Fluorescence Micropipette Aspiration Assay to Investigate Red Blood Cell Mechanosensing

Summary

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.

Abstract

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 ± 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.

Introduction

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 processes^1,2.

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 tension. These assays enable the measurement of various mechanical parameters, such as cell membrane 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 budding^3,4.

Mechanical Parameter	Description	Seminal Approaches
Cell Stiffness	Measurement of a cell's mechanical rigidity and elasticity.	Aspiration of the cell membrane and analysis of deformation response to the negative pressure20,21.
Adhesion Strength	Evaluation of how strongly cells adhere to surfaces.	Application of controlled suction to detach adhered cells from a substrate2,22.
Membrane Tension	Assessment of the tension or stress within cell membranes.	Measurement of the membrane deformation in response to applied pressure23,24.
Viscoelastic Properties	Characterization of a cell's combined viscous and elastic behavior.	Analysis of the time-dependent deformation response to aspiration23,25.
Deformability	Determination of how easily a cell can change shape.	Evaluation of the extent of deformation under controlled suction20,24.
Surface Tension	Measurement of the tension at the cell's surface.	Assessment of the pressure required to form a micropipette membrane protrusion26.
Cell-Material Interaction	Study of interactions between cells and materials or substrates.	Aspiration of cells in contact with different materials and observation of interactions2,24.
Cell-Cell Interaction	Examination of interactions between neighboring cells.	Aspiration of a group of cells and analysis of their intercellular forces27.

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 clefts^5,6. During this journey, RBCs must traverse through passages as narrow as 0.5-1.0 µm, subjecting themselves to a multitude of mechanical forces, including tension and compression^7,8,9. 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 stimuli^11,12. 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.

Protocol

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.

1. Withdraw 5 mL of blood from the median cubital vein using a 19 G butterfly needle.
2. Transfer the collected blood into a 15 mL tube containing 1:200 enoxaparin to prevent clotting.
3. Dilute 5 µL of enoxaparin-anticoagulated blood in 1 mL of carbonate/bicarbonate buffer (C-buffer, pH = 8.5-9; Table of Materials).
4. Centrifuge the diluted blood sample at 900 × g for 1 min to sediment the RBCs. Carefully decant the supernatant without disturbing the pellet.
5. Perform two washes of the RBC pellet with 1 mL of C-buffer (Table of Materials), centrifuging each time at 900 × g for 1 min.
6. Subsequently, wash the RBC pellet 2x with 1 mL of Tyrode's buffer using the same centrifugation conditions and then resuspend the final pellet in 1 mL of Tyrode's buffer to obtain the washed stock RBC suspension.

2. Calcium indicator loading

1. Adjust the concentration of the washed stock RBC solution to 10 × 10⁶ cells/mL in Tyrode's buffer, based on the cell count obtained using an automatic cell counter (Table of Materials).
2. Label the calcium inside the RBCs by incubating with 16.67 µM Cal-520 AM, a calcium-sensitive dye, while agitating on a rotary tube mixer for 1 h.
3. Dilute the RBCs in Tyrode's buffer containing 0.5% bovine serum albumin (BSA) at a 1:50 ratio. The cells are now ready for experimental use.

3. Micropipette fabrication

1. 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).
2. 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 °C.
3. Locate the micropipette using a 10x eyepiece. Move the micropipette close to the borosilicate glass bead by using the knobs for adjustment.
4. Change the eyepiece to 30x before positioning the micropipette as close to the borosilicate glass bead as possible without bending the pipette tip.
5. 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, the opening diameter, has been reached.
6. Release the foot pedal 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.
7. Gently extract the micropipette, leading to a clear straight cut on the closed micropipette. Confirm that the final diameter of the capillary is 1 µm.
   ​NOTE: CHECKPOINT (see Supplemental Table S1)

4. Cell chamber preparation

1. Use a diamond pencil to divide a standard 40 mm x 22 mm x 0.17 mm glass coverslip into three equal strips.
2. 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.
3. 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)
4. Inject 200 µL of the labeled RBC suspension between two coverslips using a 200 µL pipette gun (Figure 1).

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 µL of the cell solution in Tyrode's Buffer is seeded. Please click here to view a larger version of this figure.

5. Micropipette aspiration assembly

1. 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).
2. Lower the micropipette holder to below the fluid level of the connected water reservoir.
3. Inject either demineralized water or Tyrode'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).
4. 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)
5. Insert the micropipette into the holder tip. Tighten the holder screw to ensure the micropipette is fixed.
6. Insert the micropipette into the cell chamber and locate the micropipette and RBCs under the microscope. Use the micromanipulator to adjust the position.
7. Lower the micropipette tip further to ensure the tip is leveled with the located RBC.
   NOTE: CHECKPOINT (see Supplemental Table S1)
8. 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.

6. Perform the fluorescence-coupled micropipette aspiration assay

1. 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).
2. 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.
3. Find the micropipette under the field of view using the micromanipulator.
4. 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.
5. 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.
6. 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.
7. Start the acquisition in the camera-operating software. Switch on the fluorescence shutter.
8. 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 Δp = -5 to -40 mmHg. There should be a noticeable tongue elongation within the micropipette tip (Figure 2D).
9. Hold the pressure for a preset period; then, release the pressure.
10. Move the micropipette to pick up the next cell and repeat the experiment.

7. Fluorescence intensity analysis

1. Load the saved fluorescence images into the analysis software.
2. 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).
3. Scroll to the timeline at the bottom of the software. Locate the designated aspiration event.
4. 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.
5. 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).
6. Add an area filter to exclude background noises (see Supplemental File 1-Supplemental Figure S6).
   NOTE: This is completed in the post process stage.
7. Select the statistics tab | detailed tab | average values tab. Scroll to find and select the intensity mean (see Supplemental File 1-Supplemental Figure S7).
8. Export the fluorescence signal trace over time to a .csv file.
9. Open the exported csv file. Subtract the background signals, F_b, from all measurements.
10. Calculate the calcium intensity change, ΔF_max, using equation (1):
         (1)
    Where ΔF_max is the maximum calcium intensity change, F_b, is the background intensity, and F₀ is the resting intensity.

Figure 2: Fluorescence-coupled micropipette aspiration assembly. (A) 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) The microscope stage depicting the experiment cell chamber and micromanipulator system with a single micropipette. (C) 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) 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 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 µm. Abbreviations: fMPA = Fluorescence-coupled Micropipette Aspiration; DM = dichroic mirror; RBC = red blood cell. Please click here to view a larger version of this figure.

Representative Results

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 × 7 mm × 0.17 mm) were affixed to create a chamber filled with 200 µL of RBCs suspended in Tyrode'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 was brought closer to capture the target RBC.

For cell aspiration, the method used a pneumatic high-speed pressure clamp to fine-tune the negative aspiration pressure. Fluorescence imaging was then conducted to investigate the calcium mobilization at the different negative aspiration pressures applied.

By using the fMPA to investigate how RBCs respond to varying negative aspiration pressures, our findings reveal that there is a clear proportional relationship between the negative pressure applied and the calcium influx present in the aspirated RBC. To determine the calcium intensity change, the maximum intensity of the single aspirated cell (F_max) minus the background intensity (F_b) was divided by the resting intensity (F₀) minus F_b. There was a corresponding increase in the influx of calcium ions into the RBCs when the pressure was incrementally increased between -10 mmHg (Figure 3A) to -40 mmHg (Figure 3D). This suggests that RBCs possess the ability to sense changes in their mechanical and respond by rapid calcium-specific channel activities^14,15.

Figure 3: Fluorescence imaging with a graphical representation of normalized intensity changes against time. Horizontally across depicts fluorescence snapshots of the RBCs at rest (left) and the human RBCs being aspirated by the micropipette (middle). Representative traces of the RBCs being aspirated at multiple negative aspiration pressures (Δp) can be seen on the right. The negative aspiration pressure is incrementally increased, starting at (A) Δp = -10 mmHg, (B) Δp = -20 mmHg, (C) Δp = -30 mmHg, and (D) Δp = -40 mmHg. When the tongue of the RBC is elongated during aspiration, a significant calcium mobilization can be observed as shown by the increased Cal-520 AM fold change. F/F₀ was used to investigate the calcium mobilization inside the aspirated RBC. From the above curves, the observed trend demonstrated that the Cal-520 AM signal increased proportionally with the increase of the applied negative aspiration pressure. Scale bar = 5 µm. Abbreviation: RBCs = red blood cell. Please click here to view a larger version of this figure.

Supplemental Table S1: Checkpoints for critical steps in the fMPA preparation protocols. Please click here to download this File.

Supplemental File 1: A guided process to adjust and perform segmentation. Please click here to download this File.

Discussion

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, advancing the monitoring and detection of intracellular calcium influx in real time. This approach provides an integrative insight into the complex mechanosensing reactions of RBCs.

Importantly, the applicability of the fMPA technique extends beyond RBCs, as it can be employed with other cell types that disapply high mechanical sensitivity, such as platelets and neurons⁹. Moreover, owing to its minimal impact on cell integrity during experimental handling, fMPA guarantees the preservation of the cell's natural state, making it highly suitable for use with primary cells¹. Furthermore, the fMPA method offers versatility through the tunable geometry of the micropipettes, allowing for a broader range of experimental designs tailored to specific research questions and the effective exploration of various mechanical conditions.

Additionally, with the improvement in the optical pathway that enhances the simultaneous use of multi-fluorescence imaging, the fMPA system allows for real-time detection of intracellular calcium concentration upon aspiration. This capability opens an opportunity to explore calcium-related molecules, such as the mechanosensitive ion channel, PIEZO1, which has been shown to mediate and perceive mechanical cues from the external environments^7,14. Recent literature highlights an increase in membrane tension as a significant factor stimulating PIEZO1 channel activity, subsequently facilitating the Ca²⁺ influx⁶. This underscores a crucial application of fMPA, which is to investigate the interplay between membrane tension, PIEZO1, and calcium influx, shedding light on the intricate mechanosensing processes within cells.

From a technical perspective, it is worth mentioning that the two primary components of the fMPA system, responsible for generating the aspiration force (mechanical stimuli) and conducting real-time fluorescence imaging, are the pneumatic pressure pump and the fluorescence camera, respectively. The selection of the appropriate pump and camera for the system should be based on the specific cell type and biological process under study. The pump employed in our system operates within a steady-state pressure range of ± 200 mmHg and can respond to commands for pressure changes as significant as ± 200 mmHg. The pressure rise time and fall time should not exceed 6-8 ms for a 20 mmHg step and 15-20 ms for a 200 mmHg step. To ensure force resolution, the noise level of the HSPC system should be less than ± 0.5 mmHg, peak-to-peak. These requirements are applied in this setup; however, a range of ± 40 mmHg is sufficient to achieve a response for the experimental protocol¹⁵. Regarding the fluorescence camera, we selected features a 95% Quantum Efficiency and an 11 µm x 11 µm pixel size chip. This sensitive sensor configuration efficiently captures the fluorescence signal at high speed. Additionally, it is crucial to maintain a median read noise of 1.6^e- for an adequate signal-to-noise ratio during imaging¹⁶.

From a procedural perspective, executing fMPA requires a set of advanced skills. For example, crafting micropipettes using the micropipette cutter demands precision and dexterity, along with meticulous control of temperature and positioning. Positioning the micropipette within the microscope's field of view requires meticulous effort to avoid damage to both the micropipette tip and the cellular chamber. Additionally, one common challenge associated with fluorescence imaging is photobleaching. To mitigate this issue, it is important to minimize the sample's exposure time to the illumination system. With reference to the 'fluorescence-coupled micropipette aspiration assay' protocol, the fluorescence shutter of the excitation light source remains switched off until all parameters are inputted through the camera-operating software and the aspiration system is ready for experiments. Only then is the fluorescence shutter turned on to commence imaging. To further reduce the photobleaching, it is recommended to use the minimum light source intensity that still yields adequate fluorescence expression in the sample.

Furthermore, the current manual operation required for fMPA assays makes this technique labor-intensive. Consequentially, the inconsistencies that may arise in this assay primarily stem from operator-dependent variables, as well as the factors associated with variations in filament preheating and capillary glass characteristics. In the future, optimizing the performance of this technique necessitates consideration of potential tandem implementations. For example, when combined with microfluidic devices, the micropipette aspiration can be transformed into a high-throughput platform, significantly increasing the experimental rate from studying around 20 cells per hour to approximately 1,000 cells/h^17,18. This incorporation also improves the reproducibility of the data. Moreover, certain avenues have been successfully explored by incorporating automation and image analysis systems¹⁹. Notably, finite element analysis (FEA) is a computational tool commonly employed to model micropipette aspiration assays. FEA can predict the cellular response to mechanical stimuli and characterize their mechanical properties. It holds the potential to optimize the micropipette design and further validate the experimental results¹⁹.

In conclusion, the fMPA approach offers valuable insights into the mechanosensitive behaviors of RBCs in responding to mechanical stimuli. This study establishes a foundational framework for future investigations into the intricate mechanisms of mechanotransduction within RBCs and across broader biological systems. Such inquiries hold great promise for advancing our understanding of these mechanisms and unraveling their extensive implications in various physiological contexts.

Materials

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