The authors gratefully acknowledge MBI financial support.

1. Preparation of Silicon Primary Mold
NOTE: A photomask is required for the preparation of the primary mold. The photomask can be either purchased from independent providers or fabricated in-house through standard optical mask fabrication procedures. A photomask with bright field polarity is used in this case (Figure 2a).
<ol>
	<li>Pattern definition
	<ol>
		<li>Spin coat a 4 inch silicon wafer with SU-8 3010 negative photoresist at 2,300 rpm for 30 s.</li>
		<li>Soft-bake the photoresist on a hot plate at 65 &#176;C for 2 min and then at 95 &#176;C for 8 min.</li>
		<li>Expose the photoresist to UV light (i-line, 365 nm) through the photomask under the mask aligner for a total energy dose of 100-120 mJ/cm2; the hard contact mode is preferred to achieve a better resolution.</li>
		<li>Remove the wafer and apply a post exposure bake at 65 &#176;C for 1 min and then at 95 &#176;C for 2 min.</li>
		<li>Develop the photoresist using a SU-8 developer at room temperature, then rinse with isopropyl alcohol, and gently blow dry with nitrogen; the measured pattern thickness should be 10 &#181;m and below. See Figure 2b for the picture of the actual silicon mold.</li>
	</ol>
	</li>
	<li>Silanization of silicon mold
	<ol>
		<li>Treat the silicon mold with oxygen plasma at 60 W for 30 s with 20 sccm of oxygen flow. Set the chamber pressure to 1-10 mbar during the process.</li>
		<li>Place the mold in a vacuum jar with 50 &#181;L of silane and leave the jar in the vacuum state (1-10 mbar) for at least 2 h. 
		NOTE: The silanization process creates a hydrophobic surface coating which prevents PDMS from sticking to the mold15. Note that the primary mold can also be fabricated using an alternative method, which involves dry etching of the silicon. In this case, the photomask will be of opposite polarity (dark field), and the pattern definition in step 1.1 will use a positive photoresist.</li>
	</ol>
	</li>
</ol>
2. Preparation of PDMS Secondary Mold
<ol>
	<li>PDMS mixing
	<ol>
		<li>Mix the PDMS elastomer and curing agent, 10:1; the total amount is such that the resulting PDMS thickness is approximately 1 to 1.5 mm.</li>
		<li>After thorough mixing, degas the mixture by leaving it in a vacuum jar in the vacuum state (1-10 mbar) for approximately 15 min or until there are no visible bubbles; this is to remove any trapped air within the mixture.</li>
	</ol>
	</li>
	<li>Mold replication
	<ol>
		<li>Pour the PDMS mixture on the silicon mold fabricated in step 1 and degas the mixture to remove any trapped air with the same settings as in step 2.1.2. Heat at 70 &#176;C for 2 h on a hot plate to cure the mixture.</li>
		<li>Remove the cured PDMS from the hot plate and let it cool down to room temperature. With a razor blade, cut the PDMS along the edges of the silicon mold.</li>
		<li>With a pair of tweezers, pinch one corner of the cut PDMS and carefully peel the PDMS replica off the silicon mold; the resulting microfluidic pattern on this secondary mold is a protrusion, which is the opposite polarity of the primary mold (Figure 2c).</li>
	</ol>
	</li>
	<li>Silanization of the PDMS replica (same as in step 1.2)
	<ol>
		<li>Treat the PDMS mold with oxygen plasma at 60 W for 30 s with 20 sccm of oxygen flow. Set the chamber pressure to 1-10 mbar during the process.</li>
		<li>Place the mold in a vacuum jar with 50 &#181;L of silane and leave the jar in the vacuum state (1-10 mbar) for at least 2 h.</li>
	</ol>
	</li>
</ol>
3. Preparation of PDMS Templates
NOTE: To standardize the shape and size of the final devices and to ease the alignment of the main features in the two halves, two separate PDMS templates were used, which define the geometry of the device, the placement of the transparent window and the in- and out-let connections. The first PDMS template aids the fabrication of the patterned half of the device, while the second helps to ease the fabrication of the flat half of the device.
<ol>
	<li>Design the templates using a computer aided design (CAD) software. Figure 3a shows the lay-out and dimensions of the template used to fabricate the patterned half of the device. To fabricate the flat half of the device, remove the 1.5 mm diameter holes from the design.</li>
	<li>Acquire the templates from an external provider or through an internal mechanical workshop if available. 
	NOTE: Acrylic was used as the template material due to ease of fabrication and low cost achievable in any standard workshop. There are alternative options, such as 3D printing.</li>
	<li>Mix the PDMS elastomer and curing agent 10:1; be sure to prepare a sufficient amount of PDMS to entirely submerge the templates.</li>
	<li>After thorough mixing, degas the mixture by leaving it in a vacuum jar in the vacuum state (1-10 mbar) for approximately 15 min or until there are no visible bubbles (whichever is later); this is to remove any trapped air within the mixture.</li>
	<li>Pour the PDMS mixture on the acrylic templates until their top-most surface is submerged about 1 mm below the liquid surface. Degas the PDMS again to remove any trapped air with the same settings as in 3.4. Heat this at 60 &#176;C for 2 h on a leveled hot plate to cure the mixture.</li>
	<li>Remove the cured PDMS from the hot plate and let it cool down to room temperature. With a razor blade, cut the PDMS along the edges of the acrylic templates.</li>
	<li>With a pair of tweezers, pinch one corner of the cut PDMS and carefully peel the PDMS off the acrylic templates. 
	NOTE: Figure 3b shows the lay-out and dimensions of the PDMS replica used to fabricate the patterned half of the device.</li>
	<li>Prepare the second PDMS replica for fabricating the flat half of the device by repeating steps 3.3 to 3.7 but using the acrylic template without the 1.5 mm diameter holes.</li>
</ol>
4. Microfluidic Device Fabrication
<ol>
	<li>Fabrication of the patterned half of the device (i.e. , with device layout)
	<ol>
		<li>Treat the CaF2 window with oxygen plasma at 60 W for 30 s with 20 sccm of oxygen flow. This is done to improve the flow of NOA during the following fabrication. 
		NOTE: This step is not mandatory.</li>
		<li>Place carefully the first PDMS template (one with the 1.5 mm diameter pillars) on a flat surface, e.g., a soda lime plate (Figure 4a). Place a CaF2 window centered on top of the PDMS plug and gently press the window such that it is in good contact with the plug (Figure 4b).</li>
		<li>Take the PDMS mold made in step 2 and place a thin UV-transparent plate (in this case, a quartz plate, 500 &#181;m thick and 1.5 cm x 1.5 cm in size) on the backside of the mold, aligned with the location of the central chamber (Figure 4c). Make sure that the quartz plate is in good contact with the PDMS mold. 
		NOTE: The quartz plate prevents the unwanted area of the mold from easily coming into contact with the CaF2 window.</li>
		<li>Gently place this PDMS mold face down towards the CaF2 window with the fluidic chamber aligned to the center of the CaF2 window. Make sure that all the elements (template, mold and window) are in good contact and aligned (Figure 4c-4d).</li>
		<li>Gradually dispense drops of NOA at the in-let of the PDMS template and let it slowly fill the cavity. Once the resin comes into contact with the edge of the window, the capillary flow will fill the thin gap (~10 &#181;m) between the PDMS mold and CaF2 window (Figure 4e-4f).</li>
		<li>After the cavity is completely filled, cure the NOA by exposure to UV light (e.g., with a UV-LED exposure system, Figure 4g). 
		NOTE: The exposure time may vary accordingly with the energy of the UV source. The UV-LED exposure system, which provides a power density of 24 mW/cm2, requires around 90 s at 100% power and continuous exposure mode.</li>
		<li>Carefully remove the thin quartz plate from the back of the PDMS mold and then gently peel the PDMS mold from the top of the NOA layer (Figure 4h). Finally, remove the NOA layer from the PDMS template (Figure 4i). 
		NOTE: The resulting device layout on the cured NOA would have the same polarity of the pattern in the primary silicon mold.</li>
	</ol>
	</li>
	<li>Fabrication of the flat half of the device (i.e. , without device layout)
	<ol>
		<li>Treat the CaF2 window with oxygen plasma at 60 W for 30 s with 20 sccm of oxygen flow. 
		NOTE: This step is not mandatory.</li>
		<li>Place carefully the second PDMS template (one without the 1.5 mm diameter pillars) on a flat surface, e.g., a soda lime plate. Place a CaF2 window centered on top of the PDMS plug and gently press the window such that it is in good contact with the plug.</li>
		<li>Place a 1 mm thick PDMS sheet with 5 cm x 3.5 cm size on top of the CaF2 window, with the PDMS sheet aligned with the center of the PDMS template. Make sure that the PDMS sheet is in good contact with the window.</li>
		<li>Gradually dispense drops of NOA at the in-let of the PDMS template and let it slowly fill the cavity.</li>
		<li>After the cavity is completely filled, cure the NOA by exposure to UV light (e.g., with a UV-LED exposure system). 
		NOTE: The exposure time may vary accordingly with the UV energy source. With the UV-LED exposure system, which provides a power density of 24 mW/cm2, this requires around 50 s at 100% power and continuous exposure mode.</li>
		<li>Peel off the PDMS sheet from the top of the NOA layer and carefully remove the cured NOA layer from the PDMS template.</li>
	</ol>
	</li>
	<li>Bonding of the two halves of the device
	<ol>
		<li>Align the two halves of the device such that both CaF2 windows are aligned. Gently finger-press both halves at the corner of the NOA layers such that the position of the two halves are fixed.</li>
		<li>Cut two circular discs out of a 1 mm thick PDMS sheet using an 8 mm diameter puncher (Figure 5a).</li>
		<li>Cut two rectangles with the same size of the device (4 cm x 2.5 cm) from a 1 mm thick PDMS sheet. On both PDMS rectangles, cut openings corresponding to the channels and the in-let/out-let of the device. 
		NOTE: The pre-cut openings in the PDMS rectangles are meant to prevent the channels from collapsing during pressing.</li>
		<li>Stack in the following order from the bottom: one PDMS rectangle with pre-cut openings, one PDMS disc (in contact with the bottom window, cut in step 4.3.2), the two finger-pressed halves of the device, the second PDMS disc (sitting on the top window), and lastly the second PDMS rectangle with pre-cut openings (Figure 5b).</li>
		<li>Place this assembly in the vacuum press setup such that it is sandwiched between 2 plates and seal the plastic bag (Figure 5c). Turn on the vacuum pump and evacuate the assembly. Let the vacuum pump reach its base pressure or apply the vacuum for at least 10 min. 
		NOTE: The base pressure achieved depends on the vacuum pump used and quality of the sealing of the plastic bag.</li>
		<li>Expose the evacuated assembly to UV with a broad-band Hg gas lamp at 270 W for 15 min. Turn off the vacuum pump and let the assembly slowly vent to atmospheric pressure before removing the final device from the assembly.</li>
	</ol>
	</li>
</ol>

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A., Keese, M., Pepperkok, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=IR+microspectroscopy+of+live+cells.">IR microspectroscopy of live cells.</a> Vibrational Spectroscopy. 38 (1-2), 185-191 (2005).</li><li>Rahmelow, K., Hubner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Infrared+Spectroscopy+in+Aqueous+Solution:+Difficulties+and+Accuracy+of+Water+Subtraction.">Infrared Spectroscopy in Aqueous Solution: Difficulties and Accuracy of Water Subtraction.</a> Appl Spectrosc. 51 (2), 160-170 (1997).</li><li>Kazarian, S. G., Chan, K. 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The authors have nothing to disclose.

In order to assess and optimize the fabrication protocol, we used a simple layout for the microfluidic pattern with a large rectangular chamber (5 mm x 2.5 mm size) at the center, two small rectangular chambers (5.5 mm x 0.75 mm size) separated from the main circuit on the upper and lower sides, and 300 &#181;m wide in-let/out-let channels. The central chamber is used for the seeding and observation of the cells, while the two separated smaller chambers are used to measure the air background during FTIR experiments as a ...

Fourier Transform Infrared Spectro-microscopy (FTIR) has been extensively utilized as a label-free and non-invasive imaging technique to provide detailed chemical information of a sample. This enables the extraction of biochemical information to study the chemistry of biological samples, with a minimum amount of preparation since the absorption spectrum of the specimen carries the intrinsic fingerprints of its chemical composition1,2. Recently, FTIR has been increasingly applied to the study of live biological samples, e.g., cells3. However, water, which is the medium for living cells in most cases, shows a strong absorbance in the mid-IR region. Even as a thin layer, its presence can completely overwhelm the important structural information of the specimens.
For many years, the common approach was fixing or drying samples to completely exclude the water absorption signal in the spectrum. However, this approach does not allow for real time measurements on living cells, which is essential to study the change of their chemical composition and cellular processes with time. One way to obtain reliable absorption spectra from live biological samples, is to limit the total optical path length in the medium of the IR beam to less than 10 &#181;m4.
A well-established approach in living cell experiments has been so far, Attenuated Total Reflection (ATR)-FTIR imaging, which enables measurements independent of the sample thickness, allowing cells to be sustained in a thicker layer of aqueous medium. However, the small depth of penetration of the evanescent wave restricts measurements of samples to only the first few microns from the surface of the ATR crystal5.
Alternatively, the water absorption limitation has been circumvented with the emergence of various microfluidic systems, which are generally classified into two large groups: open channel (where one of the fluid surfaces is exposed to the atmosphere) and closed channel (where two IR-transparent windows are separated by a spacer with a defined thickness).
Loutherback et al. developed an open-channel membrane device that enables long term continuous IR measurements of live cells for up to 7 days6. The method requires high humidity in the environment to prevent evaporation of the medium from the cell surface. The system works best with cells that naturally grow at air-liquid interfaces, such as epithelial tissues of the skin, lung, and eyes, or microbial biofilms7.
A closed-channel configuration aims to create a uniform, thin layer between two parallel IR-transparent windows, where cells are maintained in their aqueous media. The thickness of this cavity is such that the water absorption signal is below saturation. Water background can then be subtracted to obtain the correct sample spectra. Most of the closed-channel methods utilize a plastic spacer separating the two windows to form a demountable liquid chamber3,8,9. An advantage of this method is that it does not require microfabrication; however, structures that are more complex than a measuring chamber with in- and out-let channels are extremely difficult to realize in the thin spacer. There is also an issue with the reproducibility of the path length between IR measurements due to its reliance on mechanical clamping. In order to achieve a more precise control of the spacing for a more reliable spectrum acquisition, optical lithography methods have been implemented to pattern photoresist on top of the IR substrate to define the spacer9,10,11,12. Even though this makes it possible for more complex structures to be defined in the spacer, the method requires access to a microfabrication facility to produce the pattern on every substrate.
In this paper, we present a simple fabrication technique of an IR-compatible microfluidic device, with the aim to reduce the fabrication cost and the requirement of accessing a microfabrication facility. The method presented here (see Figure 1) uses an established process known as soft lithography. Two molds are required in this case. The primary mold is made from a 4-inch silicon wafer using a standard UV lithography process. The secondary mold is its replica made of PDMS, which has a reversed polarity of the pattern in the silicon primary mold and serves as the master mold for subsequent device fabrication.
The device has two separate layers: a first layer with the microfluidic layout (which in the presented case consists of the microfluidic channel, in-let/out-let, and an observation chamber with a CaF2 viewport), and a second layer with a flat surface (which consists of only a CaF2 viewport).
Here an UV-curable optical adhesive, Norland optical adhesive 73 (NOA73, abbreviated as NOA henceforth), is used to form the main plastic body of the device. There are several advantages of using this optical adhesive: low fabrication cost, ease of connectivity to external systems, good optical transparency, low viscosity, and most importantly, biocompatibility13. CaF2 is a suitable choice as the viewport due to its biocompatibility and excellent IR-transparency14.
With this new approach, access to a microfabrication facility is strictly required only for the fabrication of the primary mold. Subsequent fabrication processes for the plastic microfluidic device can be carried out in any laboratory equipped with an UV-illumination source.

<table border="0" cellpadding="0" cellspacing="0" width="856">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Chemical</td>
			<td style="width:212px;"></td>
			<td style="width:112px;"></td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:291px;">Trichloro(1H,1H,2H,2H-perfluorooctyl)silane 97%</td>
			<td style="width:212px;">Sigma Aldrich</td>
			<td style="width:112px;">448931-10G</td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:291px;">Sylgard 184 Silicone Elastomer Kit</td>
			<td style="width:212px;">Dow Corning</td>
			<td style="width:112px;"></td>
			<td style="width:243px;">Polydimethylsiloxane or in short, PDMS</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Norland Optical Adhesive 73</td>
			<td style="width:212px;">Norland Products Inc.</td>
			<td style="width:112px;">7304</td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">SU8 3010 photoresist</td>
			<td style="width:212px;">MicroChem</td>
			<td style="width:112px;">Y311060</td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">SU8 developer</td>
			<td style="width:212px;">MicroChem</td>
			<td style="width:112px;">Y020100</td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Material</td>
			<td style="width:212px;"></td>
			<td style="width:112px;"></td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Silicon wafer, 4 inch, prime grade</td>
			<td style="width:212px;">Bonda Technology Pte Ltd</td>
			<td style="width:112px;"></td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">CaF2 IR-grade windows</td>
			<td style="width:212px;">Crystran, UK</td>
			<td style="width:112px;">CAFP10-1</td>
			<td style="width:243px;">10 mm diameter, 1 mm thickness</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Acrylic templates</td>
			<td style="width:212px;"></td>
			<td style="width:112px;"></td>
			<td style="width:243px;">Custom made</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Equipment</td>
			<td style="width:212px;"></td>
			<td style="width:112px;"></td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">UV-KUB 2 (UV LED exposure system)</td>
			<td style="width:212px;">KLOE</td>
			<td style="width:112px;"></td>
			<td style="width:243px;">Emission spectrum 365nm &#177; 5nm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Newport UV lamp</td>
			<td style="width:212px;">Newport</td>
			<td style="width:112px;">Model 66902</td>
			<td style="width:243px;">50-500 Watt Hg arc lamp</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">CEE Spin coater</td>
			<td style="width:212px;">Brewer Science</td>
			<td style="width:112px;">Model 200x</td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">MJB4 mask aligner</td>
			<td style="width:212px;">SUSS MicroTec</td>
			<td style="width:112px;"></td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Precision digital hot plate</td>
			<td style="width:212px;">Harry Gestigkeit GmbH</td>
			<td style="width:112px;">2860SR</td>
			<td style="width:243px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:291px;">Plasma Surface Technology</td>
			<td style="width:212px;">Diener Electronic GmbH + Co. KG</td>
			<td style="width:112px;"></td>
			<td style="width:243px;">For O2 plasma treatment</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:291px;">IDP-3 Dry Scroll Vacuum Pump</td>
			<td style="width:212px;">Agilent Technologies</td>
			<td></td>
			<td>ultimate pressure 3.3 x 10-1 mbar</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:291px;">Bruker IFS 66v/s FTIR Spectrometer</td>
			<td style="width:212px;">Bruker</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Figure 6 presents the transmittance spectra of a brand new CaF2 window, the patterned half of the device, and the complete device. All three spectra exhibit excellent transparency to mid IR with transmittance larger than 80%. The interference pattern visible in the spectrum of the full device (yellow curve in the figure) is caused by the air gap in the range of 9-10 &#181;m between the two windows. These spectra demonstrate that the fabrication app...

Watch this Scientific Journal Video about Soft Lithographic Procedure for Producing Plastic Microfluidic Devices with View-ports Transparent to Visible and Infrared Light at JoVE.com

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

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 ...

soft-lithographic-procedure-for-producing-plastic-microfluidic

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

13.0K Views.  National University of Singapore. A protocol for the fabrication of plastic microfluidic devices with transparent view-ports for visible and infrared light imaging is described.

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

Soft Lithographic Functionalization and Patterning Oxide-free Silicon and Germanium

The development of hybrid electronic devices relies in large part on the integration of (bio)organic materials and inorganic semiconductors through a stable interface that permits efficient electron transport and protects underlying substrates from oxidative degradation. Group IV semiconductors can be effectively protected with highly-ordered self-assembled monolayers (SAMs) composed of simple alkyl chains that act as impervious barriers to both organic and aqueous solutions. Simple alkyl SAMs, however, are inert and not amenable to traditional patterning techniques. The motivation for immobilizing organic molecular systems on semiconductors is to impart new functionality to the surface that can provide optical, electronic, and mechanical function, as well as chemical and biological activity. 
Microcontact printing (&mu;CP) is a soft-lithographic technique for patterning SAMs on myriad surfaces.1-9 Despite its simplicity and versatility, the approach has been largely limited to noble metal surfaces and has not been well developed for pattern transfer to technologically important substrates such as oxide-free silicon and germanium. Furthermore, because this technique relies on the ink diffusion to transfer pattern from the elastomer to substrate, the resolution of such traditional printing is essentially limited to near 1 &mu;m.10-16
In contrast to traditional printing, inkless &mu;CP patterning relies on a specific reaction between a surface-immobilized substrate and a stamp-bound catalyst. Because the technique does not rely on diffusive SAM formation, it significantly expands the diversity of patternable surfaces. In addition, the inkless technique obviates the feature size limitations imposed by molecular diffusion, facilitating replication of very small (&lt;200 nm) features.17-23 However, up till now, inkless &mu;CP has been mainly used for patterning relatively disordered molecular systems, which do not protect underlying surfaces from degradation.
Here, we report a simple, reliable high-throughput method for patterning passivated silicon and germanium with reactive organic monolayers and demonstrate selective functionalization of the patterned substrates with both small molecules and proteins. The technique utilizes a preformed NHS-reactive bilayered system on oxide-free silicon and germanium. The NHS moiety is hydrolyzed in a pattern-specific manner with a sulfonic acid-modified acrylate stamp to produce chemically distinct patterns of NHS-activated and free carboxylic acids. A significant limitation to the resolution of many &mu;CP techniques is the use of PDMS material which lacks the mechanical rigidity necessary for high fidelity transfer. To alleviate this limitation we utilized a polyurethane acrylate polymer, a relatively rigid material that can be easily functionalized with different organic moieties. Our patterning approach completely protects both silicon and germanium from chemical oxidation, provides precise control over the shape and size of the patterned features, and gives ready access to chemically discriminated patterns that can be further functionalized with both organic and biological molecules. The approach is general and applicable to other technologically-relevant surfaces.

Here we describe a simple method for patterning oxide-free silicon and germanium with reactive organic monolayers and demonstrate functionalization of the patterned substrates with small molecules and proteins. The approach completely protects surfaces from chemical oxidation, provides precise control over feature morphology, and provides ready access to chemically discriminated patterns.

The development of hybrid electronic devices relies in large part on the integration of (bio)organic materials and inorganic semiconductors through a ...

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

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 ...

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

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

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

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

Fully Automated Centrifugal Microfluidic Device for Ultrasensitive Protein Detection from Whole Blood

Enzyme-linked immunosorbent assay (ELISA) is a promising method to detect small amount of proteins in biological samples. The devices providing a platform for reduced sample volume and assay time as well as full automation are required for potential use in point-of-care-diagnostics. Recently, we have demonstrated ultrasensitive detection of serum proteins, C-reactive protein (CRP) and cardiac troponin I (cTnI), utilizing a lab-on-a-disc composed of TiO2 nanofibrous (NF) mats. It showed a large dynamic range with femto molar (fM) detection sensitivity, from a small volume of whole blood in 30 min. The device consists of several components for blood separation, metering, mixing, and washing that are automated for improved sensitivity from low sample volumes. Here, in the video demonstration, we show the experimental protocols and know-how for the fabrication of NFs as well as the disc, their integration and the operation in the following order: processes for preparing TiO2 NF mat; transfer-printing of TiO2 NF mat onto the disc; surface modification for immune-reactions, disc assembly and operation; on-disc detection and representative results for immunoassay. Use of this device enables multiplexed analysis with minimal consumption of samples and reagents. Given the advantages, the device should find use in a wide variety of applications, and prove beneficial in facilitating the analysis of low abundant proteins.

This protocol demonstrates how to achieve femto molar detection sensitivity of proteins in 10 &#181;L of whole blood within 30 min. This can be achieved by using electrospun nanofibrous mats integrated in a lab-on-a-disc, which offers high surface area as well as effective mixing and washing for enhanced signal-to-noise ratio.

Enzyme-linked immunosorbent assay (ELISA) is a promising method to detect small amount of proteins in biological samples. The devices providing a platform ...

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 ...

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic bioprinting to generate a scaffold with multilayer interlacing hydrogel microfibers. To achieve smooth bioprinting, a core-sheath microfluidic printhead containing a composite bioink formulation extruded from the core flow and the crosslinking solution carried by the sheath flow, was designed and fitted onto the bioprinter. By blending gelatin methacryloyl (GelMA) with alginate, a polysaccharide that undergoes instantaneous ionic crosslinking in the presence of select divalent ions, followed by a secondary photocrosslinking of the GelMA component to achieve permanent stabilization, a microfibrous scaffold could be obtained using this bioprinting strategy. Importantly, the endothelial cells encapsulated inside the bioprinted microfibers can form the lumen-like structures resembling the vasculature over the course of culture for 16 days. The endothelialized microfibrous scaffold may be further used as a vascular bed to construct a vascularized tissue through subsequent seeding of the secondary cell type into the interstitial space of the microfibers. Microfluidic bioprinting provides a generalized strategy in convenient engineering of vascularized tissues at high fidelity.

We provide a generalized protocol based on a microfluidic bioprinting strategy for engineering a microfibrous vascular bed, where a secondary cell type could be further seeded into the interstitial space of this microfibrous structure to generate vascularized tissues and organoids.

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic ...

Rapid Fabrication of Custom Microfluidic Devices for Research and Educational Applications

Microfluidic devices allow for the manipulation of fluids, particles, cells, micro-sized organs or organisms in channels ranging from the nano to submillimeter scales. A rapid increase in the use of this technology in the biological sciences has prompted a need for methods that are accessible to a wide range of research groups. Current fabrication standards, such as PDMS bonding, require expensive and time consuming lithographic and bonding techniques. A viable alternative is the use of equipment and materials that are easily affordable, require minimal expertise and allow for the rapid iteration of designs. In this work we describe a protocol for designing and producing PET-laminates (PETLs), microfluidic devices that are inexpensive, easy to fabricate, and consume significantly less time to generate than other approaches to microfluidics technology. They consist of thermally bonded film sheets, in which channels and other features are defined using a craft cutter. PETLs solve field-specific technical challenges while dramatically reducing obstacles to adoption. This approach facilitates the accessibility of microfluidics devices in both research and educational settings, providing a reliable platform for new methods of inquiry.

Here we present a protocol to design and fabricate custom microfluidic devices with minimal financial and time investment. The aim is to facilitate the adoption of microfluidic technologies in biomedical research laboratories and educational settings.

Microfluidic devices allow for the manipulation of fluids, particles, cells, micro-sized organs or organisms in channels ranging from the nano to ...

Bioinspired Soft Robot with Incorporated Microelectrodes

Bioinspired soft robotic systems that mimic living organisms using engineered muscle tissue and biomaterials are revolutionizing the current biorobotics paradigm, especially in biomedical research. Recreating artificial life-like actuation dynamics is crucial for a soft-robotic system. However, the precise control and tuning of actuation behavior still represents one of the main challenges of modern soft robotic systems. This method describes a low-cost, highly scalable, and easy-to-use procedure to fabricate an electrically controllable soft robot with life-like movements that is activated and controlled by the contraction of cardiac muscle tissue on a micropatterned sting ray-like hydrogel scaffold. The use of soft photolithography methods makes it possible to successfully integrate multiple components in the soft robotic system, including micropatterned hydrogel-based scaffolds with carbon nanotubes (CNTs) embedded gelatin methacryloyl (CNT-GelMA), poly(ethylene glycol) diacrylate (PEGDA), flexible gold (Au) microelectrodes, and cardiac muscle tissue. In particular, the hydrogels alignment and micropattern are designed to mimic the muscle and cartilage structure of the sting ray. The electrically conductive CNT-GelMA hydrogel acts as a cell scaffold that improves the maturation and contraction behavior of cardiomyocytes, while the mechanically robust PEGDA hydrogel provides structural cartilage-like support to the whole soft robot. To overcome the hard and brittle nature of metal-based microelectrodes, we designed a serpentine pattern that has high flexibility and can avoid hampering the beating dynamics of cardiomyocytes. The incorporated flexible Au microelectrodes provide electrical stimulation across the soft robot, making it easier to control the contraction behavior of cardiac tissue.

A bioinspired scaffold is fabricated by a soft photolithography technique using mechanically robust and electrically conductive hydrogels. The micropatterned hydrogels provide directional cardiomyocyte cell alignment, resulting in a tailored direction of actuation. Flexible microelectrodes are also integrated into the scaffold to bring electrical controllability for a self-actuating cardiac tissue.

Bioinspired soft robotic systems that mimic living organisms using engineered muscle tissue and biomaterials are revolutionizing the current biorobotics ...

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

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Chapters in this video

Soft Lithographic Functionalization and Patterning Oxide-free Silicon and Germanium

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

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

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

Fully Automated Centrifugal Microfluidic Device for Ultrasensitive Protein Detection from Whole Blood

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

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

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

Rapid Fabrication of Custom Microfluidic Devices for Research and Educational Applications

Bioinspired Soft Robot with Incorporated Microelectrodes