Name: construction of a human aorta smooth muscle cell organ-on-a-chip model for recapitulating biomechanical strain in the aortic wall
Uploaded: 2022-07-06T14:00:00.000+00:00
Duration: 11 min 47 s
Description: Watch this Scientific Journal Video about Construction of a Human Aorta Smooth Muscle Cell Organ-On-A-Chip Model for Recapitulating Biomechanical Strain in the Aortic Wall at JoVE.com

The authors acknowledge that this work was supported by grants from the Science and Technology Commission of Shanghai Municipality (20ZR1411700), the National Natural Science Foundation of China (81771971), and Shanghai Sailing Program (22YF1406600).

Human aortic specimens were utilized for primary HASMC isolation under the approval of Zhongshan Hospital, Fudan University Ethics Committee (NO. B2020-158). Aortic specimens were collected from patients who underwent ascending aorta surgery at Zhongshan Hospital, Fudan University. Written informed consent was obtained from all patients before participation.
1. Primary human aortic smooth muscle cell isolation
<ol>
	<li>Wash the right lateral region of the ascending aorta with sterile PBS, 1x-2x.</li>
	<li>Remove the intima and adventitia layers of the tissue with two ophthalmic forceps and retain the media layer to harvest the cells.</li>
	<li>Place the media layer onto a 10 cm culture dish and cut it into small pieces (2-3 mm) with scissors. Add 5 mL of smooth muscle cell culture medium (SMCM) containing 10% FBS and 1% penicillin-streptomycin.</li>
	<li>Move the small tissue into the culture bottle with a sterile dropper, spreading the tissue evenly. Discard the culture medium as much as possible, then invert the bottle upside down.</li>
	<li>Add 2 mL of SMCM into the inverted culture bottle and place it in the incubator (5% CO2) at 37 &#176;C for 1-2 h, then slowly turn it right side up and add another 2 mL of SMCM.</li>
	<li>After 5-7 days of incubation, exchange the SMCM with 4 mL of fresh SMCM. Generally, sporadic smooth muscle cells climb out in approximately 2 weeks.</li>
	<li>Slowly discard the SMCM and add it slowly when changing the medium. Transport the culture bottle slowly when moving to a microscope station; otherwise, the tissues will float, and the cells will grow slowly.</li>
	<li>When the cells reach approximately 80% confluency, wash with 2 mL of PBS, digest with 2 mL of 0.25% trypsin, and divide them into two new culture bottles with 4 mL of fresh SMCM.</li>
	<li>Identify the cells through an immunofluorescence analysis of four different specific markers for smooth muscle cells (CNN1, SM22)14.</li>
</ol>
2. Preparation of PDMS chip
<ol>
	<li>To polymerize PDMS, add curing agent (B component) to base (A component) at a weight ratio of A: B = 10:1 w/w and mix the complex completely for 5 min; the volume depends on the need of the study.</li>
	<li>Place the prepared PDMS gel in a vacuum extraction tank for 30-60 min and maintain the pressure below -0.8 mPa. 
	NOTE: High pressure will lead to insufficient extraction of small bubbles inside the PDMS gel that will affect the next step of chip fabrication.</li>
	<li>Using computer-aided design (CAD) software, design the mold with a 100 mm &#215; 40 mm &#215; 8&#160;mm external frame structure and a 70 mm &#215; 6 mm &#215; 4 mm channel.</li>
	<li>Custom make the molds of the three layers using a high-precision computer numerical control engraving machine. Carve out the frame of the molds and the microchannels using polymethyl methacrylate (PMMA) plates, and then glue them onto another PMMA plate.</li>
	<li>Pour the prepared PDMS gel onto a mold with a 100 mm &#215; 40 mm &#215; 6 mm outer frame and 70 mm &#215; 6 mm &#215; 4 mm channel, and then cross-link at 70 &#176;C for 1-2 h.</li>
	<li>Peel off the cross-linked PDMS slabs from the mold and cut the commercialized PDMS membranes into 100 mm &#215; 40 mm sections.</li>
	<li>Treat the three prepared PDMS slabs and two PDMS membranes with oxygen plasma for 5 min for PDMS surface activation. Apply the following specific settings: room air as the process gas; pressure set to -100 kPa; current set to 180--200 Ma; voltage set to 200 V; process time to 5 min.</li>
	<li>Punch holes on the three PDMS slabs using a 1 mm hole puncher to make the inlet and outlet of air and medium microchannels on the PDMS chip.</li>
	<li>Bond three PDMS slabs and two PDMS membranes together in the order: air channel PDMS slab (top layer) - PDMS membrane - medium channel PDMS slab (middle layer) - PDMS membrane- air channel PDMS slab (bottom layer). Perform this step under a stereoscopic microscope at 4x to fully overlap the air microchannels with the medium microchannels.</li>
	<li>Place the assembled PDMS chip in a 70 &#176;C incubator for 1 h.</li>
	<li>Prepare several 1 mm inner diameter and 3 cm length latex hoses. Insert a 1 mm outer diameter and 1 cm long stainless-steel needle into one end of the prepared hose, and then insert a Luer into the other end of the hose to create the tube connected to the air and medium microchannels of the PDMS chip.</li>
	<li>Insert the prepared tubes into the outlets and inlets of the air and medium microchannels on the PDMS chip.</li>
</ol>
3. PDMS chip surface treatment and sterilization
<ol>
	<li>Infuse 2 mL of 80 mg/mL mouse collagen into the medium microchannel of the PDMS chip using a 2 mL syringe and incubate at room temperature for 30 min.</li>
	<li>After incubation, remove the collagen from the channel and recollect with a 2 mL syringe. Place the collagen-coated PDMS chips in a 60 &#176;C incubator overnight.</li>
	<li>Place the PDMS chips in a UV sterilizer for more than 1 h. Place the sterilized PDMS chips on an ultraclean bench in preparation for the cell experiment.</li>
</ol>
4. Cell seeding on the PDMS chip and cell stretching process
<ol>
	<li>Culture 4 x 105 primary human smooth muscle cells (HASMCs) from patients using smooth muscle cell medium (SMCM) containing 2% fetal bovine serum (FBS) and 1% penicillin-streptomycin in an 5% CO2 incubator at 37 &#176;C.</li>
	<li>When the HASMC density reaches 80%, discard the SMCM and wash the cells with 2 mL of PBS.</li>
	<li>Digest the cells using 1 mL of 0.25% trypsin for 2 min and centrifuge at 100 x g for 5 min. Remove the supernatant and resuspend the cell pellet in 1 mL of fresh SMCM. Use 8 mL of SMCM to culture the cells on a 10 cm culture dish.</li>
	<li>Calculate the cell number using a cytometer and use the cells at a final concentration of 2 x 105 cells/mL.</li>
	<li>Slowly pour 2 mL of PBS into the collagen-coated and sterilized PDMS chip medium microchannel and later discard using a 2 mL syringe.</li>
	<li>Slowly pour a total of 2 mL of 2 x 105 cells/mL HASMC suspension into the medium microchannel of the PDMS chip using a 2 mL syringe. Then, close the Luer at the entrance and exit of the PDMS chip. Place the PDMS chip in an incubator (5% CO2) at 37 &#176;C for 1 day.</li>
	<li>After the cells are attached to the PDMS membrane in the PDMS chip, nearly after 24 h, connect the outlet of the air microchannel on the PDMS chip to the vacuum controller system. When the cell is attached, an elongated, normal cellular form can be seen under the microscope, contrasting with the suspended round cells.</li>
	<li>Turn on the solenoid valve and the vacuum pump. Open the vacuum regulator and adjust the pressure level to 10 kPa for 7.18 &#177; 0.44% strain and 15 kPa for 17.28 &#177; 0.91% strain.</li>
	<li>After the parameters of the control system are set, place the PDMS chips in an incubator (5% CO2)&#160;at 37 &#176;C and stretch for 24 h.</li>
</ol>
5. Preparation of the mechanical control system
<ol>
	<li>Prepare several solenoid valves, vacuum filters, vacuum regulators, a vacuum pump, a peristaltic pump, and a programmable logic controller (PLC) controlling the solenoid valve.</li>
	<li>Program the PLC controller and set the on/off time interval to 1 Hz. Connect the solenoid valves to the programmed controller.</li>
	<li>Connect the inlet of the vacuum pump to the vacuum filters, and then connect the outlet of the vacuum filters to the vacuum regulators. Connect the outlet of the vacuum regulators to solenoid valves, and finally, connect the outlet of the solenoid valves to the outlets of the air microchannels of the PDMS chips.</li>
	<li>Connect the outlet of the peristaltic pump to the inlet of the medium microchannel of the PDMS chip and the inlet of the peristaltic pump to the outlet of the medium microchannel PDMS chip for culture medium replacement and drug handling.</li>
	<li>Adjust the amplitude of the strain by the regulator and the strain frequency by the microcontroller.</li>
</ol>

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The authors have nothing to disclose.

With the rapid development of microfluidic technology, OOC models that can replicate the biological function and structure of one or more organs in vitro have emerged in recent years for applications in biology, medicine, and pharmacology15. OOC can simulate key functions of the human physiological microenvironment, essential for exploring disease mechanisms and promoting preclinical drug translation8,16. Although OOC is still in the early stages and more input is needed to optimize the design of OOC, much progress has been made in recent years17,18. Unlike other three-dimensional cell culture models, OOC can mimic the biomechanical parameters of the human body that are essential for tissue development and maintenance of organ function. Organs and tissues of the cardiovascular system are constantly subjected to fluid flow-induced shear and cyclic tension in vivo. Therefore, in vitro cellular experiments of the cardiovascular system need to be performed in an experimental platform that provides such biomechanical parameters to obtain realistic experimental results comparable to those obtained from in vivo studies.
In this study, we established a HASMC-OOC that can precisely control the parameters of the biomechanical strain to which HASMCs are subjected. It is possible to control the fluid velocity and fluid shear to which the cells are subjected, as well as to precisely control the amplitude, frequency, and rhythm of different patterns of cyclic strain. Of the many materials that have been used in the field of OOC, PDMS is one of the most widely adopted19,20,21. PDMS is transparent, flexible, biocompatible, and breathable, and these properties contribute to its wide usage in the field of microfluidic chips19. Applied pressure changes can deform PDMS to achieve the mechanical force of contraction and relaxation, and the permeability of PDMS ensures that the cell culture environment on the chip can be consistent with the incubation conditions of 5% CO2 at 37 &#176;C in the incubator. Therefore, PDMS not only provides controlled mechanical force to the cells but also provides a culture environment suitable for normal cell growth. Our experimental results show that the morphology and activity of cells cultured in the PDMS chip are consistent with 2D culture conditions, indicating that PDMS is biocompatible.
Under biomimetic in vivo physiological conditions, HASMCs in the aortic wall are subjected to the cyclic strain of approximately 9%22. In pathological conditions, such as hypertension, aortic dilatation, and aortic aneurysm, HASMCs are subjected to cyclic strain greater than 16%23. We characterized the tensile properties of the PDMS chip, and the experimental results show that a vacuum pressure of 10 kPa induced 7.18 &#177; 0.44% strain and 15 kPa induced 17.28 &#177; 0.91% strain. In addition, the frequency of the strain and fluid shear stress can be controlled by this PDMS chip system. Using BAV-TAAD and TAV-TAAD patient-derived primary HASMCs, the molecular mechanisms and drug screening of these aortic diseases can be conducted using this HASMC-OOC in vitro model, which can replicate the disease pathogenesis of different aortic diseases. Using this system, we have established a disease model to study the pathogenesis of BAV-TAAD and revealed that mitochondrial fusion activator could partially rescue the mitochondrial dysfunction in diseased cells24.
The HASMC-OOC was designed based on the inspiration and reference of lung-on-a-chip from Ingber&#39;s work9. Their device replicated physiological breathing movements of the alveoli in the lung, and the chip consisted of three layers: upper and lower medium channels, a porous PDMS membrane, and two side vacuum chambers. Side vacuum chambers can only be made in specialized laboratories with advanced microfluidic equipment. To enable assembly in more general laboratory conditions, we placed the vacuum chambers on the top and bottom of the chip. Both stretching approaches have similar functions, so the chip structure can be chosen according to the laboratory conditions and the research needs so thatselected. In addition, to structurally establish the tubular structure of the human aorta and obtain more biological samples from the cells for conducting further complex biological analysis and experimental assays, we established a five-layered PDMS chip with larger channels, which consisted of a middle medium channel, two PDMS membranes, and top and bottom vacuum chambers. Two PDMS membranes and the large channels of the PDMS chip increased the cell culture area up to 8.4 cm2, providing enough samples for protein analysis. The previously reported platforms mainly focused on investigating minor artery diseases, such as a cerebral artery, peripheral vessel, or other arterial diseases25,26. The aorta is a large vessel with a diameter of 25-35 mm, and the tunica media of the aorta is mainly composed of smooth muscle cells subject to high tensile strain. The pathology of aortopathy is associated with remodeling of the aortic wall, smooth muscle cell apoptosis, and phenotypic switching, all of which can be regulated by mechanical stress. Thus, patient-derived HASMCs and tensile strain are essential components for the success of HASMC-OOC. However, there are some limitations to this study. HASMC-OOC did not utilize cocultured vascular cells, including HASMCs, aortic endothelial cells, and fibroblasts, which might better simulate the in vivo microenvironment of the human aorta and can be used to study the interaction between these cells. Although HASMCs experience rhythmic strain, the cells are still cultured on a flat PDMS membrane that fails to mimic the extracellular matrix. Ia n future studies, we will overcome these limitations by combining microfluidic chips with 3D bioprinting technology.

Thoracic aortic aneurysm and dissection (TAAD) is a localized dilatation or delamination of the aortic wall that is associated with high morbidity and mortality1. Human aortic smooth muscle cells (HASMCs) play a vital role in the pathogenesis of TAAD. HASMCs are not terminally differentiated cells, and HASMCs retain high plasticity, allowing them to switch phenotypes in response to different stimuli2. HASMCs are mainly subjected to rhythmic tensile strain in vivo, and this is one of the key factors regulating smooth muscle morphological changes, differentiation and physiological functions3,4. Therefore, the role of cyclic strain in the study of HASMCs cannot be ignored. However, conventional 2D cell cultures cannot replicate the biomechanical stimulation of cyclic strain experienced by HASMCs in vivo. Additionally, the construction of an animal TAAD model is not suitable for some types of TAAD, such as bicuspid aortic valve (BAV)-related TAAD. Moreover, the species gap between clinical human studies and animal experiments cannot be ignored. It hinders pharmaceutical translation in clinical practice. Thus, there is an urgent need for more complex and physiological systems to simulate the in vivo biomechanical environment in the research of aortic diseases.
Animal experiments used in biomedical research and drug development are costly, time-consuming, and ethically questionable. In addition, the results from animal studies frequently fail to predict the results obtained in human clinical trials5,6. The lack of human preclinical models and high failure rate in clinical trials have resulted in few effective drugs for the clinic, which has increased health care costs7. Thus, it is urgent to find other experimental models to complement animal models. Microfabrication and microfluidic techniques have developed greatly in recent years, providing novel techniques for establishing organoids-on-a-chip models that remedy the drawbacks of traditional 2D cell culture techniques and establish a more realistic, low-cost, and efficient in vitro model for physiological studies and drug development. Using microfluidic devices, organs-on-chips are established to culture living cells in micrometer-sized chambers with different stimuli to replicate the key functions of a tissue or organ. The system consists of single or multiple microfluidic microchannels, with either one kind of cell cultured in a perfused chamber replicating functions of one tissue type or different cell types cultured on porous membranes to recreate interfaces between different tissues. Microfluidic-based organoids combined with patient-derived cells have the unique advantage of bridging the large species difference between mouse and human disease models and overcoming the disadvantages of traditional 2D cell culture for disease mechanism research and drug discovery. With the rapid development of microfluidics in the past few years, researchers have realized the usefulness of in vitro organ-on-a-chip (OOC) models replicating complex in vivo biological parameters8. These microfluidic organoids simulate in vitro biomechanical environments, such as cyclic strain, shear stress, and liquid pressure, providing a three-dimensional (3D) cell culture environment. To date, several OOC models have been established to simulate biomechanical stimuli in organs such as the lung9, kidney10, liver11, intestine12, and heart13, but these have not been widely applied to the study of human aortic disease.
In this study, we present a human aorta smooth muscle cell organ-on-a-chip (HASMC-OOC) model that can control the biomimetic mechanical forces and rhythms applied to TAAD patient-derived primary HASMCs. The chip consists of three-layered thick plates of polydimethylsiloxane (PDMS) etched with channels and two commercialized highly flexible PDMS membranes. HASMCs are cultured on the PDMS membranes. The channel in the middle of the chip is filled with a culture medium for cell culture. The top and bottom channels of the chip are connected to a vacuum pressure supply system that can control the rhythm and frequency of mechanical tensile strain of the PDMS membranes. Rhythmic strain experienced by HASMCs can be simulated in HASMC-OOC, replicating the biomechanical microenvironment of tissue or organ not functionally achievable with conventional 2D culture systems. With the advantage of high-resolution, real-time imaging, and biomechanical microenvironment, the biochemical, genetic and metabolic activities of living cells can be studied for tissue development, organ physiology, disease etiology, molecular mechanisms and biomarker identification ,cardiovascular disease and aortic disease. Combined with tissue-specific and patient cells, this system can be used for drug screening, personalized medicine, and toxicity testing. This HASMC-OOC model provides a novel in vitro platform for studying the pathogenesis of the aortic disease.

<table><tbody><tr><td>4% paraformaldehyde</td><td>Beyotime</td><td>P0099-100ml</td><td>Used for cell immobilization</td></tr><tr><td>Alexa Fluor 350-labeled Goat Anti-Rabbit IgG</td><td>Beyotime</td><td>A0408</td><td>Antibodies used for immunostaining</td></tr><tr><td>Bovine serum albumin</td><td>Beyotime</td><td>ST025-20g</td><td></td></tr><tr><td>Calcium AM/PI</td><td>Invitrogen</td><td>L3224</td><td></td></tr><tr><td>Cell culture flask&amp;nbsp;</td><td>Corning</td><td>430639</td><td></td></tr><tr><td>CNN1</td><td>Abcam&amp;nbsp;</td><td>Ab46794</td><td></td></tr><tr><td>Commercial flexible&lt;br/&gt; PDMS membrane</td><td>Hangzhou Bald Advanced Materials</td><td>KYQ-200</td><td></td></tr><tr><td>F-actin</td><td>Invitrogen</td><td>R415</td><td></td></tr><tr><td>FBS</td><td>Sigma</td><td>M8318</td><td></td></tr><tr><td>Hoses</td><td>Runze Fluid</td><td>96410</td><td>1 mm inner diameter; 3 mm outer diameter; 1 mm wall thickness; Official website address: https://www.runzefluidsystem.com</td></tr><tr><td>Human aortic smooth&lt;br/&gt; muscle cell line CRL1999</td><td>ATCC</td><td>Lot Number:70019189</td><td></td></tr><tr><td>Image J</td><td>Imagej.net/fiji/downloads</td><td>Free Download: https://fiji.sc</td><td>Imaging platform that is used to identify fluorescence intensity</td></tr><tr><td>Incubator</td><td>Thermo Fisher Scientific</td><td></td><td>Ensures that the temperature,&lt;br/&gt; humidity, and light exposure is&lt;br/&gt; exactly the same throughout&lt;br/&gt; experiment.</td></tr><tr><td>Luer</td><td>Runze Fluid</td><td>RH-M016</td><td>Official website address: https://www.runzefluidsystem.com.</td></tr><tr><td>Microscope</td><td>Olympus</td><td></td><td></td></tr><tr><td>mouse collagen</td><td>Sigma</td><td>C7661</td><td></td></tr><tr><td>Oxygen plasma&amp;nbsp;</td><td>Changzhou Hongming Instrument</td><td>HM-Plasma5L</td><td></td></tr><tr><td>Pasteur pipette</td><td>Biologix</td><td>30-0138A1</td><td></td></tr><tr><td>PBS</td><td>Beyotime</td><td>C0221A</td><td></td></tr><tr><td>Pen-Strep</td><td>Sigma</td><td>P4458-100ml</td><td>Antibiodics used to prevent bacterial&lt;br/&gt; contamination of cells during culture.</td></tr><tr><td>peristaltic pump</td><td>Kamoer</td><td>F01A-STP-B046</td><td></td></tr><tr><td>Petri dish</td><td>Corning</td><td>430167</td><td></td></tr><tr><td>PLC controller</td><td>Zhejiang Jun Teng (BenT) CNC factory</td><td>BR010-11T8X2M</td><td>The detailed program setting can be found in supplementary. Official website address: files.http://www.btcnc.net</td></tr><tr><td>polydimethylsiloxane (PDMS)</td><td>Dow Corning</td><td>Sylgard 184</td><td></td></tr><tr><td>SM22</td><td>Abcam&amp;nbsp;</td><td>ab14106</td><td></td></tr><tr><td>SMCM</td><td>ScienCell</td><td>Cat 1101</td><td></td></tr><tr><td>solenoid valve</td><td>SMC (China)</td><td>VQZ300</td><td></td></tr><tr><td>Syringe</td><td>Becton,Dickinson and Company</td><td>300841</td><td></td></tr><tr><td>Triton-X 100</td><td>Beyotime</td><td>ST795</td><td>To penetrate cell membranes</td></tr><tr><td>Trizol</td><td>Invitrogen</td><td>10296010</td><td>Used for RNA extraction</td></tr><tr><td>trypsin</td><td>Sigma</td><td>15400054</td><td></td></tr><tr><td>vacuum filter</td><td>SMC (China)</td><td>ZFC5-6</td><td>Official website address: https://www.smc.com.cn</td></tr><tr><td>vacuum pump</td><td>Kamoer</td><td>KVP15-KL-S</td><td></td></tr><tr><td>vacuum regulator</td><td>AirTAC</td><td>GVR-200-06</td><td></td></tr><tr><td>&lt;strong&gt;Primers&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Primer Name</td><td>Forward (5&amp;rsquo; to 3&amp;rsquo;)</td><td>Reverse (5&amp;rsquo; to 3&amp;rsquo;)</td><td></td></tr><tr><td>SM22</td><td>CCGTGGAGATCCCAACTGG</td><td>CCATCTGAAGGCCAATGACAT</td><td></td></tr><tr><td>CNN1</td><td>CTCCATTGACTCGAACGACTC</td><td>CAGGTCTGCGAAACTTCTTAGA</td><td></td></tr></tbody></table>

construction of a human aorta smooth muscle cell organ-on-a-chip model for recapitulating biomechanical strain in the aortic wall

Conventional two-dimensional cell culture techniques and animal models have been used in the study of human thoracic aortic aneurysm and dissection (TAAD). However, human TAAD sometimes cannot be characterized by animal models. There is an apparent species gap between clinical human studies and animal experiments that may hinder the discovery of therapeutic drugs. In contrast, the conventional cell culture model is unable to simulate in vivo biomechanical stimuli. To this end, microfabrication and microfluidic techniques have developed greatly in recent years, providing novel techniques for establishing organoids-on-a-chip models that replicate the biomechanical microenvironment. In this study, a human aorta smooth muscle cell organ-on-a-chip (HASMC-OOC) model was developed to simulate the pathophysiological parameters of aortic biomechanics, including the amplitude and frequency of cyclic strain experienced by human aortic smooth muscle cells (HASMCs) that play a vital role in TAAD. In this model, the morphology of HASMCs became elongated in shape, aligned perpendicularly to the strain direction, and presented a more contractile phenotype under strain conditions than under static conventional conditions. This was consistent with the cell orientation and phenotype in native human aortic walls. Additionally, using bicuspid aortic valve-related TAAD (BAV-TAAD) and tricuspid aortic valve-related TAAD (TAV-TAAD) patient-derived primary HASMCs, we established BAV-TAAD and TAV-TAAD disease models, which replicate HASMC characteristics in TAAD. The HASMC-OOC model provides a novel in vitro platform that is complementary to animal models for further exploring the pathogenesis of TAAD and discovering therapeutic targets.

The HASMC-OOC model consists of a vacuum control system, a circulating system, and PDMS chips, and the schematic design of the HASMC-OOC model (Figure 1). The vacuum control system consists of a vacuum pump, solenoid valves, and a PLC controller. To act as the circulating system, a peristaltic pump was used to refresh the cell culture medium and add drugs. The PDMS chip was composed of two vacuum chambers and a middle chamber filled with SMCM for cell growth. According to the design of the chip structure, the PMMA mold was prepared, and the three PDMS slabs were prepared by pouring into molds and cross-linking at 70 &#176;C for 2 h. After treatment with plasma, the three PDMS slabs and two PDMS membranes were bonded together (Figure 2). When the PDMS chips were prepared, the HASMCs were cultured on the PDMS membrane in the chip, and the PDMS chips were connected to the vacuum control system and the circulating system. A schematic of the entire system is shown in Supplementary Figure 1. The PDMS chip parameters are shown in Figure 3A, and the mechanical properties of the PDMS membrane are shown in Figure 3B. To quantify the tensile strains of the PDMS membrane, we captured the real-time deformations of the PDMS membranes from a cross-sectional view of the microfluidic model, with vacuum pressures of 0 kPa, 10 kPa, 15 kPa, 20 kPa, 25 kPa, and 30 kPa. We also measured the changes in length to evaluate the strain magnitude of the PDMS membrane in different culturing channel widths of 2 mm, 4 mm, and 6 mm (Figure 3C-E). Microchannels 6 mm in width were used in the PDMS chip. The results showed that a vacuum pressure of 10 kPa induced 7.18 &#177; 0.44% strain and 15 kPa induced 17.28 &#177; 0.91% strain (Figure 3E).
To verify the cell viability of the HASMC cell line (CRL1999) in the PDMS chip, a LIVE/DEAD assay was performed on days 3 and 5 after the cells were cultured. The results showed that the cell viability was higher than 90% at day 3 and day 5 (Figure 4A). Cytoskeleton (F-actin) staining of CRL1999 in the PDMS chip showed normal cell morphology and cells aligned perpendicularly to the strain after stretching for 24 h (Figure 4B-C). Immunofluorescence staining of the contractile markers SM22 and CNN1 was performed using HASMC-OOC after 24 h of rhythmic strain (Figure 4D-E). To start, 2 mL of PBS was slowly poured into the channel and discarded with a 2 mL syringe. Subsequently, 2 mL of 4% (v/v) paraformaldehyde was poured into the channel and incubated for 30 min at room temperature. After this, 2 mL of 0.2% (v/v) Triton X-100 was poured into the channel by a 2 mL syringe and incubated for 15 min at room temperature. This was followed by 2 mL of PBS being slowly poured into the channel and discarded by a 2 mL syringe after washing the cells. Next, 2 mL of 5% (w/v) bovine serum albumin was slowly poured into the channel and incubated for 30 min at room temperature. Afterward, 1 mL of SM22 or CNN1 antibody was slowly poured into the channel and incubated overnight at 4 &#176;C. After incubation, 1 mL of goat anti-rabbit secondary antibody was slowly poured into the channel and incubated for 1 h at room temperature. Finally, 1 mL of DAPI solution was slowly poured into the channel and incubated for 10 min at room temperature. The results showed that the fluorescence intensity of SM22 and CNN1 under strain conditions was higher than that under static conditions (Figure 4F). Compared with static conditions, the SM22 and CNN1 genes in HASMCs were upregulated in strain conditions (Figure 4G). These data suggested that CRL1999 aligned perpendicularly to the strain direction and that a contractile phenotype was more pronounced under strain conditions than under conventional cell culturing under static conditions.
Using bicuspid aortic valve-related TAAD (BAV-TAAD) and tricuspid aortic valve-related TAAD (TAV-TAAD) patient-derived primary HASMCs, we established BAV-TAAD and TAV-TAAD disease models. The results of F-actin staining of BAV-TAAD and TAV-TAAD patient-derived primary HASMCs showed normal cell morphology and cells aligned perpendicularly to the strain direction after stretching for 24 h (Figure 5A-B). Immunofluorescence staining showed that SM22 and CNN1 expression in BAV-TAAD and TAV-TAAD patient-derived primary HASMCs was higher under strain conditions than under static conditions (Figure 5C-G,I). The gene expression levels of SM22 and CNN1 in BAV-TAAD and TAV-TAAD patient-derived primary HASMCs were upregulated under strain conditions in contrast to static conditions (Figure 5H,J). These results in BAV-TAAD and TAV-TAAD patient-derived primary HASMCs in the PDMS chip were consistent with the results from HASMC cell line CRL1999, indicating that the combination of patient-derived primary HASMCs and the HASMC-OOC model replicate HASMC characteristics in TAAD, which provides a BAV-TAAD and TAV-TAAD in vitro disease model for further investigation of disease&#39;s molecular mechanisms and drug screening.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64122/64122fig01.jpg" /> 
Figure 1: Schematic design of the HASMC-OOC system. The system consists of a vacuum control system, a circulating system, and PDMS chips. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64122/64122fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64122/64122fig02.jpg" /> 
Figure 2: PDMS chip preparation procedure. The PDMS gel was poured into a PMMA mold and cross-linked at 70 &#176;C for 1-2 h. Then, the three PDMS slabs and two PDMS membranes were treated with plasma for 5 min and bonded together. Finally, the prepared PDMS chips were sterilized, and a concentration of 2 x 105 cells/mL HASMC suspension was slowly poured into the medium microchannel of the PDMS chip. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64122/64122fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64122/64122fig03.jpg" /> 
Figure 3: The detailed parameters and mechanical properties of the PDMS chip. (A) The detailed parameters of the PDMS chip. The PDMS slab size was 100 mm &#215; 40 mm &#215; 6 mm, and the microchannel size was 70 mm &#215; 6 mm &#215; 4 mm. (B) The parameters of the commercial PDMS membrane. The calculated stretching amplitude of the PDMS membrane at different vacuum pressures for microchannel widths of 2 mm (C), 4 mm (D), and 6 mm (E). The data shows mean &#177; standard deviation (SD). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64122/64122fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64122/64122fig04.jpg" /> 
Figure 4: Cell morphology, orientation, and phenotype alteration of HASMCs under cyclic strain conditions. (A) Representative image of the LIVE/DEAD assay at day 3 and day 5 after HASMC cell line (CRL1999) culturing on a PDMS chip. (B) F-actin staining of CRL1999 under static and strain conditions. (C) The cell orientation of CRL1999 under static and strain conditions. Representative image of the immunofluorescence staining of SM22 (D) and CNN1 (E) in CRL1999 under static and strain conditions. (F) The relative intensity of immunofluorescence staining of SM22 and CNN1. (G) SM22 and CNN1 mRNA expression in CRL1999 under static and strain conditions. n = 3, *p &#60; 0.05, **p &#60; 0.01, ***p &#60; 0.001, ****p &#60; 0.0001, two-tailed Student&#39;s t-tests were used to compare the two groups. The data shows mean &#177; SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64122/64122fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64122/64122fig05.jpg" /> 
Figure 5: Cell morphology and phenotypic alterations in BAV-TAAD and TAV-TAAD patient-derived HASMC-OOC. Representative image of F-actin staining of BAV-TAAD (A)-derived primary HASMCs and TAV-TAAD (B)-derived primary HASMCs under static and strain conditions. SM22 immunofluorescence staining in BAV-TAAD (C)-derived primary HASMCs and TAV-TAAD (D)-derived primary HASMCs under static and strain conditions. Representative image of the immunofluorescence staining of CNN1 in BAV-TAAD (E)-derived primary HASMCs and TAV-TAAD (F)-derived primary HASMCs under static and strain conditions. (G) The relative intensity of SM22 and CNN1 immunofluorescence staining in BAV-TAAD-derived primary HASMCs. (H) SM22 and CNN1 mRNA expression in BAV-TAAD-derived primary HASMCs under static and strain conditions. (I) The relative intensity of SM22 and CNN1 immunofluorescence staining in TAV-TAAD-derived primary HASMCs. (J) SM22 and CNN1 mRNA expression in TAV-TAAD-derived primary HASMCs under static and strain conditions. n = 3, *p &#60; 0.05, **p &#60; 0.01, ***p &#60; 0.001, ****p &#60; 0.0001, two-tailed Student&#39;s t-tests were used to compare the two groups. The data shows mean &#177; SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64122/64122fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Schematic instruction of equipment preparation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64122/supplementary figure 1.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Construction of a Human Aorta Smooth Muscle Cell Organ-On-A-Chip Model for Recapitulating Biomechanical Strain in the Aortic Wall at JoVE.com

Construction of a Human Aorta Smooth Muscle Cell Organ-On-A-Chip Model for Recapitulating Biomechanical Strain in the Aortic Wall

Here, we developed a human aorta smooth muscle cell organ-on-a-chip model to replicate the in vivo biomechanical strain of smooth muscle cells in the human aortic wall.

Conventional two-dimensional cell culture techniques and animal models have been used in the study of human thoracic aortic aneurysm and dissection ...

construction-human-aorta-smooth-muscle-cell-organ-on-chip-model-for

Nanyang Technological University

Research

JoVE Journal

Bioengineering

3.0K Views.  Fudan University. Dit model stimuleert biomechanische prikkels die worden ervaren door gladde spiercellen in menselijke aorta. In combinatie met van patiënten afgeleide cellen kan dit systeem worden gebruikt voor geneesmiddelenscreening en gepersonaliseerde geneeskunde van aortaziekten. Dit model kan de kale mechanische parameters van de gladde spiercellen in menselijke aorta nabootsen en nauwkeurig regelen, wat een nieuw in vitro platform biedt voor het bestuderen en pathogenese van aortaziekten.Miera...

Video: Construction of a Human Aorta Smooth Muscle Cell Organ-On-A-Chip Model for Recapitulating Biomechanical Strain in the Aortic Wall

Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do not have suitable autologous donor vessels due to disease progression or previous harvest. The aim of vascular tissue engineering is to develop a suitable alternative source for these bypass grafts. In addition, engineered vascular tissue may prove valuable as living vascular models to study cardiovascular diseases. Several promising approaches to engineering blood vessels have been explored, with many recent studies focusing on development and analysis of cell-based methods.2-5 Herein, we present a method to rapidly self-assemble cells into 3D tissue rings that can be used in vitro to model vascular tissues.
To do this, suspensions of smooth muscle cells are seeded into round-bottomed annular agarose wells. The non-adhesive properties of the agarose allow the cells to settle, aggregate and contract around a post at the center of the well to form a cohesive tissue ring.6,7 These rings can be cultured for several days prior to harvesting for mechanical, physiological, biochemical, or histological analysis. We have shown that these cell-derived tissue rings yield at 100-500 kPa ultimate tensile strength8 which exceeds the value reported for other tissue engineered vascular constructs cultured for similar durations (&lt;30 kPa).9,10 Our results demonstrate that robust cell-derived vascular tissue ring generation can be achieved within a short time period, and offers the opportunity for direct and quantitative assessment of the contributions of cells and cell-derived matrix (CDM) to vascular tissue structure and function.

This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do ...

Engineering Fibrin-based Tissue Constructs from Myofibroblasts and Application of Constraints and Strain to Induce Cell and Collagen Reorganization

Collagen content and organization in developing collagenous tissues can be influenced by local tissue strains and tissue constraint. Tissue engineers aim to use these principles to create tissues with predefined collagen architectures. A full understanding of the exact underlying processes of collagen remodeling to control the final tissue architecture, however, is lacking. In particular, little is known about the (re)orientation of collagen fibers in response to changes in tissue mechanical loading conditions. We developed an in&#160;vitro model system, consisting of biaxially-constrained myofibroblast-seeded fibrin constructs, to further elucidate collagen (re)orientation in response to i) reverting biaxial to uniaxial static loading conditions and ii) cyclic uniaxial loading of the biaxially-constrained constructs before and after a change in loading direction, with use of the Flexcell FX4000T loading device. Time-lapse confocal imaging is used to visualize collagen (re)orientation in a nondestructive manner.
Cell and collagen organization in the constructs can be visualized in real-time, and an internal reference system allows us to relocate cells and collagen structures for time-lapse analysis. Various aspects of the model system can be adjusted, like cell source or use of healthy and diseased cells. Additives can be used to further elucidate mechanisms underlying collagen remodeling, by for example adding MMPs or blocking integrins. Shape and size of the construct can be easily adapted to specific needs, resulting in a highly tunable model system to study cell and collagen (re)organization.

This model system starts from a myofibroblast-populated fibrin gel that can be used to study endogenous collagen (re)organization real-time in a nondestructive manner. The model system is very tunable, as it can be used with different cell sources, medium additives, and can be adapted easily to specific needs.

Collagen content and organization in developing collagenous tissues can be influenced by local tissue strains and tissue constraint. Tissue engineers aim ...

A Novel Stretching Platform for Applications in Cell and Tissue Mechanobiology

Tools that allow the application of mechanical forces to cells and tissues or that can quantify the mechanical properties of biological tissues have contributed dramatically to the understanding of basic mechanobiology. These techniques have been extensively used to demonstrate how the onset and progression of various diseases are heavily influenced by mechanical cues. This article presents a multi-functional biaxial stretching (BAXS) platform that can either mechanically stimulate single cells or quantify the mechanical stiffness of tissues. The BAXS platform consists of four voice coil motors that can be controlled independently. Single cells can be cultured on a flexible substrate that can be attached to the motors allowing one to expose the cells to complex, dynamic, and spatially varying strain fields. Conversely, by incorporating a force load cell, one can also quantify the mechanical properties of primary tissues as they are exposed to deformation cycles. In both cases, a proper set of clamps must be designed and mounted to the BAXS platform motors in order to firmly hold the flexible substrate or the tissue of interest. The BAXS platform can be mounted on an inverted microscope to perform simultaneous transmitted light and/or fluorescence imaging to examine the structural or biochemical response of the sample during stretching experiments. This article provides experimental details of the design and usage of the BAXS platform and presents results for single cell and whole tissue studies. The BAXS platform was used to measure the deformation of nuclei in single mouse myoblast cells in response to substrate strain and to measure the stiffness of isolated mouse aortas. The BAXS platform is a versatile tool that can be combined with various optical microscopies in order to provide novel mechanobiological insights at the sub-cellular, cellular and whole tissue levels.

We present in this article a novel stretching platform that can be used to investigate single cell responses to complex anisotropic biaxial mechanical deformation and quantify the mechanical properties of biological tissues.

Tools that allow the application of mechanical forces to cells and tissues or that can quantify the mechanical properties of biological tissues have ...

Microfluidic Model to Mimic Initial Event of Neovascularization

Neovascularization is usually initialized from an existing normal vasculature and the biomechanical microenvironment of endothelial cells (ECs) in the initial stage varies dramatically from the following process of neovascularization. Although there are plenty of models to simulate different stages of neovascularization, an in vitro 3D model that capitulates the initial process of neovascularization under the corresponding stimulations of normal vasculature microenvironments is still lacking. Here, we reconstructed an in vitro 3D model that mimics the initial event of neovascularization (MIEN). The MIEN model contains a microfluidic sprouting chip and an automatic control, highly efficient circulation system. A functional, perfusable microchannel coated with endothelium was formed and the process of sprouting was simulated in the microfluidic sprouting chip. The initially physiological microenvironment of neovascularization was recapitulated with the microfluidic control system, by which ECs would be exposed to high luminal shear stress, physiological transendothelial flow, and various vascular endothelial growth factor (VEGF) distributions simultaneously. The MIEN model can be readily applied to the study of neovascularization mechanism and holds a potential promise as a low-cost platform for drug screening and toxicology applications.

Here, we provide a microfluidic chip and an automatically controlled, highly efficient circulation microfluidic system that recapitulates the initial microenvironment of neovascularization, allowing endothelial cells (ECs) to be stimulated by high luminal shear stress, physiological level of transendothelial flow, and various vascular endothelial growth factor (VEGF) distribution simultaneously.

Neovascularization is usually initialized from an existing normal vasculature and the biomechanical microenvironment of endothelial cells (ECs) in the ...

Developing 3D Organized Human Cardiac Tissue within a Microfluidic Platform

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart muscle, the myocardium, is notoriously difficult to accomplish in vitro. Mainly, obstacles lie in the need for human cardiomyocytes (CMs) that are either adult or exhibit adult-like phenotypes and can successfully replicate the myocardium&#39;s cellular complexity and intricate 3D architecture. Unfortunately, due to ethical concerns and lack of available primary patient-derived human cardiac tissue, combined with the minimal proliferation of CMs, the sourcing of viable human CMs has been a limiting step for cardiac tissue engineering. To this end, most research has transitioned toward cardiac differentiation of human induced pluripotent stem cells (hiPSCs) as the primary source of human CMs, resulting in the wide incorporation of hiPSC-CMs within in vitro assays for cardiac tissue modeling.
Here in this work, we demonstrate a protocol for developing a 3D mature stem cell-derived human cardiac tissue within a microfluidic device. We specifically explain and visually demonstrate the production of a 3D in vitro anisotropic cardiac tissue-on-a-chip model from hiPSC-derived CMs. We primarily describe a purification protocol to select for CMs, the co-culture of cells with a defined ratio via mixing CMs with human CFs (hCFs), and suspension of this co-culture within the collagen-based hydrogel. We further demonstrate the injection of the cell-laden hydrogel within our well-defined microfluidic device, embedded with staggered elliptical microposts that serve as surface topography to induce a high degree of alignment of the surrounding cells and the hydrogel matrix, mimicking the architecture of the native myocardium. We envision that the proposed 3D anisotropic cardiac tissue-on-chip model is suitable for fundamental biology studies, disease modeling, and, through its use as a screening tool, pharmaceutical testing.

The goal of this protocol is to explain and demonstrate the development of a three-dimensional (3D) microfluidic model of highly aligned human cardiac tissue, composed of stem cell-derived cardiomyocytes co-cultured with cardiac fibroblasts (CFs) within a biomimetic, collagen-based hydrogel, for applications in cardiac tissue engineering, drug screening, and disease modeling.

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart ...

Fibroblast Derived Human Engineered Connective Tissue for Screening Applications

Fibroblasts are phenotypically highly dynamic cells, which quickly transdifferentiate into myofibroblasts in response to biochemical and biomechanical stimuli. The current understanding of fibrotic processes, including cardiac fibrosis, remains poor, which hampers the development of new anti-fibrotic therapies. Controllable and reliable human model systems are crucial for a better understanding of fibrosis pathology. This is a highly reproducible and scalable protocol to generate engineered connective tissues (ECT) in a 48-well casting plate to facilitate studies of fibroblasts and the pathophysiology of fibrotic tissue in a 3-dimensional (3D) environment. ECT are generated around the poles with tunable stiffness, allowing for studies under a defined biomechanical load. Under the defined loading conditions, phenotypic adaptations controlled by cell-matrix interactions can be studied. Parallel testing is feasible in the 48-well format with the opportunity for the time-course analysis of multiple parameters, such as tissue compaction and contraction against the load. From these parameters, biomechanical properties such as tissue stiffness and elasticity can be studied.

Presented here is a protocol to generate engineered connective tissues for a parallel culture of 48 tissues in a multi-well plate with double poles, suitable for mechanistic studies, disease modeling, and screening applications. The protocol is compatible with fibroblasts from different organs and species and is exemplified here with human primary cardiac fibroblasts.

Fibroblasts are phenotypically highly dynamic cells, which quickly transdifferentiate into myofibroblasts in response to biochemical and biomechanical ...

Reconstituting Cytoarchitecture and Function of Human Epithelial Tissues on an Open-Top Organ-Chip

Nearly all human organs are lined with epithelial tissues, comprising one or multiple layers of tightly connected cells organized into three-dimensional (3D) structures. One of the main functions of epithelia is the formation of barriers that protect the underlining tissues against physical and chemical insults and infectious agents. In addition, epithelia mediate the transport of nutrients, hormones, and other signaling molecules, often creating biochemical gradients that guide cell positioning and compartmentalization within the organ. Owing to their central role in determining organ-structure and function, epithelia are important therapeutic targets for many human diseases that are not always captured by animal models. Besides the obvious species-to-species differences, conducting research studies on barrier function and transport properties of epithelia in animals is further compounded by the difficulty of accessing these tissues in a living system. While two-dimensional (2D) human cell cultures are useful for answering basic scientific questions, they often yield poor in vivo predictions. To overcome these limitations, in the last decade, a plethora of micro-engineered biomimetic platforms, known as organs-on-a-chip, have emerged as a promising alternative to traditional in vitro and animal testing. Here, we describe an Open-Top Organ-Chip (or Open-Top Chip), a platform designed for modeling organ-specific epithelial tissues, including skin, lungs, and the intestines. This chip offers new opportunities for reconstituting the multicellular architecture and function of epithelial tissues, including the capability to recreate a 3D stromal component by incorporating tissue-specific fibroblasts and endothelial cells within a mechanically active system. This Open-Top Chip provides an unprecedented tool for studying epithelial/mesenchymal and vascular interactions at multiple scales of resolution, from single cells to multi-layer tissue constructs, thus allowing molecular dissection of the intercellular crosstalk of epithelialized organs in health and disease.

The present protocol describes the capabilities and the essential culture modalities of the Open-Top Organ-Chip for the successful establishment and maturation of full-thickness organ-on-chip cultures of primary tissues (skin, alveolus, airway, and intestine), providing the opportunity to investigate different functional aspects of the human epithelial/mesenchymal and vascular niche interface in vitro.

Nearly all human organs are lined with epithelial tissues, comprising one or multiple layers of tightly connected cells organized into three-dimensional ...

Designing a Bioreactor to Improve Data Acquisition and Model Throughput of Engineered Cardiac Tissues

Heart failure remains the leading cause of death worldwide, creating a pressing need for better preclinical models of the human heart. Tissue engineering is crucial for basic science cardiac research; in vitro human cell culture eliminates the interspecies differences of animal models, while a more tissue-like 3D environment (e.g., with extracellular matrix and heterocellular coupling) simulates in vivo conditions to a greater extent than traditional two-dimensional culture on plastic Petri dishes. However, each model system requires specialized equipment, for example, custom-designed bioreactors and functional assessment devices. Additionally, these protocols are often complicated, labor-intensive, and plagued by the failure of the small, delicate tissues.
This paper describes a process for generating a robust human engineered cardiac tissue (hECT) model system using induced pluripotent stem-cell-derived cardiomyocytes for the longitudinal measurement of tissue function. Six hECTs with linear strip geometry are cultured in parallel, with each hECT&#160;suspended from a pair of force-sensing polydimethylsiloxane (PDMS) posts attached to PDMS racks. Each post is capped with a black PDMS stable post tracker (SPoT), a new feature that improves the ease of use, throughput, tissue retention, and data quality. The shape allows for the reliable optical tracking of post deflections, yielding improved twitch force tracings with absolute active and passive tension. The cap geometry eliminates tissue failure due to hECTs slipping off the posts, and as they involve a second step after PDMS rack fabrication, the SPoTs can be added to existing PDMS post-based designs without major changes to the bioreactor fabrication process.
The system is used to demonstrate the importance of measuring hECT function at physiological temperatures and shows stable tissue function during data acquisition. In summary, we describe a state-of-the-art model system that reproduces key physiological conditions to advance the biofidelity, efficiency, and rigor of engineered cardiac tissues for in vitro applications.

Three-dimensional cardiac tissues bioengineered using stem-cell-derived cardiomyocytes have emerged as promising models for studying healthy and diseased human myocardium in vitro while recapitulating key aspects of the native cardiac niche. This manuscript describes a protocol for fabricating and analyzing high-content engineered cardiac tissues generated from human induced pluripotent stem-cell-derived cardiomyocytes.

Heart failure remains the leading cause of death worldwide, creating a pressing need for better preclinical models of the human heart. Tissue engineering ...

Murine Aortic Crush Injury: An Efficient In Vivo Model of Smooth Muscle Cell Proliferation and Endothelial Function

Arterial reconstruction, whether angioplasty or bypass surgery, involves iatrogenic trauma causing endothelial disruption and vascular smooth muscle cell (VSMC) proliferation. Common murine models study small vessels such as the carotid and femoral arteries. Herein we describe an in vivo system in which both VSMC proliferation and endothelial barrier function can be simultaneously assessed in a large vessel. We studied the infrarenal aortic response to injury in C57BL/6 mice. The aorta was injured from the left renal vein to the aortic bifurcation by 30 transmural crushes of 5-seconds duration with a cotton-tipped applicator. Morphological changes were assessed with conventional histology. Aorta wall thickness was measured from the luminal surface to the adventitia. EdU integration and counter staining with DAPI and alpha-actin was used to demonstrate VSMC proliferation. Activation of ERK1/2, a known moderator of intimal hyperplasia formation, was determined by Western Blot analysis. The effect of inflammation was determined by immunohistochemistry for B-cells, T-cells, and macrophages. En face sections of endothelium were visualized with scanning electron microscopy (SEM). Endothelial barrier function was determined with Evans Blue staining. Transmural injury resulted in aortic wall thickening. This injury induced VSMC proliferation, most prominently at 3 days after injury, and early activation of ERK1/2 and decreased p27kip1 expression. Injury did not result in increased B-cells, T-cells, or macrophages infiltration in the vessel wall. Injury caused partial endothelial cell denudation and loss of cell-cell contact. Injury resulted in a significant loss of endothelial barrier function, which returned to baseline after seven days. The murine transmural blunt aortic injury model provides an efficient system to simultaneously study both VSMC proliferation and endothelial barrier function in a large vessel.

Murine Aortic Crush Injury: An Efficient In Vivo Model of Smooth Muscle Cell Proliferation and Endothelial Function

Restenosis following cardiovascular procedures (bypass surgery, angioplasty, or stenting) is a significant problem reducing the durability of these procedures. An ideal therapy would inhibit smooth muscle cell (VSMC) proliferation while promoting regeneration of the endothelium. We describe a model for simultaneous assessment of VSMC proliferation and endothelial function in vivo.

Arterial reconstruction, whether angioplasty or bypass surgery, involves iatrogenic trauma causing endothelial disruption and vascular smooth muscle cell ...

Biology

Isolation of Primary Patient-specific Aortic Smooth Muscle Cells and Semiquantitative Real-time Contraction Measurements In Vitro

Smooth muscle cells (SMCs) are the predominant cell type in the aortic media. Their contractile machinery is important for the transmission of force in the aorta and regulates vasoconstriction and vasodilation. Mutations in genes encoding for the SMC contractile apparatus proteins are associated with aortic diseases, such as thoracic aortic aneurysms. Measuring SMC contraction in vitro is challenging, especially in a high-throughput manner, which is essential for screening patient material. Currently available methods are not suitable for this purpose. This paper presents a novel method based on electric cell-substrate impedance sensing (ECIS). First, an explant protocol is described to isolate patient-specific human primary SMCs from aortic biopsies and patient-specific human primary dermal fibroblasts for the study of aortic aneurysms. Next, a detailed description of a new contraction method is given to measure the contractile response of these cells, including the subsequent analysis and suggestion for comparing different groups. This method can be used to study the contraction of adherent cells in the context of translational (cardiovascular) studies and patient and drug screening studies.

Isolation of Primary Patient-specific Aortic Smooth Muscle Cells and Semiquantitative Real-time Contraction Measurements In Vitro

This paper describes an explant culture-based method for the isolation and culturing of primary, patient-specific human aortic smooth muscle cells and dermal fibroblasts. Furthermore, a novel method is presented for measuring cell contraction and subsequent analysis, which can be used to study patient-specific differences in these cells.

Smooth muscle cells (SMCs) are the predominant cell type in the aortic media. Their contractile machinery is important for the transmission of force in ...

Construction of a Human Aorta Smooth Muscle Cell Organ-On-A-Chip Model for Recapitulating Biomechanical Strain in the Aortic Wall

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