M. T. Holley is supported by the Graduate Fellows program of the Louisiana Board of Regents, and C. Danielson is supported by the Howard Hughes Medical Institute Professors Program. This study is supported by NSF Grant No: 1530884.

All procedures described here have been carried out using an approved protocol and in accordance with the regulations of the Institutional Animal Care and Use Committee of the University of Notre Dame.
1. Cell Seeding and Culture
<ol>
	<li>Before starting, prepare the required items: a small funnel, pipettes, and warm Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% Fetal Bovine Serum (FBS) and 1% penicillin antibiotic (DMEM complete).</li>
	<li>Take T-25 flasks along with the functionalized device (biological actuator or the biorobot) within it. Refer to section 4 of Part 1 of this two-part manuscript for details regarding device preparation, functionalization, and storage prior to cell seeding.</li>
	<li>Prepare a funnel, which can be made by rolling a small, square plastic sheet. Place it over the biological actuator or biorobot inside the T-25 flask. Adjust the diameter of the wider end to fit the entire device and the height so that it fits snugly when the top of the flask is tightened.
	<ol>
		<li>For the biorobots, use a magnet to hold the device in place at the bottom of the flask throughout the seeding process. 
		Note: Here, a single neodymium disc magnet (1.26&#34; in diameter) was used, but any magnet of similar size and strength can also be used to hold down the biorobot with the magnetic nickel-PDMS composite base.</li>
		<li>UV sterilize the plastic sheet for at least 30 min prior usage.</li>
	</ol>
	</li>
	<li>Ensure that there is not a large gap between the base of the funnel and the flask.</li>
	<li>Resuspend the cardiomyocytes in DMEM complete at a density of 1.6 x 107 cells/mL and slowly drop 400 &#181;L of suspension onto the device through the funnel. Use a hemocytometer or any other cell counter to determine the number of cells obtained.</li>
	<li>Slowly move the system back into the incubator without disturbing the device and the funnel within. Culture for 24 h at 37 &#176;C.</li>
	<li>After the incubation period, slowly remove the funnel, gently wash the sample with PBS, and refill the flask with 10 mL of fresh DMEM complete. 
	Note: For the biorobots, remove the magnet so that the device is afloat.</li>
</ol>
2. Biochemical Characterization
<ol>
	<li>Calcium flux assay 
	Note: The calcium flux assay is carried out to assess cell interconnectivity. The procedure for loading the cells with the fluorescent, calcium ion-specific dye follows the process described in a previously established protocol11.
	<ol>
		<li>First, prepare the required materials, calcium fluo-4-acetomethyl (AM) ester, nonionic surfactant polyol (see the Table of Materials), and Tyrode&#39;s salt solution.</li>
		<li>Using long tweezers, gently transfer the device from the culture flask to a 35-mm Petri dish with 2 mL of Tyrode&#39;s salt solution.</li>
		<li>In a separate centrifuge tube, take 1 mL of warm Tyrode&#39;s solution (warmed to 37 &#176;C) and add 3-5 &#181;L of stock calcium fluo-4 AM dye (working concentration: 3-5 &#181;M) and an equal part of nonionic surfactant polyol (working concentration: 0.2 %). Replace the sample solution with warm Tyrode&#39;s solution supplemented with the calcium indicator dye, fluo-4, and 0.2% nonionic surfactant. Incubate for 25 - 30 min at 37 &#176;C.</li>
		<li>Remove the dye solution and gently wash the sample with fresh Tyrode&#39;s solution. Re-incubate the sample in 2 mL of fresh DMEM complete for another 30 min at 37 &#176;C prior to imaging. 
		Note: The results of this assay and related video are provided in the previously published work7.</li>
	</ol>
	</li>
	<li>Immunofluorescence 
	Note: The double immunostaining of all the samples was performed following previously established protocol12.
	<ol>
		<li>First, prepare 10% goat serum (GS) in phosphate-buffered saline solution (PBS), 4% paraformaldehyde (PFA) in diH2O, 0.1% cell lyse detergent (see the Table of Materials) in PBS, primary antibodies (anti-mouse monoclonal antibody cardiac troponin-I and anti-rabbit monoclonal antibody connexins-43), secondary antibodies (Alexa 594 conjugate and goat anti-mouse IgG (H+L) Alexa 488 conjugate), and DAPI. 
		Caution: Paraformaldehyde is carcinogenic.</li>
		<li>Remove the sample of interest from the flask and gently wash it twice with PBS. Refer to section 4 in Part 1 of this two-part manuscript for details regarding sample preparation and functionalization.</li>
		<li>Add a droplet of PBS onto a small coverslip (diameter: 12 mm or 15 mm). Gently hold the base of the device with tweezers and cut the thin PDMS arms (cantilever, Figure 1) using scissors from the end where it connects to the top of the base. Transfer the cantilever arms onto the droplet with the cell-adhered side facing upward. The PBS droplet will prevent the cells from drying.</li>
		<li>Fix the samples with 4% PFA and perform double immunostaining of the samples, as described previously12.</li>
		<li>After immunostaining, mount the samples onto a clean glass slide using anti-fade mounting reagent and set aside, undisturbed, in the dark for 24 h.</li>
		<li>Repeat the procedure for all samples. 
		Note: The results of this assay and the related images are discussed in depth in the previously published work7.</li>
	</ol>
	</li>
</ol>
3. Imaging
<ol>
	<li>Place the T-25 flask upright in a CO2 incubator and prepare the imaging system inside the incubator. Record the device using a camera (see the Table of Materials) with a zoom lens (see the Table of Materials). For a light source, use a strip of LED lights. 
	Note: A strip of white-light LEDs was used here, but any normal LEDs will also work.</li>
	<li>Connect the camera to an operating system and open the camera-specific software (see the Table of Materials). Click on the camera image underneath the &#34;File&#34; tab, in the top panel, to open all camera options, and choose the correct camera.</li>
	<li>Choose &#34;live&#34; from the list of tabs on the top panel within the software.</li>
	<li>Manually bring the image into focus by adjusting the lens dial. Select &#34;Crop to region of interest (ROI)&#34; from the top panel. Then, manually draw a rectangle in the video frame, enclosing the biological actuator device and the cantilever arms, to mark the ROI. 
	Note: Choosing an appropriate ROI minimizes the size of the image files.
	<ol>
		<li>In the case of biorobots, capture the whole screen in order to record the swimming motion of the device. 
		NOTE: There is no need to draw an ROI for the biorobots</li>
	</ol>
	</li>
	<li>Before starting the recording, select &#34;Camera Settings&#34; from one of the tabs in the top panel on the screen. Set the frame rate by adjusting the exposure and pixel ratio of the live image by sliding the bar for each or by manually entering the values. Set the frame rates to about 30 &#177; 2 fps. 
	Note: Changing the exposure and pixel ratio changes the brightness and contrast of the live image.</li>
	<li>Click the &#34;Record&#34; button from the top panel in the software to start recording the videos of the actuators with 1,000 x 1,000 pixel resolution for precisely 30 s. Repeat the process for all samples.</li>
</ol>
4. Image Analysis of the Biological Actuators on a Stationary Base
<ol>
	<li>Analyze the images using a programming software (e.g., Matlab) running the custom script. See the Table of Materials and the Supplemental File for further details. 
	Note: The script displays each frame of the recorded videos, receives the user&#39;s mouse input to record the coordination of the points of the cantilever on the images, calculates the diameter and center of the circle that passes the inputted points via least square fitting, and exports all of the inputted and calculated data for further use.
	<ol>
		<li>Open the programming software by clicking on the icon. Click &#34;File&#34; -&#62; &#34;Open&#34; from the menu bar at the top and select the .m script file for image analysis. Ensure that recorded TIFF images are in the same folder as the .m file. Click &#34;Run&#34; to run the script. 
		Note: An interactive display will pop up for altering.</li>
		<li>Press &#34;play&#34; to start the actual program. Click the &#34;open&#34; button and locate the TIFF file that is going to be analyzed.</li>
		<li>Click the &#34;base&#34; button and then click on the point where the cantilever attaches to the base at the top; hit enter. This will place a square marker on the image for each frame to denote the location of the cantilever base.</li>
		<li>Click the &#34;Scale&#34; button and then manually click on one edge of the glass bead. Bring the mouse pointer to the opposite side of the glass bead and press &#34;Enter.&#34; 
		Note: This should draw a line that measures the diameter of the glass bead. Since the glass bead is 3 mm in diameter, this will relate 3 mm to the pixels displayed.</li>
		<li>Click the &#34;Analyze&#34; button. Click along the cantilever at a short distance from the first square marker that represents the cantilever base.</li>
		<li>Click the &#8220;Analyze&#8221; button. Then, click along the cantilever at a short distance from the first square marker that represents the cantilever base. Continue to click along the cantilever, including the tip, and press enter when done. This will place an &#8220;x&#8221; on each point clicked on the cantilever. 
		Note: Based on the coordination of the square marker and on the x markers, the center and diameter of a circle will be calculated using a least square fitting function (refer to the attached supplemental file for the script used). The circle, which passes the x markers and the square maker, will be superimposed on the image automatically.</li>
		<li>Check if the superimposed circle correctly traces the cantilever profile. 
		Note: When the cantilever is very flat, it is hard to judge if the cantilever profile is traced correctly. Refer to Figure 3.</li>
		<li>Click the &#34;next frame&#34; button. This will switch to the next frame in the TIFF file. The base and scale are already set from the previous step.</li>
		<li>Repeat steps 4.1.5 through 4.1.7 until all of the frames in the TIFF file have been completed. Once all frames have been processed, click the &#34;Export&#34; button. 
		Note: This will make a spreadsheet file with the TIFF file name for the cantilever that was just analyzed. Edit the filename to include which side (left or right) of the cantilever was analyzed.</li>
	</ol>
	</li>
	<li>Calculating stress in the spreadsheet.
	<ol>
		<li>Calculate the surface stress, &#34;&#963;,&#34; on the cantilever using the following equation: 
		<img alt="Equation 1" src="/files/ftp_upload/55643/55643eq1.jpg" /> 
		where E, R, v, and h are Young&#39;s modulus, the radius of curvature, Poisson&#39;s ratio, and the cantilever thickness, respectively. 
		Note: The thickness of the cantilever can be altered to change the sensitivity. In this study, the values were as follows: E = 750 kPa, v = 0.49, and h = 25 &#181;m13,14.</li>
		<li>Calculate the surface stress using the equation in step 4.2.1. Open the .xls spreadsheet file. Note that the output has multiple columns showing first the x and y coordinates of base and circle and then the radius of curvature. Calculate the x and y coordinates of each point clicked based on these. 
		Note: Plotting the stresses over the time-lapse image frames shows changes in the force exerted on the cantilever over time. The troughs show the stress on the cantilever during the relaxation of the cardiomyocytes or the static stress exerted on the cantilever due to cell traction forces. The peaks show the dynamic stress on the cantilever, which was exerted by the beating of the cardiomyocytes. This value corresponds to the maximum amount of force generated by the contraction of the cardiomyocytes.</li>
	</ol>
	</li>
</ol>
5. Analysis of Swimming Biorobots
<ol>
	<li>Record the position of the biorobot using image analysis software. 
	Note: See the Materials list for the software used in this section.
	<ol>
		<li>Open the image analysis software (e.g., ImageJ). Press &#34;File&#34; and &#34;Open&#34; and select the swimming biorobot video file. Click &#34;OK&#34; and let the program load the file. Open the spreadsheet software.</li>
		<li>In the loaded biorobot video, find a reference of known dimensions (e.g., the glass bead 3 mm in diameter that was embedded in the biological actuator). 
		Note: Any object with known dimension will work. This will determine the pixel-to-length ratios in each video.</li>
		<li>Use the &#34;Straight&#34; tool to draw a line across the glass bead. Click &#34;Analyze&#34; and select &#34;Set scale.&#34; Set the &#34;Known distance&#34; field to &#34;3,000 &#181;m&#34; and click &#34;Ok.&#34; 
		Note: This will set the x and y coordinates to micrometers.</li>
		<li>Choose a point on the device that does not wobble between frames to act as a marker. 
		Note: It is recommend to choose a corner of the base.</li>
		<li>Point to the selected point in 5.1.4 on the first frame. Record the x and y coordinates on the spreadsheet.</li>
		<li>Switch back to the image analysis software window and press the right arrow key to change to the next frame. Point to the marker again (from step 5.1.4) and record the x and y coordinates on the spreadsheet.</li>
		<li>Repeat step 5.1.6 for all frames.</li>
	</ol>
	</li>
	<li>Calculate the swimming parameters of the biorobot using the spreadsheet of coordinates7.
	<ol>
		<li>Calculate the period between frames from the known frame rate of each video.</li>
		<li>Calculate the change in the x and y coordinates between frames to find the distance moved, including the total distance.</li>
		<li>Calculate the amplitude of contraction from the maximum change along the y-axis. Determine the beating frequency for each biorobot from the inverse of the period between two contractions.</li>
		<li>Calculate the swimming speed of each device from the total time and distance moved in the x-direction.</li>
		<li>Repeat step 5.2 for each biorobot video that was analyzed.</li>
		<li>Normalize each measured parameter. 
		Note: Normalize all the values to better visualize the differences. This protocol demonstrates normalization with respect to a horizontal mode biorobot with low-frequency contractions (horizontal LF) (Figure 4)7.</li>
	</ol>
	</li>
</ol>
6. Analysis of Protein Expression
Note: The mounted samples prepared in steps 2.2.4 and 2.2.5 were imaged using a confocal microscope. Images were acquired at 20X, 40X, and 60X magnification sequentially in three channels simultaneously: 460 nm, 488 nm, and 594 nm. A set of 5 images were captured at 40X magnification, from different positions for each sample, and each channel was saved as an individual .TIFF file. The exposure setting was determined by the magnification of the objective used and was set constant for all the captures at that magnification.
<ol>
	<li>Open the image analysis software and select &#34;File&#34; -&#62; &#34;Open&#34; to load the images.</li>
	<li>Draw a rectangular polygon on the image frame to mark the ROI. Select &#34;Analyze&#34; -&#62; &#34;Measure&#34; to measure the mean fluorescent intensity.</li>
	<li>Repeat step 6.2 to collect intensity measurements from all the samples and calculate the respective mean intensity for each condition. 
	Note: Here, a different condition refers to different time points, as in, day 1, day 2, and up to day 6.</li>
	<li>Export the results on a spreadsheet for further statistical analysis and for the production of data plots.</li>
</ol>

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The authors have nothing to disclose.

The procedure outlined here describes a successful seeding method for PDMS-based actuators and biorobots, which facilitates the attachment of cardiomyocytes. Furthermore, the process of image acquisition and the subsequent analysis that characterizes the behavior of the cells and the performance of the devices was described.
We observed spontaneous contraction of cells on the cantilever arms after 24 h; the intensity of contractions continued to increase steadily over time and reached a maximu...

Biorobots are devices based on living cells that are incorporated within a mechanical backbone that is usually composed of soft, elastic materials, such as PDMS or hydrogels1. The cells undergo rhythmic contractions, either spontaneously or in response to stimuli, and thus function as an actuator. The power generated from cell contraction drives various biorobots. Mammalian heart cells (cardiomyocytes) and skeletal muscle cells are often used for biorobot actuation due to their contractile properties. Aside from cardiomyocyte and skeletal muscle cells, other cell types, such as insect muscle tissues2 and explanted muscle tissues3, have been used. Insect muscle tissues enable the operation of biological actuators at room temperature.
The function and performance of a biorobot are chiefly determined by the strength and consistency of the biological actuator (i.e.&#160;muscle cells), while the mechanical backbone structure primarily determines the mechanisms of locomotion, stability, and power. Since these devices are solely driven by forces generated by cells, there are no chemical pollutants or operating noises. Therefore, they form an energy-efficient alternative to other conventional robots. Various literature sources have discussed the different methods to integrate living cells and tissues into biorobots1,4,5. Advances in microfabrication and tissue engineering techniques have enabled the development of biorobots that can walk, grip, swim, or pump5,6. In general, cells are cultured directly onto the mechanical (polymeric) backbone as a confluent cell sheet or they are molded into 3-dimensional actuating structures within scaffolds such as rings and strips. Most often, biorobots are fabricated using cardiomyocyte sheets6,7, as these cells have an innate ability to exhibit spontaneous contraction without external stimuli. On the other hand, reports on skeletal muscle cell sheets are limited due to their need for stimuli to initiate contractions in vitro in order to initiate membrane depolarization8.
This protocol first describes how to seed cardiomyocytes on a functionalized biological actuator made of a thin PDMS cantilever. It then describes in detail the seeding and analysis of the swimming profiles. The cantilever is functionalized with a cell adhesive protein such as fibronectin and is seeded confluently with cardiomyocytes. As the cells seeded on the device contract, they cause the cantilever to bend and thus to act as an actuator. Over time, as the cells mature, we trace the changes in surface stress on the device by analyzing videos of the cantilever bending. The biological actuator developed here can be used to determine the contractile properties of any cell type, such as the fibroblasts or induced pluripotent stems cells, as they undergo differentiation.
Much of the earlier research on biorobots has been focused on developing biological actuators, while optimization of the biorobot architecture and functional capabilities were largely neglected. Recently, a few studies have demonstrated the implementation of swimming modes in biorobots that are inspired by nature. For example, swimming biorobots with flagella-based motion6, jellyfish propulsion9, and bio-hybrid rays4 have been engineered. Unlike other works in literature, here we focus on varying the properties of the mechanical backbone to create a self-stabilizing structure. The biorobot developed in this study is capable of maintaining a constant pitch, roll, and immersion depth as it swims. These parameters can be modified by varying the thickness of each base composite. The fabrication steps involved in developing the PDMS actuator, the submergible biorobot, and the functionalization of the device are described in detail in Part 1 of this two-part article, as well as in our recent work7.The technique developed here can pave the way for the development of novel, highly efficient biorobots for various applications, such as cargo delivery.
The isolation process followed in this study is similar to the process described in an earlier work10, as well as in recently published work7. The microfabrication methods used for fabricating the PDMS actuators and biorobot devices are described in detail in Part 1 of this two-part manuscript. The protocol section of this manuscript describes the steps involved in seeding cardiomyocytes onto the fabricated PDMS actuator and the biorobot following their functionalization with cell adhesive proteins.

<table>
	<tbody>
		<tr>
			<td>Chemicals and reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cardiomyocytes (primary cardiac cells)</td>
			<td>Charles River</td>
			<td>NA</td>
			<td>Isolated from 2-day old neonatal Sprague Dawley rats</td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s modified eagle&#8217;s media (DMEM)</td>
			<td>Hyclone Laboratories</td>
			<td>16750-074</td>
			<td>with 4500 mg/L glucose, 4.0 mM L-glutamine, and 110 mg/L sodium pyruvate</td>
		</tr>
		<tr>
			<td>Fetalclone III serum</td>
			<td>Hyclone industries, GE</td>
			<td>16777-240</td>
			<td>Fetal bovin serum (FBS)</td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s phosphate buffer (PBS)</td>
			<td>Sigma-Aldrich</td>
			<td>D1408-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-G sodium salt</td>
			<td>Sigma-Aldrich</td>
			<td>P3032</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat serum</td>
			<td>Sigma-Aldrich</td>
			<td>G9023</td>
			<td></td>
		</tr>
		<tr>
			<td>4,6-diamidino-2-phenylindole dihydrocholride powder (DAPI)</td>
			<td>Sigma-Aldrich</td>
			<td>D9542</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin from bovine plasma</td>
			<td>Sigma-Aldrich</td>
			<td>F1141</td>
			<td>Solution (1 mg/ml)</td>
		</tr>
		<tr>
			<td>Calcein-AM and ethidium homodimer-1 kit (Live/Dead Assay)</td>
			<td>Molecular Probes</td>
			<td>L3224</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Fluo-4, AM</td>
			<td>Molecular Probes</td>
			<td>F14217</td>
			<td>calcium indicator dye</td>
		</tr>
		<tr>
			<td>Tyrodes salt solution</td>
			<td>Sigma-Aldrich</td>
			<td>T2397</td>
			<td>buffer solution</td>
		</tr>
		<tr>
			<td>Pluronic F-127</td>
			<td>Molecular Probes</td>
			<td>P3000MP</td>
			<td>nonionic surfactant-20 % solution in Dimethylsiloxane (DMSO)</td>
		</tr>
		<tr>
			<td>16% Parafomaldehyde</td>
			<td>Electron microscopy</td>
			<td>15710</td>
			<td>Caution: Irritant and combustible</td>
		</tr>
		<tr>
			<td>Triton x-100</td>
			<td>Sigma-Aldrich</td>
			<td>X-100 100 mL</td>
			<td>cell lyses detergent, (4-(1,1,3,3-Tetramethylbutyl)phenyl-polyethylene glycol, t-Octylphenoxypolyethoxyethanol, Polyethylene glycol tert-octylphenyl ether)</td>
		</tr>
		<tr>
			<td>ProLong gold antifade reagent</td>
			<td>Molecular Probes</td>
			<td>P10144</td>
			<td>Mounting agent</td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 Phalloidin</td>
			<td>Molecular Probes</td>
			<td>A12381</td>
			<td>Actin filament marker</td>
		</tr>
		<tr>
			<td>Goat anti-rabbit IgG (H+L) secondary antibody, Alexa Fluor 594 conjugate</td>
			<td>Molecular Probes</td>
			<td>A-11012</td>
			<td></td>
		</tr>
		<tr>
			<td>pha</td>
			<td>Molecular Probes</td>
			<td>A-11001</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-connexin 43 antibody</td>
			<td>Abcam</td>
			<td>ab11370</td>
			<td>Gap junction marker</td>
		</tr>
		<tr>
			<td>Anti-cardiac troponin I antibody</td>
			<td>Abcam</td>
			<td>ab10231</td>
			<td>Contractile protein</td>
		</tr>
		<tr>
			<td>16% EM grade paraformaldehyde solution</td>
			<td>Electron microscopy</td>
			<td>100503-916</td>
			<td></td>
		</tr>
		<tr>
			<td>Polydimethylsiloxane (PDMS)</td>
			<td>Elsevier</td>
			<td></td>
			<td>Sylgard 184</td>
		</tr>
		<tr>
			<td>Materials and Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Camera</td>
			<td>Thor Labs</td>
			<td>DCC1545M</td>
			<td></td>
		</tr>
		<tr>
			<td>LED light strip</td>
			<td>NA</td>
			<td>NA</td>
			<td>Any white LED without spectrum emission</td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Nikkon C2</td>
			<td>NA</td>
			<td>Confocal microscope with three filter set.</td>
		</tr>
		<tr>
			<td>Zooming lens</td>
			<td>Infinity</td>
			<td>Model# 252120</td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Matlab</td>
			<td>Mathworks</td>
			<td>NA</td>
			<td>Used in Section 4) for biological actuator analysis.</td>
		</tr>
		<tr>
			<td>Image J</td>
			<td>National Institute of Health</td>
			<td>NA</td>
			<td>Java-based image processing software. Used in Section 5) for biorobot analysis. 
			Free Image Processing and Analysis software in java. (https://imagej.nih.gov/ij/)</td>
		</tr>
		<tr>
			<td>Thor Cam</td>
			<td>Thor Labs</td>
			<td>NA</td>
			<td>Camera operating software</td>
		</tr>
	</tbody>
</table>

cardiac muscle cell-based actuator and self-stabilizing biorobot - part 2

In recent years, hybrid devices that consist of a living cell or tissue component integrated with a synthetic mechanical backbone have been developed. These devices, called biorobots, are powered solely by the force generated from the contractile activity of the living component and, due to their many inherent advantages, could be an alternative to conventional fully artificial robots. Here, we describe the methods to seed and characterize a biological actuator and a biorobot that was designed, fabricated, and functionalized in the first part of this two-part article. Fabricated biological actuator and biorobot devices composed of a polydimethylsiloxane (PDMS) base and a thin film cantilever were functionalized for cell attachment with fibronectin. Following functionalization, neonatal rat cardiomyocytes were seeded onto the PDMS cantilever arm at a high density, resulting in a confluent cell sheet. The devices were imaged every day and the movement of the cantilever arms was analyzed. On the second day after seeding, we observed the bending of the cantilever arms due to the forces exerted by the cells during spontaneous contractions. Upon quantitative analysis of the cantilever bending, a gradual increase in the surface stress exerted by the cells as they matured over time was observed. Likewise, we observed movement of the biorobot due to the actuation of the PDMS cantilever arm, which acted as a fin. Upon quantification of the swimming profiles of the devices, various propulsion modes were observed, which were influenced by the resting angle of the fin. The direction of motion and the beating frequency were also determined by the resting angle of the fin, and a maximum swim velocity of 142 &#181;m/s was observed. In this manuscript, we describe the procedure for populating the fabricated devices with cardiomyocytes, as well as for the assessment of the biological actuator and biorobot activity.

The biological actuator made of a thin PDMS cantilever (25 &#181;m in thickness) and cardiomyocytes constitutes the core of the swimming biorobot, as shown in the schematic and screenshot of the devices in Figure 1. The cells start to exhibit contractions after 24 h in culture, and bending of the cantilever arms was observed by day 2. The side profile of the device was recorded every day, and the surface stress was quantified from the bending of the cantilever arms using ...

Watch this Scientific Journal Video about Cardiac Muscle Cell-based Actuator and Self-stabilizing Biorobot - Part 2 at JoVE.com

Cardiac Muscle Cell-based Actuator and Self-stabilizing Biorobot - Part 2

In this study, a biological actuator and a self-stabilizing, swimming biorobot with functionalized elastomeric cantilever arms are seeded with cardiomyocytes, cultured, and characterized for their biochemical and biomechanical properties over time.

In recent years, hybrid devices that consist of a living cell or tissue component integrated with a synthetic mechanical backbone have been developed. ...

cardiac-muscle-cell-based-actuator-self-stabilizing-biorobot-part

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

8.7K Views.  University of Notre Dame. In this study, a biological actuator and a self-stabilizing, swimming biorobot with functionalized elastomeric cantilever arms are seeded with cardiomyocytes, cultured, and characterized for their biochemical and biomechanical properties over time.

Video: Cardiac Muscle Cell-based Actuator and Self-stabilizing Biorobot - Part 2

Shock Wave Application to Cell Cultures

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to target cells or tissue without any subsequent damage. Therefore, in vitro experiments are of increasing interest. Various methods of applying shock waves onto cell cultures have been described. In general, all existing models focus on how to best apply shock waves onto cells.
However, this question remains: What happens to the waves after passing the cell culture? The difference of the acoustic impedance of the cell culture medium and the ambient air is that high, that more than 99% of shock waves get reflected! We therefore developed a model that mainly consists of a Plexiglas built container that allows the waves to propagate in water after passing the cell culture. This avoids cavitation effects as well as reflection of the waves that would otherwise disturb upcoming ones. With this model we are able to mimic in vivo conditions and thereby gain more and more knowledge about how the physical stimulus of shock waves gets translated into a biological cell signal (&#8220;mechanotransduction&#34;).

Shock waves nowadays are well known for their regenerative effects. Therefore in vitro experiments are of increasing interest. We therefore developed a model for in vitro shock wave trials (IVSWT) that enables us to mimic in vivo conditions thereby avoiding distracting physical effects.

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to ...

Engineered Vascularized Muscle Flap

One of the main factors limiting the thickness of a tissue construct and its consequential viability and applicability in vivo, is the control of oxygen supply to the cell microenvironment, as passive diffusion is limited to a very thin layer. Although various materials have been described to restore the integrity of full-thickness defects of the abdominal wall, no material has yet proved to be optimal, due to low graft vascularization, tissue rejection, infection, or inadequate mechanical properties. This protocol describes a means of engineering a fully vascularized flap, with a thickness relevant for muscle tissue reconstruction. Cell-embedded poly L-lactic acid/poly lactic-co-glycolic acid constructs are implanted around the mouse femoral artery and vein and maintained in vivo for a period of one or two weeks. The vascularized graft is then transferred as a flap towards a full thickness defect made in the abdomen. This technique replaces the need for autologous tissue sacrifications and may enable the use of in vitro engineered vascularized flaps in many surgical applications.

To date, thick tissue defects are typically reconstructed by applying autologous tissue flaps or engineered tissues. In this protocol, we present a new method for engineering vascularized tissue flap bearing an autologous pedicle, to serve as a substitute to autologous flaps.

One of the main factors limiting the thickness of a tissue construct and its consequential viability and applicability in vivo, is the control of oxygen ...

Construction of Defined Human Engineered Cardiac Tissues to Study Mechanisms of Cardiac Cell Therapy

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that replicate the biofidelity of three-dimensional native human myocardium, while also enabling a controlled level of biological complexity, and allowing non-destructive longitudinal monitoring of tissue contractile function. Initially, human engineered cardiac tissues (hECT) were created using the entire cell population obtained from directed differentiation of human pluripotent stem cells, which typically yielded less than 50% cardiomyocytes. However, to create reliable predictive models of human myocardium, and to elucidate mechanisms of heterocellular interaction, it is essential to accurately control the biological composition in engineered tissues. 
To address this limitation, we utilize live cell sorting for the cardiac surface marker SIRP&#945; and the fibroblast marker CD90 to create tissues containing a 3:1 ratio of these cell types, respectively, that are then mixed together and added to a collagen-based matrix solution. Resulting hECTs are, thus, completely defined in both their cellular and extracellular matrix composition.
Here we describe the construction of defined hECTs as a model system to understand mechanisms of cell-cell interactions in cell therapies, using an example of human bone marrow-derived mesenchymal stem cells (hMSC) that are currently being used in human clinical trials. The defined tissue composition is imperative to understand how the hMSCs may be interacting with the endogenous cardiac cell types to enhance tissue function. A bioreactor system is also described that simultaneously cultures six hECTs in parallel, permitting more efficient use of the cells after sorting.

This manuscript describes the creation of defined engineered cardiac tissues using surface marker expression and cell sorting. The defined tissues can then be used in a multi-tissue bioreactor to investigate mechanisms of cardiac cell therapy in order to provide a functional, yet controlled, model system of the human heart.

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that ...

Cardiac Muscle-cell Based Actuator and Self-stabilizing Biorobot - PART 1

Biological machines often referred to as biorobots, are living cell- or tissue-based devices that are powered solely by the contractile activity of living components. Due to their inherent advantages, biorobots are gaining interest as alternatives to traditional fully artificial robots. Various studies have focused on harnessing the power of biological actuators, but only recently studies have quantitatively characterized the performance of biorobots and studied their geometry to enhance functionality and efficiency. Here, we demonstrate the development of a self-stabilizing swimming biorobot that can maintain its pitch, depth, and roll without external intervention. The design and fabrication of the PDMS scaffold for the biological actuator and biorobot followed by the functionalization with fibronectin is described in this first part. In the second part of this two-part article, we detail the incorporation of cardiomyocytes and characterize the biological actuator and biorobot function. Both incorporate a base and tail (cantilever) which produce fin-based propulsion. The tail is constructed with soft lithography techniques using PDMS and laser engraving. After incorporating the tail with the device base, it is functionalized with a cell adhesive protein and seeded confluently with cardiomyocytes. The base of the biological actuator consists of a solid PDMS block with a central glass bead (acts as a weight). The base of the biorobot consists of two composite PDMS materials, Ni-PDMS and microballoon-PDMS (MB-PDMS). The nickel powder (in Ni-PDMS) allows magnetic control of the biorobot during cells seeding and stability during locomotion. Microballoons (in MB-PDMS) decrease the density of MB-PDMS, and enable the biorobot to float and swim steadily. The use of these two materials with different mass densities, enabled precise control over the weight distribution to ensure a positive restoration force at any angle of the biorobot. This technique produces a magnetically controlled self-stabilizing swimming biorobot.

In this two-part study, a biological actuator was developed using highly flexible polydimethylsiloxane (PDMS) cantilevers and living muscle cells (cardiomyocytes), and characterized. The biological actuator was incorporated with a base made of modified PDMS materials to build a self-stabilizing, swimming biorobot.

Biological machines often referred to as biorobots, are living cell- or tissue-based devices that are powered solely by the contractile activity of living ...

Protocol for MicroRNA Transfer into Adult Bone Marrow-derived Hematopoietic Stem Cells to Enable Cell Engineering Combined with Magnetic Targeting

While CD133+ hematopoietic stem cells (SCs) have been proven to provide high potential in the field of regenerative medicine, their low retention rates after injection into injured tissues as well as the observed massive cell death rates lead to very restricted therapeutic effects. To overcome these limitations, we sought to establish a non-viral based protocol for suitable cell engineering prior to their administration. The modification of human CD133+ expressing SCs using microRNA (miR) loaded magnetic polyplexes was addressed with respect to uptake efficiency and safety as well as the targeting potential of the cells. Relying on our protocol, we can achieve high miR uptake rates of 80&#8211;90% while the CD133+ stem cell properties remain unaffected. Moreover, these modified cells offer the option of magnetic targeting. We describe here a safe and highly efficient procedure for the modification of CD133+ SCs. We expect this approach to provide a standard technology for optimization of therapeutic stem cell effects and for monitoring of the administered cell product via magnetic resonance imaging (MRI).

This protocol illustrates a safe and efficient procedure to modify CD133+ hematopoietic stem cells. The presented non-viral, magnetic polyplex-based approach may provide a basis for the optimization of therapeutic stem cell effects as well as for monitoring the administered cell product via magnetic resonance imaging.

While CD133+ hematopoietic stem cells (SCs) have been proven to provide high potential in the field of regenerative medicine, their low retention rates ...

A Net Mold-based Method of Scaffold-free Three-Dimensional Cardiac Tissue Creation

This protocol describes a novel and easy net mold-based method to create three-dimensional (3-D) cardiac tissues without additional scaffold material. Human-induced pluripotent stem-cell-derived cardiomyocytes (iPSC-CMs), human cardiac fibroblasts (HCFs), and human umbilical vein endothelial cells (HUVECs) are isolated and used to generate a cell suspension with 70% iPSC-CMs, 15% HCFs, and 15% HUVECs. They are co-cultured in an ultra-low attachment "hanging drop" system, which contains micropores for condensing hundreds of spheroids at one time. The cells aggregate and spontaneously form beating spheroids after 3 days of co-culture. The spheroids are harvested, seeded into a novel mold cavity, and cultured on a shaker in the incubator. The spheroids become a mature functional tissue approximately 7 days after seeding. The resultant multilayered tissues consist of fused spheroids with satisfactory structural integrity and synchronous beating behavior. This new method has promising potential as a reproducible and cost-effective method to create engineered tissues for the treatment of heart failure in the future.

This protocol describes a net mold-based method to create three-dimensional scaffold-free cardiac tissues with satisfactory structural integrity and synchronous beating behavior.

This protocol describes a novel and easy net mold-based method to create three-dimensional (3-D) cardiac tissues without additional scaffold material. ...

Intra-cardiac Side-Firing Light Catheter for Monitoring Cellular Metabolism using Transmural Absorbance Spectroscopy of Perfused Mammalian Hearts

Absorbance spectroscopy of cardiac muscle provides non-destructive assessment of cytosolic and mitochondrial oxygenation via myoglobin and cytochrome absorbance respectively. In addition, numerous aspects of the mitochondrial metabolic status such as membrane potential and substrate entry can also be estimated. To perform cardiac wall transmission optical spectroscopy, a commercially available side-firing optical fiber catheter is placed in the left ventricle of the isolated perfused heart as a light source. Light passing through the heart wall is collected with an external optical fiber to perform optical spectroscopy of the heart in near real- time. The transmission approach avoids numerous surface scattering interference occurring in widely used reflection approaches. Changes in transmural absorbance spectra were deconvolved using a library of chromophore reference spectra, providing quantitative measures of all the known cardiac chromophores simultaneously. This spectral deconvolution approach eliminated intrinsic errors that may result from using common dual wavelength methods applied to overlapping absorbance spectra, as well as provided a quantitative evaluation of the goodness of fit. A custom program was designed for data acquisition and analysis, which permitted the investigator to monitor the metabolic state of the preparation during the experiment. These relatively simple additions to the standard heart perfusion system provide a unique insight into the metabolic state of the heart wall in addition to conventional measures of contraction, perfusion, and substrate/oxygen extraction.

Here we introduce a method for using an intra-ventricle optical catheter in perfused hearts to perform absorbance spectroscopy across the heart wall. The data obtained provides robust information on tissue oxygen tension as well as substrate utilization and membrane potential simultaneously with cardiac performance measures in this ubiquitous preparation.

Absorbance spectroscopy of cardiac muscle provides non-destructive assessment of cytosolic and mitochondrial oxygenation via myoglobin and cytochrome ...

Preclinical Cardiac Electrophysiology Assessment by Dual Voltage and Calcium Optical Mapping of Human Organotypic Cardiac Slices

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between animal and clinical trials. Numerous animal and cell models have been used to examine the effects of drugs, yet these responses often differ in humans. Human cardiac slices offer an advantage for drug testing in that they are directly derived from viable human hearts. In addition to having preserved multicellular structures, cell-cell coupling, and extracellular matrix environments, human cardiac tissue slices can be used to directly test the effect of innumerable drugs on adult human cardiac physiology. What distinguishes this model from other heart preparations, such as whole hearts or wedges, is that slices can be subjected to longer-term culture. As such, cardiac slices allow for studying the acute as well as chronic effects of drugs. Furthermore, the ability to collect several hundred to a thousand slices from a single heart makes this a high-throughput model to test several drugs at varying concentrations and combinations with other drugs at the same time. Slices can be prepared from any given region of the heart. In this protocol, we describe the preparation of left ventricular slices by isolating tissue cubes from the left ventricular free wall and sectioning them into slices using a high precision vibrating microtome. These slices can then either be subjected to acute experiments to measure baseline cardiac electrophysiological function or cultured for chronic drug studies. This protocol also describes dual optical mapping of cardiac slices for simultaneous recordings of transmembrane potentials and intracellular calcium dynamics to determine the effects of the drugs being investigated.

This protocol describes the procedure for sectioning and culturing human cardiac slices for preclinical drug testing and details the use of optical mapping for recording transmembrane voltage and intracellular calcium signals simultaneously from these slices.

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between ...

Fabrication of Zero Mode Waveguides for High Concentration Single Molecule Microscopy

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to nanomolar ranges, several orders of magnitude less than many physiological ligand concentrations. Optical nanostructures called zero mode waveguides (ZMWs), which are 100&#8722;200 nm in diameter apertures fabricated in a thin conducting metal such as aluminum or gold, allow imaging of individual molecules at micromolar concentrations of fluorophores by confining visible light excitation to zeptoliter effective volumes. However, the need for expensive and specialized nanofabrication equipment has precluded the widespread use of ZMWs. Typically, nanostructures such as ZMWs are obtained by direct writing using electron beam lithography, which is sequential and slow. Here, colloidal, or nanosphere, lithography is used as an alternative strategy to create nanometer-scale masks for waveguide fabrication. This report describes the approach in detail, with practical considerations for each phase. The method allows thousands of aluminum or gold ZMWs to be made in parallel, with final waveguide diameters and depths of 100&#8722;200 nm. Only common lab equipment and a thermal evaporator for metal deposition are required. By making ZMWs more accessible to the biochemical community, this method can facilitate the study of molecular processes at cellular concentrations and rates.

Described here is a nanosphere lithography method for parallel fabrication of zero mode waveguides, which are arrays of nanoapertures in a metal-clad glass microscopy coverslip for single molecule imaging at nano- to micromolar concentrations of fluorophores. The method takes advantage of colloidal crystal self-assembly to create a waveguide template.

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to ...

Fabrication of 3D Cardiac Microtissue Arrays using Human iPSC-Derived Cardiomyocytes, Cardiac Fibroblasts, and Endothelial Cells

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided a unique opportunity to study the complex interplay among different cardiovascular cell types that drives tissue development and disease. In the area of cardiac tissue models, several sophisticated three-dimensional (3D) approaches use induced pluripotent stem cell-derived cardiomyocytes (iPSC-CMs) to mimic physiological relevance and native tissue environment with a combination of extracellular matrices and crosslinkers. However, these systems are complex to fabricate without microfabrication expertise and require several weeks to self-assemble. Most importantly, many of these systems lack vascular cells and cardiac fibroblasts that make up over 60% of the nonmyocytes in the human heart. Here we describe the derivation of all three cardiac cell types from iPSCs to fabricate cardiac microtissues. This facile replica molding technique allows cardiac microtissue culture in standard multi-well cell culture plates for several weeks. The platform allows user-defined control over microtissue sizes based on initial seeding density and requires less than 3 days for self-assembly to achieve observable cardiac microtissue contractions. Furthermore, the cardiac microtissues can be easily digested while maintaining high cell viability for single-cell interrogation with the use of flow cytometry and single-cell RNA sequencing (scRNA-seq). We envision that this in vitro model of cardiac microtissues will help accelerate validation studies in drug discovery and disease modeling.

Here, we describe an easy-to-use methodology to generate 3D self-assembled cardiac microtissue arrays composed of pre-differentiated human-induced pluripotent stem cell-derived cardiomyocytes, cardiac fibroblasts, and endothelial cells. This user-friendly and low cell requiring technique to generate cardiac microtissues can be implemented for disease modeling and early stages of drug development.

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided ...