Some figures included here have been adapted by permission from the Copyright Clearance Center: Springer Nature, Annals of Biomedical Engineering. Straining 3D hydrogels with uniform z-axis strains while enabling live microscopy imaging, A. Roitblat Riba, S. Natan, A. Kolel, H. Rushkin, O. Tchaicheeyan, A. Lesman, Copyright&#169; (2019).
https://doi-org.bdigitaluss.remotexs.co/10.1007/s10439-019-02426-7

The authors have nothing to disclose.

The method and protocol presented herein are largely based on our previous study by Roitblat Riba et al.41 We include here the full computer-aided design (CAD), Python and microcontroller codes of the SCyUS device.
The major advantages of the presented method over existing approaches include the possibility to strain very soft 3D hydrogels (Elastic Modulus of ~100 Pa) from their circumference, and under live confocal imaging. Other methods are usually limited in their ability to apply strain fields in the Z-axis and are unable to provide in situ high-magnification microscopy images while stretching, mostly due to working distance limitations from the sample to the objective lens. Most available systems are based on 2D substrate-stretching, which are limited in mimicking physiological tissue environments.
Limitations of the system include potential inconsistent homogeneity of fiber density of the gel post-polymerization (Figure 7A), especially when large gel volumes or complex geometries are used. Such non-homogeneities can lead to non-uniform strains and fiber alignment in the gel. Moreover, optical restrictions at relatively deep Z-sections (&#62;30 &#956;m) may give rise to errors in analysis of fiber orientation.
Critical steps in the 3D stretching protocol include the following (i) Ensure that sample gels have achieved complete adhesion to the inner walls of the geometric cut-out within the silicone, without bubbles or tears, especially along the Z-direction. Complete adhesion and continuity of the sample with a thickness of at least 100 &#181;m (Eq. 1) is crucial to reliably stretch the sample to a desired strain (ii) Properly attach the sample to the SCyUS device prior to stretching. It is necessary to attach the silicone strip to the brackets and non-elastic fabric length extenders using the 3D printed jig (Figure 5A) and to validate that all pieces are attached in-line, ensuring that strain is induced in the gel symmetrically. This also applies to the correct loading of the device&#8217;s spindle (Figure 6B-C) (iii) Carefully capture the reference (pre-stretch) position of the sample. If the reference image is taken while the sample is already under tension (e.g., due to the attachment of the servomotor prior to this step), all strain measurements will be inaccurate.
Potential future applications and modifications of the presented protocol include the following:
(i) A cyclic stretch regime can be applied to the cell embedded hydrogels. As dynamic movement of the motor is encoded by the microcontroller software, any cyclic profile can be programmed, constrained only by the resolution and the rotation speed of the motor employed in the system (more details available in Roitblat Riba et al.41). In this work, we demonstrated that fibrin gel can sustain adhesion under dynamic stretch (Table 1). Though further testing is necessary, the application of cyclic loading would provide insights into the dynamic response of the hydrogel and embedded cells.
(ii) The geometry and volume of the silicone cut-out can be modified. In the current protocol, we focused only on gels in a circular geometry. Under such conditions, internal strains are relatively homogeneous throughout the gel, as predicted by our FE simulations (Figure 13). However, if other gel geometries will be considered by modifying the cut-out section of the silicone, gradients will emerge. For example, a &#8216;diamond&#8217; shaped cut-out could potentially lead to a gradient in the gel strains due to a gradual change in the initial length of the cut-out along the direction of stretch. This method of gel strain control is especially appealing, as different shaped cut-outs are relatively trivial to manufacture. Additionally, the thickness and the area of the silicone cut-out could be easily altered allowing for adjustments in gel volumes, from miniature volumes of a few microliters to larger volumes associated with mm-cm scale gel sizes. Moreover, it is possible to include several cut-outs in the same silicone strip, allowing for stretching of several gels simultaneously.
(iii) Cells can be embedded in the hydrogel. There are a few challenges that should be taken into consideration when embedding cells in the stretched hydrogels. Most cells embedded in 3D hydrogels apply considerable traction forces on the matrix, as well as degrading the matrix over time. This can result in global contraction and digestion of the gel, which can lead to disconnection of the gel from the silicone cut-out inner wall at prolonged times of incubation. A possible way to cope with this situation is to use low cell concentrations and limiting the time of cellular experiments to an interval that gel contraction and degradation are minimal. We have previously demonstrated the possibility of culturing fibroblast cells in the fibrin gel under static stretch of 10%, over a period of 5 hours, without any evidence of gel tearing or disconnection from the silicone cut-out41.
(iv) This setup should also be tested with other types of hydrogels, in particular synthetic ones such as polyacrylamide (PAA) or polyethylene glycol (PEG)-based hydrogels, which are commonly used for cell culture. The natural adhesion of these hydrogels to the silicone cut-out should be examined, and if found to be insufficient for gel stretching, then techniques to encourage adhesion should be used. The use of biological glues48,49 or chemical treatments50,51,52 to the cut-out surface should be considered.

Tissue response to mechanical forces is an integral part of a wide range of biological functions, including gene expression1, cell differentiation2, and tissue remodeling3. Moreover, force-induced changes in the extracellular matrix (ECM) such as fiber alignment and densification can impact cell behavior and tissue formation4,5,6. The ECM&#8217;s fibrous mesh structure has intriguing mechanical properties, such as non-linear elasticity, non-affine deformation and plastic deformations7,8,9,10,11,12. These properties impact how cells and their surrounding microenvironment respond to external mechanical forces13,14. Understanding how the ECM and tissues respond to mechanical forces will enable progress in the field of tissue engineering and in the development of more accurate computational and theoretical models.
Most common methods to mechanically stretch samples have focused on cell-laden 2D substrates to explore the effects on cell behavior. These include, for example, applying strain to polydimethylsiloxane (PDMS) substrates and analyzing cell reorientation angles in relation to the stretch direction15,16,17,18,19. Yet, methods investigating the response of 3D cell-embedded hydrogels to external stretch, a situation that more closely mimics tissue microenvironment, are more limited. Advances toward 3D stretching methods are of particular importance because cells behave differently on 2D substrates when compared to 3D matrices20. These behaviors include cellular realignment, protein expression levels, and migration patterns21,22,23.
Methods and devices that allow for 3D sample stretching include both commercially available24,25,26,27,28 and those developed for laboratory research29. These methods use distensible silicone tubes30, multi-well chambers31, clamps26,32, bioreactors11,33, cantilevers34,35,36, and magnets37,38. Some techniques generate stretch that locally deforms 3D hydrogels, for example by pulling needles from two single points in the gel5, while others allow for deformation of the entire bulk of the gel16. Moreover, most of these systems focus on analysis of the strain field in the X-Y plane, with limited information on the strain field in the Z-direction. Additionally, only a handful of these devices are capable of microscopic in situ imaging. The main challenge with in situ high-magnification imaging (e.g., confocal microscope) is the limited working distance of a few hundred microns from the objective lens to the sample. Devices that do allow live imaging during stretch sacrifice the uniformity of strain in the Z-axis or are relatively complex and difficult to reproduce in other laboratories39,40.
This approach to stretch 3D hydrogels allows for static or cyclical uniaxial strain during live confocal microscopy. The stretching device (referred to as &#8216;Smart Cyclic Uniaxial Stretcher &#8211; SCyUS&#8217;) is constructed using 3D printed parts and low-cost hardware, allowing easy reproduction in other labs. Attached to the device is a commercially available silicone rubber with a geometric cut-out in its center. Hydrogel components are polymerized to fill the cut-out. During polymerization, biological hydrogels, such as fibrin or collagen, naturally adhere to the interior walls of the cut-out. Using the SCyUS, the silicone strip is uniaxially stretched, transferring controlled strains to the embedded 3D hydrogel41.
This system allows for a unique combination of features and advantages compared to other existing methods. First, the system allows uniaxial stretching of thick 3D soft hydrogels (&#62;100 &#181;m thick, &#60;1 kPa stiffness) from their periphery, with Z-homogenous deformation throughout the hydrogel. These hydrogels are too soft to be gripped and stretched by conventional tensile techniques. Second, the stretching device can be easily replicated in other labs since 3D printing is readily available to researchers and the electronics used in the design are low-cost. Third, and perhaps the most attractive feature, the geometry and the size of the cut-out in the silicone strip can be easily manipulated, allowing for tunable strain gradients and boundary conditions as well as the use of a variety of sample volumes, down to a few microliters.
The presented protocol consists of molding fibrin gel into ~2 mm diameter disks in 0.5 mm thick silicone rubber strips proceeded by uniaxial stretch under live confocal microscopy. The following discusses in detail the experimental procedures for measuring and analyzing the strains acting on the geometric cut-out, the internal strains developed in the hydrogel, as well as resulting fiber alignment after various stretch manipulations. Finally, the possibility of embedding cells in the hydrogel and exposing them to controlled external stretch is discussed.

<table><tbody><tr><td>Alexa Fluor 546 carboxylic acid, succinimidyl ester</td><td>Invitrogen</td><td>A20002</td><td></td></tr><tr><td>Cell Medium (DMEM High Glucose)</td><td>Biological Industries</td><td>01-052-1A</td><td>Add 10% FBS, 1% PNS, 1% L-Glutamine, 1% Sodium Pyruvate</td></tr><tr><td>Cover Slip #1.5</td><td>Bar-Naor Ltd.</td><td>BN72204-30</td><td>22&amp;times;40 &lt;em&gt;mm&lt;/em&gt;</td></tr><tr><td>DIMETHYL SULPHOXIDE 99.5% GC DMSO</td><td>Sigma-Aldrich Inc.</td><td>D-5879-500 ML</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate-Buffered Saline</td><td>Biological Industries</td><td>02-023-1A</td><td></td></tr><tr><td>EVICEL Fibrin Sealant (Human)</td><td>Omrix Biopharmaceuticals</td><td>3902</td><td>Fibrinogen: 70 mg/mL, Thrombin: 800-1200 IU/&lt;em&gt;mL&lt;/em&gt;</td></tr><tr><td>Fibrinogen Buffer</td><td>N/A</td><td></td><td>Recipe for 1L: 7g NaCl, 2.94g trisodium citrate dihydrate, 9g glycine, 20g arginine hydrochloride &amp;amp; 0.15g calcium chloride dihydrate. Bring final volume to 1L with PuW (pH 7.0-7.2)</td></tr><tr><td>Fluorescent micro-beads FluoSpheres (1 &lt;em&gt;&amp;micro;m&lt;/em&gt;)</td><td>Invitrogen</td><td>F8820</td><td>Orange (540/560)&lt;br/&gt; Provided as suspension (2% solids) in water plus 2 mM sodium azide</td></tr><tr><td>High-Temperature Silicone Rubber</td><td>McMaster-Carr</td><td>3788T41</td><td>580 &lt;em&gt;&amp;micro;m&lt;/em&gt;-thick&lt;br/&gt; E = 1.5 Mpa&lt;br/&gt; Poisson Ratio = 0.48&lt;br/&gt; Tensile Strength = 4.8 MPa&lt;br/&gt; Upper limit of stretch = +300% engineering strain</td></tr><tr><td>HiTrap desalting column 5 &lt;em&gt;mL&lt;/em&gt; (Sephadex G-25 packed)</td><td>GE Healthcare</td><td>17-1408-01</td><td></td></tr><tr><td>HIVAC-G High Vacuum Sealing Compound</td><td>Shin-Etsu Chemical Co., Ltd.</td><td>HIVAC-G 100</td><td></td></tr><tr><td>ImageJ FIJI software&lt;sup&gt;39&lt;/sup&gt;</td><td>National Institute of Health, Bethesda, MD</td><td>Version 1.8.0_112</td><td></td></tr><tr><td>Microcontroller (Adruino Uno + Adafruit Motorshield v2.3)</td><td>Arduino/Adafruit</td><td>Arduino-DK001/Adafruit-1438</td><td></td></tr><tr><td>MicroVL 21R Centrifuge</td><td>Thermo Scientific</td><td>75002470</td><td></td></tr><tr><td>Parafilm</td><td>Bemis</td><td>PM-996</td><td></td></tr><tr><td>Primovert Light Microscope</td><td>Carl Zeiss Suzhou Co., Ltd.</td><td>491206-0011-000</td><td></td></tr><tr><td>SCyUS CAD (Solidworks)</td><td>Dassault Syst&amp;egrave;mes</td><td>N/A</td><td></td></tr><tr><td>SCyUS Code&lt;sup&gt;37&lt;/sup&gt;</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Servomotor - TowerPro SG-5010</td><td>Adafruit</td><td>155</td><td></td></tr><tr><td>SL 16R Centrifuge</td><td>Thermo Scientific</td><td>75004030</td><td>For 50 &lt;em&gt;mL&lt;/em&gt; tubes</td></tr><tr><td>Sterile 10 &lt;em&gt;cm&lt;/em&gt; non-culture plates</td><td>Corning</td><td>430167</td><td></td></tr><tr><td>Thrombin buffer</td><td>N/A</td><td></td><td>Recipe for 1L: 20g mannitol, 8.77g NaCl, 2.72g sodium acetate trihydrate, 24 mL 25% Human Serum Albumin, 5.88g calcium chloride. Bring final volume to 1L with PuW (pH 7.0)</td></tr><tr><td>Trypsin EDTA Solution B (0.25%), EDTA (0.05%)</td><td>Biological Industries</td><td>03-052-1B</td><td></td></tr><tr><td>USB Cable (Type B Male to Type A Male)</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Zeiss LSM 880 Confocal Microscope</td><td>Carl Zeiss AG</td><td>2811000417</td><td></td></tr><tr><td>ZEN 2.3 SP1 FP3 (black)</td><td>Carl Zeiss AG</td><td>Release Version 14.0.0.0</td><td></td></tr></tbody></table>

controlled strain of 3d hydrogels under live microscopy imaging

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized in vitro stretching methods. Various available systems use 2D substrate-based stretchers, while the accessibility of 3D techniques to strain soft hydrogels, is more restricted. Here, we describe a method that allows external stretching of soft hydrogels from their circumference, using an elastic silicone strip as the sample carrier. The stretching system utilized in this protocol is constructed from 3D-printed parts and low-cost electronics, making it simple and easy to replicate in other labs. The experimental process begins with polymerizing thick (&#62;100 &#956;m) soft fibrin hydrogels (Elastic Modulus of ~100 Pa) in a cut-out at the center of a silicone strip. Silicone-gel constructs are then attached to the printed-stretching device and placed on the confocal microscope stage. Under live microscopy the stretching device is activated, and the gels are imaged at various stretch magnitudes. Image processing is then used to quantify the resulting gel deformations, demonstrating relatively homogenous strains and fiber alignment throughout the gel&#8217;s 3D thickness (Z-axis). Advantages of this method include the ability to strain extremely soft hydrogels in 3D while executing in situ microscopy, and the freedom to manipulate the geometry and size of the sample according to the user&#8217;s needs. Additionally, with proper adaptation, this method can be used to stretch other types of hydrogels (e.g., collagen, polyacrylamide or polyethylene glycol) and can allow for analysis of cells and tissue response to external forces under more biomimetic 3D conditions.

Representative data from static stretch of increasing magnitudes applied to the silicone strip carrying a 3D fibrin hydrogel, embedded with 1 &#956;m fluorescent beads, is shown in Figure 9. The analysis demonstrates the effect of silicone stretch on geometric changes of the cut-out as well as the developed strains within the gel. Z-stack images of the entire gel are used to evaluate the deformation of the original circle shaped cut-out to the elliptical geometry (Figure 9A). These images are used to calculate &#949;xx(hole) (Equation 2). Zooming in and manually tracking several bead aggregates (Figure 9B) during gel stretching, allows for the calculation of local gel strains at the center, in the axial &#949;xx(gel) and perpendicular &#949;yy(gel) directions (Figure 9C,D). The axial strains propagate relatively linearly from the silicone cut-out edge to the center of the gel and are larger than the compressive perpendicular strains (Figure 9D).
High-magnification images of the fluorescently labeled gel allows for observation and quantification of fiber alignment, induced by the silicone stretch. Figure 11 shows representative zoom-in images of a typical un-stretched and relatively isotropic hydrogel (Figure 11A) and a hydrogel under high 80% cut-out strain depicting highly aligned fibers in the stretch direction (Figure 11B). Analysis of fiber reorientation using ImageJ revealed approximately linear dependence between fiber alignment (NOP) and the external strain on the cut-out (&#949;hole) up to strains of 40%, with moderate saturation at strains above 40% (Figure 11D). A detailed analysis of fiber alignment in different Z-slices is shown in Figure 12. Here, histograms of fiber alignment are calculated for each Z-slice (Figure 12A), as well as their corresponding NOP (Figure 12B), and the averaged NOP over all Z-slices for each external strain magnitude of the cut-out (Figure 12C). As &#949;hole increases, fiber alignment increases, following an overall non-linear curve as shown in Figure 12C. Note that the gel contracts along the Z-axis due to the Poisson effect, represented by the shorter lines at higher strains in Figure 12B.
Since it has proven difficult to perform in-depth analysis of the internal strains in the entire gel, we provide here preliminary results of finite element (FE) simulations performed on 2D continuous material given the mechanical properties of fibrin (Figure 13). The external strain of the cut-out is ~40% and the color-map represents the stretch-induced strain field. The color-map ranges from 38%-42%, indicating that gel strains are relatively homogenous throughout the circular gel area. We note that if other gel geometries are used, gradients in strain may arise, and this is a topic for future studies.
In order to demonstrate that embedded fibrin gels can sustain adhesion to the silicone strip at high strains (0 to +30%) and frequencies (up to 1 Hz), we ran preliminary tests (protocol for this test is not included in this work) showing no detachments of the gels from the inner walls of the cut-out. Three gels were tested, gel 1 for a total duration of ~87 minutes, gel 2 for 60 minutes &#38; gel 3 for 30 min total (Table 1). In all three of these cases fibrin hydrogels maintained their adhesion to the silicone.
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/61671/61671fig11.jpg" /> 
Figure 11: Gel fiber alignment in response to external stretch. Images taken at the center of a fluorescently-labeled gel while (A) unstretched and (B) after the application of &#949;hole= 80% (C) Averaged histograms of fiber orientations under increasing &#949;hole [%] strains. Grey error bars indicate differences between 40 slices in the z-stack assessed for each tested strain (D) Averaged Nematic Order Parameter (NOP) as a function of &#949;hole for three different gel samples (including &#8220;Gel 1&#8221; that is analyzed in Figure 11A-C). Error bars indicate differences between slices in the Z-stack of each stretch magnitude. This figure has been adapted with permission from Roitblat Riba et al.41 <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61671/61671fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" class="xfigimg" src="/files/ftp_upload/61671/61671fig12.jpg" /> 
Figure 12: Analysis of fiber orientation using the OrientationJ44 tool (ImageJ software)43 (A) 3D histograms of fiber orientation for different Z-slices. Shown are examples of two histograms calculated for stretched (&#949;hole = 17.6%, top) and un-stretched (bottom) gels. The histogram is normalized such that the area under the curve is equal to 1 (B) Resulting NOP versus Z-depth; the graph shows the calculations of NOP of each histogram as a function of Z-depth (C) Mean NOP as a function of &#949;hole. Error bars represent the standard deviation of NOP for all the Z-slices in an individual stretch magnitude at the same XY point in the same gel. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61671/61671fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 13" class="xfigimg" src="/files/ftp_upload/61671/61671fig13v2.jpg" /> 
Figure 13: 2D finite elements (FE) simulation of a stretched gel. Gel is stretched (&#949;hole &#8776; 40%) by a silicone carrier. Strain field is presented with values of maximum principal strain where minor gradients are observed. The circle represented by the dotted line is the pre-strained geometry of the hole, the original length (lo) is the center length along the x-axis before stretch is applied, and the stretched length (l) is the center length along the x-axis after stretch is applied. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61671/61671fig13v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:font-size="8px" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="10">Gel #1</td>
			<td>Dynamic strain amplitude (Frequency 1 Hz)</td>
			<td>Dynamic stretch duration (min)</td>
		</tr>
		<tr>
			<td rowspan="5">0-37%</td>
			<td>0.1</td>
		</tr>
		<tr>
			<td>1</td>
		</tr>
		<tr>
			<td>5</td>
		</tr>
		<tr>
			<td>15</td>
		</tr>
		<tr>
			<td>30</td>
		</tr>
		<tr>
			<td rowspan="3">0-48%</td>
			<td>1</td>
		</tr>
		<tr>
			<td>5</td>
		</tr>
		<tr>
			<td>30</td>
		</tr>
		<tr>
			<td>Total time under dynamic stretch</td>
			<td>87 min</td>
		</tr>
		<tr>
			<td rowspan="7">Gel #2</td>
			<td>Dynamic strain amplitude (Frequency 1 Hz)</td>
			<td>Dynamic stretch duration (min)</td>
		</tr>
		<tr>
			<td rowspan="4">0-30%</td>
			<td>0</td>
		</tr>
		<tr>
			<td>10</td>
		</tr>
		<tr>
			<td>20</td>
		</tr>
		<tr>
			<td>30</td>
		</tr>
		<tr>
			<td>0-50%</td>
			<td>30</td>
		</tr>
		<tr>
			<td>Total time under dynamic stretch</td>
			<td>60 min</td>
		</tr>
		<tr>
			<td rowspan="5">Gel #3</td>
			<td>Dynamic strain amplitude (Frequency 1 Hz)</td>
			<td>Dynamic stretch duration (min)</td>
		</tr>
		<tr>
			<td rowspan="3">0-30%</td>
			<td>10</td>
		</tr>
		<tr>
			<td>20</td>
		</tr>
		<tr>
			<td>30</td>
		</tr>
		<tr>
			<td>Total time under dynamic stretch</td>
			<td>30 min</td>
		</tr>
	</tbody>
</table>
Table 1: Representative results of dynamic stretch proof of concept. Three gels were embedded in silicone strips and stretched dynamically (1 Hz) for various durations. Strains ranged from zero to various magnitudes above 30%. All three gels&#160;successfully sustained their adhesion to the inner walls of the circular cut-out of the silicone strip.

Watch this Scientific Journal Video about Controlled Strain of 3D Hydrogels under Live Microscopy Imaging at JoVE.com

Controlled Strain of 3D Hydrogels under Live Microscopy Imaging

The presented method involves uniaxial stretching of 3D soft hydrogels embedded in silicone rubber while allowing live confocal microscopy. Characterization of the external and internal hydrogel strains as well as fiber alignment are demonstrated. The device and protocol developed can assess the response of cells to various strain regimes.

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized ...

controlled-strain-of-3d-hydrogels-under-live-microscopy-imaging

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

3.4K Views.  Tel-Aviv University. The presented method involves uniaxial stretching of 3D soft hydrogels embedded in silicone rubber while allowing live confocal microscopy. Characterization of the external and internal hydrogel strains as well as fiber alignment are demonstrated. The device and protocol developed can assess the response of cells to various strain regimes.

Video: Controlled Strain of 3D Hydrogels under Live Microscopy Imaging

Microfabricated Platforms for Mechanically Dynamic Cell Culture

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or fundamental cell biology studies is limited by current bioreactor technologies, which cannot simultaneously apply a variety of mechanical stimuli to cultured cells. In order to address this issue, we have developed a series of microfabricated platforms designed to screen for the effects of mechanical stimuli in a high-throughput format. In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and further demonstrate how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or ...

Live Cell Imaging during Mechanical Stretch

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in their capability for measuring responses in real time in live cells or viable tissue. A protocol was developed with the use of a cell actuator to distend live cells grown on or tissues attached to an elastic substrate while imaging with confocal and atomic force microscopy (AFM). Preliminary studies show that tonic stretching of human bronchial epithelial cells caused a significant increase in the production of mitochondrial superoxide. Moreover, using this protocol, alveolar epithelial cells were stretched and imaged, which showed direct damage to the epithelial cells by overdistention simulating one form of lung injury in vitro. A protocol to conduct AFM nano-indentation on stretched cells is also provided.

A novel imaging protocol was developed using a custom motor-driven mechanical actuator to allow the measurement of real time responses to mechanical strain in live cells. Relevant to mechanobiology, the system can apply strains up to 20% while allowing near real-time imaging with confocal or atomic force microscopy.

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in ...

Gradient Strain Chip for Stimulating Cellular Behaviors in Cell-laden Hydrogel

Artificial guidance for cellular alignment is a hot topic in the field of tissue engineering. Most of the previous research has investigated single strain-induced cellular alignment on a cell-laden hydrogel by using complex experimental processes and mass controlling systems, which are usually associated with contamination issues. Thus, in this article, we propose a simple approach to building a gradient static strain using a fluidic chip with a plastic PDMS cover and a UV transparent glass substrate for the stimulation of cellular behavior in a 3D hydrogel. Overloading photo-patternable cell prepolymer in the fluidic chamber can generate a convex curved PDMS membrane on the cover. After UV crosslinking, through a concentric circular micropattern under the curved PDMS membrane, and buffer washing, a microenvironment for investigating cell behaviors under a variety of gradient strains is self-established in a single fluidic chip, without external instruments. NIH3T3 cells were demonstrated after observing the change in the cellular alignment trend under geometry guidance, in cooperation with strain stimulation, which varied from 15 - 65% on hydrogels. After a 3-day incubation, the hydrogel geometry dominated the cell alignment under low compressive strain, where cells aligned along the hydrogel elongation direction under high compressive strain. Between these, the cells showed random alignment due to the dissipation of the radical guidance of hydrogel elongation and the geometry guidance of the patterned hydrogel.

This article introduces a simple approach to providing non-continuous gradient static strains on a concentric cell-laden hydrogel to regulate cell alignment for tissue engineering.

Artificial guidance for cellular alignment is a hot topic in the field of tissue engineering. Most of the previous research has investigated single ...

Studying the Effects of Matrix Stiffness on Cellular Function using Acrylamide-based Hydrogels

Tissue stiffness is an important determinant of cellular function, and changes in tissue stiffness are commonly associated with fibrosis, cancer and cardiovascular disease1-11. Traditional cell biological approaches to studying cellular function involve culturing cells on a rigid substratum (plastic dishes or glass coverslips) which cannot account for the effect of an elastic ECM or the variations in ECM stiffness between tissues. To model in vivo tissue compliance conditions in vitro, we and others use ECM-coated hydrogels. In our laboratory, the hydrogels are based on polyacrylamide which can mimic the range of tissue compliances seen biologically12. "Reactive" cover slips are generated by incubation with NaOH followed by addition of 3-APTMS. Glutaraldehyde is used to cross-link the 3-APTMS and the polyacrylamide gel. A solution of acrylamide (AC), bis-acrylamide (Bis-AC) and ammonium persulfate is used for the polymerization of the hydrogel. N-hydroxysuccinimide (NHS) is incorporated into the AC solution to crosslink ECM protein to the hydrogel. Following polymerization of the hydrogel, the gel surface is coated with an ECM protein of choice such as fibronectin, vitronectin, collagen, etc.
The stiffness of a hydrogel can be determined by rheology or atomic force microscopy (AFM) and adjusted by varying the percentage of AC and/or bis-AC in the solution12. In this manner, substratum stiffness can be matched to the stiffness of biological tissues which can also be quantified using rheology or AFM. Cells can then be seeded on these hydrogels and cultured based upon the experimental conditions required. Imaging of the cells and their recovery for molecular analysis is straightforward. For this article, we define soft substrata as those having elastic moduli (E) &lt;3000 Pascal and stiff substrata/tissues as those with E &gt;20,000 Pascal.

The effect of substrata stiffness on cellular function can be modeled in vitro using polyacrylamide hydrogels of varying compliances.

Tissue stiffness is an important determinant of cellular function, and changes in tissue stiffness are commonly associated with fibrosis, cancer and ...

Biology

Initial 3D Cell Cluster Control in a Hybrid Gel Cube Device for Repeatable Pattern Formations

The importance of in vitro 3D cultures is considerably emphasized in cell/tissue culture. However, the lack of experimental repeatability is one of its restrictions. Producing few repeatable results of pattern formation deteriorates the analysis of the mechanisms underlying the self-organization. Reducing variation in initial culture conditions, such as the cell density and distribution in the extracellular matrix (ECM), is crucial to enhance the repeatability of a 3D culture. In this article, we demonstrate a simple but robust procedure for controlling the initial cell cluster shape in a 3D extracellular matrix to obtain highly repeatable pattern formations. A micromold with a desired shape was fabricated by using photolithography or a machining process, and it formed a 3D pocket in the ECM contained in a hybrid gel cube (HGC). Highly concentrated cells were then injected in the pocket so that the cell cluster shape matched with the fabricated mold shape. The employed HGC allowed multi-directional scanning by its rotation, which enabled high-resolution imaging and the capture of the entire tissue structure even though a low-magnification lens was used. Normal human bronchial epithelial cells were used to demonstrate the methodology.&#160;

We present a procedure for controlling the initial cell cluster shape in a 3D extracellular matrix to obtain a repeatable pattern formation. A cubic device containing two different hydrogels is employed to achieve multi-directional imaging for tissue pattern formation.

The importance of in vitro 3D cultures is considerably emphasized in cell/tissue culture. However, the lack of experimental repeatability is one of its ...

High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the effect of applied extracellular tensile strain on cell populations in vitro via biochemical assays, a device has previously been designed which can be fabricated simply and is small enough to fit inside tissue culture incubators, as well as on top of microscope stages. However, the previous design of the polydimethylsiloxane substratum did not allow high-resolution subcellular imaging via oil-immersion objectives. This work describes a redesigned geometry of the polydimethylsiloxane substratum and a customized imaging setup that together can facilitate high-resolution subcellular imaging of live cells while under applied strain. This substratum can be used with the same, earlier designed device and, hence, has the same advantages as listed above, in addition to allowing high-resolution optical imaging. The design of the polydimethylsiloxane substratum can be improved by incorporating a grid which will facilitate tracking the same cell before and after the application of strain. Representative results demonstrate high-resolution time-lapse imaging of fluorescently labeled nuclei within strained cells captured using the method described here. These nuclear dynamics data give insights into the mechanism by which applied tensile strain promotes differentiation of oligodendrocyte progenitor cells.&#160;

Using a previously designed device to apply mechanical strain to adherent cells, this paper describes a redesigned substratum geometry and a customized apparatus for high-resolution single-cell imaging of strained cells with a 100x oil immersion objective.

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the ...

A Multi-well Format Polyacrylamide-based Assay for Studying the Effect of Extracellular Matrix Stiffness on the Bacterial Infection of Adherent Cells

Extracellular matrix stiffness comprises one of the multiple environmental mechanical stimuli that are well known to influence cellular behavior, function, and fate in general. Although increasingly more adherent cell types&#39; responses to matrix stiffness have been characterized, how adherent cells&#39; susceptibility to bacterial infection depends on matrix stiffness is largely unknown, as is the effect of bacterial infection on the biomechanics of host cells. We hypothesize that the susceptibility of host endothelial cells to a bacterial infection depends on the stiffness of the matrix on which these cells reside, and that the infection of the host cells with bacteria will change their biomechanics. To test these two hypotheses, endothelial cells were used as model hosts and Listeria monocytogenes as a model pathogen. By developing a novel multi-well format assay, we show that the effect of matrix stiffness on infection of endothelial cells by L. monocytogenes can be quantitatively assessed through flow cytometry and immunostaining followed by microscopy. In addition, using traction force microscopy, the effect of L. monocytogenes infection on host endothelial cell biomechanics can be studied. The proposed method allows for the analysis of the effect of tissue-relevant mechanics on bacterial infection of adherent cells, which is a critical step towards understanding the biomechanical interactions between cells, their extracellular matrix, and pathogenic bacteria. This method is also applicable to a wide variety of other types of studies on cell biomechanics and response to substrate stiffness where it is important to be able to perform many replicates in parallel in each experiment.

We have developed a multi-well format polyacrylamide-based assay for probing the effect of extracellular matrix stiffness on bacterial infection of adherent cells. This assay is compatible with flow cytometry, immunostaining, and traction force microscopy, allowing for quantitative measurements of the biomechanical interactions between cells, their extracellular matrix, and pathogenic bacteria.

Extracellular matrix stiffness comprises one of the multiple environmental mechanical stimuli that are well known to influence cellular behavior, ...

Immunology and Infection

Control of Cell Adhesion using Hydrogel Patterning Techniques for Applications in Traction Force Microscopy

Traction force microscopy (TFM) is the main method used in mechanobiology to measure cell forces. Commonly this is being used for cells adhering to flat soft substrates that deform under cell traction (2D-TFM). TFM relies on the use of linear elastic materials, such as polydimethylsiloxane (PDMS) or polyacrylamide (PA). For 2D-TFM on PA, the difficulty in achieving high throughput results mainly from the large variability of cell shapes and tractions, calling for standardization. We present a protocol to rapidly and efficiently fabricate micropatterned PA hydrogels for 2D-TFM studies. The micropatterns are first created by maskless photolithography using near-UV light where extracellular matrix proteins bind only to the micropatterned regions, while the rest of the surface remains non-adhesive for cells. The micropatterning of extracellular matrix proteins is due to the presence of active aldehyde groups, resulting in adhesive regions of different shapes to accommodate either single cells or groups of cells. For TFM measurements, we use PA hydrogels of different elasticity by varying the amounts of acrylamide and bis-acrylamide and tracking the displacement of embedded fluorescent beads to reconstruct cell traction fields with regularized Fourier Transform Traction Cytometry (FTTC).
To further achieve precise recording of cell forces, we describe the use of a controlled dose of patterned light to release cell tractions in defined regions for single cells or groups of cells. We call this method local UV illumination traction force microscopy (LUVI-TFM). With enzymatic treatment, all cells are detached from the sample simultaneously, whereas with LUVI-TFM traction forces of cells in different regions of the sample can be recorded in sequence. We demonstrate the applicability of this protocol (i) to study cell traction forces as a function of controlled adhesion to the substrate, and (ii) to achieve a greater number of experimental observations from the same sample.

Near-UV lithography and traction force microscopy are combined to measure cellular forces on micropatterned hydrogels. The targeted light-induced release of single cells enables a high number of measurements on the same sample.

Traction force microscopy (TFM) is the main method used in mechanobiology to measure cell forces. Commonly this is being used for cells adhering to flat ...

Pattern Generation for Micropattern Traction Microscopy

Micropattern traction microscopy allows control of the shape of single cells and cell clusters. Furthermore, the ability to pattern at the micrometer length scale allows the use of these patterned contact zones for the measurement of traction forces, as each micropatterned dot allows for the formation of a single focal adhesion that then deforms the soft, underlying hydrogel. This approach has been used for a wide range of cell types, including endothelial cells, smooth muscle cells, fibroblasts, platelets, and epithelial cells. 
This review describes the evolution of techniques that allow the printing of extracellular matrix proteins onto polyacrylamide hydrogels in a regular array of dots of prespecified size and spacing. As micrometer-scale patterns are difficult to directly print onto soft substrates, patterns are first generated on rigid glass coverslips that are then used to transfer the pattern to the hydrogel during gelation. First, the original microcontact printing approach to generate arrays of small dots on the coverslip is described. A second step that removes most of the pattern to leave islands of small dots is required to control the shapes of cells and cell clusters on such arrays of patterned dots. 
Next, an evolution of this approach that allows for the generation of islands of dots using a single subtractive patterning step is described. This approach is greatly simplified for the user but has the disadvantage of a decreased lifetime for the master mold needed to make the patterns. Finally, the computational approaches that have been developed for the analysis of images of displaced dots and subsequent cell-generated traction fields are described, and updated versions of these analysis packages are provided.

We describe improvements to a standard method for measuring cellular traction forces, based on microcontact printing with a single subtractive patterning step of dot arrays of extracellular matrix proteins on soft hydrogels. This method allows for simpler and more consistent fabrication of island patterns, essential for controlling cell cluster shape.

Micropattern traction microscopy allows control of the shape of single cells and cell clusters. Furthermore, the ability to pattern at the micrometer ...

Microengineering 3D Collagen Hydrogels with Long-Range Fiber Alignment

Aligned collagen I (COL1) fibers guide tumor cell motility, influence endothelial cell morphology, control stem cell differentiation, and are a hallmark of cardiac and musculoskeletal tissues. To study cell response to aligned microenvironments in vitro, several protocols have been developed to generate COL1 matrices with defined fiber alignment, including magnetic, mechanical, cell-based, and microfluidic methods. Of these, microfluidic approaches offer advanced capabilities such as accurate control over fluid flows and the cellular microenvironment. However, the microfluidic approaches to generate aligned COL1 matrices for advanced in vitro culture platforms have been limited to thin "mats" (&lt;40 &#181;m in thickness) of COL1 fibers that extend over distances less than 500 &#181;m and are not conducive to 3D cell culture applications. Here, we present a protocol to fabricate 3D COL1 matrices (130-250 &#181;m in thickness) with millimeter-scale regions of defined fiber alignment in a microfluidic device. This platform provides advanced cell culture capabilities to model structured tissue microenvironments by providing direct access to the micro-engineered matrix for cell culture.

This protocol demonstrates the use of a microfluidic channel with changing geometry along the fluid flow direction to generate extensional strain (stretching) to align fibers in a 3D collagen hydrogel (&lt;250 &#181;m in thickness). The resulting alignment extends across several millimeters and is influenced by the extensional strain rate.

Aligned collagen I (COL1) fibers guide tumor cell motility, influence endothelial cell morphology, control stem cell differentiation, and are a hallmark ...