<td style="width:143px;">MetalsDepot</td>
			<td style="width:119px;">PS16116</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Tartrazine</td>
			<td style="width:143px;">Sigma-Aldritch</td>
			<td style="width:119px;">T0388</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Allura Red</td>
			<td style="width:143px;">Sigma-Aldritch</td>
			<td style="width:119px;">458848</td>
			<td style="width:160px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:165px;">Erioglaucine disodium salt</td>
			<td style="width:143px;">Sigma-Aldritch</td>
			<td style="width:119px;">861146</td>
			<td style="width:160px;"></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

7.5K Views.  University of California, Riverside. 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.

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

Separating Beads and Cells in Multi-channel Microfluidic Devices Using Dielectrophoresis and Laminar Flow

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting and counting cells. However, manipulating the cell within the fluidic domain remains a challenge and requires complicated fabrication protocols for forming valves and electrodes, or demands specialty equipment like optical tweezers. Here, we demonstrate that conventional printed circuit boards (PCB) can be used for the non-contact manipulation of cells by employing dielectrophoresis (DEP) for bead and cell manipulation in laminar flow fields for bioactuation, and for cell and bead separation in multichannel microfluidic devices. First, we present the protocol for assembling the DEP electrodes and microfluidic devices, and preparing the cells for DEP. Then, we characterize the DEP operation with polystyrene beads. Lastly, we show representative results of bead and cell separation in a multichannel microfluidic device. In summary, DEP is an effective method for manipulating particles (beads or cells) within microfluidic devices.

Dielectrophoresis (DEP) is an effective method to manipulate cells. Printed circuit boards (PCB) can provide inexpensive, reusable and effective electrodes for contact-free cell manipulation within microfluidic devices. By combining PDMS-based microfluidic channels with coverslips on PCBs, we demonstrate bead and cell manipulation and separation within multichannel microfluidic devices.

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting ...

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Insertion of Flexible Neural Probes Using Rigid Stiffeners Attached with Biodissolvable Adhesive

Microelectrode arrays for neural interface devices that are made of biocompatible thin-film polymer are expected to have extended functional lifetime because the flexible material may minimize adverse tissue response caused by micromotion. However, their flexibility prevents them from being accurately inserted into neural tissue. This article demonstrates a method to temporarily attach a flexible microelectrode probe to a rigid stiffener using biodissolvable polyethylene glycol (PEG) to facilitate precise, surgical insertion of the probe. A unique stiffener design allows for uniform distribution of the PEG adhesive along the length of the probe. Flip-chip bonding, a common tool used in microelectronics packaging, enables accurate and repeatable alignment and attachment of the probe to the stiffener. The probe and stiffener are surgically implanted together, then the PEG is allowed to dissolve so that the stiffener can be extracted leaving the probe in place. Finally, an in vitro test method is used to evaluate stiffener extraction in an agarose gel model of brain tissue. This approach to implantation has proven particularly advantageous for longer flexible probes (&gt;3 mm). It also provides a feasible method to implant dual-sided flexible probes. To date, the technique has been used to obtain various in vivo recording data from the rat cortex.

Insertion of flexible neural microelectrode probes is enabled by attaching probes to rigid stiffeners with polyethylene glycol (PEG). A unique assembly process ensures uniform and repeatable attachment. After insertion into tissue, the PEG dissolves and the stiffener is extracted. An in vitro test method evaluates the technique in agarose gel.

Microelectrode  arrays for neural interface devices that are made of biocompatible thin-film  polymer are expected to have extended functional lifetime ...

Rapid and Low-cost Prototyping of Medical Devices Using 3D Printed Molds for Liquid Injection Molding

Biologically inert elastomers such as silicone are favorable materials for medical device fabrication, but forming and curing these elastomers using traditional liquid injection molding processes can be an expensive process due to tooling and equipment costs. As a result, it has traditionally been impractical to use liquid injection molding for low-cost, rapid prototyping applications. We have devised a method for rapid and low-cost production of liquid elastomer injection molded devices that utilizes fused deposition modeling 3D printers for mold design and a modified desiccator as an injection system. Low costs and rapid turnaround time in this technique lower the barrier to iteratively designing and prototyping complex elastomer devices. Furthermore, CAD models developed in this process can be later adapted for metal mold tooling design, enabling an easy transition to a traditional injection molding process. We have used this technique to manufacture intravaginal probes involving complex geometries, as well as overmolding over metal parts, using tools commonly available within an academic research laboratory. However, this technique can be easily adapted to create liquid injection molded devices for many other applications.

We have devised a method for low-cost and rapid prototyping of liquid elastomer rubber injection molded devices by using fused deposition modeling 3D printers for mold design and a modified desiccator as a liquid injection system.

Biologically inert elastomers such as silicone are favorable materials for medical device fabrication, but forming and curing these elastomers using ...

Paper-based Devices for Isolation and Characterization of Extracellular Vesicles

Extracellular vesicles (EVs), membranous particles released from various types of cells, hold a great potential for clinical applications. They contain nucleic acid and protein cargo and are increasingly recognized as a means of intercellular communication utilized by both eukaryote and prokaryote cells. However, due to their small size, current protocols for isolation of EVs are often time consuming, cumbersome, and require large sample volumes and expensive equipment, such as an ultracentrifuge. To address these limitations, we developed a paper-based immunoaffinity platform for separating subgroups of EVs that is easy, efficient, and requires sample volumes as low as 10 &#956;l. Biological samples can be pipetted directly onto paper test zones that have been chemically modified with capture molecules that have high affinity to specific EV surface markers. We validate the assay by using scanning electron microscopy (SEM), paper-based enzyme-linked immunosorbent assays (P-ELISA), and transcriptome analysis. These paper-based devices will enable the study of EVs in the clinic and the research setting to help advance our understanding of EV functions in health and disease.

This protocol details a method to isolate extracellular vesicles (EVs), small membranous particles released from cells, from as little as 10 &#956;l serum samples. This approach circumvents the need for ultracentrifugation, requires only a few minutes of assay time, and enables the isolation of EVs from samples of limited volumes.

Extracellular vesicles (EVs), membranous particles released from various types of cells, hold a great potential for clinical applications. They contain ...

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D culture of cells in collagen type I gels helps to stabilize cell morphology and function, which is necessary for creating microfluidic tissue models in microdevices. Translating traditional 3D culture techniques for tissue culture plates to microfluidic devices is often difficult because of the limited channel dimensions. In this method, we describe a technique for modifying native type I collagen to generate polycationic and polyanionic collagen solutions that can be used with layer-by-layer deposition to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These thin collagen layers stabilize cell morphology and function, as shown using primary hepatocytes as an example cell, allowing for the long term culture of microtissues in microfluidic devices.

The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D ...

Rapid Subtractive Patterning of Live Cell Layers with a Microfluidic Probe

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one particular implementation of the MFP, the hierarchical hydrodynamic flow confinement (hHFC), multiple liquids are simultaneously brought in contact with a substrate. Local chemical action and liquid shaping using the hHFC, is exploited to create cell patterns by locally lysing and removing cells. By utilizing the scanning ability of the MFP, user-defined patterns of cell monolayers are created. This protocol enables rapid, real-time and spatially controlled cell patterning, which can allow selective cell-cell and cell-matrix interaction studies.

We present a protocol to perform subtractive patterning of live cell monolayers on a surface. This is achieved by local and selective lysis of adherent cells using a microfluidic probe (MFP). The cell lysate retrieved from local regions can be used for downstream analysis, enabling molecular profiling studies.

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one ...

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to entry for non-specialists who would benefit greatly from the ability to develop their own microfluidic devices to address research questions. Particularly lacking has been the open dissemination of protocols related to photolithography, a key step in the development of a replica mold for the manufacture of polydimethylsiloxane (PDMS) devices. While the fabrication of single height silicon masters has been explored extensively in literature, fabrication steps for more complicated photolithography features necessary for many interesting device functionalities (such as feature rounding to make valve structures, multi-height single-mold patterning, or high aspect ratio definition) are often not explicitly outlined. 
Here, we provide a complete protocol for making multilayer microfluidic devices with valves and complex multi-height geometries, tunable for any application. These fabrication procedures are presented in the context of a microfluidic hydrogel bead synthesizer and demonstrate the production of droplets containing polyethylene glycol (PEG diacrylate) and a photoinitiator that can be polymerized into solid beads. This protocol and accompanying discussion provide a foundation of design principles and fabrication methods that enables development of a wide variety of microfluidic devices. The details included here should allow non-specialists to design and fabricate novel devices, thereby bringing a host of recently developed technologies to their most exciting applications in biological laboratories.

Multilayer microfluidic devices often involve the fabrication of master molds with complex geometries for functionality. This article presents a complete protocol for multi-step photolithography with valves and variable height features tunable to any application. As a demonstration, we fabricate a microfluidic droplet generator capable of producing hydrogel beads.

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to ...

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 ...

Soft Lithographic Procedure for Producing Plastic Microfluidic Devices with View-ports Transparent to Visible and Infrared Light

Infrared (IR) spectro-microscopy of living biological samples is hampered by the absorption of water in the mid-IR range and by the lack of suitable microfluidic devices. Here, a protocol for the fabrication of plastic microfluidic devices is demonstrated, where soft lithographic techniques are used to embed transparent Calcium Fluoride (CaF2) view-ports in connection with observation chamber(s). The method is based on a replica casting approach, where a polydimethylsiloxane (PDMS) mold is produced through standard lithographic procedures and then used as the template to produce a plastic device. The plastic device features ultraviolet/visible/infrared (UV/Vis/IR) -transparent windows made of CaF2 to allow for direct observation with visible and IR light. The advantages of the proposed method include: a reduced need for accessing a clean room micro-fabrication facility, multiple view-ports, an easy and versatile connection to an external pumping system through the plastic body, flexibility of the design, e.g., open/closed channels configuration, and the possibility to add sophisticated features such as nanoporous membranes.

A protocol for the fabrication of plastic microfluidic devices with transparent view-ports for visible and infrared light imaging is described.

Infrared (IR) spectro-microscopy of living biological samples is hampered by the absorption of water in the mid-IR range and by the lack of suitable ...