The authors have no acknowledgements.

An exempt approval (3002-13) to carry out research using de-identified tissue samples pursuant to the Department of Health and Human Services regulations, 45 CFR 46.101(b)(4) was granted. Full thickness skin is received from elective surgery. In this article, the tissue source is 66-year-old Caucasian female breast.
1. Preparation of Elastomer Coated Coverslips
<ol>
	<li>In a 20 mL glass vial, mix 0.107 g of Sylgard 184 curing agent with 5.893 g base. The total mixture mass is 6 g with a base to curing agent ratio of 55:1.</li>
	<li>After mixing with a glass rod to ensure homogeneity, place the glass vial in a vacuum chamber and degas to remove all bubbles.</li>
	<li>Place a glass cover-slip (55 mm x 25 mm) in the center of a spin coater. Add ~1 mL of the mixture onto the center of the cover-slip. Use a 5,000 &#181;L pipette with the end cut off with scissors. Spin coat the cover-slip at 2,000 rpm for 60 s.
	<ol>
		<li>Repeat this process to create 5-6 substrates.</li>
	</ol>
	</li>
	<li>Cure the cover-slips in an oven for 12 h at 60 &#176;C.</li>
	<li>Use a razor blade to partially remove the elastomer film from a sacrificial substrate. Use an indelible marker to mark the topside of the elastomer film and the exposed glass.</li>
	<li>Mount the sample on an inverted microscope and use a remote focus accessory to record the difference in z-height between the focal planes of the two marks. This corresponds to the elastomer substrate thickness, h.</li>
</ol>
2. Preparation of the Stratum Corneum
<ol>
	<li>Use a water bath or heated stir plate to heat a glass beaker half filled with deionized water (DW) to 60 &#176;C. Inside a biological safety cabinet, immerse the full thickness human skin in the water for 4 min.</li>
	<li>Immediately transfer the skin sample into a beaker containing DW cooled to &#60;10 &#176;C for 4 min. Half filling the beaker minimizes splashing of biohazardous material.</li>
	<li>Remove the skin from the beaker, place in a Petri dish, and gently isolate the epidermis using a pair of bent nosed tissue tweezers.</li>
	<li>Place the isolated epidermis basal side down in a Petri dish lined with gauze. Ensure the basal layer is fully in contact with the gauze.
	<ol>
		<li>Soak the gauze in a 0.25% (wt/vol) type IX-S porcine pancreas trypsin solution dissolved in 0.1 M phosphate buffered saline for 6-8 h at room temperature. Add only enough trypsin in the container to wet the gauze.</li>
	</ol>
	</li>
	<li>Lift the gauze with tissue tweezers and float it in a container partially filled with DW. Gently pull the SC to separate it from the gauze.</li>
	<li>Wash the stratum corneum 3-4 times in DW to remove residual epidermal tissue that remains attached to the SC.</li>
	<li>Float the isolated SC in a solution of 0.4% glycine max(soybean) trypsin inhibitor in DW. Use a plate shaker to agitate the tissue for 10 min.</li>
	<li>Float SC in a Petri dish partially filled with DW. Use a plate shaker to agitate the tissue for 10 min.</li>
	<li>Dry the isolated SC sheet on an ultra-fine plastic mesh for 48 h at room temperature (25 &#176;C, 40% relative humidity (R.H.)).</li>
	<li>Separate the SC from the mesh and cut out individual circular R=3 mm radius samples using a circular hole punch. Mark the center of the outmost face with a small spiral mark using an indelible marker. This provides a visual cue for recognizing the topside of the SC. 
	NOTE: The indelible mark should be applied in the center of the sample, where the drying deformations will be smallest. This will minimize the impact of the marker on recorded drying displacement profiles.</li>
</ol>
3. Sample Treatment and Deposition
<ol>
	<li>Agitate SC samples for 30 min in 15 mL DW containing 90 &#181;L fluorescent marker beads (505/515 nm, 1 &#181;m diameter, carboxylate-modified). This deposits beads onto the SC surface 
	NOTE: While deposition of large numbers of beads on the SC may marginally slow drying relative to samples without beads present12, it will maximize the spatial resolution of the in-plane deformation fields that can be subsequently obtained. The choice of bead volume added should therefore be made ad hoc.</li>
	<li>Remove SC samples and place them in a Petri dish partially filled with DW.</li>
	<li>Partially immerse a substrate in the DW at a shallow angle of 15-30&#176;.</li>
	<li>Pin an edge of the floating SC sample at the contact line between the substrate and water interface. Vertically withdrawing the substrate from the water will smoothly laminate the SC sample to the substrate without wrinkles or entrapped air bubbles.</li>
	<li>Repeat step 3.4 to place up to 6 SC samples onto each substrate. Leave at least a 2-3 mm gap between samples and avoid sample lamination close to the substrate edge. This prevents drying of one sample influencing drying displacements in another.</li>
	<li>Dry the mounted SC samples in laboratory conditions for 60 min. This allows residual water between the SC and substrate to evaporate and ensures complete tissue adhesion. 
	NOTE: At this juncture, SC samples can be treated with a chemical or cosmetic formulations7,8 by placing the substrates upside down in a desired solution for a requisite period of time. Repeat step 3.6 once the treatment step is performed. After drying, incomplete adhesion of SC samples to the substrate can be verified using transmitted light microscopy. Trapped bubbles under the SC sample or delaminated edges will form clear contrast variations in the sample with well-defined edges.</li>
	<li>Create a humidity chamber by placing a Petri dish partially filled with water into a hermetically sealed container.</li>
	<li>Place substrates into the chamber for 24 h to equilibrate to a relative humidity of 99%. Do not place the substrates into the Petri dish.</li>
</ol>
4. Microscope Environmental Control
<ol>
	<li>Achieve control of environmental conditions through a humidity control system connected to a microscope mountable perfusion chamber. Details of the humidity control system are provided in German et al. (2013)7 and Liu and German (2015)8.</li>
	<li>Mount the substrate on the microscope, place the perfusion chamber over the substrate and seal edges of perfusion chamber to the elastomer using vacuum grease.</li>
	<li>Once mounted, equilibrate internal air to 99% R.H. prior to experimentation. This prevents evaporation of water prior to experimentation. Once imaging in sections 5 or 7 has begun, reduce internal air humidity to desired value. 
	NOTE: In this article, SC samples are dried to 25% R.H.</li>
</ol>
5. Imaging in Plane Drying Displacements
<ol>
	<li>Acquire images of SC samples using an inverted microscope with 1X objective lens. Excite fluorescent beads using a light engine with FITC filter (503-530 nm emission bandpass). Multiple samples can be imaged sequentially throughout drying using an automated x-y stage.</li>
	<li>Record fluorescent and transmitted light images using a digital CCD camera at a resolution of 1,392 x 1,040 pixels. The field of view of each image is 8.98 x 6.71 mm, allowing a single image to capture a full SC sample. Take images with a frequency of 10 min for 16 h.</li>
</ol>
6. Substrate Preparation for Thickness Measurement
<ol>
	<li>In a chemical fume hood, place 1 mL silane (3-aminopropyltriethoxysilane; &#8805;98%) in some small plastic cap. Place elastomer substrates from section 1 and the cap in a sealed container for 5 h. Do not allow substrates to come directly in contact with the silane.</li>
	<li>Add 5 mg of EDC (N-(3-Dimethylaminopropyl)-N-ethylcarbodiimide hydrochloride; &#8805;99%) in a 1.5 mL tube. Add 500 &#181;L of DW to the EDC. Agitate the solution for 10 s with a vortex mixer.</li>
	<li>Add 0.076 g sodium tetraborate and 0.1 g boric acid to 20 mL DW. Mix using a magnetic stirrer at 70 &#176;C (1 h). Add boric acid until the pH is 7.4.</li>
	<li>Add 20 mL of borate buffer to a 50 mL centrifuge tube. Add 60 &#181;L of 1 &#181;m beads (535/575 nm, carboxylate-modified) to the borate buffer. Finally, add 200 &#181;L of EDC solution to bottle. Shake the tube to mix bead solution and then pour into a 10 cm diameter Petri dish.</li>
	<li>Remove the silanated substrates from the container and place them elastomer film-side down into the bead solution. Do so slowly to prevent bubbles from becoming trapped. Two substrates will fit in each Petri dish.</li>
	<li>Leave substrates to float in the bead solution for 45 min.</li>
	<li>Use tweezers to remove the substrates from the bead solution, then rinse in DW to remove unbound beads.</li>
	<li>Air dry the substrates. Blowing compressed air over the elastomer film surface reduces the formation of water spots.</li>
	<li>Seal the substrates in an opaque box to prevent photo-bleaching of the beads until SC sample deposition.</li>
</ol>
7. Imaging Thickness of SC
<ol>
	<li>Deposit SC samples on to a substrate using section 3. However, perform step 3.1 without adding fluorescent beads to the DW. Additionally, apply a 5 &#181;L drop of undiluted fluorescent marker bead solution (505/515 nm, 0.1 &#181;m diameter) to the surface of each deposited SC sample with a pipette before completing step 3.6.</li>
	<li>Establish measurements of SC thickness using the microscope with 40X objective lens. Measure the thickness of SC samples over time using a remote focus accessory to record the difference in z-height between the two bead layer focal planes located at the SC-substrate interface and the topside of the SC.</li>
	<li>Measure the thickness of 3 regions of each SC sample over a 3 h drying period. The thickness of SC samples reaches a steady state value within this time frame8.</li>
</ol>
8. Quantifying and Modeling Tissue Deformation
<ol>
	<li>Use particle image velocimetry13 to obtain spatially resolved in-plane drying displacements from the fluorescent images at each recorded time step.</li>
	<li>Use MATLAB to obtain azimuthally averaged radial and azimuthal displacement profiles from the displacement field of each radially symmetric SC sample. 
	NOTE: An example dataset (entitled &#39;d.mat&#39;) and MATLAB code (entitled &#39;PIV-processing.m&#39;) that performs both this step and step 8.3 has been provided in the supplemental information.</li>
	<li>Fit radial displacement profiles to a model7,8,14,15,16 describing drying SC as a shrinking linear elastic circular disk of time varying thickness, hSC, radius, R, and elastic modulus, ESC, adhered to a deformable elastic substrate with elastic modulus, E. Assume SC has a well-defined and constant Poisson&#39;s ratio, &#957;SC = 0.47,8. Obtain best fits using a minimum least squares approach. 
	NOTE: The model used for fitting describes radial displacements in terms of modified Bessel functions are: 
	<img alt="Equation" src="/files/ftp_upload/55336/55336eq2.jpg" />&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; (1) 
	with <img alt="Equation" src="/files/ftp_upload/55336/55336eq3.jpg" />,&#160;<img alt="Equation" src="/files/ftp_upload/55336/55336eq4.jpg" /> and <img alt="Equation" src="/files/ftp_upload/55336/55336eq13.jpg" /> 
	The term <img alt="Equation" src="/files/ftp_upload/55336/55336eq6.jpg" /> corresponds to a penetration depth given by, 
	<img alt="Equation" src="/files/ftp_upload/55336/55336eq7.jpg" />. 
	<img alt="Equation" src="/files/ftp_upload/55336/55336eq8.jpg" /> denotes a substrate rigidity parameter; valid when sample sizes are much greater than the substrate thickness. Here, the parameters, <img alt="Equation" src="/files/ftp_upload/55336/55336eq9.jpg" /> and <img alt="Equation" src="/files/ftp_upload/55336/55336eq10.jpg" /> respectively denote the substrate thickness Poisson&#39;s ratio. The Poisson&#39;s ratio of the silicone elastomer substrate17 is &#957; = 0.5.</li>
	<li>Obtain model parameters &#945; and &#946; at each time step from the least squares fit of Equation (1) to the radial displacement profile.
	<ol>
		<li>Employ the fitting parameter &#946; to obtain SC elastic modulus, ESC, using the expression, 
		<img alt="Equation" src="/files/ftp_upload/55336/55336eq11.jpg" /></li>
		<li>Use fitting parameter &#945; to obtain the time varying contractile drying stress, PSC, using the expression, 
		<img alt="Equation" src="/files/ftp_upload/55336/55336eq12.jpg" /></li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

In this article, we describe a technique that can be used to measure the dynamic drying behavior and mechanical properties of human SC. Previous studies have demonstrated that this technique can be used to quantify the effects of environmental conditions and chemical products commonly used in cosmetic cleansers and moisturizers on the dynamic drying behavior of SC7,8. There are a number of key steps in the protocol. Firstly, SC swells notably with water content; therefore, measurements of SC thickness as well as in-plane displacements are essential for accurately predicting the elastic modulus and drying stress magnitude. Secondly, samples need to be fully adhered to the substrate. Incomplete adhesion, non-radially symmetric samples or samples with small tears or holes should be avoided because they will significantly impact the distribution of drying deformations and the radial displacement profiles used for model fitting.
The technique can be used if a humidity control system is unavailable. Without environmental control, tissue samples will dry in laboratory conditions12. As such, the laboratory environment should be continuously monitored and maintained, as drying behavior and the repeatability of results will be impacted by both diurnal and seasonal variations in temperature and humidity.
Currently, the technique is limited only to samples that can adhere to the substrate and induce deformations within the elastomer film. While the technique can be readily adapted to test samples that undergo smaller in-plane displacements, by reducing the substrate elastic modulus12, results from samples that simply slip over the substrate will lack meaning.
Numerous in-vivo and ex-vivo techniques that can assess the drying behavior and mechanical properties of SC have been reported3,8,9,10,18,19,20,21. However, in-vivo techniques cannot fully distinguish mechanical changes in SC from the underlying epidermal and dermal layers. Moreover, ex-vivo techniques can typically only assess one sample per experiment. The method we report in this article allows up to 6 SC samples to be assessed per experiment. The size of the substrate and environmental chamber however could be scaled up to allow more samples to be assessed simultaneously. We estimate for n=6 SC samples, a timescale of ~13 h is required for preparation and testing, excluding substrate curing and tissue equilibration. In comparison, we estimate uniaxial tensometry testing would require more than twice this period. Significantly less SC tissue is also required per individual sample (0.28 cm2) in comparison with those required for tensometry9 (2.5 cm2). This technique further enables more physiologically relevant drying by preventing evaporation from the underside of the SC tissue. In addition to assessing drying behavior and mechanics in SC, we believe this technique could also be applied to studies of polymeric or colloidal systems that form a cohesive film upon drying.

The outer most layer of the epidermis, or stratum corneum (SC) consists of cohesive corneocyte cells surrounded by a lipid rich matrix1,2. The composition and structural integrity of SC is essential for maintaining correct barrier functionality3, which prevents invasion from microorganisms and resists both mechanical forces and excessive water loss4. The capacity of personal care products to maintain or degrade skin barrier function is of great interest to skin healthcare and the cosmetic industry5. The daily application of personal care products is known to alter the mechanical properties of the SC6,7,8. For example, surfactants contained in cosmetic cleansers can cause significant increases in the elastic modulus and a build-up of drying stresses in SC, increasing the tissue&#39;s propensity to crack7,9. Glycerol contained in nearly all cosmetic moisturizers can soften SC and decrease the build-up of drying stresses8,10,11, reducing the likelihood of tissue rupture.
The method detailed in this article is capable of quantifying the dynamic drying behavior and mechanical properties of SC drying in controlled environments7,8. Previously, this technique has been demonstrated to be capable of elucidating the effect of different cosmetic products on changes in the dynamic drying behavior and mechanical properties of SC tissue. This is achieved by quantifying drying-induced shrinkage of human SC tissue adhered to a soft elastomer substrate, fitting drying displacements with a simple contractility model, and then extracting the elastic modulus and drying stress from the fitted profile. When testing of multiple SC samples is required, this method offers a more rapid alternative to uniaxial tensometry, utilizes significantly less tissue and provides more physiologically relevant drying by preventing evaporation from the sample underside.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Silicone elastomer base</td>
			<td>Dow-Corning</td>
			<td>1064291</td>
		</tr>
		<tr>
			<td>Silicone elastomer Curing Agent</td>
			<td>Dow-Corning</td>
			<td>1015311</td>
		</tr>
		<tr>
			<td>FluoSpheres Carboxylate 0.1 &#181;m yellow green fluorescent 505/515&#160;</td>
			<td>Thermo Fisher</td>
			<td>F8803</td>
		</tr>
		<tr>
			<td>FluoSpheres Carboxylate 1 &#181;m yellow green fluorescent 505/515&#160;</td>
			<td>Thermo Fisher</td>
			<td>F8823</td>
		</tr>
		<tr>
			<td>FluoSpheres Carboxylate 1 &#181;m nile red fluorescent 535/575&#160;</td>
			<td>Thermo Fisher</td>
			<td>F8819</td>
		</tr>
		<tr>
			<td>Trypsin from porcine pancreas</td>
			<td>Sigma-Aldrich</td>
			<td>T6567</td>
		</tr>
		<tr>
			<td>Trypsin inhibitor type II-s</td>
			<td>Sigma-Aldrich</td>
			<td>T9128</td>
		</tr>
		<tr>
			<td>(3-aminopropyl)triethoxysilane</td>
			<td>Sigma-Aldrich</td>
			<td>440140</td>
		</tr>
		<tr>
			<td>Sodium tetraborate</td>
			<td>Sigma-Aldrich</td>
			<td>221732</td>
		</tr>
		<tr>
			<td>Boric acid</td>
			<td>Sigma-Aldrich</td>
			<td>B0294</td>
		</tr>
		<tr>
			<td>Phosphate buffered saline</td>
			<td>Sigma-Aldrich</td>
			<td>P7059</td>
		</tr>
		<tr>
			<td>N-(3-Dimethylaminopropyl)-N-ethylcarbodiimide hydrochloride&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>E7750</td>
		</tr>
		<tr>
			<td>Vortexer mixer</td>
			<td>VWR</td>
			<td>58816-123</td>
		</tr>
		<tr>
			<td>6mm diameter hole punch</td>
			<td>Sigma-Aldrich</td>
			<td>Z708860</td>
		</tr>
		<tr>
			<td>SOLA 6-LCR-SB&#160;</td>
			<td>Lummencor light engine</td>
			<td>No.3526</td>
		</tr>
		<tr>
			<td>Cfi Plan Achro Uw 1x Objective</td>
			<td>Nikon Plan UW</td>
			<td>MRL00012</td>
		</tr>
		<tr>
			<td>CFI Plan Fluor 40x Oil Objective 1.3 na - 0.20mm wd</td>
			<td>Nikon Plan Fluor</td>
			<td>MRH01401</td>
		</tr>
		<tr>
			<td>Nikon Eclipse Ti-U inverted microscope&#160;</td>
			<td>Nikon</td>
			<td>MEA53200</td>
		</tr>
		<tr>
			<td>Clara-E Camera</td>
			<td>Andor</td>
			<td>DR-328G-C02-SIL</td>
		</tr>
		<tr>
			<td>Remote Focus Attachment E-RFA Ergo Design</td>
			<td>Nikon</td>
			<td>99888</td>
		</tr>
		<tr>
			<td>Ti-S-E Motorized Stage</td>
			<td>Nikon</td>
			<td>MEC56110</td>
		</tr>
	</tbody>
</table>

measuring and modeling contractile drying in human stratum corneum

Stratum corneum (SC) is the most superficial skin layer. Its contact with the external environment means that this tissue layer is subjected to both cleansing agents and daily variations in ambient moisture; both of which can alter the water content of the tissue. Reductions in water content from severe barrier dysfunction or low humidity environments can alter SC stiffness and cause a build-up of drying stresses. In extreme conditions, these factors can cause mechanical rupture of the tissue. We have established a high throughput method of quantifying dynamic changes in the mechanical properties of SC upon drying. This technique can be employed to quantify changes in the drying behavior and mechanical properties of SC with cosmetic cleanser and moisturizer treatments. This is achieved by measuring dynamic variations in spatially resolved in-plane drying displacements of circular tissue samples adhered to an elastomer substrate. In-plane radial displacements acquired during drying are azimuthally averaged and fitted with a profile based on a linear elastic contractility model. Dynamic changes in drying stress and SC elastic modulus can then be extracted from the fitted model profiles.

Figure 1(a) shows a representative fluorescent image of an SC sample coated with fluorescent beads (section 3). The corresponding transmitted light image of the sample is shown in Figure 1(b) overlaid with a quiver plot of spatially resolved drying displacements that form after 16 h drying at 25% R.H. Due to the circular symmetry of the samples, these displacements can be azimuthally averaged. Figure 1(c) shows radial (ur, solid red line) and azimuthal (u&#952;, dashed blue line) displacement profiles plotted against the dimensionless radial position, r/R. Here, R denotes the mean SC sample radius, r/R = 0 denotes the sample center and r/R = 1 denotes the edge. Standard deviations at each radial position are denoted by the shaded regions around the mean. These variations are primarily caused by the structural heterogeneity of the SC3,7,12. Throughout drying, azimuthal displacements remain small. Radial displacement profiles however increase monotonically from center to edge and grow in magnitude until an equilibrium is reached.
<img alt="Figure 1" src="/files/ftp_upload/55336/55336fig1.jpg" /> 
Figure 1: Circular SC sample (6.2 mm diameter) adhered to an elastomer substrate with elastic modulus E = 16 &#177; 1 kPa after drying for 15 h in a 25 &#177; 1% R.H. environment. (a) Fluorescent image of the SC sample highlighting the deposited fluorescent marker beads used for tracking spatially resolved in-plane drying displacements. (b) Quiver plot of spatially resolved in-plane drying displacements overlaid on a transmitted light image of the SC sample. (c) Azimuthally averaged radial (ur, solid red line) and azimuthal (u&#952;, blue dashed line) displacements of the sample plotted against dimensionless radial position, r/R. Positive values of ur correspond to contractile displacements. Shaded regions surrounding the lines indicate the standard deviation about the mean at each radial position. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55336/55336fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Profiles recorded at 30 min intervals are plotted in Figure 2(a) and show the time evolution of in-plane displacements. The average SC thickness, hSC, is plotted in Figure 2(b). Decreases of SC during drying primarily occur over the first 2 h.
<img alt="Figure 2" src="/files/ftp_upload/55336/55336fig2.jpg" /> 
Figure 2: (a) Overlay of radial displacement profiles (ur, solid red lines) at 30 min intervals over a 15 h drying period in 25% R.H. conditions plotted against dimensionless radial position r/R for a typical SC sample. Positive values of ur correspond to contractile displacements. (b) Average SC sample thickness, (hSC, n=3), plotted against drying time. (c) Radial displacement profiles from (a) overlaid with minimum least squares fits (blue dashed line) of Equation (1) for the first and last recorded radial displacement profile. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55336/55336fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Fitting displacement profiles with the linear elastic contractility model described by Equation (1) provides further insight into the mechanical properties of drying SC. Displacement profiles at each time step are fitted with the model using a minimum least squares approach, as shown in Figure 2(c). The contractile drying stress, PSC, and elastic modulus, ESC, are subsequently extracted from the model at each time step. Average changes in these parameters (based on 3 individual SC samples) are shown respectively in Figures 3(a) and 3(b). Both parameters increase rapidly over the first 2 h drying period and reach a plateau within 5 h.
<img alt="Figure3" src="/files/ftp_upload/55336/55336fig3.jpg" /> 
Figure 3: (a) Averaged SC elastic modulus, ESC, plotted against drying time over a 15 h period. (b) Average contractile drying stress, PSC, plotted against drying time over a 15 h period. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55336/55336fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Measuring and Modeling Contractile Drying in Human Stratum Corneum at JoVE.com

Measuring and Modeling Contractile Drying in Human Stratum Corneum

This article describes a method of quantifying the dynamic drying behavior and mechanical properties of stratum corneum by measuring spatially resolved in-plane drying displacements of circular tissue samples adhered to an elastomer substrate. This technique can be used to measure how different chemical treatments alter drying and tissue mechanical properties.

Stratum corneum (SC) is the most superficial skin layer. Its contact with the external environment means that this tissue layer is subjected to both ...

measuring-and-modeling-contractile-drying-in-human-stratum-corneum

Nanyang Technological University

Research

JoVE Journal

Bioengineering

6.5K Views.  Binghamton University. This article describes a method of quantifying the dynamic drying behavior and mechanical properties of stratum corneum by measuring spatially resolved in-plane drying displacements of circular tissue samples adhered to an elastomer substrate. This technique can be used to measure how different chemical treatments alter drying and tissue mechanical properties.

Video: Measuring and Modeling Contractile Drying in Human Stratum Corneum

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable around embryonic day (E) 8.51. Systolic left ventricular (LV) pressure is 2 mmHg at E9.5 and 11 mmHg at E14.52. At these mid-embryonic stages, the LV is clearly visible through the chest wall for invasive pressure measurements because the ribs and skin are not fully developed. Between E14.5 and birth (approximately E21) imaging methods must be used to view the LV. After birth, mean arterial pressure increases from 30 - 70 mmHg from postnatal day (P) 2 - 353. Beyond P20, arterial pressure can be measured with solid-state catheters (i.e. Millar or Scisense). Before P20, these catheters are too big for developing mouse arteries and arterial pressure must be measured with custom pulled plastic catheters attached to fluid-filled pressure transducers3 or glass micropipettes attached to servo null pressure transducers4.
Our recent work has shown that the greatest increase in blood pressure occurs during the late embryonic to early postnatal period in mice5-7. This large increase in blood pressure may influence smooth muscle cell (SMC) phenotype in developing arteries and trigger important mechanotransduction events. In human disease, where the mechanical properties of developing arteries are compromised by defects in extracellular matrix proteins (i.e. Marfan's Syndrome8 and Supravalvular Aortic Stenosis9) the rapid changes in blood pressure during this period may contribute to disease phenotype and severity through alterations in mechanotransduction signals. Therefore, it is important to be able to measure blood pressure changes during late embryonic and neonatal periods in mouse models of human disease.
We describe a method for measuring LV pressure in late embryonic (E18) and early postnatal (P1 - 20) mice. A needle attached to a fluid-filled pressure transducer is inserted into the LV under ultrasound guidance. Care is taken to maintain normal cardiac function during the experimental protocol, especially for the embryonic mice. Representative data are presented and limitations of the protocol are discussed.

Measuring left ventricular pressure (LV) in embryonic and neonatal mice is described. Pressure is measured by inserting a needle connected to a fluid-filled transducer into the LV under ultrasound guidance. Care must be taken to maintain normal cardiac function during the experimental protocol.

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable ...

CometChip: A High-throughput 96-Well Platform for Measuring DNA Damage in Microarrayed Human Cells

DNA damaging agents can promote aging, disease and cancer and they are ubiquitous in the environment and produced within human cells as normal cellular metabolites. Ironically, at high doses DNA damaging agents are also used to treat cancer. The ability to quantify DNA damage responses is thus critical in the public health, pharmaceutical and clinical domains. Here, we describe a novel platform that exploits microfabrication techniques to pattern cells in a fixed microarray. The &#8216;CometChip&#8217; is based upon the well-established single cell gel electrophoresis assay (a.k.a. the comet assay), which estimates the level of DNA damage by evaluating the extent of DNA migration through a matrix in an electrical field. The type of damage measured by this assay includes abasic sites, crosslinks, and strand breaks. Instead of being randomly dispersed in agarose in the traditional assay, cells are captured into an agarose microwell array by gravity. The platform also expands from the size of a standard microscope slide to a 96-well format, enabling parallel processing. Here we describe the protocols of using the chip to evaluate DNA damage caused by known genotoxic agents and the cellular repair response followed after exposure. Through the integration of biological and engineering principles, this method potentiates robust and sensitive measurements of DNA damage in human cells and provides the necessary throughput for genotoxicity testing, drug development, epidemiological studies and clinical assays.

We describe here a platform that allows comet assay detection of DNA damage with unprecedented throughput. The device patterns mammalian cells into a microarray and enables parallel processing of 96 samples. The approach facilitates analysis of base level DNA damage, exposure-induced DNA damage and DNA repair kinetics.

DNA damaging agents can promote aging, disease and cancer and they are ubiquitous in the environment and produced within human cells as normal cellular ...

Quantification of Global Diastolic Function by Kinematic Modeling-based Analysis of Transmitral Flow via the Parametrized Diastolic Filling Formalism

Quantitative cardiac function assessment remains a challenge for physiologists and clinicians.&#160;Although historically invasive methods have comprised the only means available, the development of noninvasive imaging modalities (echocardiography, MRI, CT) having high temporal and spatial resolution provide a new window for quantitative diastolic function assessment. Echocardiography is the agreed upon standard for diastolic function assessment, but indexes in current clinical use merely utilize selected features of chamber dimension (M-mode) or blood/tissue motion (Doppler) waveforms without incorporating the physiologic causal determinants of the motion itself.&#160;The recognition that all left ventricles (LV) initiate filling by serving as mechanical suction pumps allows global diastolic function to be assessed based on laws of motion that apply to all chambers. What differentiates one heart from another are the parameters of the equation of motion that governs filling. Accordingly, development of the Parametrized Diastolic Filling (PDF) formalism&#160;has shown that the entire range of clinically observed early transmitral flow (Doppler E-wave) patterns are extremely well fit by the laws of damped oscillatory motion.&#160;This permits analysis of individual E-waves in accordance with a causal mechanism (recoil-initiated suction) that yields three (numerically) unique lumped parameters whose physiologic analogues are chamber stiffness (k), viscoelasticity/relaxation (c), and load (xo).&#160;The recording of transmitral flow (Doppler E-waves) is standard practice in clinical cardiology and, therefore, the echocardiographic recording method is only briefly reviewed. Our focus is on determination of the PDF parameters from routinely recorded E-wave data.&#160;As the highlighted results indicate, once the PDF parameters have been obtained from a suitable number of load varying E-waves, the investigator is free to use the parameters or construct indexes from the parameters (such as stored energy 1/2kxo2, maximum A-V pressure gradient kxo, load independent index of diastolic function, etc.) and select the aspect of physiology or pathophysiology to be quantified.

Accurate, causality-based quantification of global diastolic function has been achieved by kinematic modeling-based analysis of transmitral flow via the Parametrized Diastolic Filling (PDF) formalism. PDF generates unique stiffness, relaxation,&#160;and load parameters and elucidates &#39;new&#39; physiology while providing sensitive and specific indexes of dysfunction.

Quantitative cardiac function assessment remains a challenge for physiologists and clinicians.&#160;Although historically invasive methods have comprised ...

Utilization of Microscale Silicon Cantilevers to Assess Cellular Contractile Function In Vitro

The development of more predictive and biologically relevant in vitro assays is predicated on the advancement of versatile cell culture systems which facilitate the functional assessment of the seeded cells. To that end, microscale cantilever technology offers a platform with which to measure the contractile functionality of a range of cell types, including skeletal, cardiac, and smooth muscle cells, through assessment of contraction induced substrate bending. Application of multiplexed cantilever arrays provides the means to develop moderate to high-throughput protocols for assessing drug efficacy and toxicity, disease phenotype and progression, as well as neuromuscular and other cell-cell interactions. This manuscript provides the details for fabricating reliable cantilever arrays for this purpose, and the methods required to successfully culture cells on these surfaces. Further description is provided on the steps necessary to perform functional analysis of contractile cell types maintained on such arrays using a novel laser and photo-detector system. The representative data provided highlights the precision and reproducible nature of the analysis of contractile function possible using this system, as well as the wide range of studies to which such technology can be applied. Successful widespread adoption of this system could provide investigators with the means to perform rapid, low cost functional studies in vitro, leading to more accurate predictions of tissue performance, disease development and response to novel therapeutic treatment.

Utilization of Microscale Silicon Cantilevers to Assess Cellular Contractile Function In Vitro

This protocol describes the use of microscale silicon cantilevers as pliable culture surfaces for measuring the contractility of muscle cells in vitro. Cellular contraction causes cantilever bending, which can be measured, recorded, and converted into readouts of force, providing a non-invasive and scalable system for measuring contractile function in vitro.

The development of more predictive and biologically relevant in vitro assays is predicated on the advancement of versatile cell culture systems which ...

Production, Characterization and Potential Uses of a 3D Tissue-engineered Human Esophageal Mucosal Model

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore esophageal adenocarcinoma generally has a poor prognosis, with little improvement in survival rates in recent years. These are difficult conditions to study and there has been a lack of suitable experimental platforms to investigate disorders of the esophageal mucosa.
A model of the human esophageal mucosa has been developed in the MacNeil laboratory which, unlike conventional 2D cell culture systems, recapitulates the cell-cell and cell-matrix interactions present in vivo and produces a mature, stratified epithelium similar to that of the normal human esophagus. Briefly, the model utilizes non-transformed normal primary human esophageal fibroblasts and epithelial cells grown within a porcine-derived acellular esophageal scaffold. Immunohistochemical characterization of this model by CK4, CK14, Ki67 and involucrin staining demonstrates appropriate recapitulation of the histology of the normal human esophageal mucosa.
This model provides a robust, biologically relevant experimental model of the human esophageal mucosa. It can easily be manipulated to investigate a number of research questions including the effectiveness of pharmacological agents and the impact of exposure to environmental factors such as alcohol, toxins, high temperature or gastro-esophageal refluxate components. The model also facilitates extended culture periods not achievable with conventional 2D cell culture, enabling, inter alia, the study of the impact of repeated exposure of a mature epithelium to the agent of interest for up to 20 days. Furthermore, a variety of cell lines, such as those derived from esophageal tumors or Barrett&#8217;s Metaplasia, can be incorporated into the model to investigate processes such as tumor invasion and drug responsiveness in a more biologically relevant environment.

This manuscript describes the production, characterization and potential uses of a tissue engineered 3D esophageal construct prepared from normal primary human esophageal fibroblast and squamous epithelial cells seeded within a de-cellularized porcine scaffold. The results demonstrate the formation of a mature stratified epithelium similar to the normal human esophagus.

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore ...

Methods for the Self-integration of Megamolecular Biopolymers on the Drying Air-LC Interface

Living organisms that use water are always prone to drying in the environment. Their activities are driven by biopolymer-based micro- and macro-structures, as seen in the cases of moving water in vascular bundles and moisturizing water in skin layers. In this study, we developed a method for assessing the effect of aqueous liquid crystalline (LC) solutions composed of biopolymers on drying. As LC biopolymers have megamolecular weight, we chose to study polysaccharides, cytoskeletal proteins, and DNA. The observation of biopolymer solutions during drying under polarized light reveals milliscale self-integration starting from the unstable air-LC interface. The dynamics of the aqueous LC biopolymer solutions can be monitored by evaporating water from a one-side-open cell. By analyzing the images taken using cross-polarized light, it is possible to recognize the spatio-temporal changes in the orientational order parameter. This method can be useful for the characterization of not only artificial materials in various fields, but also natural living tissues. We believe that it will provide an evaluation method for soft materials in the biomedical and environmental fields.

A method for the drying-induced self-integration of megamolecular biopolymers on the air-liquid crystalline interface is provided here. This methodology will be useful not only for understanding the macroscopic potentials of biopolymers, but also as an evaluation method for soft materials in biomedical and environmental fields.

Living organisms that use water are always prone to drying in the environment. Their activities are driven by biopolymer-based micro- and ...

Microhoneycomb Monoliths Prepared by the Unidirectional Freeze-drying of Cellulose Nanofiber Based Sols: Method and Extensions

Monolithic honeycomb structures have been attractive to multidisciplinary fields due to their high strength-to-weight ratio. Particularly, microhoneycomb monoliths (MHMs) with micrometer-scale channels are expected as efficient platforms for reactions and separations because of their large surface areas. Up to now, MHMs have been prepared by a unidirectional freeze-drying (UDF) method only from very limited precursors. Herein, we report a protocol from which a series of MHMs consisting of different components can be obtained. Recently, we found that cellulose nanofibers function as a distinct structure-directing agent towards the formation of MHMs through the UDF process. By mixing the cellulose nanofibers with water soluble substances which do not yield MHMs, a variety of composite MHMs can be prepared. This significantly enriches the chemical constitution of MHMs towards versatile applications.

Here, we present a general protocol to prepare a variety of microhoneycomb monoliths (MHMs) in which fluid can pass through with an extremely low pressure drop. MHMs obtained are expected to be used as filters, catalyst supports, flow-type electrodes, sensors and scaffolds for biomaterials.

Monolithic honeycomb structures have been attractive to multidisciplinary fields due to their high strength-to-weight ratio. Particularly, microhoneycomb ...

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) culture systems. However, measurement of cell function in 3D culture is often more challenging. Many biological assays require retrieval of cellular material which can be difficult in 3D cultures. One way to address this challenge is to develop new materials that enable measurement of cell function within the material. Here, a method is presented for measurement of cellular matrix metalloproteinase (MMP) activity in 3D hydrogels in a 96-well format. In this system, a poly(ethylene glycol) (PEG) hydrogel is functionalized with a fluorogenic MMP cleavable sensor. Cellular MMP activity is proportional to fluorescence intensity and can be measured with a standard microplate reader. Miniaturization of this assay to a 96-well format reduced the time required for experimental set up by 50% and reagent usage by 80% per condition as compared to the previous 24-well version of the assay. This assay is also compatible with other measurements of cellular function. For example, a metabolic activity assay is demonstrated here, which can be conducted simultaneously with MMP activity measurements within the same hydrogel. The assay is demonstrated with human melanoma cells encapsulated across a range of cell seeding densities to determine the appropriate encapsulation density for the working range of the assay. After 24 h of cell encapsulation, MMP and metabolic activity readouts were proportional to cell seeding density. While the assay is demonstrated here with one fluorogenic degradable substrate, the assay and methodology could be adapted for a wide variety of hydrogel systems and other fluorescent sensors. Such an assay provides a practical, efficient and easily accessible 3D culturing platform for a wide variety of applications.

Here, a protocol is presented for encapsulating and culturing cells in poly(ethylene glycol) (PEG) hydrogels functionalized with a fluorogenic matrix metalloproteinase (MMP)-degradable peptide. Cellular MMP and metabolic activity are measured directly from the hydrogel cultures using a standard microplate reader.

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) ...

A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of cellular function. The ECM plays a particularly important role in adipose tissue function in obesity, and alterations in adipose tissue ECM deposition and composition are associated with metabolic disease in mice and humans. Tractable in vitro models that permit dissection of the roles of the ECM and cells in contributing to global tissue phenotype are sparse. We describe a novel 3D in vitro model of human ECM-adipocyte culture that permits study of the specific roles of the ECM and adipocytes in regulating adipose tissue metabolic phenotype. Human adipose tissue is decellularized to isolate ECM, which is subsequently repopulated with preadipocytes that are then differentiated within the ECM into mature adipocytes. This method creates ECM-adipocyte constructs that are metabolically active and retain characteristics of the tissues and patients from which they are derived. We have used this system to demonstrate disease-specific ECM-adipocyte crosstalk in human adipose tissue. This culture model provides a tool for dissecting the roles of the ECM and adipocytes in contributing to global adipose tissue metabolic phenotype and permits study of the role of the ECM in regulating adipose tissue homeostasis.

We describe a 3D human extracellular matrix-adipocyte in vitro culture system that permits dissection of the roles of the matrix and adipocytes in contributing to adipose tissue metabolic phenotype.

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of ...

Electric and Magnetic Field Devices for Stimulation of Biological Tissues

Electric fields (EFs) and magnetic fields (MFs) have been widely used by tissue engineering to improve cell dynamics such as proliferation, migration, differentiation, morphology, and molecular synthesis. However, variables such stimuli strength and stimulation times need to be considered when stimulating either cells, tissues or scaffolds. Given that EFs and MFs vary according to cellular response, it remains unclear how to build devices that generate adequate biophysical stimuli to stimulate biological samples. In fact, there is a lack of evidence regarding the calculation and distribution when biophysical stimuli are applied. This protocol is focused on the design and manufacture of devices to generate EFs and MFs and implementation of a computational methodology to predict biophysical stimuli distribution inside and outside of biological samples. The EF device was composed of two parallel stainless-steel electrodes located at the top and bottom of biological cultures. Electrodes were connected to an oscillator to generate voltages (50, 100, 150 and 200 Vp-p) at 60 kHz. The MF device was composed of a coil, which was energized with a transformer to generate a current (1 A) and voltage (6 V) at 60 Hz. A polymethyl methacrylate support was built to locate the biological cultures in the middle of the coil. The computational simulation elucidated the homogeneous distribution of EFs and MFs inside and outside of biological tissues. This computational model is a promising tool that can modify parameters such as voltages, frequencies, tissue morphologies, well plate types, electrodes and coil size to estimate the EFs and MFs to achieve a cellular response.

This protocol describes the step-by-step process to build both electrical and magnetic stimulators used to stimulate biological tissues. The protocol includes a guideline to simulate computationally electric and magnetic fields and manufacture of stimulator devices.

Electric fields (EFs) and magnetic fields (MFs) have been widely used by tissue engineering to improve cell dynamics such as proliferation, migration, ...