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 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.
Most cell phenotypes exert traction forces on their environment. These traction forces are generated by a cell'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'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.
1. Creation of silicone masters
NOTE: Most of the process of the design, creation, and troubleshooting of silicon masters for the repeated molding of PDMS stamps has been covered previously21, so only key differences in this new approach will be described here.
2. Subtractive microcontact printing
3. Activated coverslips
NOTE: The bottom coverslips for use in the experimental chamber for PAA gels are made in this step. This bottom coverslip is specially treated to allow the PAA gel to remain securely adhered to as the top patterned coverslip is removed during the patterning process. Similar techniques are also described elsewhere10,12,15,28.
4. PAA gel fabrication and pattern transfer
NOTE: Once patterned coverslips are made, they must be used to transfer those protein patterns to the PAA hydrogel soon afterward (<24 h)1,29,30. The following recipe is for a PAA gel with a Young's modulus of 3.6 kPa. The amounts of bis-acrylamide, acrylamide, and DI water can be varied to adjust the stiffness of the PAA gels12.
5. Imaging
6. Image analysis
NOTE: A system has been developed that can measure the deformation of the patterned PAA gels by determining the location of the traction points, interpolating the initial locations of the deformed points, and then calculating the cellular traction forces at each location. Any software system capable of performing image processing and numerical calculations can be used. The program aims to determine traction forces rapidly, eliminating user input and preprocessing procedures that would contribute to user-related errors. The code used here is available here as Supplemental Files 2-10, and these files, along with a pair of practice images, can be accessed at www.bu.edu/mml/downloads.
PAA hydrogels with the Young's modulus of E = 3.6 kPa and the Poisson's ratio of ν = 0.445 were made for use by this subtractive micropatterning method. The hydrogels were made to be ~100 µm 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 × 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 µm 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 °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 ≤2 µm20), 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 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.
Figure 1: Design of photomask. (A) A representation of the first half of the photomask design used here, a discrete grid of 2 µm diameter dots spaced at 6 µm 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 µm 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 µm. The dots in each island are 2 µm in diameter and spaced 6 µm center-to-center. The two sections of the mask described in A and B are separated by 0.75 cm. Please click here to view a larger version of this figure.
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) 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) An example of a discrete island pattern made up of evenly spaced dots on a PAA hydrogel with a Young's modulus of 3.6 kPa. Scale bar = 20 µm. Abbreviations: PDMS = polydimethylsiloxane; PAA = polyacrylamide; NHS = N-hydroxysuccinimide. Please click here to view a larger version of this figure.
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's modulus of 3.6 kPa is shown. Dots are 2 µm in diameter and are separated by 6 µm center-to-center. (B) 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. Please click here to view a larger version of this figure.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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. Please click here to download this File.
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 µm-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 µm-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'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 µm circular dots spaced 6 µm center-to-center are used. The reason for this spacing is that a displacement at the 6 µm 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 µm. 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.
The authors have nothing to disclose.
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.
(3-aminopropyl)trimethoxysilane | Sigma Aldrich | #281778 | |
1.5 mL Microcentrifuge tube | Fisher Scientific | #05-408-129 | |
15 mL conical tube | Fisher Scientific | #05-539-12 | |
4 x 4 in 0.060 Quartz LR Chrome Photomask | Advance Reproductions Corporation | N/A | Custom-designed mask |
6 Well Plates | Fisher Scientific | #07-200-83 | |
Acetone | Fisher Scientific | #A18P-4 | |
Acrylamide Solution, 40% | Sigma Aldrich | #A4058 | |
AlexaFluor 488 | Thermo Fisher | #A20000 | |
Aminonium Persulfate | Fisher Scientific | #BP179-25 | |
Bisacrylamide | Fisher Scientific | #PR-V3141 | |
Ethanol | Greenfield Global | #111000200C1GL | |
Glass Coverslips, 25 mm round | Fisher Scientifc | #12-545-102 | |
Glass Coverslips, 30 mm round | Warner | #64-1499 | |
Hamamatsu ORCA-R2 Camera | Hamamatsu | #C10600-10B | |
Human Plasma | Valley Biomedical | #HP1051P | Used to isolate fibronectin |
Hydrochloric Acid, 1.0 N | Millipore Sigma | #1.09057 | |
ImageJ | Wayne Rasband | #1.53n | |
Interchangeable Coverslip Dish Set | Bioptechs | #190310-35 | |
Kim Wipes | Fisher Scientific | #06-666-11C | |
Mask Alinger | Karl Suss | #MA6 | |
Matlab 2021 | Mathworks | #R2021a | |
MetaMorph Basic | Molecular Devices | #v7.7.1.0 | |
N-hydroxysuccinimide ester | Sigma Aldrich | #130672-5G | |
NucBlue Live Cell Stain | Thermo Fisher | #R37605 | |
Olympus IX2-ZDC Inverted Microscope | Olympus | #IX81 | |
PD-10 Desalting Columns | GE Healthcare | #52-1308-00 | |
Photoresist Spinner Hood | Headway Research | #PWM32 | |
Plasma Cleaner | Harrick | #PDC-001 | |
Plasma Etcher | TePla | #M4L | |
Prior Lumen 200Pro Light Source | Prior Scientific | #L200 | |
Silicon Wafers, 100 mm | University Wafer | #809 | |
SU-8 2005 | Kayaku Advanced Materials Inc. | #NC9463827 | |
SU-8 Developer | Kayaku Advanced Materials Inc. | #NC9901158 | |
Sylgard 184 Silicone Elastomer | Essex Brownell | #DC-184-1.1 | |
Tetramethylethylenediamine | Fisher Scientific | #BP150-20 | |
Trichloro(1H,1H,2H,2H-perfluorooctyl)silane | Sigma Aldrich | #448931 | |
UAPON-40XW340 Objective | Olympus | #N2709300 | |
UV Flood Exposure | Newport | #69910 | |
Wafer Carrier Tray, 110 x 11 mm | Ted Pella, Inc. | #19395-40 |