This study was supported by the National Natural Science Foundation of China (Grant numbers 12075330 and U1932165) and the Natural Science Foundation of Guangdong Province, China (Grant number 2020A1515010270).

1. Fabricating and assembly of the acoustofluidic microdevice
<ol>
	<li>Fabrication of the microfluidic chip.
	<ol>
		<li>Design a single-channel chip with only one inlet and outlet as shown in Figure 1. For measuring cells, keep the rectangular cross-section of the microchannel at 740 &#181;m wide and 100 &#181;m deep. For measuring cell nucleus, change the width and depth of the microchannel to 250 &#181;m and 100 &#181;m, respectively.</li>
		<li>Prepare the microchannel on silicon wafer via reactive ion etching. Seal the top of the microchannel with a piece of transparent heat-resistant glass by anodic bonding15. Wash the chips with an ultrasonic cleaner for 10 min. Dry them in a drying oven at 50 &#176;C for later use.</li>
	</ol>
	</li>
	<li>Fabricate polydimethylsiloxane (PDMS) blocks.
	<ol>
		<li>Add 30 mL of pre-polymer to a 100 mm (in diameter) glass dish. Add 3 mL of curing agent to the pre-polymer with a syringe. 
		NOTE: The volume ratio of the curing agent and pre-polymer is 1:10.</li>
		<li>Vigorously mix the PDMS pre-polymer and curing agent with a glass rod for about 10 min. Look for small and uniformly separated air bubbles in the solution, which indicate that the PDMS pre-polymer and curing agent are well mixed.</li>
		<li>Place the glass dish in a vacuum desiccator and evacuate for 15-25 s. Repeat this process until there are no air bubbles in the mixture.</li>
		<li>Place the glass dish in a drying oven set at 50 &#176;C for 1 h to allow the mixture to cure. After the incubation, use a scalpel to cut the PDMS into blocks of suitable size about 1.2 cm long and 1 cm wide. 
		NOTE: The length of the PDMS block is consistent with the width of the chip, and the width is selected to ensure that there is enough space in the middle for the piezoelectric ceramic when two PDMS blocks are adhered on the chip.</li>
	</ol>
	</li>
	<li>Bind the PDMS block to the chip.
	<ol>
		<li>Punch holes in the PDMS block for inlet and outlet ports with a 1 mm diameter hollow needle. Put the PDMS blocks and chip (back side up) in a plasma cleaner for 1 min.</li>
		<li>Align the holes on the PDMS blocks with the chip inlet and outlet. Gently press the PDMS blocks to the chip for 15 s. This should cause bonding to occur between the PDMS blocks and the surface of the chip.</li>
	</ol>
	</li>
	<li>Connect the polytetrafluoroethylene (PTFE) catheter to the chip (Figure 2B).
	<ol>
		<li>Cut two pieces of PTFE catheter with an inner diameter of 0.8 mm and a length of 10 cm. Bend a stainless-steel needle with an inner diameter of 0.7 mm and a length of 1.5 cm by 90&#176; into an L shape. Connect it to one end of the catheter. Prepare two such catheters with needles.</li>
		<li>Insert the stainless-steel needles into the holes of the PDMS blocks. For the inlet, connect a 19 G dispensing needle to the other end of the catheter as a connector for a syringe.</li>
		<li>After completing the above steps, inject deionized water to test the tightness of the overall channel. Impervious to water means a good seal.</li>
	</ol>
	</li>
	<li>Piezoelectric ceramic assembly (Figure 2C)
	<ol>
		<li>Use a diamond wire cutter to cut piezoelectric ceramic sheets with a diameter of 2 cm into four strips with a width of 5 mm.</li>
		<li>Ensure that the resonant frequency of the piezoelectric ceramic matches the width of the chip microchannel. For the 740 &#181;m and 250 &#181;m wide microchannel, use piezoelectric ceramics with resonance frequencies of 1 MHz and 3 MHz, respectively.</li>
		<li>Weld wires on both sides of the piezoelectric ceramic at one end.</li>
		<li>Glue the piezoelectric ceramic to the middle of the back of the chip with cyanoacrylate glue.</li>
		<li>To spread the glue evenly, place a drop of glue on the piezoelectric ceramic, smooth the glue with a toothpick and remove the excess glue. Then, quickly press it on the chip and continue to press for about 1 min. Ensure that the piezoelectric ceramic and the chip are firmly bonded and evenly contacted.</li>
	</ol>
	</li>
	<li>Mount the microdevice (Figure 2D).
	<ol>
		<li>Cut a piece of PDMS (about 1.5 cm long and 1 cm wide) as the base of the microdevice. Using double-sided tape, stick one side of the base to the inlet and outlet PDMS blocks, and the other side to a transparent glass slide. Fix the whole microdevice to the microscope stage to keep the chip in one focal plane.</li>
	</ol>
	</li>
</ol>
2. Sample preparation
<ol>
	<li>Preparation of polystyrene standard particle solutions.
	<ol>
		<li>Add 0.05 mL of polystyrene particle (6 &#181;m in diameter) solution (2.1 x 108 particle/mL) to 10 mL of phosphate buffered saline (PBS) and mix well. 
		NOTE: In order to reduce the measurement error caused by the change of the acoustic energy density, the polystyrene particle solution was mixed with the sample solution in each experiment as calibration.</li>
	</ol>
	</li>
	<li>Preparation of cell suspensions.
	<ol>
		<li>Wash the adherent cells (e.g., MCF7, MDA-MB-231, HCT116) at 90% confluency (~5 x 105 cells) with PBS. Add 500 &#181;L 0.25% trypsin (1x) for 1-2 min at room temperature (25 &#176;C). Remove the trypsin, add 1 mL complete medium and form a cell suspension by pipetting.</li>
		<li>Centrifuge the cell suspension at 100 x g for 5 min. Remove the supernatant and resuspend in 0.5-1 mL of PBS in order to obtain a cell suspension. Cells were counted with a hemocytometer and the concentration was about 3-5 x 105 cells/mL.</li>
	</ol>
	</li>
	<li>Preparation of cell nucleus suspension
	<ol>
		<li>Carry out step 2.2. Then, remove the supernatant and add 200 &#181;L of cytoplasmic protein extraction reagent A (supplemented with 1% PMSF) per 20 &#181;L cell pellet (approximately 5 million cells) and mix well.</li>
		<li>Vortex the above mixture at 220 x g for 5 s, and then place on ice bath for 10 min. After incubation, add 10 &#181;L of cytoplasmic protein extraction reagent B to the solution.</li>
		<li>Vortex at 220 x g for 5 s. Place on ice bath for 1 min and vortex again at 220 x g for 5 s. Then, finally centrifuge at 1,000 x g for 5 min at 4 &#176;C. 
		NOTE: The volume ratio of cytoplasmic protein extraction reagents A and B is 20:1.</li>
		<li>Remove the supernatant and resuspend the pellet in 1 mL of PBS. Then, centrifuge at 1,000 x g at 4 &#176;C for 4 min. Remove the supernatant and resuspend in 100 &#181;L of PBS as cell nucleus suspension.</li>
		<li>Add trypan blue to the above cell nucleus suspension and stain at room temperature (25 &#176;C) for 4 min. The volume ratio of trypan blue solution to nucleus suspension is 1:1. Count the number of nuclei under the inverted microscope with a 10x objective. 
		NOTE: To clearly identify the cell nuclei under the microscope, trypan blue staining is required. Trypan blue solution needs to be in a 37 &#176;C water bath for 10 min before use for effective staining.</li>
		<li>Dilute the above cell nucleus suspension with PBS buffer to a concentration of 2-3 x 105 nucleus/mL. Filter the cell nucleus suspension through a 70 &#181;m sieve.</li>
	</ol>
	</li>
</ol>
3. Measuring the compressibility of cell and nucleus
<ol>
	<li>Set up the measurement system (Figure 3)
	<ol>
		<li>Turn on the light source of the microscope and open the camera software. Use the 4x objective to find the middle position of the microchannel, i.e., the position of the piezoelectric ceramic.</li>
		<li>Connect the wires and weld them to the positive and negative terminals of the signal generator output on the piezoelectric ceramic, respectively.</li>
		<li>Place the syringe on the microinjection pump and connect it to the inlet catheter. Place a small container at the end of the outlet catheter to hold the fluid flowing out of the microchannel.</li>
	</ol>
	</li>
	<li>Determine measurement parameters
	<ol>
		<li>Aspirate the polystyrene particle solution with the syringe and inject it into the chip microchannel. Avoid air bubbles in the chip microchannel to ensure accurate measurement. Ensure that particles are evenly distributed in the chip microchannel. 
		NOTE: Measurement can be conducted without flow or syringe pump. If needed, the flow rate of the microinjection pump should be set to a proper value. Here, the range of flow rate is 0-20 &#181;L/h.</li>
		<li>Set the output of the signal generator to a sine signal with a frequency of 1 MHz (3 MHz for cell nucleus measurement) and peak-to-peak voltage (Vpp) of 10 V.</li>
		<li>Fine tune the frequency of the signal until it&#39;s observed that the particles move toward the midline of the microchannel and remain in forward motion along the midline after reaching the midline (Figure 4). 
		NOTE: The speed of the particles moving toward the midline is determined by the voltage amplitude, which can be adjusted between 5 Vpp and 20 Vpp.</li>
	</ol>
	</li>
	<li>Measure cells and nuclei
	<ol>
		<li>Mix 1 mL cell or nucleus suspension with the standard particle solution at the ratio of 1:1 and inject it into the microchannel with a syringe.</li>
		<li>Start recording with CCD camera when the cells or nuclei enter the field of view. Then, turn on the signal generator. Stop recording when the cells or nuclei reach the midline.</li>
		<li>Rinse the microchannel with deionized water, 75% alcohol, and deionized water in sequence for later use.</li>
	</ol>
	</li>
</ol>
4. Data processing
<ol>
	<li>Map particle or cell trajectories.
	<ol>
		<li>Import the captured video into ImageJ software: File &#62; Open&#62; Select Folder. Click on the ellipse shape in the toolbar of the ImageJ software to pick a cell of interest and its adjacent particle (Figure 5).</li>
		<li>As shown in Figure 5, preset measurement parameters in ImageJ software as Analyze&#8594; Set Measurement &#8594;Area, Centroid, Display Label.</li>
		<li>Taking the frame where the target cell or particle undergoes longitudinal displacement as the start frame; record the pixel position and size of the cell or particle in each frame until it reaches the midline of the microchannel. Export the data as a spreadsheet file and repeat the step until trajectories for all cells of interest are obtained.</li>
	</ol>
	</li>
	<li>Coordinate transformation and correction.
	<ol>
		<li>Record the pixel coordinates of the four corners of the microchannel in this field of view as (0, y1), (0, y2), (x3, y3), (x4, y4). Here x3 = x4.</li>
		<li>For each measurement point (xi, yi), calculate the new coordinate (xi&#39;, yi&#39;) after rotation correction using the following formulas: 
		<img alt="Equation 1" src="/files/ftp_upload/64225/64225eq01.jpg" style="margin: auto;" /> 
		<img alt="Equation 2" src="/files/ftp_upload/64225/64225eq02.jpg" style="margin: auto;" /> 
		<img alt="Equation 3" src="/files/ftp_upload/64225/64225eq03.jpg" style="margin: auto;" /></li>
		<li>Convert pixel coordinates to real size coordinates. The actual coordinates can be obtained by multiplying pixel coordinates by the ratio. The ratio was the actual width of the microchannel divided by the pixel width (H) of the microchannel.</li>
		<li>Transform and correct the pixel coordinates of the cells and particles obtained in step 4.1 to the final motion trajectory data. All coordinates minus the coordinates of the lower-left corner, i.e., (0, y2). The frame rate of the video is 40 frames per second, so multiply the number of frames corresponding to each coordinate by 0.025 s to obtain the time of particle movement, thereby obtaining the change of the position in the y direction with time.</li>
	</ol>
	</li>
	<li>Calculate the acoustic energy density (Figure 6A,B).
	<ol>
		<li>The motion of the cell or particle in the Y direction is driven by the acoustic force Fac and hydrodynamic force FD. Calculate the motion trajectory using the following formulas: 
		<img alt="Equation 4" src="/files/ftp_upload/64225/64225eq04.jpg" style="margin: auto;" />&#160; &#160; &#160;(1) 
		<img alt="Equation 5" src="/files/ftp_upload/64225/64225eq05.jpg" style="margin: auto;" />&#160; &#160; &#160;(2) 
		<img alt="Equation 6" src="/files/ftp_upload/64225/64225eq06.jpg" style="margin: auto;" />&#160; &#160; &#160;(3) 
		where r and D are the radius and diameter of the cell or particle, &#961; and &#946; are the density and compressibility,&#160;&#957;&#160;is the velocity vector. The subscripts c and f denote the cell and fluid, respectively. d is the distance from the nearest acoustic pressure node, &#956; is the dynamic viscosity of the fluid, k is the wave number, and E is the acoustic energy density. 
		NOTE: The density of MCF7, HCT116, A549 and cell nuclei was 1068 kg/m3, 1077 kg/m3, 1073 kg/m3, and 1155 kg/m3, respectively 12,16,17.</li>
		<li>According to the formulas described in step 4.3.1, use the MATLAB software to obtain the numerical solution for the standard particle trajectory under the acoustic field with finite difference method.</li>
		<li>Within the preset acoustic field range, change the acoustic energy density and fit the numerical solution (obtained in step 4.3.2) and the measured motion trajectory (obtained in step 4.2) for the standard particle. Select the best fitting result according to the fitting mean square error. The acoustic energy density obtained here is used as a parameter for subsequent calculation of cell compressibility.</li>
	</ol>
	</li>
	<li>Calculate the cell compressibility (Figure 6C,D).
	<ol>
		<li>Set the acoustic energy density to the value obtained in step 4.3.3.</li>
		<li>According to the formulas described in step 4.3.1, use the MATLAB software to obtain the numerical solution for the cell trajectory under the acoustic field with finite difference method.</li>
		<li>Similar to step 4.3.3, within the preset compressibility range, change the compressibility and fit the numerical solution (obtained in step 4.4.2) and the measured motion trajectory for the cell (obtained in step 4.2). Use the compressibility coefficient corresponding to the best fitting result as the measured cell compressibility.</li>
	</ol>
	</li>
</ol>

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The authors have no competing financial interests or other conflicts of interest.

Commonly used cell mechanics measurement methods are AFM, micropipette aspiration, microfluidics methods, parallel-plate technique, optical tweezers, optical stretcher, and acoustic methods20. Microfluidics methods can work with three approaches: micro-constriction, extensional flow, and shear flow. Among them, optical stretcher, optical tweezers, acoustic methods, extensional flow, and shear flow approaches are non-contact measurements. In contrast to contact measurements, non-contact measurement...

Cell mechanical properties play an important role in tumor metastasis, malignant transformation of cells, and radiosensitivity1,2. To gain an in-depth understanding of the role of cell mechanical properties in the above process, accurate measurement of cellular mechanics is critical, and the measurement should not cause damage to the cells for subsequent culture and analysis. The measurement process should be as fast as possible, otherwise cell viability may be affected if cells are removed from the cultivation environment for a long time.
Existing cell mechanics measurement methods face some limitations. Some methods, such as magnetic twisting cytometry, magnetic tweezers and particle-tracking microrheology, cause cell damage due to the introduction of particles into cells3,4,5. Methods that measure by contact with cells, such as atomic force microscope (AFM), micropipette aspiration, micro-constriction, and parallel-plate technique, are also prone to cell damage and the throughput is difficult to increase6,7,8. In addition, ionizing radiation will flatten cells and increase their adhesion9; it is therefore necessary to measure whole cell mechanics in suspension.
In response to the above challenges, a cell mechanics measurement system based on acoustofluidic method10,11,12,13,14 has been developed. The channel width is matched to the acoustic half wavelength, thus creating a standing wave node at the midline of the microchannel. Under the action of acoustic radiation force, the cells or standard beads can move to the acoustic pressure node. Since the physical properties of the standard beads (size, density, and compressibility) are known, the acoustic energy density can be determined. Then, the cell compressibility can be obtained by recording the motion trajectories of cells in the acoustic field. Non-destructive high-throughput measurement of cells in suspension state can be achieved. This paper will introduce the design of the microfluidic chip, the establishment of the system and the measurement steps. Measurement of various types of tumor cells has been carried out to verify the accuracy of the method. The application scope of this method had been extended to subcellular structures (such as nucleus) by adjusting the resonance frequency of the piezoelectric ceramic and the width of the microchannel. In addition, the changes in cell compressibility after drug-induced EMT or X-ray irradiation with different doses were investigated. The results demonstrate the broad applicability of this method as a powerful tool for studying the correlation between biochemical changes and cellular mechanical properties.

<table><tbody><tr><td>0.25% trypsin(1x)</td><td>GIBCO</td><td>15050-065</td><td></td></tr><tr><td>502 glue</td><td>Evo-bond</td><td></td><td>cyanoacrylate glue</td></tr><tr><td>A549</td><td>ATCC</td><td>CCL-185</td><td>lung adenocarcinoma</td></tr><tr><td>Cytonucleoprotein and cytoplasmic protein extraction kit</td><td>Beyotime</td><td>P0027</td><td>Contains cytoplasmic protein extraction reagents A and B</td></tr><tr><td>Dulbecco&amp;rsquo;s modified Eagle medium (DMEM)&amp;nbsp;</td><td>corning</td><td>10-013-CVRC</td><td></td></tr><tr><td>Fetal Bovine Srum(FBS)</td><td>AUSGENEX</td><td>FBS500-S</td><td></td></tr><tr><td>HCT116</td><td>ATCC</td><td>CCL247</td><td>colorectal carcinoma</td></tr><tr><td>Heat-resistant glass</td><td>Pyrex</td><td></td><td></td></tr><tr><td>Leibovitz&amp;rsquo;s L-15 medium&amp;nbsp;</td><td>GIBCO</td><td>11415-064</td><td></td></tr><tr><td>MCF-7</td><td>ATCC</td><td>HTB-22&amp;nbsp;</td><td>breast Adenocarcinoma</td></tr><tr><td>MDA-MB-231</td><td>ATCC</td><td>HTB-26&amp;nbsp;</td><td>breast Adenocarcinoma</td></tr><tr><td>Minimum Essential Medium (MEM)</td><td>corning</td><td>10-010-CV</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>GIBCO</td><td>15140-122</td><td></td></tr><tr><td>Phosphate buffer</td><td>corning</td><td>21-040-cvc</td><td></td></tr><tr><td>PMSF</td><td>Beyotime</td><td>ST506</td><td>100mM</td></tr><tr><td>Polybead Polystyrene Red Dyed Microsphere&amp;nbsp;</td><td>polysciences</td><td>15714</td><td>The diameter of microshpere is 6.00&amp;micro;m</td></tr><tr><td>propidium iodide(PI)</td><td>Sigma-Aldrich</td><td>P4170</td><td></td></tr><tr><td>SYLGARD 184Silicone ELASTOMER</td><td>Dow-Corning</td><td>1673921</td><td>Contains prepolymers and curing agents</td></tr><tr><td>Trypan Blue</td><td>Beyotime</td><td>C0011</td><td></td></tr></tbody></table>

measurement of the compressibility of cell and nucleus based on acoustofluidic microdevice

Cell mechanics play an important role in tumor metastasis, malignant transformation of cells, and radiosensitivity. During these processes, studying the mechanical properties of the cells is often challenging. Conventional measurement methods based on contact such as compression or stretching are prone to cause cell damage, affecting measurement accuracy and subsequent cell culture. Measurements in adherent state can also affect accuracy, especially after irradiation since ionizing radiation will flatten cells and enhance adhesion. Here, a cell mechanics measurement system based on acoustofluidic method has been developed. The cell compressibility can be obtained by recording the cell motion trajectory under the action of the acoustic force, which can realize fast and non-destructive measurement in suspended state. This paper reports in detail the protocols for chip design, sample preparation, trajectory recording, parameter extraction and analysis. The compressibility of different types of tumor cells was measured based on this method. Measurement of the compressibility of nucleus was also achieved by adjusting the resonance frequency of the piezoelectric ceramic and the width of the microchannel. Combined with the molecular level verification of immunofluorescence experiments, the cell compressibility before and after drug-induced epithelial to mesenchymal transition (EMT) were compared. Further, the change of cell compressibility after X-ray irradiation with different doses was revealed. The cell mechanics measurement method proposed in this paper is universal and flexible and has broad application prospects in scientific research and clinical practice.

Here, the work presented a protocol for the construction of a fast and non-destructive cell compressibility measuring system based on acoustofluidic microdevice and demonstrated its advantages for measuring cell and nucleus under different situations. Figure 1 shows the schematic of the microfluidic channel. The components and assembly of the acoustofluidic microdevice are shown in Figure 2. Figure 3 shows the setup of the measureme...

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Measurement of the Compressibility of Cell and Nucleus Based on Acoustofluidic Microdevice

Here a protocol is presented to build a fast and non-destructive system for measuring cell or nucleus compressibility based on acoustofluidic microdevice. Changes in mechanical properties of tumor cells after epithelial-mesenchymal transition or ionizing radiation were investigated, demonstrating the application prospect of this method in scientific research and clinical practice.

Cell mechanics play an important role in tumor metastasis, malignant transformation of cells, and radiosensitivity. During these processes, studying the ...

measurement-compressibility-cell-nucleus-based-on-acoustofluidic

Research

JoVE Journal

Cancer Research

1.6K Views.  Sun Yat-sen University. Celmechanica speelt een belangrijke rol bij tumormetastase, kwaadaardige transformatie van cellen en radiosensitiviteit. Acoustofluidische methode kan snelle en niet-destructieve metingen realiseren in zwevende toestanden. Deze techniek kan worden gebruikt voor het spiegelen van de verandering van zelfdrukbaarheid tijdens epitheliale naar mesenchymale overgang en het scheiden van circulerende tumorcellen, wat de toepassingsvooruitzichten in kankerdiagnostiek laat zien.Bind om te beginn...

Video: Measurement of the Compressibility of Cell and Nucleus Based on Acoustofluidic Microdevice

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Bioengineering

A Direct Force Probe for Measuring Mechanical Integration Between the Nucleus and the Cytoskeleton

The mechanical properties of the nucleus determine its response to mechanical forces generated in cells. Because the nucleus is molecularly continuous with the cytoskeleton, methods are needed to probe its mechanical behavior in adherent cells. Here, we discuss the direct force probe (DFP) as a tool to apply force directly to the nucleus in a living adherent cell. We attach a narrow micropipette to the nuclear surface with suction. The micropipette is translated away from the nucleus, which causes the nucleus to deform and translate. When the restoring force is equal to the suction force, the nucleus detaches and elastically relaxes. Because the suction pressure is precisely known, the force on the nuclear surface is known. This method has revealed that nano-scale forces are sufficient to deform and translate the nucleus in adherent cells, and identified cytoskeletal elements that enable the nucleus to resist forces. The DFP can be used to dissect the contributions of cellular and nuclear components to nuclear mechanical properties in living cells.

In this protocol, we describe a micropipette method to directly apply a controlled force to the nucleus in a living cell. This assay allows interrogation of nuclear mechanical properties in the living, adherent cell.

The mechanical properties of the nucleus determine its response to mechanical forces generated in cells. Because the nucleus is molecularly continuous ...

Biochemistry

A Microfluidic Technique to Probe Cell Deformability

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an efficient manner. Typically, data for ~102 cells can be acquired within a 1 hr experiment. An automated image analysis program enables efficient post-experiment analysis of image data, enabling processing to be complete within a few hours. Our device geometry is unique in that cells must deform through a series of micron-scale constrictions, thereby enabling the initial deformation and time-dependent relaxation of individual cells to be assayed. The applicability of this method to human promyelocytic leukemia (HL-60) cells is demonstrated. Driving cells to deform through micron-scale constrictions using pressure-driven flow, we observe that human promyelocytic (HL-60) cells momentarily occlude the first constriction for a median time of 9.3 msec before passaging more quickly through the subsequent constrictions with a median transit time of 4.0 msec per constriction. By contrast, all-trans retinoic acid-treated (neutrophil-type) HL-60 cells occlude the first constriction for only 4.3 msec before passaging through the subsequent constrictions with a median transit time of 3.3 msec. This method can provide insight into the viscoelastic nature of cells, and ultimately reveal the molecular origins of this behavior.

We demonstrate a microfluidics-based assay to measure the timescale for cells to transit through a sequence of micron-scale constrictions.

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an ...

A Microfluidic Platform for Precision Small-volume Sample Processing and Its Use to Size Separate Biological Particles with an Acoustic Microdevice

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. However, to achieve robust sample manipulation it is necessary to address device integration with the macroscale environment. To realize repeatable, sensitive particle separation with microfluidic devices, this protocol presents a complete automated and integrated microfluidic platform that enables precise processing of 0.15&#8211;1.5 ml samples using microfluidic devices. Important aspects of this system include modular device layout and robust fixtures resulting in reliable and flexible world to chip connections, and fully-automated fluid handling which accomplishes closed-loop sample collection, system cleaning and priming steps to ensure repeatable operation. Different microfluidic devices can be used interchangeably with this architecture. Here we incorporate an acoustofluidic device, detail its characterization, performance optimization, and demonstrate its use for size-separation of biological samples. By using real-time feedback during separation experiments, sample collection is optimized to conserve and concentrate sample. Although requiring the integration of multiple pieces of equipment, advantages of this architecture include the ability to process unknown samples with no additional system optimization, ease of device replacement, and precise, robust sample processing.

This protocol describes a system architecture for performing automated small volume (0.15&#8211;1.5 ml) particle separations using a microfluidic device, and discusses methods to optimize acoustofluidic device performance and operation.

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. ...

Atomic Force Microscopy Cantilever-Based Nanoindentation: Mechanical Property Measurements at the Nanoscale in Air and Fluid

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by the probe tip and its contact area with the sample can be quantified, it is possible to determine the nanoscale mechanical properties (e.g., elastic or Young&#39;s modulus) of the surface being probed. A detailed procedure for performing quantitative AFM cantilever-based nanoindentation experiments is provided here, with representative examples of how the technique can be applied to determine the elastic moduli of a wide variety of sample types, ranging from kPa to GPa. These include live mesenchymal stem cells (MSCs) and nuclei in physiological buffer, resin-embedded dehydrated loblolly pine cross-sections, and Bakken shales of varying composition.
Additionally, AFM cantilever-based nanoindentation is used to probe the rupture strength (i.e., breakthrough force) of phospholipid bilayers. Important practical considerations such as method choice and development, probe selection and calibration, region of interest identification, sample heterogeneity, feature size and aspect ratio, tip wear, surface roughness, and data analysis and measurement statistics are discussed to aid proper implementation of the technique. Finally, co-localization of AFM-derived nanomechanical maps with electron microscopy techniques that provide additional information regarding elemental composition is demonstrated.

Quantifying the contact area and force applied by an atomic force microscope (AFM) probe tip to a sample surface enables nanoscale mechanical property determination. Best practices to implement AFM cantilever-based nanoindentation in air or fluid on soft and hard samples to measure elastic modulus or other nanomechanical properties are discussed.

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by ...

Engineering

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Developmental Biology

Biophysical Assays to Probe the Mechanical Properties of the Interphase Cell Nucleus: Substrate Strain Application and Microneedle Manipulation

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the physical properties of the nucleus contribute significantly to the overall biomechanical behavior of cells under physiological and pathological conditions. For example, in migrating neutrophils and invading cancer cells, nuclear stiffness can pose a major obstacle during extravasation or passage through narrow spaces within tissues.1 On the other hand, the nucleus of cells in mechanically active tissue such as muscle requires sufficient structural support to withstand repetitive mechanical stress. Importantly, the nucleus is tightly integrated into the cellular architecture; it is physically connected to the surrounding cytoskeleton, which is a critical requirement for the intracellular movement and positioning of the nucleus, for example, in polarized cells, synaptic nuclei at neuromuscular junctions, or in migrating cells.2 Not surprisingly, mutations in nuclear envelope proteins such as lamins and nesprins, which play a critical role in determining nuclear stiffness and nucleo-cytoskeletal coupling, have been shown recently to result in a number of human diseases, including Emery-Dreifuss muscular dystrophy, limb-girdle muscular dystrophy, and dilated cardiomyopathy.3 To investigate the biophysical function of diverse nuclear envelope proteins and the effect of specific mutations, we have developed experimental methods to study the physical properties of the nucleus in single, living cells subjected to global or localized mechanical perturbation. Measuring induced nuclear deformations in response to precisely applied substrate strain application yields important information on the deformability of the nucleus and allows quantitative comparison between different mutations or cell lines deficient for specific nuclear envelope proteins. Localized cytoskeletal strain application with a microneedle is used to complement this assay and can yield additional information on intracellular force transmission between the nucleus and the cytoskeleton. Studying nuclear mechanics in intact living cells preserves the normal intracellular architecture and avoids potential artifacts that can arise when working with isolated nuclei. Furthermore, substrate strain application presents a good model for the physiological stress experienced by cells in muscle or other tissues (e.g., vascular smooth muscle cells exposed to vessel strain). Lastly, while these tools have been developed primarily to study nuclear mechanics, they can also be applied to investigate the function of cytoskeletal proteins and mechanotransduction signaling.

We present two independent, microscope-based tools to measure the induced nuclear and cytoskeletal deformations in single, living adherent cells in response to global or localized strain application. These techniques are used to determine nuclear stiffness (i.e., deformability) and to probe intracellular force transmission between the nucleus and the cytoskeleton.

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the ...

Biology

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record forces curves on 1000 cells in 4 hours automatically. To maintain a 4 hour analysis time, the number of force curves per cell is reduced to 9 or 16. The method combines a Jython based program and a strategy for assembling cells on defined patterns. The program, implemented on a commercial Bio-AFM, can center the tip on the first cell of the array and then move, automatically, from cell to cell while recording force curves on each cell. Using this methodology, it is possible to access the biophysical parameters of the cells such as their rigidity, their adhesive properties, etc. With the automation and the large number of cells analyzed, one can access the behavior of the cell population. This is a breakthrough in the Bio-AFM field where data have, so far, been recorded on only a few tens of cells.

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

This protocol aims to automate AFM measurements on hundreds of microbial cells. First, microbes are immobilized into PDMS stamp microstructures and then force spectroscopy measurements are performed automatically on hundreds of immobilized cells.

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record ...

Assembly and Operation of an Acoustofluidic Device for Enhanced Delivery of Molecular Compounds to Cells

Efficient intracellular delivery of biomolecules is required for a broad range of biomedical research and cell-based therapeutic applications. Ultrasound-mediated sonoporation is an emerging technique for rapid intracellular delivery of biomolecules. Sonoporation occurs when cavitation of gas-filled microbubbles forms transient pores in nearby cell membranes, which enables rapid uptake of biomolecules from the surrounding fluid. Current techniques for in vitro sonoporation of cells in suspension are limited by slow throughput, variability in the ultrasound exposure conditions for each cell, and high cost. To address these limitations, a low-cost acoustofluidic device has been developed which integrates an ultrasound transducer in a PDMS-based fluidic device to induce consistent sonoporation of cells as they flow through the channels in combination with ultrasound contrast agents. The device is fabricated using standard photolithography techniques to produce the PDMS-based fluidic chip. An ultrasound piezo disk transducer is attached to the device and driven by a microcontroller. The assembly can be integrated inside a 3D-printed case for added protection. Cells and microbubbles are pushed through the device using a syringe pump or a peristaltic pump connected to PVC tubing. Enhanced delivery of biomolecules to human T cells and lung cancer cells is demonstrated with this acoustofluidic system. Compared to bulk treatment approaches, this acoustofluidic system increases throughput and reduces variability, which can improve cell processing methods for biomedical research applications and manufacturing of cell-based therapeutics.

This protocol describes the assembly and operation of a low-cost acoustofluidic device for rapid molecular delivery to cells via sonoporation induced by ultrasound contrast agents.

Efficient intracellular delivery of biomolecules is required for a broad range of biomedical research and cell-based therapeutic applications. ...

Injection of Porcine Adipose Tissue-Derived Stroma Cells via Waterjet Technology

Urinary incontinence (UI) is a highly prevalent condition characterized by the deficiency of the urethral sphincter muscle. Regenerative medicine branches, particularly cell therapy, are novel approaches to improve and restore the urethral sphincter function. Even though injection of active functional cells is routinely performed in clinical settings by needle and syringe, these approaches have significant disadvantages and limitations. In this context, needle-free waterjet (WJ) technology is a feasible and innovative method that can inject viable cells by visual guided cystoscopy in the urethral sphincter. In the present study, we used WJ to deliver porcine adipose tissue-derived stromal cells (pADSCs) into cadaveric urethral tissue and subsequently investigated the effect of WJ delivery on cell yield and viability. We also assessed the biomechanical features (i.e., elasticity) by atomic force microscopy (AFM) measurements. We showed that WJ delivered pADSCs were significantly reduced in their cellular elasticity. The viability was significantly lower compared to controls but is still above 80%.

We present a method of cell injection via needle free waterjet technology coupled with a sequela of post-delivery investigations in terms of cellular viability, proliferation, and elasticity measurements.

Measurement of the Compressibility of Cell and Nucleus Based on Acoustofluidic Microdevice

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