This work was supported by the National Institutes of Health Grants R01 DK091526 (JO), NSF 0852416(DTE), and Chicago Diabetes Project.

1. Preparing the Mouse Islets
<ol>
	<li>Dissect C57BL/6 mice and isolate islets by collagenase digestion and Ficoll density gradient separation. (Refer to JOVE articles referenced in2,3).</li>
	<li>Incubate islets in RPMI-1640 medium containing 10% FBS, 1% penicillin/streptomycin, and 20 mM HEPES in Petri&#160;dishes (37 &#176;C, 5% CO2). Post-isolation, culture islets for 24 hr prior to use in experiments. Use the islets within 1-2 days to ensure consistent results.</li>
</ol>
2. Making the Microfluidic Platform
<ol>
	<li>Generate the microstructure geometries on transparency photomasks for each device layer: 1) inlet and outlet, 2) Glucose microfluidic layer with perifusion chamber (8 mm diameter x 3 mm, 150 &#956;l), 3) 200 &#956;m membrane with microwells (500 &#956;m diameter, 100 &#956;m deep), and 4) gas microfluidic layers.</li>
	<li>To fabricate the Inlet and outlet layer, pour degassed, premixed PDMS to a height of 1.5 mm in a blank Petri dish and cure at 80 &#176;C for 2 hr. Punch 2 mm diameter input/out ports.</li>
	<li>To fabricate the glucose microfluidic layer, spin two 350 &#956;m layers of SU8-2150 to form a single 700 &#956;m layer on a 4 in&#160;silicon wafer.
	<ol>
		<li>Expose UV light through the appropriate photomask to transfer microchannel pattern onto the SU8 after development, producing a 700 &#956;m tall master.</li>
		<li>Pour degassed, premixed PDMS onto this master to a height of 3 mm and cure at 80 &#176;C for 2 hr. Cut this PDMS to shape with a razor blade.</li>
		<li>Punch 2 mm diameter input/output ports as well as the 8 mm diameter chamber.</li>
	</ol>
	</li>
	<li>To fabricate the 200 &#956;m membrane with microwells, spin SU8-2100 to 100 &#956;m. Apply the same UV-lithography to transfer microwell patterns onto this 100 &#956;m tall master.
	<ol>
		<li>Spin degassed, premixed PDMS onto this master at 900 rpm for 30 sec and cure at 80 &#176;C for 10 min. Spin a second layer on top of the first using the same conditions, resulting in a 200 &#956;m PDMS membrane containing microwell patterns.</li>
		<li>Punch 2 mm diameter input/output ports through this membrane</li>
	</ol>
	</li>
	<li>To fabricate the gas microfluidic layer, spin SU8-2100 to 100 &#956;m. Apply the same UV-lithography to transfer gas microfluidic patterns onto this 100 &#956;m tall master.
	<ol>
		<li>Pour degassed, premixed PDMS onto this master to a height of 1.5 mm and cure at 80 &#176;C for 2 hr. Cut the PDMS to shape with a razor blade.</li>
		<li>Punch 2 mm diameter input/output ports through this layer</li>
	</ol>
	</li>
	<li>To bond the multiple layers, prepare them by cleaning with scotch tape, exposing to corona arc, and then aligning by hand in the following order.
	<ol>
		<li>Bond the membrane to the bottom gas layer with microwells facing up.</li>
		<li>Bond the glucose microfluidic on top of the microwell membrane.</li>
		<li>Bond the inlet and outlet layer on the very top, encapsulating the whole assembly.</li>
		<li>Reinforce the bonding by adding 1 kg weight on top and baking at 100 &#176;C for 3 hr.</li>
	</ol>
	</li>
	<li>Leak-test the completed device by loading water into the aqueous layer, then submerging the entire device under water. Then, flow air through the gas layer with an empty syringe.</li>
	<li>Sterilize leak-free devices with 70% ethanol through the aqueous layer then flush with PBS buffer, in which the device is stored at 4 &#176;C until use.</li>
</ol>
3. Microdispenser Setup
<ol>
	<li>Obtain 2 microdispensers, 5 V and 20 V DC power supplies, and a digital I/O board to construct the oxygen mixing and delivery setup.
	<ol>
		<li>Connect the microdispensers&#39; control leads to the included driver units.</li>
		<li>Connect the drivers to the digital I/O board at ports corresponding to Labview controls.</li>
		<li>Connect the 5 V and 20 V power supplies to corresponding contacts on the drive units.</li>
		<li>Connect the digital I/O to a laptop to execute Labview codes for gas control (Appendix).</li>
	</ol>
	</li>
	<li>Connect one microdispenser to nitrogen while another to compressed air, both with 5% CO2. Set both gases to 2 psi. Hook up the dispensers&#39; outputs in a T-junction prior to entering the microfluidic device.</li>
	<li>Using a leak-tested device, characterize the transient response of oxygen modulation by cycling the microdispensers between 5-21% oxygen, while measuring dissolved oxygen in water with a fiber optic oxygen sensor (see equipment list and representative results).</li>
</ol>
4. Setting Up at the Microscopy
<ol>
	<li>Mount the device to the heated stage on the inverted microscope.</li>
	<li>Connect 0% and 21% O2 gases to the device and calibrate with the oxygen sensor.</li>
	<li>Connect the output port of the glucose microfluidics to a fraction collector.</li>
	<li>Prepare the Krebs Ringer bicarbonate buffer (KRB): 129 mM NaCl, 5 mM NaHCO3, 4.7 mM KCl, 1.2 mM KH2PO4, 1 mM CaCl2&#183;2H2O, 1.2 mM MgSO4&#183;7H2O, 10 mM HEPES pH 7.35-7.40, 2 mM basal glucose, and 5% FBS.</li>
	<li>Prepare two buffer solutions containing 2 mM and 14 mM glucose respectively in 50 ml conical tubes bathed in 37 &#176;C water bath.</li>
	<li>Connect a peristaltic pump to draw buffer at 250 &#956;l/min into tubing. Flow the tubing over a 37 &#176;C hotplate before entering the microfluidic device.</li>
	<li>To stain the islets, add Fura-2 AM in DMSO and Rh123 in 100% ethanol to 2 ml of KRB to reach final dye concentrations of 5 &#956;M and 2.5 &#956;M, respectively.</li>
	<li>Pick up islets with a 10 &#956;l pipette and incubate in dyes for 30 min at 37 &#176;C.</li>
	<li>Load approximately 20 islets into the device via the glucose microchannel inlet. Direct the islets into the chamber by priming buffer from the outlet back into the inlet.</li>
	<li>Perfuse the islets in buffer for 10 min to wash away excess dyes.</li>
</ol>
5. Running the Simultaneous Oxygen and Glucose Stimulation
<ol>
	<li>Deliver a glucose stimulation pulse with 5 min of baseline established by KRB2, followed by 15 min of 14 mM stimulation, then wash for 15 min in KRB2.</li>
	<li>Record 340/380 nm excitation wavelength for Fura-2 and 534 emission for Rh123.</li>
	<li>Make measurements of a normal pulse (14 mM normoxic) before each experiment.</li>
	<li>For oxygen modulation (hypoxia/intermittent hypoxia), only apply buffer flow during stimulation and washing steps and no flow in other steps, to minimize convective disturbances.</li>
	<li>Collect effluents from glucose outlet at 1 min intervals for ELISA insulin assay.</li>
</ol>

<ol>
	<li>Lo, J. F., Wang, Y., 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=Islet+Preconditioning+via+Multimodal+Microfluidic+Modulation+of+Intermittent+Hypoxia.">Islet Preconditioning via Multimodal Microfluidic Modulation of Intermittent Hypoxia.</a> Anal. Chem. 84 (4), 1987-1993 (2012).</li><li>Qi, M., Barbaro, B., Wang, S., Wang, Y., Hansen, M., Oberholzer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+Pancreatic+Islet+Isolation:+Part+I:+Digestion+and+Collection+of+Pancreatic+Tissue.">Human Pancreatic Islet Isolation: Part I: Digestion and Collection of Pancreatic Tissue.</a> J. Vis. Exp.. , e1125 (2009).</li><li>Qi, M., Barbaro, B., Wang, S., Wang, Y., Hansen, M., Oberholzer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+Pancreatic+Islet+Isolation:+Part+II:+Purification+and+Culture+of+Human+Islets.">Human Pancreatic Islet Isolation: Part II: Purification and Culture of Human Islets.</a> J. Vis. Exp.. , e1343 (2009).</li><li>Shapiro, A. M., 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=Islet+Transplantation+in+Seven+Patients+with+Type+1+Diabetes+Mellitus+Using+a+Glucocorticoid-Free+Immunosuppressive+Regimen.">Islet Transplantation in Seven Patients with Type 1 Diabetes Mellitus Using a Glucocorticoid-Free Immunosuppressive Regimen.</a> N. Engl. J. Med. 343 (4), 230-238 (2000).</li><li>Adewola, A. F., Wang, Y., Harvat, T., Eddington, D. T., Lee, D., Oberholzer, 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+Multi-Parametric+Islet+Perifusion+System+within+a+Microfluidic+Perifusion+Device.">A Multi-Parametric Islet Perifusion System within a Microfluidic Perifusion Device.</a> J. Vis. Exp.. , e1649 (2010).</li><li>Mohammed, J. S., Wang, Y., Harvat, T. A., Oberholzer, J., Eddington, D. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+device+for+multimodal+characterization+of+pancreatic+islets.">Microfluidic device for multimodal characterization of pancreatic islets.</a> Lab Chip. 9, 97-106 (2009).</li><li>Carreras, A., Kayali, F., Zhang, J., Hirotsu, C., Wang, Y., Gozal, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metabolic+Effects+Of+Intermittent+Hypoxia+In+Mice:+Steady+Versus+High+Frequency+Applied+Hypoxia+Daily+During+The+Rest+Period.">Metabolic Effects Of Intermittent Hypoxia In Mice: Steady Versus High Frequency Applied Hypoxia Daily During The Rest Period.</a> AJP - Regu Physiol. 303 (7), 700-709 (2012).</li><li>Lee, E. J., 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=Time-dependent+changes+in+glucose+and+insulin+regulation+during+intermittent+hypoxia+and+continuous+hypoxia.">Time-dependent changes in glucose and insulin regulation during intermittent hypoxia and continuous hypoxia.</a> Eur. J. Appl. Physiol. , (2012).</li><li>Kane, B. J., Zinner, M. J., Yarmush, M. L., Toner, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Liver-specific+functional+studies+in+a+microfluidic+array+of+primary+mammalian+hepatocytes.">Liver-specific functional studies in a microfluidic array of primary mammalian hepatocytes.</a> Anal. Chem. 78, 4291-4298 (2006).</li><li>Lam, R. H. W., Kim, M. C., Thorsen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culturing+aerobic+and+anaerobic+bacteria+and+mammalian+cells+with+a+microfluidic+differential+oxygenator.">Culturing aerobic and anaerobic bacteria and mammalian cells with a microfluidic differential oxygenator.</a> Anal. Chem. 81, 5918-5924 (2009).</li><li>Polinkovsky, M., Gutierrez, E., Levchenko, A., Groisman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fine+temporal+control+of+the+medium+gas+content+and+acidity+and+on-chip+generation+of+series+of+oxygen+concentrations+for+cell+cultures.">Fine temporal control of the medium gas content and acidity and on-chip generation of series of oxygen concentrations for cell cultures.</a> Lab Chip. 9, 1073-1084 (2009).</li><li>Mehta, G., 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=Quantitative+measurement+and+control+of+oxygen+levels+in+microfluidic+poly(dimethylsiloxane)+bioreactors+during+cell+culture.">Quantitative measurement and control of oxygen levels in microfluidic poly(dimethylsiloxane) bioreactors during cell culture.</a> Biomed. Microdev. 9 (2), 123-134 (2007).</li><li>Vollmer, A. P., Probstein, R. F., Gilbert, R., Thorsen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+an+integrated+microfluidic+platform+for+dynamic+oxygen+sensing+and+delivery+in+a+flowing+medium.">Development of an integrated microfluidic platform for dynamic oxygen sensing and delivery in a flowing medium.</a> Lab Chip. 5, 1059-1066 (2005).</li><li>Chen, Y., 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=Generation+of+oxygen+gradients+in+microfluidic+devices+for+cell+culture+using+spatially+confined+chemical+reactions.">Generation of oxygen gradients in microfluidic devices for cell culture using spatially confined chemical reactions.</a> Lab Chip. 11, 3626-3633 (2011).</li><li>Lo, J. F., Sinkala, E., Eddington, D. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxygen+gradients+for+open+well+cellular+cultures+via+microfluidic+substrates.">Oxygen gradients for open well cellular cultures via microfluidic substrates.</a> Lab Chip. 10, 2394-2401 (2010).</li></ol>

The authors declare that they have no competing financial interests.

The multiple modalities integrated in this islet hypoxia technique present several points noted here for troubleshooting. First the isolated islets continue to degrade and disintegrate in culture due to digestive enzymes from acinar cells. Standardizing experiments to 1-2 days after islet isolation is thus critical in obtaining consistent results. Second, the aqueous flow was stopped during hypoxia and intermittent hypoxia to prevent convective clearance at the boundary between laminar flow and diffusion. This seems to l...

Dynamic hypoxia is important in biology, specifically for islet transplants
Dynamic hypoxia is an important physiological as well as pathophysiological parameter in many biological tissues. Change in oxygen, for example, is a potent developmental signal in angiogenesis. Moreover, spatial and temporal patterns in hypoxia modulate HIF1-alpha and play roles in diseases like pancreatic cancer. Hypoxia is also a confounding factor affecting islet transplant outcomes. Recently, temporally oscillations of hypoxia, or intermittent hypoxia (IH) have demonstrated benefits in &#34;preconditioning&#34; islets1. However, both static and transient hypoxia effects on islet physiology have yet to be well understood or studied, primarily due to the lack of appropriate tools to control islet&#39;s microenvironment.
Islets are well vascularized in vivo
Pancreatic islets are 50-400 &#956;m spheroidal aggregates of endocrine cells, including beta-cells and alpha-cells that are responsible for glucose homeostasis. When islets are exposed to stimulatory glucose in the blood, uptake and glycolysis lead to ATP production, which opens up ATP-sensitive potassium (KATP) channels and results in calcium influx that triggers the exocytosis of insulin granules. Oxygen is important to drive this heavily metabolic process and insulin secretion is significantly influenced by the dynamics of blood flow and oxygen supply in addition to glucose gradients. Islets readily perform this glucose-insulin response in vivo as they are highly perfused in the pancreas, each within one cell length from a capillary vessel. However, the dense network of intraislet capillaries is removed by collagenase during islet isolation2,3. Consequently, both oxygen and nutrient supplies are constrained to a 100 &#956;m perimeter due to diffusion limitations.
Current techniques have limited success in recreating islet microenvironment
Recreating islet&#39;s native oxygen and glucose dynamics, key to modeling physiological and pathophysiological conditions, is difficult to achieve with standard hypoxic chambers that require elaborate flow and lack continuous monitoring of islet functions. Moreover, transplant therapies for Type I diabetes expose isolated islets to hypoxia in the hepatic portal system4 which has much lower pO2 (&#60;2%, 5-15 mmHg) compared to physiological pancreas (5.6%, 40 mmHg). Post-transplant, the islet grafts take two weeks or more to be revascularized. It has been demonstrated that hypoxic exposure impairs islet&#39;s glucose-insulin coupling mechanism. Among the stimulus-secretion coupling factors, calcium signaling, mitochondrial potentials, and insulin kinetics can be easily monitored using microfluidics. Our previous microfluidic technique demonstrated this real-time monitoring with precise modulation of the aqueous microenvironment around single islet5,6. However, quantification of islet&#39;s hypoxic impairment is stymied by the lack of simultaneous stimulation and monitoring techniques. Therefore, combining microfluidic control of oxygen and islet monitoring can improve islet hypoxia studies.
Microfluidics can recreate and modulate the aqueous and oxygen microenvironment
The standard technique for tissue and culture hypoxia studies has been based on hypoxic chambers. In general, the hypoxic chambers provide single oxygen concentrations with equilibration times in ~10-30 min, incompatible with minute-scaled dynamic hypoxia. Two recent studies used small custom chambers for intermittent hypoxia exposure on whole mice, with conflicting results on glucose-induced insulin response7,8. Bear in mind that at the whole animal level, the respired oxygen is not directly translated to islet capillary pO2, due to controls in the respiratory system. Furthermore, these studies do not have standardized oxygen levels, nor do they provide real-time measures at the tissue level of islets.
On the other hand, oxygen microfluidics can surpass these limitations by controlling oxygen via gas channel networks. Moreover, microfluidics is compatible with live imaging during oxygen modulation, a feat currently not possible with standard hypoxic chambers. A number of these novel microfluidics approaches utilize the gas permeability of polydimethylsiloxane to dissolve oxygen concentrations into microchannels that flow media over target cells9-14. These devices have also integrated multiple discrete oxygen concentrations, fluorescence based oxygen sensors, and even chemical oxygen generation on-chip.
Liquid solvation-based microfluidics have a hard time maintaining stable, continuous gradients as it depends on convective mixing which is sensitive to flow conditions. In comparison, the technique we use here focuses on decreasing the diffusion path of oxygen delivery. The gas solvation and shear flow are eliminated by directly diffusing oxygen across a membrane seeded with cells or islet tissues. This removes the extra microfluidics required to control solvation and prevents unnecessary shear stress to the islets, which itself can trigger insulin release. This platform has been used to demonstrate reactive oxygen species (ROS) up-regulation at both hyperoxic and hypoxic extremes (2-97% O2) in cell culture1,15. Because of the direct delivery of oxygen and removal of shear flow, our diffusion-based platform provides the optimal microfluidic solution for studying islet hypoxia.
Multimodal stimulation and monitoring
Diffusion-based microfluidics also brings additional benefits when adapted for studying islet microphysiology. By using a membrane as a diffusion barrier, the liquid can be isolated from the oxygen modulations, enabling controls of aqueous glucose stimulations independently from hypoxic stimulations. This creates a sandwich-like simultaneous stimulation that spatially pin-points delivery to the islets. Furthermore, as the gas is temporally modulated via computerized microinjectors, we can modulate the oxygen concentration from 21-0% digitally with transient time less than 60 sec. The dynamic controls of the oxygen and glucose microenvironment at the microscope allow a real-time multimodal protocol that would not be possible or extraordinarily cumbersome using standard hypoxic chambers. Using this device, calcium signaling (Fura-AM), mitochondrial potentials (Rhodamine 123), and insulin kinetics (ELISA) were monitored to provide a complete picture of the dynamic glucose-insulin response under hypoxia.

<table border="1">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagent/Material</td>
		</tr>
		<tr>
			<td>Spinner</td>
			<td>Laurell</td>
			<td>WS-400</td>
			<td></td>
		</tr>
		<tr>
			<td>SU8</td>
			<td>MicroChem</td>
			<td>SU8-2150/SU8-2100</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital Hotplate</td>
			<td>PMC Dataplate</td>
			<td>722A</td>
			<td></td>
		</tr>
		<tr>
			<td>UV Curing Lamp</td>
			<td>OmniCure</td>
			<td>S1000</td>
			<td></td>
		</tr>
		<tr>
			<td>PMDS</td>
			<td>Dow Chemical</td>
			<td>Sylgard 184</td>
			<td></td>
		</tr>
		<tr>
			<td>Corona Wand</td>
			<td>ETP</td>
			<td>BD-20AC</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum Chamber</td>
			<td>Bel-Art</td>
			<td>420220000</td>
			<td></td>
		</tr>
		<tr>
			<td>Microdispensers</td>
			<td>The Lee Company</td>
			<td>IKTX0322000A</td>
			<td></td>
		</tr>
		<tr>
			<td>5 V and 20 V DC Power</td>
			<td>Radio Shack</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NI USB</td>
			<td>National Instrument</td>
			<td>NI USB-6501</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermometer</td>
			<td>Omega Engineering, Inc.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Peristaltic Pump</td>
			<td>Gilson</td>
			<td>Minipulse 2</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygen Sensor</td>
			<td>Ocean Optics</td>
			<td>NeoFox</td>
			<td></td>
		</tr>
		<tr>
			<td>Fraction Collector</td>
			<td>Gilson</td>
			<td>203</td>
			<td></td>
		</tr>
		<tr>
			<td>Pippette</td>
			<td>Fisher Scientific</td>
			<td>Finnpipette II 100&#956;l</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted Epifluorescence Microscope</td>
			<td>Leica</td>
			<td>DMI 4000B</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml Conical Tubes</td>
			<td>Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fura-2 Fluorescence Dye</td>
			<td>Molecular Probes, Life Technologies</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Rhodamine 123 Fluorescence Dye</td>
			<td>Molecular Probes, Life Technologies</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Culture Media</td>
			<td>Sigma-Aldrich</td>
			<td>RPMI-1640</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma-Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>30 in Silicone Tubings</td>
			<td>Cole-Parmer</td>
			<td></td>
			<td>1/16 in x 1/8 in</td>
		</tr>
		<tr>
			<td>1.5 ml Eppendorf Tubes</td>
			<td>Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Y-connectors</td>
			<td>Cole-Parmer</td>
			<td></td>
			<td>1/16 in and 4 mm</td>
		</tr>
		<tr>
			<td>Syringe Connectors</td>
			<td>Cole-Parmer</td>
			<td></td>
			<td>female Luer plug 1/16 in</td>
		</tr>
		<tr>
			<td>Straight Connectors</td>
			<td>Cole-Parmer</td>
			<td></td>
			<td>1/16 in</td>
		</tr>
		<tr>
			<td>Elbow Connector</td>
			<td>Cole-Parmer</td>
			<td></td>
			<td>1/16 in</td>
		</tr>
	</tbody>
</table>

quantitative and temporal control of oxygen microenvironment at the single islet level

Simultaneous oxygenation and monitoring of glucose stimulus-secretion coupling factors in a single technique is critical for modeling pathophysiological states of islet hypoxia, especially in transplant environments. Standard hypoxic chamber techniques cannot modulate both stimulations at the same time nor provide real-time monitoring of glucose stimulus-secretion coupling factors. To address these difficulties, we applied a multilayered microfluidic technique to integrate both aqueous and gas phase modulations via a diffusion membrane. This creates a stimulation sandwich around the microscaled islets within the transparent polydimethylsiloxane (PDMS) device, enabling monitoring of the aforementioned coupling factors via fluorescence microscopy. Additionally, the gas input is controlled by a pair of microdispensers, providing quantitative, sub-minute modulations of oxygen between 0-21%. This intermittent hypoxia is applied to investigate a new phenomenon of islet preconditioning. Moreover, armed with multimodal microscopy, we were able to look at detailed calcium and KATP channel dynamics during these hypoxic events. We envision microfluidic hypoxia, especially this simultaneous dual phase technique, as a valuable tool in studying islets as well as many ex vivo tissues.

Central to this islet hypoxia technique is the ability to modulate aqueous and gaseous phase stimulation in the same microfluidic chamber with minute-scale transients. Figure 1 is a representative result of the a) dual stimulations and b) fast modulations measured within the islet chamber. Aqueous modulation, shown by introduction of fluorescein into the chamber, achieves equilibrium in three to four minutes of mixing. Furthermore, oxygen can be stepped from 5-21% with fast transients, enabling cycling o...

Watch this Scientific Journal Video about Quantitative and Temporal Control of Oxygen Microenvironment at the Single Islet Level at JoVE.com

Quantitative and Temporal Control of Oxygen Microenvironment at the Single Islet Level

Microfluidic oxygen control confers more than just convenience and speed over hypoxic chambers for biological experiments. Especially when implemented via diffusion through a membrane, microfluidic oxygen can provide simultaneous liquid and gas phase modulations at the microscale-level. This technique enables dynamic multi-parametric experiments critical for studying islet pathophysiology.

Simultaneous oxygenation and monitoring of glucose stimulus-secretion coupling factors in a single technique is critical for modeling pathophysiological ...

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JoVE Journal

Bioengineering

9.1K Views.  University of Michigan-Dearborn. Microfluidic oxygen control confers more than just convenience and speed over hypoxic chambers for biological experiments. Especially when implemented via diffusion through a membrane, microfluidic oxygen can provide simultaneous liquid and gas phase modulations at the microscale-level. This technique enables dynamic multi-parametric experiments critical for studying islet pathophysiology.

Video: Quantitative and Temporal Control of Oxygen Microenvironment at the Single Islet Level

Spatio-Temporal Manipulation of Small GTPase Activity at Subcellular Level and on Timescale of Seconds in Living Cells

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular functions and events1, 2. Their spatiotemporally dynamic nature has been revealed by visualization of their activity and localization in real time3. In order to gain deeper understanding of their roles in diverse cellular functions at the molecular level, the next step should be perturbation of protein activities at a precise subcellular location and timing. 
To achieve this goal, we have developed a method for light-induced, spatio-temporally controlled activation of small GTPases by combining two techniques: (1) rapamycin-induced FKBP-FRB heterodimerization and (2) a photo-caging method of rapamycin. With the use of rapamycin-mediated FKBP-FRB heterodimerization, we have developed a method for rapidly inducible activation or inactivation of small GTPases including Rac4, Cdc424, RhoA4 and Ras5, in which rapamycin induces translocation of FKBP-fused GTPases, or their activators, to the plasma membrane where FRB is anchored. For coupling with this heterodimerization system, we have also developed a photo-caging system of rapamycin analogs. A photo-caged compound is a small molecule whose activity is suppressed with a photocleavable protecting group known as a caging group. To suppress heterodimerization activity completely, we designed a caged rapamycin that is tethered to a macromolecule such that the resulting large complex cannot cross the plasma membrane, leading to virtually no background activity as a chemical dimerizer inside cells6. Figure 1 illustrates a scheme of our system. With the combination of these two systems, we locally recruited a Rac activator to the plasma membrane on a timescale of seconds and achieved light-induced Rac activation at the subcellular level6.

A method for spatio-temporal control of small GTPase activity by light is described. This method is based on rapamycin-induced FKBP-FRB heterodimerization and photo-caging systems. Optimization of light-irradiation enables the spatio-temporally controlled activation of small GTPases at the subcellular level.

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular ...

Engineering a Bilayered Hydrogel to Control ASC Differentiation

Natural polymers over the years have gained more importance because of their host biocompatibility and ability to interact with cells in vitro and in vivo. An area of research that holds promise in regenerative medicine is the combinatorial use of novel biomaterials and stem cells. A fundamental strategy in the field of tissue engineering is the use of three-dimensional scaffold (e.g., decellularized extracellular matrix, hydrogels, micro/nano particles) for directing cell function. This technology has evolved from the discovery that cells need a substrate upon which they can adhere, proliferate, and express their differentiated cellular phenotype and function 2-3. More recently, it has also been determined that cells not only use these substrates for adherence, but also interact and take cues from the matrix substrate (e.g., extracellular matrix, ECM)4. Therefore, the cells and scaffolds have a reciprocal connection that serves to control tissue development, organization, and ultimate function. Adipose-derived stem cells (ASCs) are mesenchymal, non-hematopoetic stem cells present in adipose tissue that can exhibit multi-lineage differentiation and serve as a readily available source of cells (i.e. pre-vascular endothelia and pericytes). Our hypothesis is that adipose-derived stem cells can be directed toward differing phenotypes simultaneously by simply co-culturing them in bilayered matrices1. Our laboratory is focused on dermal wound healing. To this end, we created a single composite matrix from the natural biomaterials, fibrin, collagen, and chitosan that can mimic the characteristics and functions of a dermal-specific wound healing ECM environment.

This protocol focuses on utilizing the inherent ability of stem cells to take cue from their surrounding extracellular matrix and be induced to differentiate into multiple phenotypes. This methods manuscript extends our description and characterization of a model utilizing a bilayered hydrogel, composed of PEG-fibrin and collagen, to simultaneously co-differentiate adipose-derived stem cells1.

Natural polymers over the years have gained more importance because of their host biocompatibility and ability to interact with cells in vitro and in ...

Nanomanipulation of Single RNA Molecules by Optical Tweezers

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be increasingly important. As RNA function often depends on its ability to adopt alternative structures, it is difficult to predict RNA three-dimensional structures directly from sequence. Single-molecule approaches show potentials to solve the problem of RNA structural polymorphism by monitoring molecular structures one molecule at a time. This work presents a method to precisely manipulate the folding and structure of single RNA molecules using optical tweezers. First, methods to synthesize molecules suitable for single-molecule mechanical work are described. Next, various calibration procedures to ensure the proper operations of the optical tweezers are discussed. Next, various experiments are explained. To demonstrate the utility of the technique, results of mechanically unfolding RNA hairpins and a single RNA kissing complex are used as evidence. In these examples, the nanomanipulation technique was used to study folding of each structural domain, including secondary and tertiary, independently. Lastly, the limitations and future applications of the method are discussed.

Optical tweezers have been used to study RNA folding by stretching individual molecules from their 5&#8217; and 3&#8217; ends. Here common procedures are described to synthesize RNA molecules for tweezing, calibration of the instrument, and methods to manipulate single molecules.

A large portion of the human genome is transcribed but not translated. In this post genomic era, regulatory functions of RNA have been shown to be ...

A Label-free Technique for the Spatio-temporal Imaging of Single Cell Secretions

Inter-cellular communication is an integral part of a complex system that helps in maintaining basic cellular activities. As a result, the malfunctioning of such signaling can lead to many disorders. To understand cell-to-cell signaling, it is essential to study the spatial and temporal nature of the secreted molecules from the cell without disturbing the local environment. Various assays have been developed to study protein secretion, however, these methods are typically based on fluorescent probes which disrupt the relevant signaling pathways. To overcome this limitation, a label-free technique is required.
In this paper, we describe the fabrication and application of a label-free localized surface plasmon resonance imaging (LSPRi) technology capable of detecting protein secretions from a single cell. The plasmonic nanostructures are lithographically patterned onto a standard glass coverslip and can be excited using visible light on commercially available light microscopes. Only a small fraction of the coverslip is covered by the nanostructures and hence this technique is well suited for combining common techniques such as fluorescence and bright-field imaging.
A multidisciplinary approach is used in this protocol which incorporates sensor nanofabrication and subsequent biofunctionalization, binding kinetics characterization of ligand and analyte, the integration of the chip and live cells, and the analysis of the measured signal. As a whole, this technology enables a general label-free approach towards mapping cellular secretions and correlating them with the responses of nearby cells.

Inter-cellular communication is critical for controlling various physiological activities within and outside the cell. This paper describes a protocol for measuring the spatio-temporal nature of single cell secretions. To achieve this, a multidisciplinary approach is used which integrates label-free nanoplasmonic sensing with live cell imaging.

Inter-cellular communication is an integral part of a complex system that helps in maintaining basic cellular activities. As a result, the malfunctioning ...

Engineering Molecular Recognition with Bio-mimetic Polymers on Single Walled Carbon Nanotubes

Semiconducting single-wall carbon nanotubes (SWNTs) are a class of optically active nanomaterial that fluoresce in the near infrared, coinciding with the optical window where biological samples are most transparent. Here, we outline techniques to adsorb amphiphilic polymers and polynucleic acids onto the surface of SWNTs to engineer their corona phases and create novel molecular sensors for small molecules and proteins. These functionalized SWNT sensors are both biocompatible and stable. Polymers are adsorbed onto the nanotube surface either by direct sonication of SWNTs and polymer or by suspending SWNTs using a surfactant followed by dialysis with polymer. The fluorescence emission, stability, and response of these sensors to target analytes are confirmed using absorbance and near-infrared fluorescence spectroscopy. Furthermore, we demonstrate surface immobilization of the sensors onto glass slides to enable single-molecule fluorescence microscopy to characterize polymer adsorption and analyte binding kinetics.

We present a protocol for engineering the corona phase of near infrared fluorescent single walled carbon nanotubes (SWNTs) using amphiphilic polymers and DNA to develop sensors for molecular targets without known recognition elements.

Semiconducting single-wall carbon nanotubes (SWNTs) are a class of optically active nanomaterial that fluoresce in the near infrared, coinciding with the ...

A Microfluidic System with Surface Patterning for Investigating Cavitation Bubble(s)&#8211;Cell Interaction and the Resultant Bioeffects at the Single-cell Level

In this manuscript, we first describe the fabrication protocol of a microfluidic chip, with gold dots and fibronectin-coated regions on the same glass substrate, that precisely controls the generation of tandem bubbles and individual cells patterned nearby with well-defined locations and shapes. We then demonstrate the generation of tandem bubbles by using two pulsed lasers illuminating a pair of gold dots with a few-microsecond time delay. We visualize the bubble-bubble interaction and jet formation by high-speed imaging and characterize the resultant flow field using particle image velocimetry (PIV). Finally, we present some applications of this technique for single cell analysis, including cell membrane poration with macromolecule uptake, localized membrane deformation determined by the displacements of attached integrin-binding beads, and intracellular calcium response from ratiometric imaging. Our results show that a fast and directional jetting flow is produced by the tandem bubble interaction, which can impose a highly localized shear stress on the surface of a cell grown in close proximity. Furthermore, different bioeffects can be induced by altering the strength of the jetting flow by adjusting the standoff distance from the cell to the tandem bubbles.

A Microfluidic System with Surface Patterning for Investigating Cavitation Bubble(s)&amp;#8211;Cell Interaction and the Resultant Bioeffects at the Single-cell Level

A microfluidic chip was fabricated to produce pairs of gold dots for tandem bubble generation and fibronectin-coated islands for single-cell patterning nearby. The resultant flow field was characterized by particle image velocimetry and was employed to study various bioeffects, including cell membrane poration, membrane deformation, and intracellular calcium response.

In this manuscript, we first describe the fabrication protocol of a microfluidic chip, with gold dots and fibronectin-coated regions on the same glass ...

An Ultrahigh-throughput Microfluidic Platform for Single-cell Genome Sequencing

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of cellular heterogeneity in biological systems. However, single-cell sequencing of large populations has been hampered by limitations in processing genomes for sequencing. In this paper, we describe a method for single-cell genome sequencing (SiC-seq) which uses droplet microfluidics to isolate, amplify, and barcode the genomes of single cells. Cell encapsulation in microgels allows the compartmentalized purification and tagmentation of DNA, while a microfluidic merger efficiently pairs each genome with a unique single-cell oligonucleotide barcode, allowing &gt;50,000 single cells to be sequenced per run. The sequencing data is demultiplexed by barcode, generating groups of reads originating from single cells. As a high-throughput and low-bias method of single-cell sequencing, SiC-seq will enable a broader range of genomic studies targeted at diverse cell populations.

Single-cell sequencing reveals genotypic heterogeneity in biological systems, but current technologies lack the throughput necessary for the deep profiling of community composition and function. Here, we describe a microfluidic workflow for sequencing &gt;50,000 single-cell genomes from diverse cell populations.

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of ...

Patterning the Geometry of Human Embryonic Stem Cell Colonies on Compliant Substrates to Control Tissue-Level Mechanics

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that correspond to primary germ layer formation during embryogenesis. Thus, these cells represent a powerful tool with which to examine the mechanisms that drive early human development. We have developed a method to culture human embryonic stem cells in confined colonies on compliant substrates that provides control over both the geometry of the colonies and their mechanical environment in order to recapitulate the physical parameters that underlie embryogenesis. The key feature of this method is the ability to generate polyacrylamide hydrogels with defined patterns of extracellular matrix ligand at the surface to promote cell attachment. This is achieved by fabricating stencils with the desired geometric patterns, using these stencils to create patterns of extracellular matrix ligand on glass coverslips, and transferring these patterns to polyacrylamide hydrogels during polymerization. This method is also compatible with traction force microscopy, allowing the user to measure and map the distribution of cell-generated forces within the confined colonies. In combination with standard biochemical assays, these measurements can be used to examine the role mechanical cues play in fate specification and morphogenesis during early human development.

Extracellular matrix ligands can be patterned onto polyacrylamide hydrogels to enable the culture of human embryonic stem cells in confined colonies on compliant substrates. This method can be combined with traction force microscopy and biochemical assays to examine the interplay between tissue geometry, cell-generated forces, and fate specification.

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that ...

Finite Element Modelling of a Cellular Electric Microenvironment

Clinical studies show electrical stimulation (ES) to be a potential therapy for the healing and regeneration of various tissues. Understanding the mechanisms of cell response when exposed to electrical fields can therefore guide the optimization of clinical applications. In vitro experiments aim to help uncover those, offering the advantage of wider input and output ranges that can be ethically and effectively assessed. However, the advancements in in vitro experiments are difficult to reproduce directly in clinical settings. Mainly, that is because the ES devices used in vitro differ significantly from the ones suitable for patient use, and the path from the electrodes to the targeted cells is different. Translating the in vitro results into in vivo procedures is therefore not straightforward. We emphasize that the cellular microenvironment's structure and physical properties play a determining role in the actual experimental testing conditions and suggest that measures of charge distribution can be used to bridge the gap between in vitro and in vivo. Considering this, we show how in silico finite element modelling (FEM) can be used to describe the cellular microenvironment and the changes generated by electric field (EF) exposure. We highlight how the EF couples with geometric structure to determine charge distribution. We then show the impact of time dependent inputs on charge movement. Finally, we demonstrate the relevance of our new in silico model methodology using two case studies: (i) in vitro fibrous Poly(3,4-ethylenedioxythiophene) poly(styrenesulfonate) (PEDOT-PSS) scaffolds and (ii) in vivo collagen in extracellular matrix (ECM).

This paper presents a strategy for building finite element models of fibrous conductive materials exposed to an electric field (EF). The models can be used to estimate the electrical input that cells seeded in such materials receive and assess the impact of changing the scaffold's constituent material properties, structure or orientation.

Clinical studies show electrical stimulation (ES) to be a potential therapy for the healing and regeneration of various tissues. Understanding the ...

Establishing an Octopus Ecosystem for Biomedical and Bioengineering Research

Many developments in biomedical research have been inspired by discovering anatomical and cellular mechanisms that support specific functions in different species. The octopus is one of these exceptional animals that has given scientists new insights into the fields of neuroscience, robotics, regenerative medicine, and prosthetics. Research with this species of cephalopods requires the set-up of complex facilities and intensive care for both the octopus and its ecosystem that is critical for the project&#39;s success. This system requires multiple mechanical and biological filtering systems to provide a safe and clean environment for the animal. Along with the control system, specialized routine maintenance and cleaning are required to effectively keep the facility operating long term. It is advised to provide an enriched environment to these intelligent animals by changing the tank&#39;s landscape, incorporating a variety of prey, and introducing challenging tasks for them to work through. Our results include MRI and a whole-body autofluorescence imaging as well as behavioral studies to better understand their nervous system. Octopuses possess unique physiology that can impact many areas of biomedical research. Providing them with a sustainable ecosystem is the first crucial step in uncovering their distinct capabilities.

Understanding the unique physiological and anatomical structures of octopuses can greatly impact biomedical research. This guide demonstrates how to set-up and maintain a marine environment to accommodate this species and includes state-of-the-art imaging and analytical approaches to visualize octopus' nervous system anatomy and function.

Many developments in biomedical research have been inspired by discovering anatomical and cellular mechanisms that support specific functions in different ...