Introduction
Between 2011 and 2018, the tissue engineering industry reported over $9 billion in sales of tissue engineering related products. Biomaterials-based companies are generating over 99% of this income, with cell-oriented companies accounting for the remaining sales. Perhaps more interesting is that the tissue engineering industry also reported over $35 billion in spending, with approximately $10.9 billion representing investment in research and development.1
Tissue engineering treatments use scaffolds, cells, and biologically active molecules to develop tissues that can replace damaged or diseased parts of the body. Regenerative medicine, a branch of tissue engineering, focuses on the concept of self-healing and allowing the body, with the help of medical therapies, to regenerate damaged tissues and organs.2 Despite the substantial financial investments and research effort in this area, tissue engineering therapies play a relatively small role in patient treatment. Most research activity has been focused on biologic and immunologic factors, which are vital to the potential success of a tissue-engineered therapy. However, failures of tissue engineering therapies may also be related to mechanical factors. For example, complications may arise as a direct consequence of compliance mismatch, which occurs when there is a sudden discontinuity in mechanical properties. This can happen when dissimilar materials are bonded together and can result in premature failure. Problems may also be indirectly related to mechanical factors. For example, failure to maintain a physiologically relevant mechanical environment may result in aberrant mechanical cues adversely affecting cellular processes driven by mechanical factors. While considerable research has been performed on the materials involved in tissue engineering applications, the effects of mechanical factors on integration of engineered tissue grafts requires more investigation. Our research aims to fill this gap in knowledge and optimize the design of tissue engineered scaffolds to improve the development of functional tissues. These tissues can then be used to replace damaged and diseased tissues in clinical settings.
Figure 1. Scaffold degradation process.
THE MECHANICAL ENVIRONMENT IN WHICH [CELLS] GROW AFFECTS HOW THEY DEVELOP AND DIFFERENTIATE |
Cells are mechanotransducers, meaning they respond to external mechanical forces biochemically through gene expression and protein production. Therefore, the mechanical environment in which they grow affects how they develop and differentiate.3 Most healthy, growing cells produce an extracellular matrix (ECM), which provides mechanical support for cells and is the primary structural component of most tissues. The ECM also contains important cues that influence how cells differentiate and grow. There are numerous strategies for producing engineered tissue grafts, though there are common elements in each. In one approach, cells are seeded, or chemically attached, onto a biodegradable synthetic scaffold, which is then implanted in a patient. After implantation, the cells begin their normal activity which includes production of ECM.4 Ultimately, the synthetic scaffold completely degrades, and the ECM remains as a biologic scaffold to support the tissue. Alternatively, the graft can be grown in vitro or outside of the patient. The process is similar with the exception being that the cells used to produce the ECM are removed from the scaffold and replaced with autologous cells, i.e. cells from the patient, prior to implant (Figure 1). Different scaffolds can be constructed by varying parameters such as material, porosity, surface properties, mechanical properties, etc. Because of the cells’ mechanotransductive properties, which dictate their behavior based on their mechanical surroundings, the mechanical design of the scaffold influences how the cells grow and what biochemical signals they produce. To explore this phenomenon on both 2D and 3D structures, we 3D-printed scaffolds, tested their mechanical properties, seeded cells onto them, and analyzed the cellular response to mechanical stimulation. |
MethodsOur study thus far has been divided into four main components: scaffold printing, mechanical analysis, cell culture, and mechanical stimulation. Initially, 3D-printed silicone scaffolds were printed using an envisionTEC®3D-Bioplotter™. Silicone (polydimethyl siloxane, PDMS) was selected for its printability and biocompatibility, meaning that it could safely be used in cell culture. By using a pressure-based extrusion printer and adjusting factors such as printing needle diameter, extrusion pressure, print speed, and strand spacing, the scaffold dimensions could easily be altered, creating a variety of microarchitectures to test. In particular, scaffolds of various structures were printed, including a 90-degree grid pattern, a 60-degree grid pattern, and a wave pattern (Figure 2). We anticipate that scaffolds of varying architectures and dimensions will produce different mechanical environments in which cells would grow, allowing us to better direct the way cells will behave once implanted in the body. Through later testing, we will gain an understanding of which scaffold features induce which cell behaviors, allowing us to better direct cell-to-tissue development. 
Figure 3. Eulerian strain calculated with Ncorr.
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Figure 2. Scaffold print patterns.
The stretch characteristics of each scaffold design were characterized using uniaxial tensile testing (Figure 3). This method utilizes a device that clamps the scaffold on both ends, then stretches the material in one direction at a specified rate. To observe the scaffold responses, we used Ncorr digital image correlation (DIC) MATLAB software, an optical measurement technique for quantifying local displacements and strains from imaging data. Each scaffold was stretched, increasing one percent in length per second, with photos for DIC taken at one-second intervals, until the scaffold had doubled in length. Using this data, we can calculate the localized strains on the scaffold at any point in time. Cells were then cultured onto the 3D scaffolds we had printed, along with 2D PDMS (silicone) membranes. These membranes were used to acquire a baseline understanding of the scaffolds by characterizing simpler, single-layered versions of the 3D membranes. Both the scaffolds and membranes were treated with surface protein to encourage cell adhesion. Scaffolds were treated with either 2%-gelatin, 4%-gelatin, or fibronectin, and membranes were treated with fibronectin only. All surfaces were introduced to 3T3 fibroblasts, a line of mouse connective tissue cells, which were incubated to confluence, when they successfully cover the surface of the structure. The 3D scaffolds were analyzed through phase contrast imaging and the 2D membranes through staining and fluorescence microscopy. Both analyses allowed us to visualize cell morphology, or physical shape, and detect the presence of key features. |
Finally, stretched cell microscopy was performed to examine cell physical behavior in response to a mechanical stimulus. In this process, cell-seeded 2D membranes were loaded into a bioreactor for mechanical testing. This bioreactor contained clamps to hold the membrane sample in place and was filled with cell media that enabled cell survival throughout the mechanical testing process. A stepper motor attached to the bioreactor was used to induce a cyclic load on the membranes that would repeatedly stretch and relax samples. The membranes were stretched for six hours under a sinusoidal stretch to ten percent strain at a frequency of 1 Hz. After a second incubation period, the membranes were fixed and stained through the aforementioned process. Performing fluorescent microscopy on the mechanically stimulated membranes as well allowed us to compare changes between membranes with and without loading in the nuclei and F-actin fibers.
ResultsFibroblasts cultured on both the 2D PDMS membranes and the 3D scaffolds successfully grew to confluence, covering a wide area of each surface and maintaining healthy morphology. Our results pertaining to the 3D scaffolds provide initial insight on the viability of our testing methods and will inform future experiments. The ability to culture fibroblasts onto these surfaces shows the potential for 3D bioprinting to provide a stable environment for cellular development. After cells on the membranes were fixed and stained with 4′,6-diamidino-2-phenylindole (DAPI) to show nuclei5 and rhodamine phalloidin to indicate F-actin protein fibers, observation under a fluorescent microscope revealed the presence of both nuclei and F-actin fibers (Figure 4). |
Figure 4. Fibroblasts on 2D membrane stained to show presence of cell nuclei (blue) and F-actin fibers (red).
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Because these key cell structures appeared, we verified that PDMS membranes are a viable option for future cell culture tests. Additionally, analysis of the cell-seeded 3D scaffolds under phase contrast imaging revealed that the cells adhered best to the untreated scaffolds, independent of any other factors. This surprising result will require further investigation and will help us to better understand the conditions that promote cell growth (Figure 5). Lastly, staining and imaging the membranes subjected to mechanical loading revealed that the F-actin fibers aligned in a uniform direction when stretched (Figure 6). This supported our hypothesis that physically manipulating the scaffolds affects the physicochemical properties of the cells. Further understanding of how different mechanical factors affect cell behavior will allow us to better predict how they may differentiate or what tissues they may form (e.g., muscle, bone, skin). This will aid us in being able to properly characterize the specific effects that the scaffold microarchitectures have on cell development.
With our experimentation, we successfully developed a protocol for testing various scaffold designs and analyzing the response within the cells. This can be used in future trials to validate and verify the accuracy of our data. Through our study, we demonstrated that varying the structures of 2D and 3D scaffolds could provide feasible ways to induce changes in the physicochemical properties of the cells. This knowledge will enable us to create higher-performing engineered tissues in the future. Better engineered tissues allow for a higher degree of specificity in patient treatments, and subsequently a greater likelihood of treatment success.
Figure 5. Enlarged phase contrast image of untreated, Bioplotter-produced 3D structure seeded with fibroblasts.
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Figure 6. Fluorescent images of F-actin with and without loading.
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Future Directions & ConclusionIn the future, we plan to produce more complicated scaffold structures, potentially using soluble support structures and different materials like collagen. Here, we could alter grid angles to explore different microarchitectures, which would affect the surface area to which cells could adhere.6 Additionally, printing new geometric structures would modify the scaffold mechanical properties, providing different mechanical environments in which cells would develop (Figure 7). Different cell types will be seeded onto the various scaffold designs then observed and tested as they differentiate. We will also begin using RT-PCR, reverse transcriptase polymerase chain reaction, to measure the type and quantity of genes that the cells are expressing. By identifying what chemicals the cells are releasing, we will be able to better investigate the effects induced by the mechanical environments on cells. Tissue engineering technologies could potentially address many of the diseases and injuries that are difficult to treat with the limits of current methods, particularly because replacement tissues may offer a better alternative to some current medical solutions. For instance, engineered tissues could be used to enhance hernia repair or skin regeneration techniques. In a greater context, our research could contribute to advancing the medical field to be capable of solving a greater variety of medical problems through highly-personalized solutions. |
Figure 7. Future scaffold pattern.
ENGINEERED TISSUES COULD BE USED TO ENHANCE HERNIA REPAIR OR SKIN REGENERATION TECHNIQUES |
Acknowledgments
This work was supported by the Biomechanical Environments Laboratory at Texas A&M University. We would like to thank our graduate mentor, Raghuveer Lalitha Sridhar, and our research advisors, Dr. Andrew B. Robbins and Dr. Michael R. Moreno, for their unwavering encouragement and advice. Without their constant guidance and support, this project would not have been possible.
References
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DAPI (4’,6-diamidino-2-phenylindole), https://www.thermofisher.com/us/en/home/life-science/cell-analysis/fluorophores/dapi-stain.html.
L. Phillip and S. Ramille, “3D-Printed gelatin scaffolds of differing pore size and geometry modulate hepatocyte function and gene expression,” Frontiers in Bioengineering and Biotechnology 4, (2016), doi:10.3389/conf.fbioe.2016.01.01756.
Erica Michelle Huebner ‘20Erica Michelle Huebner ‘20 is a senior biomedical engineering major with a minor in mathematics from Seabrook, Texas. Erica has been a part of the Biomechanical Environments Laboratory working under Dr. Michael Moreno since summer 2018 and continued her research through the Undergraduate Summer Research Grant Program in 2019. Upon graduation, Erica hopes to attend graduate school and eventually pursue a career researching tissue engineering and stem cell therapies. |
Darby Jane Ballard ‘20Darby Jane Ballard ‘20 is a senior biomedical engineering major from Del Rio, Texas. Ballard has conducted research in the Biomechanical Environments Laboratory at Texas A&M and plans to pursue her interest in autoimmune disease therapies at the graduate level once she receives her undergraduate degree. |