The authors would like to thank Dr. Paul Barbone from the Boston University Department of Mechanical Engineering for helpful discussions and assistance with data analysis. This study was supported by NSF grant CMMI-1910401.

The authors have no conflicts of interest to disclose.

An improved method of indirectly patterning PAA hydrogels is described in this paper. This approach builds on methods that have been used previousely20,35,36,37,38,39,40,41,42. The primary change is that PDMS stamps are now used to remove protein and leave the desired pattern behind on the intermediate substrate rather than directly stamping the pattern down onto it. This allows for much more consistent creation of high-fidelity micropatterns and the creation of isolated micropatterned islands that previously necessitated two production steps. The shape and size of the island patterns made with this method are also more easily controlled than those made with the previous two-step method. The new technique is less susceptible to the amount of pressure applied during microprinting than the old technique. A second advantage is that this removal method can make island patterns of controlled shape and size in only one step. In contrast, previous methods required two steps to make islands, including stamping for deposition and subsequent stamping for removal, and the shapes of these islands are less precise than those made with the removal method.
The main disadvantage of this method is that the lifetime of the masters used to mold the new style of PDMS stamps seems to be shorter than those used in the previous stamping method. This can likely be attributed to the shape of the new masters used in the removal method. The old masters were composed of a 1.5 x 1.5 cm area of SU-8 composed of equally spaced 5 &#181;m-deep circular holes, which when cast in PDMS, would create stamps made up of evenly spaced cylindrical posts of the same height. Conversely, the new masters are made up of a 1.5 x 1.5 cm area of 5 &#181;m-tall SU-8 cylindrical posts, which, when cast in PDMS, make stamps that are made up of evenly spaced holes.
With this change in the structure of the masters, it has been found that the SU-8 tends to become delaminated from the surface much more easily than in the old method. Precautions were taken to prevent this, such as surface-treating the silicon wafers in a plasma asher to make them more amenable to binding to SU-8. The surface of the wafers was also silanized before the first casting in PDMS to prevent the SU-8 from sticking to the PDMS upon removal of the stamps. Despite this, SU-8 delamination from the silicon wafers has been observed after repeated casting in PDMS, and caution should be taken to ensure that new silicon wafers be made prior to loss of the current master. One possible way to avoid this delamination is to use the PDMS double-cast method, which has been described previously43, though this other method has its limitations.
Another shortcoming of this method (and other methods that use deformable hydrogels to measure traction forces44) is that the traction field is not in mechanical equilibrium due to experimental noise. Thus, a postprocessing analysis is needed to obtain an equilibrated traction field after traction forces are calculated. One way to balance traction forces is to obtain the forces closest to the measurements that satisfy equilibrium using a least-squares method. As a result, the magnitude and orientation of measured traction forces become altered45.
There are limitations to this method of calculating cellular traction forces. To accurately determine cellular traction forces using Eq. (1) (step 5.4), the pattern must be designed such that the displacement of one adhesion dot does not significantly affect the displacement of those directly adjacent to it. Theoretically, the displacement of a circular adhesion region on the surface of an infinite half-space due to a tangential force acting at the center decreases as a radial distance from the center of the circle increases23. Specifically, for a substrate whose Poisson&#39;s ratio is ~0.445 (such as those described here), displacement at the edge of the circular region is approximately two-thirds the displacement at the center of the region. However, theoretical predictions do not extend beyond the circular region. Thus, it is assumed that the decreasing trend in displacement magnitude, determined theoretically23, continues beyond the edge of the adhesion circle. In the micropattern design described here, 2 &#181;m circular dots spaced 6 &#181;m center-to-center are used. The reason for this spacing is that a displacement at the 6 &#181;m distance from the center of a dot is estimated to be approximately 1/12th the displacement at the center of the dot, which is assumed to be small enough to affect displacement values of adjacent dots. However, it is possible that the traction forces exerted by cells could displace the adhesion dots on a substrate so much so that the center-to-center distance between them becomes less than 2 &#181;m. In this case, the assumption about the displacement of adjacent adhesion dots not interfering with one another does not hold, and traction forces cannot be accurately calculated with the equation given in step 6.4 (Eq (1)).
The proposed technique provides a powerful tool for measuring cellular traction forces. These forces give insight into the mechanical environment of both individual cells and clusters and can help understand if and how different types of cells maintain mechanical stability. Maintaining a homeostatic level of cell tension is essential for many cellular processes, and loss of this tensional homeostasis has been linked to various diseases, such as atherosclerosis, asthma, and cancer9,10,12. Tensional homeostasis is defined as the ability of a cell or a cluster of cells to maintain a consistent level of tension, with a low temporal variability around a set point46.
This indirect micropatterning method can be used to determine the ability of various cell types to maintain tensional homeostasis, both at the individual and multicellular levels. This is done by tracking changes in the values of the cellular traction field over time and then quantifying temporal fluctuation of the traction field using the coefficient of variation (CV), which represents that ratio of the standard deviation of the magnitude of the traction field to its mean value. The sum of the magnitudes of traction forces and the magnitude of the contractile moment (the first moment of the traction forces) are used as scalar metrics of the magnitude of the traction field46. If the CV of the traction field remains close to zero throughout a timelapse experiment, it shows that the cell/cluster maintained tensional homeostasis over time46.
In summary, this new method for indirect micropatterning of soft hydrogels offers a simpler and more efficient method of creating patterned hydrogels than previous methods. There are certain steps that can be taken to improve and expand upon this method. Primarily, improving the fabrication process of the silicon masters in a way that extends their lifetime would eliminate the primary disadvantage of this micropatterning method. As for ways to expand upon this method, exploring different island shapes beyond just the square islands described here would improve the versatility of this method.

Most cell phenotypes exert traction forces on their environment. These traction forces are generated by a cell&#39;s contractile cytoskeleton, which is a network of actin and myosin, and other filamentous biopolymers and crosslinking proteins1,2,3,4. Forces generated within the cell can be transmitted to the extracellular environment or adjacent cells, primarily via transmembrane proteins such as integrins and cadherins, respectively5,6. How a cell spreads or contracts-and the magnitudes of the traction forces associated with those movements-is the result of an intimate conversation with its environment, which largely depends on the type and quantity of protein present in the extracellular matrix (ECM)7,8 and the stiffness of the ECM. Indeed, traction force microscopy has become an invaluable tool for understanding cell responsiveness to local stimuli such as substrate stiffness, imposed mechanical stresses and strains, or contact with other cells. This information is directly relevant to the understanding of diseases such as cancer and asthma9,10,11,12.
A system that can be used to measure force-induced deformation of a substrate of known material properties is required to calculate traction forces. These changes must be tracked over time, requiring both imaging and image processing techniques. One of the first methods used to determine cellular traction forces was the observation and analysis of the contraction of collagen hydrogels seeded with cells, though this method was only semiquantitative13. Another, more refined method was to measure the traction forces exerted by single cells by determining the forces resulting from the deformation of a thin sheet of silicone14. Later on, more quantitative measurement techniques were developed, and these methods also allowed for the use of soft hydrogels such as polyacrylamide (PAA)12,15,16. When using these soft materials, traction forces could be determined from the force-induced displacement of randomly displaced beads embedded in the hydrogel and the mechanical properties of the gel16,17. Another advancement came with the development of micropost arrays made of soft polydimethylsiloxane (PDMS) so that their deflection could be measured and converted to force using the beam theory18.
Finally, methods for micropatterning soft hydrogels were developed as these approaches allow control of the contact areas for cell adhesion. By measuring the deformation of the micropattern within a cell&#39;s contact area, traction forces could easily be calculated because a force-free reference image is not required19. This method has been widely adopted as it allows for the indirect patterning of a regular array of micron-sized, discrete fluorescent protein adhesion points onto PAA gels for the measurement of cellular traction forces20. To calculate these forces, an image-processing algorithm, which can track the movements of each micropatterned dot without requiring user input, has been developed21.
While this method is simple for creating entire grids of dot patterns, it is more complicated when patterns of isolated patches (or islands) of dots are desired. Micropatterned islands are useful when control of shape, and to some extent of size, of clusters of cells is needed. To create these islands, the aforementioned method of microcontact printing necessitates two distinct steps: i) using one PDMS stamp to create a high-fidelity pattern of dots on a coverslip, and then ii) using a second different PDMS stamp to remove most of those dots, leaving behind isolated islands of dots21. The difficulty in creating islands with this original method is compounded by the fact that making consistent grid patterns in the first step of the process is challenging on its own. Microprinting stamps are composed of an array of circular microposts, the diameter of which corresponds to the desired dot size. These stamps are then coated with an even layer of protein and then stamped with a precise amount of pressure onto treated coverslips to create the desired pattern. On the one hand, applying too much pressure to the stamp can result in uneven protein transfer and poor pattern fidelity due to pillar buckling or sagging between pillars, leading to contact with the glass. On the other hand, applying too little pressure results in little to no protein transfer and poor pattern fidelity. For these reasons, a transfer process that can be used to consistently create high-quality micropatterns of isolated islands of dots in just one step is desired.
Herein, a method is described for the indirect micropatterning of islands of micron-sized fluorescent protein adhesion points onto a PAA gel that is more consistent and versatile than previously developed methods. Whereas older indirect micropatterning methods rely on the transfer of protein patterns from a PDMS stamp to an intermediate substrate, the method introduced here uses PDMS stamps instead as a vessel for protein removal, not addition. This is done by first fundamentally changing the structure of the PDMS stamps used. Rather than making stamps that are composed of a pattern of evenly spaced circular pillars, stamps are made up of a pattern of evenly spaced circular holes in this method.
With this new structure, the surface of these PDMS stamps can then be treated with glutaraldehyde as described previously20,29,30, making the stamp able to bond covalently with protein. When used on a glass coverslip evenly coated with fluorescent protein, these glutaraldehyde-treated PDMS stamps are used to remove most of the protein on the surface of the coverslip, leaving behind only the desired pattern of dots predetermined by the location of micron-sized holes on the stamp. This change increases the success rate for generating patterns made up of a near-continuous grid of dots and for creating isolated islands of dots through only one step.

<table><tbody><tr><td>(3-aminopropyl)trimethoxysilane</td><td>Sigma Aldrich</td><td>#281778</td><td></td></tr><tr><td>1.5 mL Microcentrifuge tube</td><td>Fisher Scientific</td><td>#05-408-129</td><td></td></tr><tr><td>15 mL conical tube</td><td>Fisher Scientific</td><td>#05-539-12</td><td></td></tr><tr><td>4 x 4 in 0.060 Quartz LR Chrome Photomask</td><td>Advance Reproductions Corporation</td><td>N/A</td><td>Custom-designed mask</td></tr><tr><td>6 Well Plates</td><td>Fisher Scientific</td><td>#07-200-83</td><td></td></tr><tr><td>Acetone</td><td>Fisher Scientific</td><td>#A18P-4</td><td></td></tr><tr><td>Acrylamide Solution, 40%</td><td>Sigma Aldrich</td><td>#A4058</td><td></td></tr><tr><td>AlexaFluor 488</td><td>Thermo Fisher</td><td>#A20000</td><td></td></tr><tr><td>Aminonium Persulfate</td><td>Fisher Scientific</td><td>#BP179-25</td><td></td></tr><tr><td>Bisacrylamide</td><td>Fisher Scientific</td><td>#PR-V3141</td><td></td></tr><tr><td>Ethanol</td><td>Greenfield Global</td><td>#111000200C1GL</td><td></td></tr><tr><td>Glass Coverslips, 25 mm round</td><td>Fisher Scientifc</td><td>#12-545-102</td><td></td></tr><tr><td>Glass Coverslips, 30 mm round</td><td>Warner</td><td>#64-1499</td><td></td></tr><tr><td>Hamamatsu ORCA-R2 Camera</td><td>Hamamatsu</td><td>#C10600-10B</td><td></td></tr><tr><td>Human Plasma</td><td>Valley Biomedical</td><td>#HP1051P</td><td>Used to isolate fibronectin</td></tr><tr><td>Hydrochloric Acid, 1.0 N</td><td>Millipore Sigma</td><td>#1.09057</td><td></td></tr><tr><td>ImageJ</td><td>Wayne Rasband</td><td>#1.53n</td><td></td></tr><tr><td>Interchangeable Coverslip Dish Set</td><td>Bioptechs</td><td>#190310-35</td><td></td></tr><tr><td>Kim Wipes</td><td>Fisher Scientific</td><td>#06-666-11C</td><td></td></tr><tr><td>Mask Alinger</td><td>Karl Suss</td><td>#MA6</td><td></td></tr><tr><td>Matlab 2021</td><td>Mathworks</td><td>#R2021a</td><td></td></tr><tr><td>MetaMorph Basic</td><td>Molecular Devices</td><td>#v7.7.1.0</td><td></td></tr><tr><td>N-hydroxysuccinimide ester</td><td>Sigma Aldrich</td><td>#130672-5G</td><td></td></tr><tr><td>NucBlue Live Cell Stain</td><td>Thermo Fisher</td><td>#R37605</td><td></td></tr><tr><td>Olympus IX2-ZDC Inverted Microscope</td><td>Olympus</td><td>#IX81</td><td></td></tr><tr><td>PD-10 Desalting Columns</td><td>GE Healthcare</td><td>#52-1308-00</td><td></td></tr><tr><td>Photoresist Spinner Hood</td><td>Headway Research</td><td>#PWM32</td><td></td></tr><tr><td>Plasma Cleaner</td><td>Harrick</td><td>#PDC-001</td><td></td></tr><tr><td>Plasma Etcher</td><td>TePla</td><td>#M4L</td><td></td></tr><tr><td>Prior Lumen 200Pro Light Source</td><td>Prior Scientific</td><td>#L200</td><td></td></tr><tr><td>Silicon Wafers, 100 mm</td><td>University Wafer</td><td>#809</td><td></td></tr><tr><td>SU-8 2005</td><td>Kayaku Advanced Materials Inc.</td><td>#NC9463827</td><td></td></tr><tr><td>SU-8 Developer</td><td>Kayaku Advanced Materials Inc.</td><td>#NC9901158</td><td></td></tr><tr><td>Sylgard 184 Silicone Elastomer</td><td>Essex Brownell</td><td>#DC-184-1.1</td><td></td></tr><tr><td>Tetramethylethylenediamine</td><td>Fisher Scientific</td><td>#BP150-20</td><td></td></tr><tr><td>Trichloro(1H,1H,2H,2H-perfluorooctyl)silane</td><td>Sigma Aldrich</td><td>#448931</td><td></td></tr><tr><td>UAPON-40XW340 Objective</td><td>Olympus</td><td>#N2709300</td><td></td></tr><tr><td>UV Flood Exposure</td><td>Newport</td><td>#69910</td><td></td></tr><tr><td>Wafer Carrier Tray, 110 x 11 mm</td><td>Ted Pella, Inc.</td><td>#19395-40</td><td></td></tr></tbody></table>

pattern generation for micropattern traction microscopy

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

PAA hydrogels with the Young&#39;s modulus of E = 3.6 kPa and the Poisson&#39;s ratio of &#957; = 0.445 were made for use by this subtractive micropatterning method. The hydrogels were made to be ~100 &#181;m thick, which allows them to be imaged with the imaging setup used here while also preventing the cells from sensing the rigid coverslip below the gel, which would cause problems in studies focused on cellular rigidity sensing23,33. Gels of many other stiffness levels (up to 30 kPa) have been successfully made and imaged using the indirect micropatterning method34, so the method is not limited to the use of only 3.6 kPa hydrogels. The coverslips are treated ahead of time with glutaraldehyde to allow the gel to remain fixed to the lower coverslip when the top patterned coverslip is removed (Figure 2C,D).
To visualize and measure cellular traction forces within clusters, the 3.6 kPa hydrogels were indirectly patterned with fluorescent fibronectin (Figure 2A-D) to create island patterns of predetermined size and shape (Figure 2E). Other types of proteins can also be patterned onto these soft hydrogels21,35,36,37,38. The quality of the transferred pattern is related directly to the fidelity of the master mold from which the PDMS stamp is cast. Molds that have had SU-8 delamination will see a deterioration in the quality of the patterns made from stamps cast from these delaminated molds. For example, molds with delaminated SU-8 will often create micropatterns containing sections of unpatterned fibronectin between the predetermined islands. If cast on a gel, these patterns permit cell attachment to the gel in areas outside the island micropattern. Because of this, imaging cell clusters attached to fully or mostly intact island patterns was the priority.
Bovine vascular smooth muscle cells (BVSMCs) were seeded onto these hydrogels at a density of approximately 60-80 &#215; 103 cells/gel to promote cluster formation and allowed to adhere for 18-24 h prior to imaging. Just prior to the experiments, cells were stained with a live-cell nucleus stain to allow living cells to be identified and determine the number of cells in each cluster. Micropattern islands with and without cells were imaged and analyzed. Imaging islands without cells allowed for the calculation of the noise present in traction force calculations for 3.6 kPa gels. These false-positive displacement measurements can then be used to determine an appropriate threshold value below which displacements should be excluded.
It has been found that 0.3 &#181;m is usually sufficient to eliminate pattern infidelity that leads to false-positive displacement measurements. Doing so may cause the loss of low-force tractions but is essential for removing false-positive tractions. When imaging, cell clusters on mostly intact island patterns were prioritized (i.e., those not missing many if any dots) and mostly intact patterns without cells. Each ROI was imaged once every 5 min for 2 h. The microscope used to capture images of both cells and the patterned PAA gels during extended timelapse experiments was a fluorescence microscope with an automatic stage, a fluorescence light source, a camera, and standard microscope software. This microscope has a custom set of filters to observe many different colors of fluorescence. The objective used to view the cells and the patterned hydrogel is a 40x water immersion objective, NA = 1.15. This microscope is also equipped with a customized temperature (37 &#176;C)-, humidity (70%)-, and CO2 (5%)-regulated system to keep cells viable during long experiments.
To calculate the traction forces from the images of the micropattern islands, image analysis software was used to track the displacements of the fluorescent fibronectin dots over time. Knowing the displacements of the fluorescent dots and the stiffness of the PAA gel onto which the dots are patterned, the program can determine the values of the traction forces imparted by both individual cells and clusters on the PAA substrate. However, there are two ways by which the program becomes unable to determine traction values. If too many dots within a given frame of interest are deformed, the program struggles to calculate forces, as the program relies on most of the dots in an analyzed image to be undeformed. Moreover, if two dots are too close to one another (center-to-center distance of &#8804;2 &#181;m20), the displacements of the individual dots could interfere with each other, which prevents accurate determination of their actual displacement values and thus their traction values. As long as the micropatterns are designed with dots that are well spaced apart, cells should not be able to displace the dots so much that they become that close together, even on softer substrates.
Figure 3&#160;shows the capabilities of the removal patterning method. Here, the ability of this method to fabricate isolated, well-defined island patterns of predetermined shape is shown in Figure 3B-E, where the fibronectin adhesion dots are present only within the desired area of the island. These isolated islands of adhesion dots further allow for better control of cluster shape, as shown in Figure 3A. Because the shape and size of the islands are consistent, the cell clusters that attach and grow onto them will also tend to have consistent shape, as the island patterns limit the cluster growth area. Finally, Figure 3B-E also show the ability of these islands to be used to calculate cellular traction forces and the tendency of these traction forces to change over time. Here, the traction forces of the cluster are the largest around the edges of the island. This is expected because the cells at the edges of a cluster have fewer interactions with other cells in the cluster than those in the interior of the cluster. Thus, these cells at the edges exert greater forces on the substrate in response to cytoskeletal contraction than the cells in the interior.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63628/63628fig01.jpg" /> 
Figure 1: Design of photomask. (A) A representation of the first half of the photomask design used here, a discrete grid of 2 &#181;m diameter dots spaced at 6 &#181;m center-to-center. While the image shown here is just a small portion of the design, this grid fills a space of 1.5 x 1.5 cm total on the photomask. (B) A representation of the second half of the photomask design; shown here are six small islands of 6 x 6 dots. On the photomask, there are multiple different island sizes: 6 x 6 (shown here), 12 x 12, 25 x 25, and 42 x 42 dots. An equally spaced (50 &#181;m between islands) array of each island size takes up a space of 1.5 x 0.375 cm, and the arrays of different islands are separated by 50 &#181;m. The dots in each island are 2 &#181;m in diameter and spaced 6 &#181;m center-to-center. The two sections of the mask described in A and B are separated by 0.75 cm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/63628fig01large.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/63628/63628fig02.jpg" /> 
Figure 2: Subtractive microcontact printing. (A) A PDMS stamp is treated with glutaraldehyde and put in contact with a glass coverslip evenly coated with fluorescent fibronectin solution. (B)&#160;Upon removal from the coverslip, the PDMS stamp strips away most of the fluorescent fibronectin on the surface of the coverslip, leaving micron-sized dots of protein only in locations predetermined by the design of the PDMS stamp. (C) The patterned coverslip is placed in contact with a PAA prepolymer and NHS solution. (D) Once the PAA gel has been allowed to fully polymerize, the top coverslip is removed, and a pattern of fibronectin is printed onto the PAA gel surface. (E)&#160;An example of a discrete island pattern made up of evenly spaced dots on a PAA hydrogel with a Young&#39;s modulus of 3.6 kPa. Scale bar = 20 &#181;m. Abbreviations: PDMS = polydimethylsiloxane; PAA = polyacrylamide; NHS = N-hydroxysuccinimide. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/63628fig02large.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/63628/63628fig03.jpg" /> 
Figure 3: Analysis of cells on island micropatterns. (A) A brightfield image of a cluster of 3 BVSMCs overlaid onto a fluorescent fibronectin island micropattern on a PAA hydrogel with a Young&#39;s modulus of 3.6 kPa is shown. Dots are 2 &#181;m in diameter and are separated by 6 &#181;m center-to-center. (B)&#160;The fluorescent pattern shows that a number of the fibronectin points in the island have been displaced due to the application of forces by the BVSMCs. (C) The traction forces applied on the adhesion points at time point 1 (start of a 2 h experiment). Direction of these force vectors is indicated by direction of the colored arrows. Their magnitude is indicated by the arrow color and its corresponding value on the color bar (all force values are in nN). Vector length is relative based on the minimum and maximum forces calculated by the program for each time point. (D) Traction forces of the same cluster at time point 13 (halfway through a 2 h experiment) (E) Traction forces of the same cluster at time point 25 (end of a 2 h experiment). Abbreviations = BVSMCs = bovine vascular smooth muscle cells; PAA = polyacrylamide. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/63628fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental File 1: Calculator for labeling fibronectin This is a tool for calculating the correct amount of Alexa488 fluorescent dye to use when labeling fibronectin. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/JoVE_Fn_Alexa488_Calculator.xlsx" target="_blank">Please click here to download this File.</a>
Supplemental File 2: CTFTimelapse.m This is the image processing file, which allows for the calculation of cellular traction forces as described in Section 5 (Image Analysis). This includes selecting a desired grid of dots (Steps 6.3-6.3.2) and choosing the region of interest for the CTF calculations (Steps 6.5 and 6.5.1). All of the following supplemental files are called directly by this script or one of the other functions used within it. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/CTF_Timelapse.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 3: analyze_initial_image_4.m This function is called by CTFTimelapse.m, and its purpose is to locate and align the fluorescent dots on a grid from the first frame of the image stack of the deformed grid pattern to match the original undeformed grid pattern. This allows for traction force calculations for the first frame in the image stack. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/analyze_initial_image_4.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 4: analyze_subsequent_images.m This function is called by CTFTimelapse.m, and its purpose is to locate and align the fluorescent dots on a grid from all frames of the image stack of the deformed grid pattern after the first. This allows for traction force calculations for all frames in the image stack after the first. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/analyze_subsequent_images.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 5: bpass.m This function is called by both analyze_initial_image_4.m and analyze subsequent_images.m, and its purpose is to implement a real-space bandpass filter that processes the image stack of the deformed grid pattern. This filter suppresses pixel noise and long-wavelength image variations while retaining information of a characteristic size. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/bpass.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 6: CellBoundary.m This function is called by CTFTimelapse.m, and its purpose is that it allows the program user to draw a region of interest around the cell/cluster for which traction forces are to be calculated. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/CellBoundary.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 7: pkfnd.m This function is called by both analyze_initial_image_4.m and analyze_subsequent_images.m, and its purpose is to find local maxima in an image with pixel-level accuracy. These peaks are used by cntrd.m to locate the fluorescent grid pattern for CTFTimelapse.m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/pkfnd.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 8: cntrd.m This function is called by both analyze_initial_image_4.m and analyze_subsequent_images.m, and its purpose is to locate the centroid of bright spots in an image to sub-pixel accuracy. This allows for location of the fluorescent grid pattern by CTFTimelaspe.m, as described in Step 6.3. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/cntrd.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 9: FourCorners.m This function is called by both analyze_initial_image_4.m and analyze_subsequent_images.m, and its purpose is to take the grid chosen by the user (Steps 6.3-6.3.2) and calculate the distance between each corner dot in two orthogonal axes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/FourCorners.zip" target="_blank">Please click here to download this File.</a>
Supplemental File 10: track.m This function is called by both analyze_initial_image_4.m and analyze_subsequent_images.m, and its purpose is to track the movement of the dots in the fluorescent grid pattern between frames, which is essential for calculating the corresponding traction forces. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63628/track.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Pattern Generation for Micropattern Traction Microscopy at JoVE.com

Pattern Generation for Micropattern Traction Microscopy

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

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

pattern-generation-for-micropattern-traction-microscopy

Nanyang Technological University

Research

JoVE Journal

Bioengineering

2.1K Views.  Boston University. We describe improvements to a standard method for measuring cellular traction forces, based on microcontact printing with a single subtractive patterning step of dot arrays of extracellular matrix proteins on soft hydrogels. This method allows for simpler and more consistent fabrication of island patterns, essential for controlling cell cluster shape.

Video: Pattern Generation for Micropattern Traction Microscopy

Micropunching Lithography for Generating Micro- and Submicron-patterns on Polymer Substrates

Conducting polymers have attracted great attention since the discovery of high conductivity in doped polyacetylene in 19771. They offer the advantages of low weight, easy tailoring of properties and a wide spectrum of applications2,3. Due to sensitivity of conducting polymers to environmental conditions (e.g., air, oxygen, moisture, high temperature and chemical solutions), lithographic techniques present significant technical challenges when working with these materials4. For example, current photolithographic methods, such as ultra-violet (UV), are unsuitable for patterning the conducting polymers due to the involvement of wet and/or dry etching processes in these methods. In addition, current micro/nanosystems mainly have a planar form5,6. One layer of structures is built on the top surfaces of another layer of fabricated features. Multiple layers of these structures are stacked together to form numerous devices on a common substrate. The sidewall surfaces of the microstructures have not been used in constructing devices. On the other hand, sidewall patterns could be used, for example, to build 3-D circuits, modify fluidic channels and direct horizontal growth of nanowires and nanotubes.
A macropunching method has been applied in the manufacturing industry to create macropatterns in a sheet metal for over a hundred years. Motivated by this approach, we have developed a micropunching lithography method (MPL) to overcome the obstacles of patterning conducting polymers and generating sidewall patterns. Like the macropunching method, the MPL also includes two operations (Fig. 1): (i) cutting; and (ii) drawing. The "cutting" operation was applied to pattern three conducting polymers4, polypyrrole (PPy), Poly(3,4-ethylenedioxythiophen)-poly(4-styrenesulphonate) (PEDOT) and polyaniline (PANI). It was also employed to create Al microstructures7. The fabricated microstructures of conducting polymers have been used as humidity8, chemical8, and glucose sensors9. Combined microstructures of Al and conducting polymers have been employed to fabricate capacitors and various heterojunctions9,10,11. The "cutting" operation was also applied to generate submicron-patterns, such as 100- and 500-nm-wide PPy lines as well as 100-nm-wide Au wires. The "drawing" operation was employed for two applications: (i) produce Au sidewall patterns on high density polyethylene (HDPE) channels which could be used for building 3D microsystems12,13,14, and (ii) fabricate polydimethylsiloxane (PDMS) micropillars on HDPE substrates to increase the contact angle of the channel15.

A micropunching lithography approach is developed to generate micro- and submicron-patterns on top, sidewall and bottom surfaces of polymer substrates. It overcomes the obstacles of patterning conducting polymers and generating sidewall patterns. This method allows rapid fabrication of multiple features and is free of aggressive chemistry.

Conducting polymers have attracted great attention since the discovery of high conductivity in doped polyacetylene in 19771. They offer the advantages of ...

Engineering

Stretching Micropatterned Cells on a PDMS Membrane

Mechanical forces exerted on cells and/or tissues play a major role in numerous processes. We have developed a device to stretch cells plated on a PolyDiMethylSiloxane (PDMS) membrane, compatible with imaging. This technique is reproducible and versatile. The PDMS membrane can be micropatterned in order to confine cells or tissues to a specific geometry. The first step is to print micropatterns onto the PDMS membrane with a deep UV technique. The PDMS membrane is then mounted on a mechanical stretcher. A chamber is bound on top of the membrane with biocompatible grease to allow gliding during the stretch. The cells are seeded and allowed to spread for several hours on the micropatterns. The sample can be stretched and unstretched multiple times with the use of a micrometric screw. It takes less than a minute to apply the stretch to its full extent (around 30%). The technique presented here does not include a motorized device, which is necessary for applying repeated stretch cycles quickly and/or computer controlled stretching, but this can be implemented. Stretching of cells or tissue can be of interest for questions related to cell forces, cell response to mechanical stress or tissue morphogenesis. This video presentation will show how to avoid typical problems that might arise when doing this type of seemingly simple experiment.

This manuscript presents a technique to apply or release forces on adherent cells or tissues using unidirectional stretching.

Mechanical forces exerted on cells and/or tissues  play a major role in numerous processes. We have developed a device to stretch  cells plated on a ...

Mapping the Emergent Spatial Organization of Mammalian Cells using Micropatterns and Quantitative Imaging

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro when stem cells are spatially confined onto micropatterns, thus setting up experimental models which offer unique opportunities to identify, in vitro, the fundamental principles of biological organisation.
Here we describe our own implementation of the methodology. We adapted a photo-patterning technique to reduce the need for specialized equipment to make it easier to establish the method in a standard cell biology laboratory. We also developed a free, open-source and easy to install image analysis framework in order to precisely measure the preferential positioning of sub-populations of cells within colonies of standard shapes and sizes. This method makes it possible to reveal the existence of patterning events even in seemingly disorganized populations of cells. The technique provides quantitative insights and can be used to decouple influences of the environment (e.g., physical cues or endogenous signaling), on a given patterning process.

The method presented here uses micropatterning together with quantitative imaging to reveal spatial organization within mammalian cultures. The technique is easy to establish in a standard cell biology laboratory and offers a tractable system to study patterning in vitro.

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro ...

Developmental Biology

Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules present in the cellular context and preserved in a near-native frozen-hydrated state. However, there are challenges associated with culturing and/or adhering cells onto TEM grids in a manner that is suitable for tomography while retaining the cells in their physiological state. Here, a detailed step-by-step protocol is presented on the use of micropatterning to direct and promote eukaryotic cell growth on TEM grids. During micropatterning, cell growth is directed by depositing extra-cellular matrix (ECM) proteins within specified patterns and positions on the foil of the TEM grid while the other areas remain coated with an anti-fouling layer. Flexibility in the choice of surface coating and pattern design makes micropatterning broadly applicable for a wide range of cell types. Micropatterning is useful for studies of structures within individual cells as well as more complex experimental systems such as host-pathogen interactions or differentiated multi-cellular communities. Micropatterning may also be integrated into many downstream whole-cell cryo-ET workflows, including correlative light and electron microscopy (cryo-CLEM) and focused-ion beam milling (cryo-FIB).

The goal of this protocol is to direct cell adhesion and growth to targeted areas of grids for cryo-electron microscopy. This is achieved by applying an anti-fouling layer that is ablated in user-specified patterns followed by deposition of extra-cellular matrix proteins in the patterned areas prior to cell seeding.

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules ...

Biochemistry

Patterning of Microorganisms and Microparticles through Sequential Capillarity-assisted Assembly

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, including studies of microbial physiology and interactions. At the simplest level, accurate spatial patterning of microorganisms would enable reliable, long-term imaging of large numbers of individual cells and transform the ability to quantitatively study distance-dependent microbe-microbe interactions. More uniquely, coupling accurate spatial patterning and full control over environmental conditions, as offered by microfluidic technology, would provide a powerful and versatile platform for single-cell studies in microbial ecology. 
This paper presents a microfluidic platform to produce versatile and user-defined patterns of microorganisms within a microfluidic channel, allowing complete optical access for long-term, high-throughput monitoring. This new microfluidic technology is based on capillarity-assisted particle assembly and exploits the capillary forces arising from the controlled motion of an evaporating suspension inside a microfluidic channel to deposit individual microsized objects in an array of traps microfabricated onto a polydimethylsiloxane (PDMS) substrate. Sequential depositions generate the desired spatial layout of single or multiple types of micro-sized objects, dictated solely by the geometry of the traps and the filling sequence. 
The platform has been calibrated using colloidal particles of different dimensions and materials: it has proven to be a powerful tool to generate diverse colloidal patterns and perform surface functionalization of trapped particles. Furthermore, the platform was tested on microbial cells, using Escherichia coli cells as a model bacterium. Thousands of individual cells were patterned on the surface, and their growth was monitored over time. In this platform, the coupling of single-cell deposition and microfluidic technology allows both geometric patterning of microorganisms and precise control of environmental conditions. It thus opens a window into the physiology of single microbes and the ecology of microbe-microbe interactions, as shown by preliminary experiments.

We present a technology that uses capillarity-assisted assembly in a microfluidic platform to pattern micro-sized objects suspended in a liquid, such as bacteria and colloids, into prescribed arrays on a polydimethylsiloxane substrate.

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, ...

Fabrication and Implementation of a Reference-Free Traction Force Microscopy Platform

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their microenvironment. While many approaches exist for measuring cell-induced material strain, here we provide a methodology for monitoring strain with sub-micron resolution in a reference-free manner. Using a two-photon activated photolithographic patterning process, we demonstrate how to generate mechanically and bio-actively tunable synthetic substrates containing embedded arrays of fluorescent fiducial markers to easily measure three-dimensional (3D) material deformation profiles in response to surface tractions. Using these substrates, cell tension profiles can be mapped using a single 3D image stack of a cell of interest. Our goal with this methodology is to make traction force microscopy a more accessible and easier to implement tool for researchers studying cellular mechanotransduction processes, especially newcomers to the field.

This protocol provides instructions for implementing multiphoton lithography to fabricate three-dimensional arrays of fluorescent fiducial markers embedded in poly(ethylene glycol)-based hydrogels for use as reference-free, traction force microscopy platforms. Using these instructions, measurement of 3D material strain and calculation of cellular tractions is simplified to promote high-throughput traction force measurements.

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their ...

Traction Microscopy Integrated with Microfluidics for Chemotactic Collective Migration

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical gradient, known as chemotaxis, plays an important role in development, the immune response, wound healing, and cancer metastasis. While chemotaxis modulates the migration of single cells as well as collections of cells in vivo, in vitro research focuses on single-cell chemotaxis, partly due to the lack of the proper experimental tools. To fill that gap, described here is a unique experimental system that combines microfluidics and micropatterning to demonstrate the effects of chemical gradients on collective cell migration. Furthermore, traction microscopy and monolayer stress microscopy are incorporated into the system to characterize changes in cellular force on the substrate as well as between neighboring cells. As proof-of-concept, the migration of micropatterned circular islands of Madin-Darby canine kidney (MDCK) cells is tested under a gradient of hepatocyte growth factor (HGF), a known scattering factor. It is found that cells located near the higher concentration of HGF migrate faster than those on the opposite side within a cell island. Within the same island, cellular traction is similar on both sides, but intercellular stress is much lower on the side of higher HGF concentration. This novel experimental system can provide new opportunities to studying the mechanics of chemotactic migration by cellular collectives.

Collective cell migration in development, wound healing, and cancer metastasis is often guided by the gradients of growth factors or signaling molecules. Described here is an experimental system combining traction microscopy with a microfluidic system and a demonstration of how to quantify the mechanics of collective migration under biochemical gradient.

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical ...

Control of Cell Geometry through Infrared Laser Assisted Micropatterning

Micropatterning is an established technique in the cell biology community used to study connections between the morphology and function of cellular compartments while circumventing complications arising from natural cell-to-cell variations. To standardize cell shape, cells are either confined in 3D molds or controlled for adhesive geometry through adhesive islands. However, traditional micropatterning techniques based on photolithography and deep UV etching heavily depend on clean rooms or specialized equipment. Here we present an infrared laser assisted micropatterning technique (microphotopatterning) modified from Doyle et al. that can be conveniently set up with commercially available imaging systems. In this protocol, we use a Nikon A1R MP+ imaging system to generate micropatterns with micron precision through an infrared (IR) laser that ablates preset regions on poly-vinyl alcohol coated coverslips. We employ a custom script to enable automated pattern fabrication with high efficiency and accuracy in systems not equipped with a hardware autofocus. We show that this IR laser assisted micropatterning (microphotopatterning) protocol results in defined patterns to which cells attach exclusively and take on the desired shape. Furthermore, data from a large number of cells can be averaged due to the standardization of cell shape. Patterns generated with this protocol, combined with high resolution imaging and quantitative analysis, can be used for relatively high throughput screens to identify molecular players mediating the link between form and function.

The protocol presented here enables automated fabrication of micropatterns that standardizes cell shape to study cytoskeletal structures within mammalian cells. This user-friendly technique can be set up with commercially available imaging systems and does not require specialized equipment inaccessible to standard cell biology laboratories.

Micropatterning is an established technique in the cell biology community used to study connections between the morphology and function of cellular ...

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

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

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

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

Generation of Multicue Cellular Microenvironments by UV-Photopatterning of Three-Dimensional Cell Culture Substrates

The extracellular matrix is an important regulator of cell function. Environmental cues existing in the cellular microenvironment, such as ligand distribution and tissue geometry, have been increasingly shown to play critical roles in governing cell phenotype and behavior. However, these environmental cues and their effects on cells are often studied separately using in vitro platforms that isolate individual cues, a strategy that heavily oversimplifies the complex in vivo situation of multiple cues. Engineering approaches can be particularly useful to bridge this gap, by developing experimental setups that capture the complexity of the in vivo microenvironment, yet retain the degree of precision and manipulability of in vitro systems.
This study highlights an approach combining ultraviolet (UV)-based protein patterning and lithography-based substrate microfabrication, which together enable high-throughput investigation into cell behaviors in multicue environments. By means of maskless UV-photopatterning, it is possible to create complex, adhesive protein distributions on three-dimensional (3D) cell culture substrates on chips that contain a variety of well-defined geometrical cues. The proposed technique can be employed for culture substrates made from different polymeric materials and combined with adhesive patterned areas of a broad range of proteins. With this approach, single cells, as well as monolayers, can be subjected to combinations of geometrical cues and contact guidance cues presented by the patterned substrates. Systematic research using combinations of chip materials, protein patterns, and cell types can thus provide fundamental insights into cellular responses to multicue environments.

Traditionally, cell culture is performed on planar substrates that poorly mimic the natural environment of cells in vivo. Here we describe a method to produce cell culture substrates with physiologically relevant curved geometries and micropatterned extracellular proteins, allowing systematic investigations into cellular sensing of these extracellular cues.