This work was funded by the Dorothy J. Wingfield Phillips Chancellor Faculty Fellowship. Emily Kight was funded by NSF GRFP.

1. Fabricate and assemble the urinalysis device
<ol>
	<li>Fabricate the base plate (Figure 1A).
	<ol>
		<li>Use a computer-aided design (CAD) software to draw a rectangular area with dimensions 2.1641 in x 0.0547 in x 6.3828 in (W x H x L) using the polyline tool.</li>
		<li>Measure the test area (rectangular area encompassing the distance between the first and last pad and the width of the pads) on the dipstick. 
		NOTE: This information is needed to draw the through-holes that hold the dipstick in place and separate the liquid in between the pads (to prevent cross-contamination).</li>
		<li>Add through-holes that mimic the size and position of each test pad in the test area.</li>
		<li>Draw two raised side ledges that measure 2.1641 in x 0.6797 in (W x L).</li>
		<li>Draw a stop (0.1172 in by 0.2109 in (W x L)) using the polyline tool to facilitate alignment between the base plate and the slide. The stop should be perpendicular to the ledges and physically stops the slide from moving passed the urine dipstick pads.</li>
		<li>Select the lines for the stop and ledge to make one region using the Region command. Use the Extrude command to raise the region up to a height of 0.0703 in. Repeat this step on the other side of the device.</li>
		<li>Create a notch (0.1895 in by 0.3500 in (W x L)) on both ledges to facilitate alignment with the box. Position it 0.466 in from the bottom edge of the ledge. Use the Region command to create a rectangle and make the extrusion height 0.1250 in.</li>
		<li>Use the Solid Subtract command, select device, press Enter, select the notch region and press Enter. Repeat on the other side of the device. 
		NOTE: The shape will be removed from the device.</li>
		<li>Print the base plate on a 3D printer and sand the top face area between the ledges with sandpaper to roughen the surface. 
		NOTE: Sanding is important so that the hydrophobic coating can adhere to the base plate securely.</li>
		<li>Tape the ledges with adhesive tape (to avoid spraying the ledges) and spray the base plate with a hydrophobic spray. Apply several (4-8) coats of the basecoat to the base plate. Hold the can approximately 8-12 inches away from the base plate when spraying. The device should have a milky white appearance upon drying. 
		CAUTION: Follow manufacturer instructions for appropriate location and PPE for spraying.</li>
		<li>Wait 30 minutes before applying the topcoat several times (6-8x). Allow the base plate to dry for 12 hours before use. Remove the tape from the ledges.</li>
	</ol>
	</li>
	<li>Fabricate the top plate (Figure 1B).
	<ol>
		<li>Draw a rectangular area to measure 2.05 in x 5.470 in (W x L) in a CAD software using the polyline tool.</li>
		<li>Add a rectangular through-hole (the &#8220;viewing through-hole&#8221;) slightly larger than the size of the test area of the dipstick (e.g., 0.230 in x 3.147 in (W x L)). Place it 0.921 in from the top, 1.165 in from the left, and 1.165 in from the right edges of the top plate.</li>
		<li>Draw a second through-hole (the &#8220;Inlet-hole&#8221;) sized 0.075 in x 3.146 in (W x L). Place it 0.236 in from the bottom edge, 1.737 in from the top edge, and 1.162 in from the left and right edges of the top plate.</li>
		<li>Cut the top plate from a piece of clear acrylic with a laser cutter. Wipe off any remaining dust or debris.</li>
	</ol>
	</li>
	<li>Fabricate the inlet cover (Figure 1B).
	<ol>
		<li>Draw a rectangular area with dimensions 0.247 in x 3.3378 in (W x L) in a CAD software using the polyline tool. Add two circular through-holes with a diameter of 0.127 in approximately 0.073 in from the two edges of the inlet cover, one on either side.</li>
		<li>Cut the inlet cover from a piece of clear acrylic with a laser cutter.</li>
	</ol>
	</li>
	<li>Fabricate the slide (Figure 1C)
	<ol>
		<li>Draw a rectangular area in CAD software that measures measure 2.771 in x 0.0625 in x 5.000 in (W x H x L) using the polyline tool.</li>
		<li>Add through-holes that match the position of each test pad in the test area. Draw the first 0.105 in square through-hole to overlap with the placement of the first test pad: 1.096 in from the left and right edges of the slide, 0.960 in from the top edge, and 1.681 in from the bottom edge. Add more through-holes as needed (usually 10 total) for the selected dipstick brand of choice. Space each next through-hole by measuring the distance between test pads on the dipstick. 
		NOTE: The size of the through-holes is important in order to deposit the correct volume of liquid onto the dipstick pad. For our brand of dipstick, we created holes which deposit 15 ul onto each dipstick pad.</li>
		<li>Cut the slide from a piece of clear acrylic using a laser cutter. Wipe off any remaining dust or debris.</li>
		<li>Spray the front of the slide with a hydrophobic spray. Apply several coats (6-8x) of basecoat to the slide. Hold the can approximately 8 -12 in away from the slide when spraying.</li>
		<li>Wait 30 minutes before applying the topcoat several times (8-12x). Allow the slide to dry for 12 hours before use.</li>
		<li>Download a QR code from an online QR code generator and print the desired code on paper with sticky adhesive backing. Place the QR code 0.17 in from the right of the first through-hole along the same row as all the through-holes. 
		NOTE: As long as the QR code is adjacent to the through-holes, accurate placement is not important.</li>
		<li>Use clear tape to cover the QR code and secure it to the slide.</li>
	</ol>
	</li>
	<li>Assemble the inlet and plate sleeve (Figure 1D).
	<ol>
		<li>Fabricate the inlet by using acrylic cement to glue the inlet cover onto the top plate where the inlet-hole is located. Wait 24-48 hours to securely bond the pieces.</li>
		<li>Spray the back of the top plate with a hydrophobic spray once the inlet cover is securely bonded to the top plate. Place the top plate upside down. Apply the first basecoat several times (4-8x).</li>
		<li>Hold the spray 8-12 inches away from the top plate and wait 30 minutes for it to dry. Apply the topcoat several times (6-8x). Allow the top plate to dry for 12 hours before use.</li>
		<li>Assemble the plate sleeve (combined top plate and base plate) by gluing the completed top plate to the ledges of the base plate with acrylic cement. The two pieces are easy to align by visual inspection, as the bottom edge of the top plate will align with that of the base plate. Apply a clamp to the base plate ledges to secure it during drying and wait 24-48 hours before use, as per the manufacturer&#8217;s instructions.</li>
	</ol>
	</li>
	<li>Create the chart sticker.
	<ol>
		<li>Download the color chart for the brand of dipstick from the manufacturer&#8217;s website.</li>
		<li>Open the downloaded file in a graphics editor software.</li>
		<li>Open the digital file for the top plate template previously used for the laser cutter (Step 1.2 of this protocol) in a graphics editor software.</li>
		<li>Create the color boxes for the chart sticker by matching color boxes from the manufacturer color chart. Select the first block of color on the manufacturer&#8217;s chart with the dropper tool in the graphics editor software and then use the box shape tool to make a box shape in the same color on the top plate template, in the same row where the dipstick pad will be located. Repeat this for each color block corresponding to each pad row.</li>
		<li>Delete the layers associated with the top plate template.</li>
		<li>Print the chart sticker as a vinyl sticker with an online sticker print service. Place the chart sticker onto the plate sleeve and align it with each through-hole.</li>
	</ol>
	</li>
	<li>Fabricate the box (Figure 1E).
	<ol>
		<li>Draw the two long-sided box pieces (parts &#8220;a&#8221; and &#8220;b&#8221;) in the CAD software as rectangles with dimensions of 4.92 in x 6.63 in (W x L). Add a cut-out to part &#8220;a&#8221; centered on the bottom edge measuring 0.2 in x 6.11 in (W x L).</li>
		<li>Draw the two narrow-sided box pieces (parts &#8220;d&#8221; and &#8220;e&#8221;) in the CAD software as rectangles with dimensions measuring 1.805 in x 6.63 in (W x L).</li>
		<li>Draw the box top (part &#8220;c&#8221;) as a rectangle with dimensions 1.805 in x 6.63 (W x L). Draw the &#8220;imaging through-hole&#8221; on the top: 0.74 in x 0.910 in (W x L), positioned 3.17 in from the bottom, 2.53 in from the top, 0.65 in from the right edge, and 0.42 in from the left edge. 
		NOTE: The exact position of the imaging through-hole should be selected on the basis of the cell phones that will be used for the analysis.</li>
		<li>Draw each box piece to feature a pattern of interlocking edges that will allow all the box sides to snap together on each edge as described in Figure 1D. To make an interlock edge pattern, alternate an extrusion/intrusion pattern on the long edge with 0.135 in by 1.17 in (W x L) protrusions. Draw two extrusions on each long edge for every side of the box. Use the same extrusion/intrusion pattern for the short edge, but with intrusions measuring 0.135 in by 0.460 in (W x L).</li>
		<li>Cut the five pieces with a laser cutter or print them with a 3D printer. 
		NOTE: A laser-cut component using acrylic pieces will be cheap to manufacture and can be flattened for easy shipping. Use black acrylic as it is helpful to absorb scattered light during testing.</li>
		<li>Add black color construction paper to the box interior to prevent scatter from the flash during image analysis if the box material has a gloss finish.</li>
	</ol>
	</li>
</ol>
2. Prepare the test
<ol>
	<li>Download the UrineTest mobile application from GitHub (https://github.com/Iftak/UrineTestApp).&#160;</li>
	<li>Install the app onto a mobile phone.&#160; 
	NOTE: This step only has to be done once for all future uses of a given phone. If needed, enable developer status on the phone to do this.&#160;</li>
	<li>Launch the UrineTest application in the phone (Figure 2A).</li>
	<li>Read the instructions to change the analyte names and reading timings (Figure 2B) to match those for the dipstick of interest (based on the manufacturer&#8217;s specifications) and insert new input via the text holder window on the screen (Figure 2C). 
	NOTE: The necessary readout time for each dipstick pad will depend on the brand of the dipstick used.</li>
	<li>Assemble the various components together and insert the dipstick into the through-holes underneath the plate sleeve (Figure 1F).&#160;</li>
	<li>Place the plate sleeve inside the box so that its notch is aligned with the box gap.&#160;</li>
	<li>Place the slide inside the plate sleeve so that its through-holes align with the inlet.</li>
	<li>Place the phone on the top of the box with the back-camera lens facing the viewing through-hole to enable imaging. Ensure the camera visibility is not occluded by checking for the image on the phone screen prior to testing. The app will enable the flashlight on the phone automatically.</li>
	<li>Read the instruction for phone alignment (Figure 2D) and align the phone accordingly so that the dipstick coincides with the boundaries of the black rectangular overlay on screen (Figure 2E).</li>
	<li>Click the Start button on the app window to begin the test. 
	NOTE: This will open the phone camera to read the QR code once in view (Figure 2F).</li>
</ol>
3. Conduct the test
<ol>
	<li>Deposit urine into the inlet hole with a disposable polyethylene transfer pipet containing approximately 0.5 mL of urine (Figure 3). 
	NOTE: The exact amount of liquid is not important, but it should be at least 0.5 mL to ensure that all the through-holes receive sufficient urine. Upon adding the liquid, observe that it moves across inlet and is deposited in each through-hole of the slide.</li>
	<li>Initiate the test by pushing the slide into the plate sleeve until it is stopped by the base plate stop. 
	NOTE: Urine should make contact with dipstick pad when the QR code is in the field of view of the cell phone. After reading the QR code, the application will open a window to analyze the color changes (Figure 2G) and show the results automatically within the same window (Figure 2H).</li>
	<li>Discard the urine appropriately and clean the plate sleeve and slide with 10% bleach solution and rinse again with de-ionized water. Allow it to dry before additional use.</li>
</ol>

<ol>
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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Technology+Behind+Diagnostic+Reagent+Strips.">Technology Behind Diagnostic Reagent Strips.</a> Laboratory Medicine. 31, 92-96 (2000).</li><li>Dungchai, W., Chailapakul, O., Henry, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrochemical+detection+for+paper-based+microfluidics.">Electrochemical detection for paper-based microfluidics.</a> Analytical Chemistry. 81, 5821-5826 (2009).</li><li>Van Delft, S., 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=Prospective,+observational+study+comparing+automated+and+visual+point-of-care+urinalysis+in+general+practice.">Prospective, observational study comparing automated and visual point-of-care urinalysis in general practice.</a> BMJ Open. 6, 1-7 (2016).</li><li>. Urisys 1100 Analyzer Available from: <a target="_blank" href="https://diagnostics.roche.com/us/en/products/instruments/urisys-1100.html">https://diagnostics.roche.com/us/en/products/instruments/urisys-1100.html</a> (2020)</li><li>Filippini, D., Lundstr&#246;m, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+strategy+and+instrumental+performance+of+a+computer+screen+photo-assisted+technique+for+the+evaluation+of+a+multi-parameter+colorimetric+test+strip.">Measurement strategy and instrumental performance of a computer screen photo-assisted technique for the evaluation of a multi-parameter colorimetric test strip.</a> Analyst. 131, 111-117 (2006).</li><li>Shen, L., Hagen, J. 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The authors have nothing to disclose.

Traditional dipstick urinalysis is affordable and convenient but requires manual attention to detail to yield accurate results. Manual dipstick urinalysis is subject to variable lighting conditions, individual color perception differences and cross-contamination. Many clinics and hospitals already have instruments to automate urine dipstick analysis, but the instruments are usually bulky, expensive, and still rely on proper performance of the dip-wipe method. Additionally, these instruments require yearly calibration and...

Urine is a non-invasive source of multiple metabolic indicators of disease or health. Urinalysis, the physical and/or chemical analysis of urine, can be performed quickly to detect renal disease, urinary tract disease, liver disease, diabetes mellitus, and general hydration1. Urinalysis dipsticks are affordable, semi-quantitative diagnostic tools that rely on colorimetric changes to indicate approximate physiological levels. Each dipstick can perform a wide variety of assays including testing for pH, osmolality, hemoglobin/myoglobin, hematuria, leukocyte esterase, glucose, proteinuria, nitrite, ketone, and bilirubin2. The principle of dipstick urinalysis relies on the occurrence of a timed reaction through which a color change on the dipstick pad can be compared to a chart to determine analyte concentration3. Given their affordability and ease of use, dipsticks are one of the most common tools for urinalysis in healthcare.
Traditionally, dipstick urinalysis relies on a trained nurse or medical technician to manually insert the dipstick into a cup of urine sample, wipe off excess urine, and compare the color pads to chart colors at specific times. While the dip-and-wipe method is the gold standard for dipstick analysis, its reliance on human visual assessment limits the quantitative information that can be obtained. Moreover, the two manual steps of dipstick urinalysis &#8211; the dip-wipe step and colorimetric result comparison &#8211; require accurate technique, which limits the possibility of reliable testing in home settings by patients directly. Cross-contamination of the sample pads due to wiping can cause inaccurate color changes. Additionally, inconsistent volumes resulting from the lack of volume control during wiping can result in improper measurement of analyte concentrations. Importantly, the time between dipping the urine (i.e., the start of the assay) and comparison to a chart is critical for accurate analysis of the results and is a huge potential source of human error. The difficulty in manual colorimetric comparison is that many pads must be read at the same time, while some pads are read at different times. Even perfectly timed color comparisons still depend on the visual acuity of the human reader, who may suffer from color blindness or perceive different colors in different lighting environments4. These challenges underscore why clinicians can only rely on dipstick urinalysis performed by trained personnel. However, an automated urinalysis system could address all the aforementioned concerns by eliminating the need for manual dip-wipe steps, incorporating timing controls, and enabling simultaneous color comparisons with calibrated color references. This, in turn, would reduce user error, allowing for possible adoption in home settings.
In the last 20 years, automatic analyzers have been employed to read the results of dipstick urine tests with the same accuracy as or exceeding visual analysis5. Many clinics and doctor offices use such machines to rapidly analyze and print traditional dipstick results. Most urinalysis machines minimize visual inspection errors and ensure consistency in results6. They are easy to use and more efficient than manual inspection but still require the user to perform the dip-wipe method correctly. Hence, these machines have limited ability to be operated by untrained persons such as at-home users; moreover, they are extremely expensive.
Recently, cell phones have emerged as a resourceful tool for various biological colorimetric measurements7,8,9,10, including for urinalysis11,12,13. Given their remote sensing capabilities and high imaging resolution, cell phones have become effective healthcare analytical devices14,15. Indeed, the FDA has cleared several smartphone-based home urine tests16,17,18. Some of the new smartphone-based commercial products incorporate established urinalysis dipsticks, while others feature proprietary colorimetric pads. All such products feature proprietary methods to calibrate for different lighting conditions across different phones types. Still, a problem with these solutions is that the user must manually take a picture at the right time in addition to carrying out a proper manual dip-wipe method (i.e., without cross-contamination). Notably, none of these tests control the volume deposited onto the dipsticks, which we have found can affect the color change19 and interpreted physiological result. The present gaps and costs in the workflows of these devices suggest an additional need to enable a human-free, volume-controlled urine deposition procedure and hands-free dipstick photography.
We describe a protocol for volume-controlled, automated dipstick urinalysis without the need for a manual dip-wipe step. The key to the automated process is a device19 whose underlying principle is based on the SlipChip20 and that transfers liquid between different layers using surface chemistry effects. In brief, the hydrophobic coating on the transfer slide and surrounding plate sleeve force the liquid to move effortlessly through the device and to release onto the dipstick pad once the slide is in its final position, at which point the bottom hydrophobic barrier is replaced with air. Additionally, the coordinated light-blocking box standardizes the lighting conditions, camera angle of view, and the distance for camera focus to ensure accurate and repeatable results that are not influenced by ambient lighting conditions. An accompanying software app automates the capture of images and colorimetric analysis. Following description of the protocol, we provide representative results of the urinalysis test under different conditions. Comparisons with the standard dip-wipe method demonstrate reliability of the proposed method.

<table>
	<tbody>
		<tr>
			<td>Black Cast Acrylic Sheet 
			12&#34; x 24&#34; x 1/8&#34;</td>
			<td>McMaster Carr</td>
			<td>8505K742</td>
			<td>$14.27</td>
		</tr>
		<tr>
			<td>Chart sticker</td>
			<td>Stickeryou.com</td>
			<td></td>
			<td>$12.39</td>
		</tr>
		<tr>
			<td>Clear Scratch- and UV-Resistant Cast Acrylic Sheet 
			12&#34; x 24&#34; x 1/16&#34;</td>
			<td>McMaster Carr</td>
			<td>8560K172</td>
			<td>$9.52</td>
		</tr>
		<tr>
			<td>disposable polyethylene transfer pipet</td>
			<td>Fischer Brand</td>
			<td>13-711-9AM</td>
			<td>lot# 14311021</td>
		</tr>
		<tr>
			<td>Fortus ABS-M30</td>
			<td>Stratasys</td>
			<td>345-42207</td>
			<td>lot# : 108078</td>
		</tr>
		<tr>
			<td>Githut: https://github.com/Iftak/UrineTestApp</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Innovating Science - Replacement Fluids for Urinalysis Diagnostic Test Kit (IS3008)</td>
			<td>Amazon</td>
			<td></td>
			<td>$49</td>
		</tr>
		<tr>
			<td>Nonwhitening Cement for Acrylic 
			Scigrip 4, 4 oz. Can</td>
			<td>MCM</td>
			<td>7517A1</td>
			<td>$9.22</td>
		</tr>
		<tr>
			<td>Rust-Oleum 274232 Repelling treatment base coat-9 oz and top-coat 9-oz , Frosted Clear</td>
			<td>Amazon</td>
			<td>Color: Frosted Clear</td>
			<td>$6.99</td>
		</tr>
		<tr>
			<td>Urinalysis Reagent Strips 10 Panel (100 Tests) MISSION BRAND</td>
			<td>Medimpex United, Inc</td>
			<td>MUI-MS10</td>
			<td>$10.59</td>
		</tr>
	</tbody>
</table>

low-cost, volume-controlled dipstick urinalysis for home-testing

Dipstick urinalysis provides quick and affordable estimations of multiple physiological conditions but requires good technique and training to use accurately. Manual performance of dipstick urinalysis relies on good human color vision, proper lighting control, and error-prone, time-sensitive comparisons to chart colors. By automating the key steps in the dipstick urinalysis test, potential sources of error can be eliminated, allowing self-testing at home. We describe the steps necessary to create a customizable device to perform automated urinalysis testing in any environment. The device is cheap to manufacture and simple to assemble. We describe the key steps involved in customizing it for the dipstick of choice and for customizing a mobile phone app to analyze the results. We demonstrate its use to perform urinalysis and discuss the critical measurements and fabrication steps necessary to ensure robust operation. We then compare the proposed method to the dip-and-wipe method, the gold standard technique for dipstick urinalysis.

Figure 4 demonstrates how the urine is transferred to the dipstick during a urinalysis test. During a typical test, the transfer of urine is not observable because the box occludes the view. Once the sample is deposited in the inlet using a pipette (Step 3.1), it will fill the holes on the slide (Figure 4A). Figure 4B and Figure 4C, respectively, show the progressive movement of the urine across the pla...

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Low-Cost, Volume-Controlled Dipstick Urinalysis for Home-Testing

Dipstick urinalysis is a quick and affordable method of assessing one&#8217;s personal state of health. We present a method to perform accurate, low-cost dipstick urinalysis that removes the primary sources of error associated with traditional dip-and-wipe protocols and is simple enough to be performed by lay users at home.

Dipstick urinalysis provides quick and affordable estimations of multiple physiological conditions but requires good technique and training to use ...

low-cost-volume-controlled-dipstick-urinalysis-for-home-testing

Research

JoVE Journal

Bioengineering

5.4K Views.  Vanderbilt University. Dipstick urinalysis is a quick and affordable method of assessing one’s personal state of health. We present a method to perform accurate, low-cost dipstick urinalysis that removes the primary sources of error associated with traditional dip-and-wipe protocols and is simple enough to be performed by lay users at home.

Video: Low-Cost, Volume-Controlled Dipstick Urinalysis for Home-Testing

Procedure for Lung Engineering

Lung tissue, including lung cancer and chronic lung diseases such as chronic obstructive pulmonary disease, cumulatively account for some 280,000 deaths annually; chronic obstructive pulmonary disease is currently the fourth leading cause of death in the United States1. Contributing to this mortality is the fact that lungs do not generally repair or regenerate beyond the microscopic, cellular level. Therefore, lung tissue that is damaged by degeneration or infection, or lung tissue that is surgically resected is not functionally replaced in vivo. To explore whether lung tissue can be generated in vitro, we treated lungs from adult rats using a procedure that removes cellular components to produce an acellular lung extracellular matrix scaffold. This scaffold retains the hierarchical branching structures of airways and vasculature, as well as a largely intact basement membrane, which comprises collagen IV, laminin, and fibronectin. The scaffold is mounted in a bioreactor designed to mimic critical aspects of lung physiology, such as negative pressure ventilation and pulsatile vascular perfusion. By culturing pulmonary epithelium and vascular endothelium within the bioreactor-mounted scaffold, we are able to generate lung tissue that is phenotypically comparable to native lung tissue and that is able to participate in gas exchange for short time intervals (45-120 minutes). These results are encouraging, and suggest that repopulation of lung matrix is a viable strategy for lung regeneration. This possibility presents an opportunity not only to work toward increasing the supply of lung tissue for transplantation, but also to study respiratory cell and molecular biology in vitro for longer time periods and in a more accurate microenvironment than has previously been possible.

We have developed a decellularized lung extracellular matrix and novel biomimetic bioreactor that can be used to generate functional lung tissue. By seeding cells into the matrix and culturing in the bioreactor, we generate tissue that demonstrates effective gas exchange when transplanted in vivo for short periods of time.

Lung tissue, including lung cancer and chronic lung diseases such as chronic obstructive pulmonary disease, cumulatively account for some 280,000 deaths ...

Low-Cost Cryo-Light Microscopy Stage Fabrication for Correlated Light/Electron Microscopy

The coupling of cryo-light microscopy (cryo-LM) and cryo-electron microscopy (cryo-EM) poses a number of advantages for understanding cellular 
dynamics and ultrastructure. First, cells can be imaged in a near native environment for both techniques. Second, due to the vitrification process, samples are
 preserved by rapid physical immobilization rather than slow chemical fixation. Third, imaging the same sample with both cryo-LM and cryo-EM provides correlation of
 data from a single cell, rather than a comparison of "representative samples". While these benefits are well known from prior studies, the widespread use of correlative 
 cryo-LM and cryo-EM remains limited due to the expense and complexity of buying or building a suitable cryogenic light microscopy stage. Here we demonstrate the 
 assembly, and use of an inexpensive cryogenic stage that can be fabricated in any lab for less than $40 with parts found at local hardware and grocery stores. This 
 cryo-LM stage is designed for use with reflected light microscopes that are fitted with long working distance air objectives. For correlative cryo-LM and cryo-EM 
 studies, we adapt the use of carbon coated standard 3-mm cryo-EM grids as specimen supports. After adsorbing the sample to the grid, previously established protocols 
 for vitrifying the sample and transferring/handling the grid are followed to permit multi-technique imaging. As a result, this setup allows any laboratory with a 
 reflected light microscope to have access to direct correlative imaging of frozen hydrated samples.

We demonstrate the fabrication of a low-cost cryogenic stage designed to fit most reflected light microscopes. This lab-built cryogenic stage enables efficient and reliable correlative imaging between cryo-light and cryo-electron microscopy.

The coupling of cryo-light microscopy (cryo-LM) and cryo-electron microscopy (cryo-EM) poses a number of advantages for understanding cellular 
dynamics ...

Graphene Coatings for Biomedical Implants

Atomically smooth graphene as a surface coating has potential to improve implant properties. This demonstrates a method for coating nitinol alloys with nanometer thick layers of graphene for applications as a stent material. Graphene was grown on copper substrates via chemical vapor deposition and then transferred onto nitinol substrates. In order to understand how the graphene coating could change biological response, cell viability of rat aortic endothelial cells and rat aortic smooth muscle cells was investigated. Moreover, the effect of graphene-coatings on cell adhesion and morphology was examined with fluorescent confocal microscopy. Cells were stained for actin and nuclei, and there were noticeable differences between pristine nitinol samples compared to graphene-coated samples. Total actin expression from rat aortic smooth muscle cells was found using western blot. Protein adsorption characteristics, an indicator for potential thrombogenicity, were determined for serum albumin and fibrinogen with gel electrophoresis. Moreover, the transfer of charge from fibrinogen to substrate was deduced using Raman spectroscopy. It was found that graphene coating on nitinol substrates met the functional requirements for a stent material and improved the biological response compared to uncoated nitinol. Thus, graphene-coated nitinol is a viable candidate for a stent material.

Graphene offers potential as a coating material for biomedical implants. In this study we demonstrate a method for coating nitinol alloys with nanometer thick layers of graphene and determine how graphene may influence implant response.

Atomically smooth graphene as a surface coating has potential to improve implant properties. This demonstrates a method for coating nitinol alloys with ...

High Throughput Microfluidic Rapid and Low Cost Prototyping Packaging Methods

In this work, 3 different packaging and assembly techniques are presented. They can be classified into two categories: one-time use and reusable packaging techniques.
The one-time use packaging technique employs UV-based and temperature curing epoxies to connect microtubes to access holes, wire-bonding for integrated circuit connections, and silver epoxy for electrical connections. This method is based on a robust assembly technique that can support relatively high pressure close to 1 psi and does not need any support to strengthen the microfluidic architecture.
Reusable packaging techniques consist of PDMS-based microtube interconnectors and anisotropic adhesive films for electrical connections. These devices are more sensitive and fragile. Consequently, Plexiglas support is added to the microfluidic structure to improve the electrical contact when anisotropic adhesive films are used, and also to strengthen the microfluidic architecture. In addition, a micromanipulator is needed to maintain tubes while using a thin PDMS layer to connect them to the access holes. Different PDMS layer thicknesses, ranging from 0.45-3 mm, are tested to compare the best adherence versus injection rates. Applied injection rates are varied from 50-300 &#956;l/hr for 0.45-3 mm PDMS layers, respectively. These techniques are mainly applicable for low-pressure applications. However, they can be extended for high-pressure ones through plasma-oxygen process to permanently seal the PDMS to glass substrates. The main advantage of this technique, besides the fact that it is reusable, consists of keeping the device observable when the microchannel length is very short (in the range of 3 mm or lower).

In this article we describe different techniques for microfluidic rapid prototyping platforms. The proposed techniques are based on ultraviolet (UV) sensitive and temperature curing epoxies, polydimethylsiloxane (PDMS) based tubing, wire-bonding, and anisotropic adhesive films. The assembling procedures presented are developed for both one-time use devices as well as reusable microfluidic systems.

In this work, 3 different packaging and assembly techniques are presented. They can be classified into two categories: one-time use and reusable packaging ...

Characterization of Complex Systems Using the Design of Experiments Approach: Transient Protein Expression in Tobacco as a Case Study

Plants provide multiple benefits for the production of biopharmaceuticals including low costs, scalability, and safety. Transient expression offers the additional advantage of short development and production times, but expression levels can vary significantly between batches thus giving rise to regulatory concerns in the context of good manufacturing practice. We used a design of experiments (DoE) approach to determine the impact of major factors such as regulatory elements in the expression construct, plant growth and development parameters, and the incubation conditions during expression, on the variability of expression between batches. We tested plants expressing a model anti-HIV monoclonal antibody (2G12) and a fluorescent marker protein (DsRed). We discuss the rationale for selecting certain properties of the model and identify its potential limitations. The general approach can easily be transferred to other problems because the principles of the model are broadly applicable: knowledge-based parameter selection, complexity reduction by splitting the initial problem into smaller modules, software-guided setup of optimal experiment combinations and step-wise design augmentation. Therefore, the methodology is not only useful for characterizing protein expression in plants but also for the investigation of other complex systems lacking a mechanistic description. The predictive equations describing the interconnectivity between parameters can be used to establish mechanistic models for other complex systems.

We describe a design of experiments approach that can be used to determine and model the influence of transgene regulatory elements, plant growth and development parameters, and incubation conditions on the transient expression of monoclonal antibodies and reporter proteins in plants.

Plants provide multiple benefits for the production of biopharmaceuticals including low costs, scalability, and safety. Transient expression offers the ...

Harmonic Nanoparticles for Regenerative Research

In this visualized experiment, protocol details are provided for in vitro labeling of human embryonic stem cells (hESC) with second harmonic generation nanoparticles (HNPs). The latter are a new family of probes recently introduced for labeling biological samples for multi-photon imaging. HNPs are capable of doubling the frequency of excitation light by the nonlinear optical process of second harmonic generation with no restriction on the excitation wavelength.
Multi-photon based methodologies for hESC differentiation into cardiac clusters (maintained as long term air-liquid cultures) are presented in detail. In particular, evidence on how to maximize the intense second harmonic (SH) emission of isolated HNPs during 3D monitoring of beating cardiac tissue in 3D is shown. The analysis of the resulting images to retrieve 3D displacement patterns is also detailed.

Protocol details are provided for in vitro labeling human embryonic stem cells with second harmonic generating nanoparticles. Methodologies for hESC investigation by multi-photon microscopy and their differentiation into cardiac clusters are also presented.

In this visualized experiment, protocol details are provided for in vitro labeling of human embryonic stem cells (hESC) with second harmonic generation ...

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

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

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

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

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Techniques for the Evolution of Robust Pentose-fermenting Yeast for Bioconversion of Lignocellulose to Ethanol

Lignocellulosic biomass is an abundant, renewable feedstock useful for production of fuel-grade ethanol and other bio-products. Pretreatment and enzyme saccharification processes release sugars that can be fermented by yeast. Traditional industrial yeasts do not ferment xylose (comprising up to 40% of plant sugars) and are not able to function in concentrated hydrolyzates. Concentrated hydrolyzates are needed to support economical ethanol recovery, but they are laden with toxic byproducts generated during pretreatment. While detoxification methods can render hydrolyzates fermentable, they are costly and generate waste disposal liabilities. Here, adaptive evolution and isolation techniques are described and demonstrated to yield derivatives of the native Scheffersomyces stipitis strain NRRL Y-7124 that are able to efficiently convert hydrolyzates to economically recoverable ethanol despite adverse culture conditions. Improved individuals are enriched in an evolving population using multiple selection pressures reliant on natural genetic diversity of the S. stipitis population and mutations induced by exposures to two diverse hydrolyzates, ethanol or UV radiation. Final evolution cultures are dilution plated to harvest predominant isolates, while intermediate populations, frozen in glycerol at various stages of evolution, are enriched on selective media using appropriate stress gradients to recover most promising isolates through dilution plating. Isolates are screened on various hydrolyzate types and ranked using a novel procedure involving dimensionless relative performance index (RPI) transformations of the xylose uptake rate and ethanol yield data. Using the RPI statistical parameter, an overall relative performance average is calculated to rank isolates based on multiple factors, including culture conditions (varying in nutrients and inhibitors) and kinetic characteristics. Through application of these techniques, derivatives of the parent strain had the following improved features in enzyme saccharified hydrolyzates at pH 5-6: reduced initial lag phase preceding growth, reduced diauxic lag during glucose-xylose transition, significantly enhanced fermentation rates, improved ethanol tolerance and accumulation to 40 g/L.

Adaptive evolution and isolation techniques are described and demonstrated to yield derivatives of Scheffersomyces stipitis strain NRRL Y-7124 that are able to rapidly consume hexose and pentose mixed sugars in enzyme saccharified undetoxified hydrolyzates and to accumulate over 40 g/L ethanol.

Lignocellulosic biomass is an abundant, renewable feedstock useful for production of fuel-grade ethanol and other bio-products. Pretreatment and enzyme ...

A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current regenerative medicine strategies have focused on transplanting stem cells or genetic manipulation of surrounding support cells in the inner ear to encourage replacement of damaged stem cells to correct hearing loss. Yet, the extracellular matrix (ECM) may play a vital role in inducing and maintaining function of hair cells, and has not been well investigated. Using the cochlear ECM as a scaffold to grow adult stem cells may provide unique insights into how the composition and architecture of the extracellular environment aids cells in sustaining hearing function. Here we present a method for isolating and decellularizing cochleae from mice to use as scaffolds accepting perfused adult stem cells. In the current protocol, cochleae are isolated from euthanized mice, decellularized, and decalcified. Afterward, human Wharton&#39;s jelly cells (hWJCs) that were isolated from the umbilical cord were carefully perfused into each cochlea. The cochleae were used as bioreactors, and cells were cultured for 30 days before undergoing processing for analysis. Decellularized cochleae retained identifiable extracellular structures, but did not reveal the presence of cells or noticeable fragments of DNA. Cells perfused into the cochlea invaded most of the interior and exterior of the cochlea and grew without incident over a duration of 30 days. Thus, the current method can be used to study how cochlear ECM affects cell development and behavior.

The goal of this protocol is to demonstrate an effective method to decellularize and decalcify mouse cochleae for utilization as scaffolds for tissue engineering applications.

In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current ...