We thank Nurul Aisha Zainal Abidin and Laura Moldovan for additional donor recruitment, blood collection, and phlebotomy support. We thank Tomas Anderson and Arian Nasser for organizing the equipment and reagents. This research was funded by the Australian Research Council (ARC) Discovery Project (DP200101970-L.A.J.); the National Health and Medical Research Council (NHMRC) of Australia Ideas Grant (APP2003904-L.A.J.); NHMRC Equipment Grant-L.A.J.; NSW Cardiovascular Capacity Building Program (Early-Mid Career Researcher Grant-L.A.J.); NSW CVRN-VCCRI Research Innovation Grant; Office of Global and Research Engagement (Sydney-Glasgow Partnership Collaboration Award-L.A.J.); L.A.J. is a National Heart Foundation Future Leader Fellow Level 2 (105863), and a Snow Medical Research Foundation Fellow (2022SF176).

This protocol follows the guidelines of and has been approved by the Human Research Ethics Committee of the University of Sydney. Informed consent was obtained from the donors for this study.
1. Human RBC isolation
NOTE: Step 1.1 should be performed by a trained phlebotomist using a protocol that has been approved by the Institutional Review Board.
<ol>
	<li>Withdraw 5 mL of blood from the median cubital vein using a 19 G butterfly needle.</li>
	<li>Transfer the collected blood into a 15 mL tube containing 1:200 enoxaparin to prevent clotting.</li>
	<li>Dilute 5 &#181;L of enoxaparin-anticoagulated blood in 1 mL of carbonate/bicarbonate buffer (C-buffer, pH = 8.5-9; Table of Materials).</li>
	<li>Centrifuge the diluted blood sample at 900 &#215; g for 1 min to sediment the RBCs. Carefully decant the supernatant without disturbing the pellet.</li>
	<li>Perform two washes of the RBC pellet with 1 mL of C-buffer (Table of Materials), centrifuging each time at 900 &#215; g for 1 min.</li>
	<li>Subsequently, wash the RBC pellet 2x with 1 mL of Tyrode&#39;s buffer using the same centrifugation conditions and then resuspend the final pellet in 1 mL of Tyrode&#39;s buffer to obtain the washed stock RBC suspension.</li>
</ol>
2. Calcium indicator loading
<ol>
	<li>Adjust the concentration of the washed stock RBC solution to 10 &#215; 106 cells/mL in Tyrode&#39;s buffer, based on the cell count obtained using an automatic cell counter (Table of Materials).</li>
	<li>Label the calcium inside the RBCs by incubating with 16.67 &#181;M Cal-520 AM, a calcium-sensitive dye, while agitating on a rotary tube mixer for 1 h.</li>
	<li>Dilute the RBCs in Tyrode&#39;s buffer containing 0.5% bovine serum albumin (BSA) at a 1:50 ratio. The cells are now ready for experimental use.</li>
</ol>
3. Micropipette fabrication
<ol>
	<li>Mount the borosilicate glass capillary tube (1 mm outer diameter x 0.6 mm inner diameter) onto the P-1000 micropipette puller to produce two corresponding micropipettes with closed tips at the pulling site using the preset pulling program. For this setup, use the following pulling program values: heat 516, pull 150, velocity 75, time 250, and pressure 500. 
	NOTE: Heating and pulling parameters set in the pulling program can be customized and are dependent on the desired settings of the experimental design12. CHECKPOINT (see Supplemental Table S1).</li>
	<li>Open the closed tip by mounting one of the close-ended micropipettes procured after pulling onto the micropipette cutter. Adjust the heating temperature to approximately 50-60 &#176;C.</li>
	<li>Locate the micropipette using a 10x eyepiece. Move the micropipette close to the borosilicate glass bead by using the knobs for adjustment.</li>
	<li>Change the eyepiece to 30x before positioning the micropipette as close to the borosilicate glass bead as possible without bending the pipette tip.</li>
	<li>Soften the borosilicate glass bead using heat by stepping onto the heating pedal. Gently insert the raw closed micropipette tip into the softened bead until the desired endpoint,&#160;the opening diameter, has been reached.</li>
	<li>Release the foot pedal&#160;and let the glass bead cool down. Make sure the tip of the micropipette always remains inside the bead. 
	NOTE: Inserting the tip further leads to larger opening diameters.</li>
	<li>Gently extract the micropipette, leading to a clear straight cut on the closed micropipette. Confirm that the final diameter of the capillary is 1 &#181;m. 
	&#8203;NOTE: CHECKPOINT (see Supplemental Table S1)</li>
</ol>
4. Cell chamber preparation
<ol>
	<li>Use a diamond pencil to divide a standard 40 mm x 22 mm x 0.17 mm glass coverslip into three equal strips.</li>
	<li>Adhere one piece of the cut glass coverslip to the bottom of a homemade chamber holder with vacuum grease. 
	NOTE: The chamber holder consists of two metal (copper/aluminum) squares that are linked by a curved handle. The distance between the metal blocks must span less than 40 mm for the cut coverslip to adhere to the holder to form a parallel chamber.</li>
	<li>Adhere the second piece of the cut glass coverslip to the top of the homemade chamber holder with vacuum grease . 
	NOTE: CHECKPOINT (see Supplemental Table S1)</li>
	<li>Inject 200 &#181;L of the labeled RBC suspension between two coverslips using a 200 &#181;L pipette gun (Figure 1).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66265/66265fig01.jpg" /> 
Figure 1: Illustration of the cell chamber. Two cut pieces of a 40 mm x 22 mm x 0.17 mm glass cover slip are adhered to the chamber holder using grease. Between the two cut glass coverslips, approximately 200 &#181;L of the cell solution in Tyrode&#39;s Buffer is seeded. <a href="https://www.jove.com/files/ftp_upload/66265/66265fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Micropipette aspiration assembly
<ol>
	<li>Mount the cell chamber onto the holder stage present on the microscope platform. Adjust the position so that the cell chamber is directly above the objective (Figure 2B).</li>
	<li>Lower the micropipette holder to below the fluid level of the connected water reservoir.</li>
	<li>Inject either demineralized water or Tyrode&#39;s buffer into the fabricated micropipette and carefully remove all air bubbles using a syringe coupled with a 34 G needle (see Table of Materials).</li>
	<li>Unscrew the end of the micropipette holder halfway and allow the water to drip from the micropipette holder for a few seconds. 
	NOTE: CHECKPOINT (see Supplemental Table S1)</li>
	<li>Insert the&#160;micropipette into the holder tip. Tighten the holder screw to ensure the micropipette is fixed.</li>
	<li>Insert the micropipette into the cell chamber and locate the micropipette and RBCs under the microscope. Use the micromanipulator to adjust the position.</li>
	<li>Lower the micropipette tip further to ensure the tip is leveled with the located RBC. 
	NOTE: CHECKPOINT (see Supplemental Table S1)</li>
	<li>Zero the hydraulic pressure at the micropipette tip by adjusting the height of the water reservoir. Then, slightly raise the water reservoir to generate a subtle positive pressure at the tip.</li>
</ol>
6. Perform the fluorescence-coupled micropipette aspiration assay
<ol>
	<li>Turn on the 488 nm fluorescent excitation light source. Do not switch on the fluorescence shutter at this stage to avoid photobleaching (Figure 2C). Turn on the fluorescence camera and the transmitted camera. 
	NOTE: Both cameras are operated using the appropriate software (see Table of Materials).</li>
	<li>Set up the desired exposure time (100 ms for both cameras in this study), region of interest (ROI), binning size (none for this study) for both cameras in the software. Open up the multi-dimension acquisition panel to set up the acquisition frame number, 2,000 for this study, and saving directory. 
	NOTE: The acquisition frame number is dependent on the desired number of aspiration events that are to be recorded. For 1 aspiration event, the range of the acquisition number should be set within 100-500, which is approximately 10-50 s.</li>
	<li>Find the micropipette under the field of view using the micromanipulator.</li>
	<li>Turn on the pneumatic pressure clamp, including the control box and the clamp system (Figure 2A). Make sure the control box is in the EXTRNL mode. Compensate any offset pressure inside the system by slowly rotating the knob.</li>
	<li>Turn on the separate software that controls the pneumatic clamp. The software has an electrical control panel to control the discrete analog input to the clamp system. The pressure is controlled with a 20 mV/mmHg conversion factor.</li>
	<li>Zero the pressure inside the system. Carefully relocate the micropipette close to the RBCs. Adjust the water reservoir position until a subtle positive pressure is noticed at the micropipette tip.</li>
	<li>Start the acquisition in the camera-operating software. Switch on the fluorescence shutter.</li>
	<li>Aspirate an RBC by typing in the calculated voltage magnitude into the control panel to reach the desired pressure. 
	NOTE: The pressure for aspirating an RBC is typically in the range of &#916;p = -5 to -40 mmHg. There should be a noticeable tongue elongation within the micropipette tip (Figure 2D).</li>
	<li>Hold the pressure for a preset period; then, release the pressure.</li>
	<li>Move the micropipette to pick up the next cell and repeat the experiment.</li>
</ol>
7. Fluorescence intensity analysis
<ol>
	<li>Load the saved fluorescence images into the analysis software.</li>
	<li>Adjust the intensity threshold using the display adjustment tab. Do this by either manually inputting the values or using the slider to ensure the fluorescence images show a clear contrast of the cell in the analysis software (see Supplemental File 1-Supplemental Figure S1).</li>
	<li>Scroll to the timeline at the bottom of the software. Locate the designated aspiration event.</li>
	<li>Click Add new surfaces. Define the analysis ROI. 
	NOTE: The software provides a guided five-step process to adjust and complete the segmentation (see Supplemental File 1-Supplemental Figure S2 and Supplemental Figure S3). 
	NOTE: Keep the ROI as small as possible to save computational resources.</li>
	<li>Use the background subtraction slider and adjust the segmentation threshold using the slider to obtain the best segmentation outcome. 
	NOTE: This means that apart from the aspiration event, the background should be segmented as accurately as possible (see Supplemental File 1-Supplemental Figure S4 and Supplemental Figure S5).</li>
	<li>Add an area filter to exclude background noises (see Supplemental File 1-Supplemental Figure S6). 
	NOTE: This is completed in the post process stage.</li>
	<li>Select the statistics tab | detailed tab | average values tab. Scroll to find and select the intensity mean (see Supplemental File 1-Supplemental Figure S7).</li>
	<li>Export the fluorescence signal trace over time to a .csv file.</li>
	<li>Open the exported csv file. Subtract the background signals, Fb, from all measurements.</li>
	<li>Calculate the calcium intensity change, &#916;Fmax, using equation (1): 
	<img alt="Equation 1" src="/files/ftp_upload/66265/66265eq01.jpg" style="margin: auto;" />&#160; &#160; &#160;(1) 
	Where&#160;&#916;Fmax is the maximum calcium intensity change, Fb, is the background intensity, and F0 is the resting intensity.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66265/66265fig02.jpg" /> 
Figure 2: Fluorescence-coupled micropipette aspiration assembly. (A)&#160;An overview of the fMPA hardware system incorporating the inverted microscope combined with the brightfield and fluorescence cameras. The left side of the image depicts the homemade water manometer and the control box that allows to precisely tune the pressure of the pneumatic pressure pump. (B)&#160;The microscope stage depicting the experiment cell chamber and micromanipulator system with a single micropipette. (C)&#160;Schematic of the fMPA system setup. Concurrent imaging of brightfield (yellow) and fluorescence (blue emission, green excitation) signals utilizing two dichroic mirrors to direct the light paths from the fluorescence light source (blue) to the target, then to the cameras for imaging (green). (D)&#160;The top row depicts the brightfield images whereas the bottom row demonstrates the fluorescence images. The left represents the position of the micropipette before aspiration when the RBC&#160;is at rest.The middle column snapshots the aspiration process where the RBC experiences a negative pressure of -40 mmHg. The right depicts the cell morphology after experiencing the negative aspiration pressure. Scale bar = 5 &#181;m. Abbreviations: fMPA = Fluorescence-coupled Micropipette Aspiration; DM = dichroic mirror; RBC = red blood cell. <a href="https://www.jove.com/files/ftp_upload/66265/66265fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Single-Cell Mechanobiology Using Fluorescence Micropipette Aspiration

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I., Husson, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+cell+adhesion+by+using+micropipette+aspiration.">Characterizing cell adhesion by using micropipette aspiration.</a> Biophysical Journal. 109 (2), 209-219 (2015).</li><li>Henriksen, J. R., Ipsen, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+membrane+elasticity+by+micro-pipette+aspiration.">Measurement of membrane elasticity by micro-pipette aspiration.</a> The European Physical Journal E. 14 (2), 149-167 (2004).</li><li>Hochmuth, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Micropipette+aspiration+of+living+cells.">Micropipette aspiration of living cells.</a> Journal of Biomechanics. 33 (1), 15-22 (2000).</li><li>Pu, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Micropipette+aspiration+of+single+cells+for+both+mechanical+and+electrical+characterization.">Micropipette aspiration of single cells for both mechanical and electrical characterization.</a> IEEE Transactions on Biomedical Engineering. 66 (11), 3185-3191 (2019).</li><li>Guevorkian, K., Ma&#238;tre, J. -. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+10+-+Micropipette+aspiration:+A+unique+tool+for+exploring+cell+and+tissue+mechanics+in+vivo.">Chapter 10 - Micropipette aspiration: A unique tool for exploring cell and tissue mechanics in vivo.</a> Methods in Cell Biology. , 187-201 (2017).</li><li>Biro, M., Ma&#238;tre, J. -. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+14+-+Dual+pipette+aspiration:+A+unique+tool+for+studying+intercellular+adhesion.">Chapter 14 - Dual pipette aspiration: A unique tool for studying intercellular adhesion.</a> Methods in Cell Biology. 125, 255-267 (2015).</li></ol>

The authors declare that they have no competing interests to report regarding the present study.

Micropipette aspiration assays embody a refined methodology, deploying substantial pressure modulation, exact spatial orchestration, and reliable temporal discernment to probe the profound intricacies of cellular biomechanics. This study places particular emphasis on the application of fMPA as a crucial tool for unveiling the nuanced mechanosensitive responses showcased by RBCs under varying stimuli. The concurrent use of brightfield and fluorescence signals enabled a multifaceted exploration of cellular phenomena, advan...

The unfolding discoveries in the world of cellular behaviors have accentuated the role of mechanical stimuli, such as tension, fluid shear stress, compression, and substrate stiffness, in dictating dynamic cellular activities such as adhesion, migration, and differentiation. These mechanobiological aspects are of paramount importance in elucidating how cells interact with and respond to their physiological environments, impacting various biological processes1,2.
Over the past decade, micropipette-based aspiration assays have stood out as a versatile tool in studying diverse cellular responses to mechanical stimuli. This technique offers valuable insights into the intrinsic mechanical properties of living cells at the single-cell level, including cellular elastic modulus, stiffness, and cortical&#160;tension. These assays enable the measurement of various mechanical parameters, such as cell membrane&#160;tension, pressure exerted on the cell membrane, and cortical tension (summarized in Table 1). Studying the aspirational forces has enriched our understanding of how they influence cellular functions and processes, particularly in the realm of membrane dynamics, including fragmentation, elongation, and budding3,4.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Mechanical Parameter</td>
			<td>Description</td>
			<td>Seminal Approaches</td>
		</tr>
		<tr>
			<td>Cell Stiffness</td>
			<td>Measurement of a cell&#39;s mechanical rigidity and elasticity.</td>
			<td>Aspiration of the cell membrane and analysis of deformation response to the negative pressure20,21.</td>
		</tr>
		<tr>
			<td>Adhesion Strength</td>
			<td>Evaluation of how strongly cells adhere to surfaces.</td>
			<td>Application of controlled suction to detach adhered cells from a substrate2,22.</td>
		</tr>
		<tr>
			<td>Membrane Tension</td>
			<td>Assessment of the tension or stress within cell membranes.</td>
			<td>Measurement of the membrane deformation in response to applied pressure23,24.</td>
		</tr>
		<tr>
			<td>Viscoelastic Properties</td>
			<td>Characterization of a cell&#39;s combined viscous and elastic behavior.</td>
			<td>Analysis of the time-dependent deformation response to aspiration23,25.</td>
		</tr>
		<tr>
			<td>Deformability</td>
			<td>Determination of how easily a cell can change shape.</td>
			<td>Evaluation of the extent of deformation under controlled suction20,24.</td>
		</tr>
		<tr>
			<td>Surface Tension</td>
			<td>Measurement of the tension at the cell&#39;s surface.</td>
			<td>Assessment of the pressure required to form a micropipette membrane protrusion26.</td>
		</tr>
		<tr>
			<td>Cell-Material Interaction</td>
			<td>Study of interactions between cells and materials or substrates.</td>
			<td>Aspiration of cells in contact with different materials and observation of interactions2,24.</td>
		</tr>
		<tr>
			<td>Cell-Cell Interaction</td>
			<td>Examination of interactions between neighboring cells.</td>
			<td>Aspiration of a group of cells and analysis of their intercellular forces27.</td>
		</tr>
	</tbody>
</table>
Table 1: Mechanical parameters characterized by the micropipette aspiration assay.
The micropipette-based aspiration technique has been widely used to study red blood cells (RBCs), assessing the deformability and various mechanical characteristics of RBCs, which is essential in understanding their function in the circulatory system. RBCs exhibit remarkable adaptability, preserving their mechanical versatility against deformation when navigating through the intricate capillary network and inter-endothelial clefts5,6. During this journey, RBCs must traverse through passages as narrow as 0.5-1.0 &#181;m, subjecting themselves to a multitude of mechanical forces, including tension and compression7,8,9. They also have high sensitivity to the shear stress generated by blood flow during circulation10. These processes promote the activation of regulatory mechanisms involving calcium influx, a crucial signaling event with well-established roles in cellular responses to mechanical stimuli11,12. The complex mechanisms governing the calcium-mediated mechanosensing remain compelling subjects of ongoing investigation.
In this context, the fMPA stands as an effective approach to reveal the extent of calcium mobilization under precisely controlled mechanical forces, allowing for the simultaneous application of mechanical modulation (using the micropipette aspiration system) and visualization of calcium intensity (using fluorescent indicators). It particularly mimics the physiological scenario when the RBC travels through narrowing blood vessels. It is worth noting that the fMPA system we developed can generate pressure with a resolution of 1 mmHg. The implemented high-speed camera can achieve a temporal resolution of 100 ms and a spatial resolution at the submicron-meter level. These configurations ensure the precise application of mechanical forces to live cells and simultaneously capture the resulting cellular signaling. Moreover, due to the integrative engineered nature of this setup, the micropipette aspiration assay can be readily adapted to complement other equipment or techniques, enabling further exploration of the intricacies of cell mechanics. This versatility stands as an additional advantage of this approach.

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			<td>&#181;Manager</td>
			<td>Micro-Manager</td>
			<td></td>
			<td>Version 2.0.0</td>
		</tr>
		<tr>
			<td>1 mL Syringe&#160;</td>
			<td>Terumo</td>
			<td>210320D</td>
			<td>Cooperate with the Microfil&#160;</td>
		</tr>
		<tr>
			<td>200 &#181;L Pipette&#160;</td>
			<td>Eppendorf&#160;</td>
			<td>3123000055</td>
			<td>Red clood cell preparation</td>
		</tr>
		<tr>
			<td>22 x 40 mm Cover Slips</td>
			<td>Knittel Glass&#160;</td>
			<td>MS0014</td>
			<td>Cell chamber assembly</td>
		</tr>
		<tr>
			<td>50 mL Syringe&#160;</td>
			<td>Terumo</td>
			<td>220617E</td>
			<td>Connect to the water tower</td>
		</tr>
		<tr>
			<td>Calcium Chloride (CaCl2)</td>
			<td>Sigma-Aldrich</td>
			<td>C1016</td>
			<td>Tryode&#39;s&#160; buffer preparation - 12 mM NaHCO3, 10 mM HEPES, 0.137 M NaCl, 2.7 mM KCl, and 5.5 mM D-glucose supplemented with 1 mM CaCl2. Final pH = 7.2</td>
		</tr>
		<tr>
			<td>Centrifuge 5425</td>
			<td>Eppendorf&#160;</td>
			<td>5405000280</td>
			<td>Red clood cell preparation</td>
		</tr>
		<tr>
			<td>Clexane</td>
			<td>Sigma-Aldrich</td>
			<td>1235820</td>
			<td>To prevent clotting of the collected blood. 10,000 U/mL</td>
		</tr>
		<tr>
			<td>DAQami</td>
			<td>Diligent</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence light source</td>
			<td>CoolLED</td>
			<td>pE-300</td>
			<td>Micropipette aspiration hardware system</td>
		</tr>
		<tr>
			<td>Glass capillary</td>
			<td>Narishige</td>
			<td>G-1</td>
			<td>Micropipette manufacture</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G8270</td>
			<td>Tryode&#39;s&#160; buffer preparation - 12 mM NaHCO3, 10 mM HEPES, 0.137 M NaCl, 2.7 mM KCl, and 5.5 mM D-glucose supplemented with 1 mM CaCl2. Final pH = 7.2</td>
		</tr>
		<tr>
			<td>Hepes</td>
			<td>Thermo Fisher</td>
			<td>15630080</td>
			<td>Tryode&#39;s&#160; buffer preparation - 12 mM NaHCO3, 10 mM HEPES, 0.137 M NaCl, 2.7 mM KCl, and 5.5 mM D-glucose supplemented with 1 mM CaCl2. Final pH = 7.2</td>
		</tr>
		<tr>
			<td>High speed GigE camera</td>
			<td>Manta</td>
			<td>G-040B</td>
			<td>Micropipette aspiration hardware system</td>
		</tr>
		<tr>
			<td>High speed pressure clamp</td>
			<td>Scientific Instrument</td>
			<td>HSPC-2-SB</td>
			<td>Cooperate with the pressure pump</td>
		</tr>
		<tr>
			<td>High speed pressure clamp head stage&#160;</td>
			<td>Scientific Instrument</td>
			<td>HSPC-2-SB</td>
			<td>Cooperate with the pressure pump</td>
		</tr>
		<tr>
			<td>Imaris</td>
			<td>Oxford Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted Microscopy&#160;</td>
			<td>Olympus&#160;</td>
			<td>Olympus IX83</td>
			<td>Micropipette aspiration hardware system</td>
		</tr>
		<tr>
			<td>Microfil&#160;</td>
			<td>World Precision Instruments&#160;</td>
			<td>MF34G-5</td>
			<td>34 G (67 mm Long) 
			Revome air bubble in the cut micropipette and test the opening of the pipette tip&#160;</td>
		</tr>
		<tr>
			<td>Micropipette Puller&#160;</td>
			<td>Sutter instrument</td>
			<td>P1000</td>
			<td>Micropipette manufacture&#160;</td>
		</tr>
		<tr>
			<td>Milli Q EQ 7000 Ultrapure Water Purification System</td>
			<td>Merck Millipore</td>
			<td>ZEQ7000T0C</td>
			<td>Carbonate/bicarbonate buffer &#38; Tryode&#39;s buffer preparation</td>
		</tr>
		<tr>
			<td>Pipette microforge&#160;</td>
			<td>Narishige</td>
			<td>MF-900</td>
			<td>Micropipette manufacture</td>
		</tr>
		<tr>
			<td>Potassium Chloride (KCl)</td>
			<td>Sigma-Aldrich</td>
			<td>P9541</td>
			<td>Tryode&#39;s&#160; buffer preparation - 12 mM NaHCO3, 10 mM HEPES, 0.137 M NaCl, 2.7 mM KCl, and 5.5 mM D-glucose supplemented with 1 mM CaCl2. Final pH = 7.2</td>
		</tr>
		<tr>
			<td>Pressue Pump&#160;</td>
			<td>Scientific Instrument</td>
			<td>PV-PUMP</td>
			<td>Induce controlled pressure during experiment</td>
		</tr>
		<tr>
			<td>Prime 95B Camera&#160;</td>
			<td>Photometrics</td>
			<td>Prime 95B sCMOS</td>
			<td>Flourscent imaging</td>
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		<tr>
			<td>Rotary wheel remote unit&#160;</td>
			<td>Sensapex&#160;</td>
			<td>uM-RM3</td>
			<td>Control panel for micropipette position adjustment&#160;</td>
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		<tr>
			<td>Scepter 3.0 Handheld Cell Counter</td>
			<td>Merck Millipore</td>
			<td>PHCC340KIT</td>
			<td>Automatic cell counter</td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate (NaHCO3)</td>
			<td>Sigma-Aldrich</td>
			<td>S5761</td>
			<td>Carbonate/bicarbonate buffer preparation - 2.65 g of NaHCO3 with 2.1 g of Na2CO3 in 250 mL of Mili Q water - Final pH = 8-9.</td>
		</tr>
		<tr>
			<td>Sodium Carbonate (Na2CO3)</td>
			<td>Sigma-Aldrich</td>
			<td>S2127</td>
			<td>Carbonate/bicarbonate buffer preparation - 2.65 g of NaHCO3 with 2.1 g of Na2CO3 in 250 mL of Mili Q water - Final pH = 8-9.</td>
		</tr>
		<tr>
			<td>Sodium Chloride (NaCl)</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td>Tryode&#39;s&#160; buffer preparation - 12 mM NaHCO3, 10 mM HEPES, 0.137 M NaCl, 2.7 mM KCl, and 5.5 mM D-glucose supplemented with 1 mM CaCl2. Final pH = 7.2</td>
		</tr>
		<tr>
			<td>Sodium Phosphate Monobasic 
			Monohydrate (NaH2PO4 &#8226; H2O)</td>
			<td>Sigma-Aldrich</td>
			<td>S9638</td>
			<td>Tryode&#39;s&#160; buffer preparation - 12 mM NaHCO3, 10 mM HEPES, 0.137 M NaCl, 2.7 mM KCl, and 5.5 mM D-glucose supplemented with 1 mM CaCl2. Final pH = 7.2</td>
		</tr>
		<tr>
			<td>Touch screen control unit&#160;</td>
			<td>Sensapex&#160;</td>
			<td>uM-TSC</td>
			<td>Control panel for micropipette position adjustment&#160;</td>
		</tr>
		<tr>
			<td>X dry Objective&#160;</td>
			<td>Olympus&#160;</td>
			<td>Olympus 60x/0.70 LUCPlanFL</td>
			<td>Micropipette aspiration hardware system</td>
		</tr>
	</tbody>
</table>

fluorescence micropipette aspiration assay to investigate red blood cell mechanosensing

Micropipette aspiration assays have long been a cornerstone for the investigation of live-cell mechanics, offering insights into cellular responses to mechanical stress. This paper details an innovative adaptation of the fluorescence-coupled micropipette aspiration (fMPA) assay. The fMPA assay introduces the capability to administer precise mechanical forces while concurrently monitoring the live-cell mechanotransduction processes mediated by ion channels. The sophisticated setup incorporates a precision-engineered borosilicate glass micropipette connected to a finely regulated water reservoir and pneumatic aspiration system, facilitating controlled pressure application with increments as refined as &#177; 1 mmHg. A significant enhancement is the integration of epi-fluorescence imaging, allowing for the simultaneous observation and quantification of cell morphological changes and intracellular calcium fluxes during aspiration. The fMPA assay, through its synergistic combination of epi-fluorescence imaging with micropipette aspiration, sets a new standard for the study of cell mechanosensing within mechanically challenging environments. This multifaceted approach is adaptable to various experimental setups, providing critical insights into the single-cell mechanosensing mechanisms.

To establish micropipette aspiration assays, we first constructed a custom cell chamber comprising two metal squares (copper/aluminum) connected by a handle. Two third-cut glass coverslips (40 mm &#215; 7 mm &#215; 0.17 mm) were affixed to create a chamber filled with 200 &#181;L of RBCs suspended in Tyrode&#39;s Buffer. After introducing RBCs into the chamber, a tailored borosilicate micropipette was secured on a holder and carefully positioned within the chamber using a micro-manipulator. Subsequently, the micropipette...

Watch this Scientific Journal Video about Advancing Live-Cell Mechanobiology Through Fluorescence Microaspiration at JoVE.com

Author Spotlight: Advancing Live-Cell Mechanobiology Through Fluorescence Microaspiration

The exploration of cellular behavior under mechanical stress is pivotal for advances in cellular mechanics and mechanobiology. We introduce the Fluorescence Micropipette Aspiration (fMPA) technique, a novel method combining controlled mechanical stimulation with comprehensive analysis of intracellular signaling in single cells. This technique investigates new in-depth studies of live-cell mechanobiology.

Fluorescence Micropipette Aspiration Assay to Investigate Red Blood Cell Mechanosensing

Micropipette aspiration assays have long been a cornerstone for the investigation of live-cell mechanics, offering insights into cellular responses to ...

fluorescence-micropipette-aspiration-assay-to-investigate-red-blood

Research

JoVE Journal

Biology

767 Views.  The University of Sydney. The exploration of cellular behavior under mechanical stress is pivotal for advances in cellular mechanics and mechanobiology. We introduce the Fluorescence Micropipette Aspiration (fMPA) technique, a novel method combining controlled mechanical stimulation with comprehensive analysis of intracellular signaling in single cells. This technique investigates new in-depth studies of live-cell mechanobiology.

Video: Author Spotlight: Advancing Live-Cell Mechanobiology Through Fluorescence Microaspiration

A Behavioral Assay to Measure Responsiveness of Zebrafish to Changes in Light Intensities 

The optokinetic reflex (OKR) is a basic visual reflex exhibited by most vertebrates and plays an important role in stabilizing the eye relative to the visual scene. However, the OKR requires that an animal detect moving stripes and it is possible that fish that fail to exhibit an OKR may not be completely blind. One zebrafish mutant, the no optokinetic response c (nrc) has no OKR under any light conditions tested and was reported to be completely blind. Previously, we have shown that OFF-ganglion cell activity can be recorded in these mutants. To determine whether mutant fish with no OKR such as the nrc mutant can detect simple light increments and decrements we developed the visual motor behavioral assay (VMR). In this assay, single zebrafish larvae are placed in each well of a 96-well plate allowing the simultaneous monitoring of larvae using an automated video-tracking system. The locomotor responses of each larva to 30 minutes light ON and 30 minutes light OFF were recorded and quantified. WT fish have a brief spike of motor activity upon lights ON, known as the startle response, followed by return to lower-than baseline activity, called a freeze. WT fish also sharply increase their locomotor activity immediately following lights OFF and only gradually (over several minutes) return to baseline locomotor activity. The nrc mutants respond similarly to light OFF as WT fish, but exhibit a slight reduction in their average activity as compared to WT fish. Motor activity in response to light ON in nrc mutants is delayed and sluggish. There is a slow rise time of the nrc mutant response to light ON as compared to WT light ON response. The results indicate that nrc fish are not completely blind. Because teleosts can detect light through non-retinal tissues, we confirmed that the immediate behavioral responses to light-intensity changes require intact eyes by using the chokh (chk) mutants, which completely lack eyes from the earliest stages of development. In our VMR assay, the chk mutants exhibit no startle response to either light ON or OFF, showing that the lateral eyes mediate this behavior. The VMR assay described here complements the well-established OKR assay, which does not test the ability of zebrafish larvae to respond to changes in light intensities. Additionally, the automation of the VMR assay lends itself to high-throughput screening for defects in light-intensity driven visual responses.

A Behavioral Assay to Measure Responsiveness of Zebrafish to Changes in Light Intensities

We developed the Visual-Motor Response to quantitate the motor output of larval zebrafish in response to light increments and decrements. We also examined zebrafish vision mutants, including the no optokinetic response (nrc) mutants, which were thought to be completely blind when tested by another vision assay, the optokinetic reflex.

The optokinetic reflex (OKR) is a basic visual reflex exhibited by most vertebrates and plays an important role in stabilizing the eye relative to the ...

Preparation and Using Phantom Lesions to Practice Fine Needle Aspiration Biopsies

Currently, health workers including residents and fellows do not have a suitable phantom model to practice the fine- needle aspiration biopsy (FNAB) procedure. In the past, we standardized a model consisting of latex glove containing fresh cattle liver for practicing FNAB. However, this model is difficult to organize and prepare on short notice, with the procurement of fresh cattle liver being the most challenging aspect. Handling of liver with contamination-related problems is also a significant draw back. In addition, the glove material leaks after a few needle passes, with resulting mess.
We have established a novel simple method of embedding a small piece of sausage or banana in a commercially available silicone rubber caulk. This model allows the retention of vacuum seal and aspiration of material from the embedded specimen, resembling an actual FNAB procedure on clinical mass lesions.
The aspirated material in the needle hub can be processed similar to the specimens procured during an actual FNAB procedure, facilitating additional proficiency in smear preparation and staining.

Practicing of fine needle aspiration biopsies (FNAB) by trainees is relatively challenging, due to the lack of an easily available, appropriate lesion. Preparation of an AV phantom lesion for practicing the FNAB procedure and mastering proficiency is relatively easy.

Currently, health workers including residents and fellows do not have a suitable phantom model to practice the fine- needle aspiration biopsy (FNAB) ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Bimolecular Fluorescence Complementation (BiFC) Assay for Protein-Protein Interaction in Onion Cells Using the Helios Gene Gun

Investigation of gene function in diverse organisms relies on knowledge of how the gene products interact with each other in their normal cellular environment. The Bimolecular Fluorescence Complementation (BiFC) Assay1 allows researchers to visualize protein-protein interactions in living cells and has become an essential research tool. This assay is based on the facilitated association of two fragments of a fluorescent protein (GFP) that are each fused to a potential interacting protein partner. The interaction of the two protein partners would facilitate the association of the N-terminal and C-terminal fragment of GFP, leading to fluorescence. For plant researchers, onion epidermal cells are an ideal experimental system for conducting the BiFC assay because of the ease in obtaining and preparing onion tissues and the direct visualization of fluorescence with minimal background fluorescence. The Helios Gene Gun (BioRad) is commonly used for bombarding plasmid DNA into onion cells. We demonstrate the use of Helios Gene Gun to introduce plasmid constructs for two interacting Arabidopsis thaliana transcription factors, SEUSS (SEU) and LEUNIG HOMOLOG (LUH)2 and the visualization of their interactions mediated by BiFC in onion epidermal cells.

Bimolecular Fluorescence Complementation BiFC Assay for Protein-Protein Interaction in Onion Cells Using the Helios Gene Gun

This article illustrates how to properly use the BioRad Helios Gene Gun to introduce plasmid DNA into onion epidermal cells and how to test for protein-protein interactions in onion cells based on the principle of Bimolecular Fluorescence Complementation (BiFC)

Investigation of gene function in diverse organisms relies on knowledge of how the gene products interact with each other in their normal cellular ...

Cell Electrofusion Visualized with Fluorescence Microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

A Cell-to-cell Macromolecular Transport Assay in Planta Utilizing Biolistic Bombardment

Here, we present a simple and rapid protocol to detect and assess the extent of cell-to-cell macromolecular transport in planta. In this protocol, a fluorescently tagged-protein of interest is transiently expressed in plant tissue following biolistic delivery of its encoding DNA construct. The intra- and intercellular distribution of the tagged protein is then analyzed by confocal microscopy. We describe this technology in detail, providing step-by-step protocols to assay and evaluate the extent of symplastic protein transport in three plant species, Arabidopsis thaliana, Nicotiana benthamiana and N. tabacum (tobacco).

A Cell-to-cell Macromolecular Transport Assay in Planta Utilizing Biolistic Bombardment

Macromolecular trafficking between plant cells can be assessed by transiently expressing a fluorescently-tagged protein of interest and analyzing its intra- and intercellular distribution by confocal microscopy.

Here, we present a simple and rapid protocol to detect and assess the extent of cell-to-cell macromolecular transport in planta. In this protocol, a ...

Whole-mount Immunohistochemical Analysis for Embryonic Limb Skin Vasculature: a Model System to Study Vascular Branching Morphogenesis in Embryo

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching morphogenesis. We have developed the limb skin vasculature model to study vascular development in which a pre-existing primitive capillary plexus is reorganized into a hierarchically branched vascular network. Whole-mount confocal microscopy with multiple labelling allows for robust imaging of intact blood vessels as well as their cellular components including endothelial cells, pericytes and smooth muscle cells, using specific fluorescent markers. Advances in this limb skin vasculature model with genetic studies have improved understanding molecular mechanisms of vascular development and patterning. The limb skin vasculature model has been used to study how peripheral nerves provide a spatial template for the differentiation and patterning of arteries. This video article describes a simple and robust protocol to stain intact blood vessels with vascular specific antibodies and fluorescent secondary antibodies, which is applicable for vascularized embryonic organs where we are able to follow the process of vascular development.

We introduce a whole-mount immunohistochemistry and laser scanning confocal microscopy with multiple labelling for analyzing intricate vascular network formation in mouse embryonic limb skin.

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching ...

A Simplified System for Evaluating Cell Mechanosensing and Durotaxis In Vitro

The composition and mechanical properties of the extracellular matrix are highly variable between tissue types. This connective tissue stroma diversity greatly impacts cell behavior to regulate normal and pathologic processes including cell proliferation, differentiation, adhesion signaling and directional migration. In this regard, the innate ability of certain cell types to migrate towards a stiffer, or less compliant matrix substrate is referred to as durotaxis. This phenomenon plays an important role during embryonic development, wound repair and cancer cell invasion. Here, we describe a straightforward assay to study durotaxis, in vitro, using polydimethylsiloxane (PDMS) substrates. Preparation of the described durotaxis chambers creates a rigidity interface between the relatively soft PDMS gel and a rigid glass coverslip. In the example provided, we have used these durotaxis chambers to demonstrate a role for the cdc42/Rac1 GTPase activating protein, cdGAP, in mechanosensing and durotaxis regulation in human U2OS osteosarcoma cells. This assay is readily adaptable to other cell types and/or knockdown of other proteins of interests to explore their respective roles in mechanosignaling and durotaxis.

A Simplified System for Evaluating Cell Mechanosensing and Durotaxis In Vitro

Many mammalian cells preferentially migrate towards a more rigid matrix or substrate through durotaxis. The goal of this protocol is to provide a simple in vitro system that can be used to study and manipulate cell durotaxis behaviors by incorporating polydimethylsiloxane (PDMS) substrates of defined rigidity, interfacing with glass coverslips.

The composition and mechanical properties of the extracellular matrix are highly variable between tissue types. This connective tissue stroma diversity ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

A Simple Fluorescence-based Reporter Assay to Identify Cellular Components Required for Ricin Toxin A Chain (RTA) Trafficking in Yeast

Bacterial and plant A/B toxins exploit the natural trafficking pathways in eukaryotic cells to reach their intracellular target(s) in the cytosol and to ultimately kill. Such A/B toxins generally consist of an enzymatically active Asubunit (e.g., ricin toxin A (RTA)) and one or more cell binding Bsubunit(s), which are responsible for toxin binding to specific cell surface receptors. Our current knowledge of how A/B toxins are capable of efficiently intoxicating cells helped scientists to understand fundamental cellular mechanisms, like endocytosis and intracellular protein sorting in higher eukaryotic cells. From a medical point of view, it is likewise important to identify the major toxin trafficking routes to find adequate treatment solutions for patients or to eventually develop therapeutic toxin-based applications for cancer therapy.
Since genome-wide analyses of A/B toxin trafficking in mammalian cells is complex, time-consuming, and expensive, several studies on A/B toxin transport have been performed in the yeast model organism Saccharomyces cerevisiae. Despite being less complex, fundamental cellular processes in yeast and higher eukaryotic cells are similar and very often results obtained in yeast can be transferred to the mammalian situation.
Here, we describe a fast and easy to use reporter assay to analyze the intracellular trafficking of RTA in yeast. An essential advantage of the new assay is the opportunity to investigate not only RTA retro-translocation from the endoplasmic reticulum (ER) into the cytosol, but rather endocytosis and retrograde toxin transport from the plasma membrane into the ER. The assay makes use of a reporter plasmid that allows indirect measurement of RTA toxicity through fluorescence emission of the green fluorescent protein (GFP) after in vivo translation. Since RTA efficiently prevents the initiation of protein biosynthesis by 28S rRNA depurination, this assay allows the identification of host cell proteins involved in intracellular RTA transport through the detection of changes in fluorescence emission.

A Simple Fluorescence-based Reporter Assay to Identify Cellular Components Required for Ricin Toxin A Chain RTA Trafficking in Yeast

In the manuscript, we describe the use of a yeast-based fluorescence reporter assay to identify cellular components involved in the trafficking and killing processes of the cytotoxic A subunit of the plant toxin ricin (RTA).

Bacterial and plant A/B toxins exploit the natural trafficking pathways in eukaryotic cells to reach their intracellular target(s) in the cytosol and to ...