Name: molecular spring constant analysis by biomembrane force probe spectroscopy
Uploaded: 2021-11-20T13:00:00.000+00:00
Duration: 8 min 10 s
Description: Watch this Scientific Journal Video about Molecular Spring Constant Analysis by Biomembrane Force Probe Spectroscopy at JoVE.com

We thank Guillaume Troadec for helpful discussion, Zihao Wang for hardware consultation, and Sydney Manufacturing Hub, Gregg Suaning and Simon Ringer for support of our lab startup. This work was supported by Australian Research Council Discovery Project (DP200101970 - L.A.J.), NSW Cardiovascular Capacity Building Program (Early-Mid Career Researcher Grant - L.A.J.), Sydney Research Accelerator prize (SOAR - L.A.J.), Ramaciotti Foundations Health Investment Grant (2020HIG76 - L.A.J.), National Health and Medical Research Council Ideas Grant (APP2003904 - L.A.J.), and The University of Sydney Faculty of Engineering Startup Fund and Major Equipment Scheme (L.A.J.). Lining Arnold Ju is an Australian Research Council DECRA fellow (DE190100609).

1. Obtain Analyzable DFS Events
<ol>
	<li>Start the experiment in the software (e.g., LabVIEW VI) for the BFP control and parameter setting (Figure S1A).</li>
	<li>Observe the repetitive probe bead-target bead/cell touches in the software for BFP Monitor (Figure S1B).</li>
	<li>Test and achieve the adhesion frequency &#8804; 20% within the first 50 touches by tuning the impingement force and contact time, by which it ensures that &#8805; 89% of DFS adhesion event are mediated by single bonds12,13,14. 
	NOTE: For each Bead-Cell/Bead pair, we perform 200 repetitive touch cycles. To obtain publishable data quality, we usually perform n &#8805; 3 Bead-Bead or Bead-Cell pairs.
	<ol>
		<li>Save data, in the form of Force vs. Time, to the user directed folder by the end of each pair, prompted by the software for BFP control and parameter setting.</li>
	</ol>
	</li>
	<li>Collect the Force vs. Time raw data of Bond events, as exemplified in the Figure 2A, using the BFP acquisition platform (Figure S1C).
	<ol>
		<li>Open the BFP data analysis software. Click on the yellow folder icon and select the corresponding raw data file by double clicking on them.</li>
	</ol>
	</li>
	<li>Run the program, and then click the up and down button to switch between events. Use the outlier exclusion criteria (Figure S2) to screen out invalid events. Select the exporting data type as Force vs. Time format and click on the Export Plots Data button.</li>
</ol>
2. Convert the Force vs. Time Curve to the Force vs. Displacement Curve
<ol>
	<li>Export the data segment corresponding to the Retract stage to a spreadsheet (Figure 2A, square marquee), which is relevant to the spring constant analysis.</li>
	<li>Plot the Force vs. Time Curve using spreadsheet software. To obtain the Force vs. Displacment curve, convert the time values (Figure 2A, x-axis) to the total displacement values (&#916;xtot) by multiplying time values with piezo movement velocity (i.e., 4,000 nm/s by preset).</li>
	<li>Zero the first data point by subtracting the smallest displacement value from each acquired displacement value. This horizontal transformation does not affect the ascending slopes of the Retract stage nor the subsequent spring constant calculation.</li>
	<li>Notably, the BFP is considered as a serial spring system in which &#916;xtot (Table 1) sum deformations of the RBC,&#160;&#916;xRBC&#160;(Table 1), the molecular bond, &#916;xmol (Table 1), and the Target cell,&#160;&#916;xcell (Table 1), as the Eq. 2: 
	<img alt="Equation 2" src="/files/ftp_upload/62490/62490eq02.jpg" /> (Eq. 2)</li>
	<li>Plot the Force (F) vs. Displacement (&#916;xtot) curve as shown in the Figure 2B.</li>
</ol>
3. Spring Cnstant Analysis of Bead- Cell Mode
<ol>
	<li>In the Force vs. Displacement curve, two distinct slop can be identified, where each can represent the compressive phase and the tensile phase. Fit a regression line to each data group (Figure 2B), where the larger linear fit slope represents the total spring constant at compressive phase (Figure 2B, red), denoted as k1 (Table 1); and the smaller linear fit slope represents the total spring constant at tenslile phase (Figure 2B, blue), denotated as k2 (Table 1).</li>
	<li>For springs connected in series per step 2.2 description, express the reciprocal of the total spring constant, ktot (Table 1), as the sum of the spring constant inverses of RBC, kRBC&#160;(Table 1), the molecular bond, kmol&#160;(Table 1), and the Target cell, kcell (Table 1). During the compressive phase of the Bead-Cell mode, the molecular bond is not stretched, therefore kmol is not taken into consideration. The reciprocal of the ktot in this scenario (1/k1) is expressed as 
	<img alt="Equation 3" src="/files/ftp_upload/62490/62490eq03.jpg" /> (Eq. 3). 
	In the example data, kRBC&#160;is pre-determined (0.25 pN/nm by default). kcell can be derived from the Eq. 3 with the acquired k1 and kRBC (Figure 3B).</li>
	<li>During the tensile phase, adhesion is formed between the ligand-receptor pair. Express the reciprocal of the ktot in this scenario (1/k2) as 
	<img alt="Equation 4" src="/files/ftp_upload/62490/62490eq04.jpg" /> (Eq. 4) 
	where k2 (Table 1) represents the total spring constant during the tensile phase.</li>
	<li>Derive kmol from subtracting 1/k1 from 1/k2 (compare Eq. 3 vs. Eq. 4).</li>
</ol>
4. Spring Constant Analysis of Bead- Bead Mode
<ol>
	<li>Fit a regression line to the compressive phase data to obtain k1&#160;(similar to the Figure 2B, red). Of note, in the Bead-Bead mode, the Target cell is replaced by a glass bead coated with the receptor of interest (Figure 1C). Since bead deformation is negligible, the 1/kcell term can be removed from the Eq. 3 and Eq. 4 accordingly. The reciprocal ktot of the compressive phase (1/k1) can be expressed as: 
	<img alt="Equation 5" src="/files/ftp_upload/62490/62490eq05.jpg" /> (Eq. 5)</li>
	<li>Fit a regression line to the tensile phase data to obtain k2&#160;(similar to the Figure 2B, blue). The reciprocal ktot of the tensile phase (1/k2) can be expressed as: 
	<img alt="Equation 6" src="/files/ftp_upload/62490/62490eq06.jpg" /> (Eq. 6)</li>
	<li>Derive kmol from subtracting 1/k1 from 1/k2 (compare Eq. 5 vs. Eq. 6).</li>
</ol>

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

In summary, we have provided a detailed data analysis protocol for preprocessing the DFS raw data and deriving molecular spring constants in the BFP Bead-Bead and Bead-Cell analysis modes. Biomechanical models and equations required for determining molecular and cellular spring constants are presented. Albeit different integrins are studied, the kmol measured by the Bead-Bead mode and the Bead-Cell mode possesses significant range differences (Figure 3A vs. Figure 3C). Of note, with the Bead-Bead mode, the receptor is covalently linked to the glass bead. In contrast, with the Bead-Cell mode, the surface receptor is adapted by the underlying plasma membrane and cytoskeletons, which most likely influence the measured kmol.
Data quality control is crucial to ensure the reproducibility. To this end, we have implemented the DFS data pre-screening and outlier exclusion criteria on the Force vs. Time plots. To demonstrate this, a representative dataset was selected, in which we categorized the DFS raw data into three levels of quality: Good (Figure S2A), Acceptable with noise (Figure S2B), and Poor unacceptable (Figure S2C). For beginners of using the BFP, we recommend the strict criteria to pre-screen the data with the Good quality (Figure S2A). Of note, based on the data pre-screening criteria, the regression fit line of the compressive phase should be steeper than that of the tensile phase, specifically k1 &#62; k2 (Figure S2C, vii). When measured k1 &#60; k2 (Figure S2C, vii), the derived kmol &#60; 0 is against the rationale per calculation in the step 4. Such events should then be considered as the invalid outliers and discarded.
To favor the BFP measurement on single-molecular level during data acquisition, multiple experimental configurations have been implemented according to previous study12. Firstly, protein coating density on beads is usually titrated down to a minimal level (e.g. 60 &#181;m-2) by strictly control the solution concentration, quantity of the protein and reaction conditions15. The average spatial distance between proteins on the bead is thereby estimated much larger than the linear dimensions of the protein, favoring our measurements on single-molecular level12,13,14. Secondly, we control the adhesion frequency for each ligand-receptor pair &#8804; 20%, under which molecular binding events will follow Poisson distribution predicting &#8805; 89% of events would be single-molecular binding14,15. To achieve so, impingement force and contact time are set accordingly and need to be consistent throughout the experiment12. Nevertheless, it is still possible that multiple bonds occur sequentially (Figure S2C, vi).&#160;In such cases, we will discard the events with signatures of multiple bonds. Last but not least, negative control experiments will be performed with beads coated with bovine serum albumin (Table of Materials) or SA alone to ensure the non-specific adhesion frequency is &#8804; 2%16,17.
Although BFP is powerful to investigate protein dynamics on living cell surface10,11,12, there are technical limitations. In BFP, only one ligand-receptor pair can be investigated at a time. It would be time consuming to obtain sufficient data with statistical significance. Besides, the experimental procedures are labor intensive with steep learning curves. Implementation wise, the current BFP system is susceptible to the environmental drifting and surrounding mechanical vibration. As a result, continuous manual adjustment is required to ensure the DFS data quality. To this end, one of our recent studies has introduced ultra-stable BFP feedback control algorithms to improve the stability of the BFP force-clamp DFS assays4. This technical advance enables measurements of stronger molecular interaction such as antigen-antibody binding with ultra-long bond lifetime (&#62;50 s). Nevertheless, we foresee future efforts will be made to automate and integrate the BFP data acquisition and DFS analysis into one computerized program, making the entire BFP operation and data analysis more user friendly and high-throughput.

As a live-cell DFS technique, BFP engineers a human red blood cell (RBC; Figure 1) into an ultrasensitive and tunable force transducer with a compatible spring constant range at 0.1-3 pN/nm1,2,3. To probe ligand-receptor interaction, BFP enables DFS measurements at ~1 pN (10-12 N), ~3 nm (10-9 m), and ~0.5 ms (10-3 s) in force, spatial, and temporal resolution4,5. Its experimental configuration consists of two opposing micropipettes, namely the Probe and the target. The Probe micropipette aspirates a RBC and a bead is glued at its apex via a biotin-streptavidin interaction. The bead is coated with the ligand of interest (Figure 1A). The Target micropipette aspirates either a cell or a bead bearing the receptor of interest, corresponding to the Bead-Cell (Figure 1B) and Bead-Bead (Figure 1C) modes, respectively5.
BFP construction, assembly and the DFS experimental protocols were described in detail previously1,6. Briefly, a BFP touch cycle consists of 5 stages: Approach, Impinge, Contact, Retract and Dissociate (Figure 1D). The horizontal RBC apex position is denoted as &#916;xRBC. At the beginning, the unstressed (zero-force) RBC deformation &#916;xRBC is 0 (Table 1). The Target is then driven by a piezotranslator to impinge on and retract from the Probe bead (Figure 1D). The RBC probe is first compressed by the Target with negative RBC deformation &#916;xRBC&#160;&#60; 0. In a Bond event, the Retract stage transitions from a compressive to a tensile phase with positive RBC deformation &#916;xRBC&#160;&#62; 0 (Figure 2C and D). According to Hooke&#39;s law, the BFP bearing force is able to be measured as F = kRBC&#160;&#215;&#160;&#916;xRBC, where kRBC (Table 1) is the RBC spring constant of the BFP. Upon bond rupture and the completion of one touch cycle, the probe bead returns to zero-force position with &#916;xRBC&#160;= 0 (Figure 1D).
To determine the kRBC, we measure and record the radii of the probe micropipette inner orifice (Rp), the RBC (R0) and the circular contact area (Rc) between the RBC and the probe bead (Figure 1A). Then kRBC is calculated according to the Evan&#39;s model (Eq. 1)7,8 using a LabVIEW program that acts as a virtual instrument (VI) to operate the BFP (Figure S1A)8,9.
<img alt="Equation 1" src="/files/ftp_upload/62490/62490eq01.jpg" /> (Eq. 1)
With a BFP established and DFS raw data obtained, hereby we present how to analyze the spring constant of ligand-receptor pair or cells. The DFS raw data on the interaction of the glycosylated protein Thy-1 and K562 cell bearing integrin &#945;5&#946;1 (Thy-1-&#945;5&#946;1; Figures 3A and 3B)10 and that of the fibrinogen and bead coated integrin &#945;IIb&#946;3 (FGN-&#945;IIb&#946;3; Figure 3C)11,12 have been used to demonstrate the Bead-Cell and Bead-Bead analysis modes, respectively.
BFP Experimental Preparation 
For details of BFP experimental preparation and instrumentation, please refer to the previously published protocols3. In brief, human RBC has been biotinylated using the Biotin-PEG3500-NHS in the carbon/bicarbonate buffer. Proteins of interest have been covalently coupled to the borosilicate glass beads using MAL-PEG3500-NHS in the phosphate buffer. To attach to the biotinylated RBC, the probe bead is also coated with streptavidin (SA) using the MAL-SA. Please see the Table of Materials and Table 2.
To assemble the BFP (Figure 1, left), the third micropipette termed &#39;Helper&#39; will be used to deliver the probe bead and glue it to the RBC&#39;s apex1,3. The covalent interaction between the SA coated probe bead and biotinylated RBC is much stronger than the ligand-receptor bond of interest. Thus, the Dissociate stage can be interpreted as the ligand-receptor bond rupture rather than the detachment of Probe bead from the RBC.

<table><tbody><tr><td>3-Mercaptopropyltrimethoxysilane (MPTMS)</td><td>Uct, Specialties, llc</td><td>4420-74-0</td><td>Glass bead functionalization</td></tr><tr><td>Anhy. Sodium Phosphate Dibasic (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S7907</td><td>Phosphate buffer preparation</td></tr><tr><td>BFP data acquisition VI</td><td>LabVIEW</td><td></td><td>BFP control and parameter setting</td></tr><tr><td>BFP data analysis VI</td><td>LabVIEW</td><td></td><td>BFP raw data analysis</td></tr><tr><td>Biotin-PEG3500-NHS</td><td>JenKem</td><td>A5026-1</td><td>RBC biotinylation</td></tr><tr><td>Borosilicate Glass beads</td><td>Distrilab Particle Technology, Netherlands</td><td>9002</td><td>Glass bead functionalization</td></tr><tr><td>Bovine serum albumin</td><td>Sigma-Aldrich</td><td>A0336</td><td>Ligand functionalization</td></tr><tr><td>Camera VI</td><td>LabVIEW</td><td></td><td>BFP monitoring</td></tr><tr><td>&lt;sub&gt;D&lt;/sub&gt;-glucose</td><td>Sigma-Aldrich</td><td>G7021</td><td>Tyrode&amp;rsquo;s buffer preparation</td></tr><tr><td>Hepes</td><td>Sigma-Aldrich</td><td>H3375</td><td>Tyrode&amp;rsquo;s buffer preparation</td></tr><tr><td>MAL-PEG3500-NHS</td><td>JenKem</td><td>A5002-1</td><td>Glass bead functionalization</td></tr><tr><td>Potassium Chloride (KCl)</td><td>Sigma-Aldrich</td><td>P9541</td><td>Tyrode&amp;rsquo;s buffer preparation</td></tr><tr><td>Sodium Bicarbonate (NaHCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S5761</td><td>Carbonate/bicarbonate buffer preparation; Tyrode&amp;rsquo;s buffer preparation</td></tr><tr><td>Sodium Carbonate (Na&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S2127</td><td>Carbonate/bicarbonate buffer preparation</td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Sigma-Aldrich</td><td>S7653</td><td>Tyrode&amp;rsquo;s buffer preparation</td></tr><tr><td>Sodium Phosphate Monobasic Monohydrate (NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;&amp;bull;H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>S9638</td><td>Phosphate buffer preparation</td></tr><tr><td>Streptavidin-Maleimide</td><td>Sigma-Aldrich</td><td>S9415</td><td>Glass bead functionalization</td></tr></tbody></table>

molecular spring constant analysis by biomembrane force probe spectroscopy

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure single-molecular binding kinetics, assess mechanical properties of ligand-receptor interactions, visualize protein dynamic conformational changes and more excitingly elucidate receptor mediated cell mechanosensing mechanisms. More recently, BFP has been used to measure the spring constant of molecular bonds. This protocol describes the step-by-step procedure to perform molecular spring constant DFS analysis. Specifically, two BFP operation modes are discussed, namely the Bead-Cell and Bead-Bead modes. This protocol focuses on deriving spring constants of the molecular bond and cell from DFS raw data.

In this work, we have demonstrated the protocol of the BFP spring constant analysis. For the Bead-Cell analysis mode, we analyzed the kmol&#160;of the molecular bond between the glycosylated protein Thy-1 coated on the Probe bead and the integrin &#945;5&#946;1 expressed on the Target K562 cell (Thy-1-integrin &#945;5&#946;1; Figure 3A)10. The kcell is also derived from the Bead-Cell mode (K562 Cell; Figure 3B). For the Bead-Bead mode, the molecular bond formed between fibrinogen and integrin &#945;IIb&#946;3 (FGN-integrin &#945;IIb&#946;3; Figure 3C)11,12 is used to demonstrate the Bead-Bead analysis mode.
For the Bead-Cell mode, we measured the spring constants of Thy-1-integrin &#945;5&#946;1 bond and K562 cell as kmol = 0.45 &#177; 0.28 pN/nm (Figure 3A) and kcell = 0.18&#160;&#177; 0.07 pN/nm (Figure 3B) from 27 pre-screened analyzable events. For the Bead-Bead mode, we measured the spring constants of FGN-integrin &#945;IIb&#946;3 bond as kmol = 0.53 &#177; 0.29 pN/nm (Figure 3C) from 33 pre-screened analyzable events.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Symbol</td>
			<td>Definition</td>
			<td>Symbol</td>
			<td>Definition</td>
		</tr>
		<tr>
			<td>&#916;xtot</td>
			<td>The total displacement of the piezo, which can also be interpreted as the total deformation of RBC, target cell and molecular bond.</td>
			<td>&#916;xRBC</td>
			<td>The RBC deformation, which can also be interpreted as the displacement of the Probe bead.</td>
		</tr>
		<tr>
			<td>ktot</td>
			<td>The total spring constant of the entire BFP serial spring system.</td>
			<td>kRBC</td>
			<td>The spring constant of the aspirated RBC by the Probe micropipette.</td>
		</tr>
		<tr>
			<td>kmol</td>
			<td>The spring constant of the BFP detected molecular bond</td>
			<td>kcell</td>
			<td>The spring constant of the Target cell.</td>
		</tr>
		<tr>
			<td>k1</td>
			<td>ktot of the compressive phase in the Retract stage.</td>
			<td>k2</td>
			<td>ktot of the tensile phase in the Retract stage.</td>
		</tr>
		<tr>
			<td>&#916;F1</td>
			<td>The increment of force sensed by the Probe bead in the compressive phase.</td>
			<td>&#916;F2</td>
			<td>The increment of force sensed by the Probe bead in the tensile phase.</td>
		</tr>
		<tr>
			<td>&#916;x1</td>
			<td>The increment of displacement in the compressive phase.</td>
			<td>&#916;x2</td>
			<td>The increment of displacement in the tensile phase.</td>
		</tr>
	</tbody>
</table>
Table 1. Symbol definitions for the BFP molecular spring constant analysis.&#160;Horizontal positions of all objects are defined as x, while &#916;x&#160;[nm]&#160;refers to the deformation relative to the original position. &#916;F [pN] refers to the force increment measured by BFP. k [pN/nm] refers to the spring constant. Subscripts 1 and 2 correspond to the compressive and tensile phases, respectively. The molecular spring constant is derived from the Force (F) vs. Displacement (&#916;xtot) curve.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62490/62490fig01.jpg" /> 
Figure 1: BFP configuration and DFS touch cycle. (A) The BFP system assembles two opposing micropipettes, namely the Probe (left) and the Target (right). The Probe micropipette aspirates a RBC (red) with a glass bead glued at its apex to serve as a force transducer. The Target micropipette aspirates a receptor-bearing cell (blue). The RBC spring constant (kRBC) is determined by the aspiration pressure (&#916;p)&#160;and the radii of aspirated RBC (R0), Probe micropipette (Rp) and circular contact area (Rc) between the RBC and the Probe bead. (B and C) Micrographs of the Bead-Cell (B) and Bead-Bead (C) BFP modes. Scale bars = 5 &#181;m. (D) A BFP touch cycle that consists of Approach, Impinge, Contact, Retract and Dissociate stages. The &#916;xRBC = 0 dash line indicates the BFP unstressed, or zero-force, position. The Retract stage includes compressive phase (&#916;xRBC &#60; 0, red), zero-force position (&#916;xRBC = 0, black) and tensile phase (&#916;xRBC &#62; 0, blue) in sequence. Black arrow indicates the position of Bond event. <a href="https://www.jove.com/files/ftp_upload/62490/62490fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62490/62490fig02.jpg" /> 
Figure 2: Derive molecular spring constants from DFS raw data. (A) Representative Force (F) vs. Time (t) curve&#160;of DFS raw data in one BFP touch cycle. (B)&#160;Representative converted Force (F) vs. Displacement (&#916;xtot) curve that depicts the Retract stage. k1 and k2 represent the fit slope of the consecutive compressive and tensile phases respectively. &#916;F1 and&#160;&#916;F2 represent the increments of force in the compressive phase and the tensile phase data, respectively, where &#916;x1 and&#160;&#916;x2 represent the increments of displacement in the compressive phase and tensile phase data, respectively. R2 values for spring constant during compressive stage (R12) and tensile phase (R22) are labelled on the graph to indicate good statistical fitness. (C and D) Illustrations of the Retract stage in the Bead-Cell (C) and Bead-Bead (D) experimental modes. kRBC represents the spring constant of RBC; kcell and kmol represent the spring constants of the Target cell and the molecular bond, respectively. During the tensile phase, adhesion is formed between the ligand-receptor pair, the RBC deflects in the same direction as piezo retracts beyond zero-force position (&#916;xRBC &#62; 0). <a href="https://www.jove.com/files/ftp_upload/62490/62490fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62490/62490fig03.jpg" /> 
Figure 3:&#160;Representative histograms of BFP measured spring constants. The event number (left y-axis) and frequency distribution (right y-axis) of measured spring constants for Thy-1-integrin &#945;5&#946;1 bond (A) and K562 Target cell (B) in the Bead-Cell mode and FGN-integrin &#945;IIb&#946;3 bond (C) in the Bead-Bead mode. Histograms are fit with Gaussian distribution curve (pink) and the statistical parameter, R2, is used to indicate the strength of fitness. <a href="https://www.jove.com/files/ftp_upload/62490/62490fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Figure S1: The homemade BFP interface. (A)&#160;BFP control and parameter setting interface. The parameters to determine RBC spring constant are entered from the panel of biophysical parameters. (B) BFP monitoring.&#160;Live BFP touch cycles will be observed from this camera view. (C) BFP DFS analysis interface where the Force (F) vs. Time (t) curves are offline reviewed and pre-processed for subsequent molecular spring constant analysis. <a href="https://www.jove.com/files/ftp_upload/62490/Fig S1.tif" target="_blank">Please click here to download this File.</a>
Figure S2. BFP analyzable data quality control and pre-screening criteria. (A) Good DFS events with high quality: (i) Bead-Cell rupture force event; (ii) Bead-Cell lifetime event; (iii) Bead-Bead rupture force event; (iv) Bead-Bead lifetime event. (B)&#160;Acceptable DFS events with some noise: (i) Data drifting but the zoom-in Retract stage remains valid; (ii)Slight data drifting after bond dissociation; (iii) Data kink at the zero-force regime; (iv) Holding force is small (&#60; 10 pN). (C)Poor-quality events that should be discarded: (i) No adhesion; (ii) Data oscillation; (iii) Data drifting all the time; (iv) Discontinuous data; (v) Compressive force too small (&#8776; 0 pN); (vi) Multiple bonds; (vii) Invalid data with derived kmol &#60; 0; (viii) Signal error. Zero force is indicated by the grey intermittent line. <a href="https://www.jove.com/files/ftp_upload/62490/Fig S2.tif" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Molecular Spring Constant Analysis by Biomembrane Force Probe Spectroscopy at JoVE.com

Molecular Spring Constant Analysis by Biomembrane Force Probe Spectroscopy

A biomembrane force probe (BFP) is an in situ dynamic force spectroscopy (DFS) technique. BFP can be used to measure the spring constant of molecular interactions on living cells. This protocol presents spring constant analysis for molecular bonds detected by BFP.

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure ...

molecular-spring-constant-analysis-biomembrane-force-probe

Nanyang Technological University

Research

JoVE Journal

Bioengineering

2.9K Views.  The University of Sydney. A biomembrane force probe (BFP) is an in situ dynamic force spectroscopy (DFS) technique. BFP can be used to measure the spring constant of molecular interactions on living cells. This protocol presents spring constant analysis for molecular bonds detected by BFP. 

Video: Molecular Spring Constant Analysis by Biomembrane Force Probe Spectroscopy

Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

The generation and detection of mechanical forces is a ubiquitous aspect of cell physiology, with direct relevance to cancer metastasis1, atherogenesis2 and wound healing3. In each of these examples, cells both exert force on their surroundings and simultaneously enzymatically remodel the extracellular matrix (ECM). The effect of forces on ECM has thus become an area of considerable interest due to its likely biological and medical importance4-7.
	Single molecule techniques such as optical trapping8, atomic force microscopy9, and magnetic tweezers10,11 allow researchers to probe the function of enzymes at a molecular level by exerting forces on individual proteins. Of these techniques, magnetic tweezers (MT) are notable for their low cost and high throughput. MT exert forces in the range of ~1-100 pN and can provide millisecond temporal resolution, qualities that are well matched to the study of enzyme mechanism at the single-molecule level12. Here we report a highly parallelizable MT assay to study the effect of force on the proteolysis of single protein molecules. We present the specific example of the proteolysis of a trimeric collagen peptide by matrix metalloproteinase 1 (MMP-1); however, this assay can be easily adapted to study other substrates and proteases.

In this article we describe the use of magnetic tweezers to study the effect of force on enzymatic proteolysis at the single molecule level in a highly parallelizable manner.

The  generation and detection of mechanical forces is a ubiquitous aspect of cell physiology,  with direct relevance to cancer metastasis1, atherogenesis2 ...

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. Atomic force microscopy (AFM) can address this problem by enabling stretching and relaxation of single protein molecules, which gives direct evidence of specific unfolding and refolding characteristics. AFM-based single-molecule force-spectroscopy (AFM-SMFS) provides a means to consistently measure high-energy conformations in proteins that are not possible in traditional bulk (biochemical) measurements. Although numerous papers were published to show principles of AFM-SMFS, it is not easy to conduct SMFS experiments due to a lack of an exhaustively complete protocol. In this study, we briefly illustrate the principles of AFM and extensively detail the protocols, procedures, and data analysis as a guideline to achieve good results from SMFS experiments. We demonstrate representative SMFS results of single protein mechanical unfolding measurements and we provide troubleshooting strategies for some commonly encountered problems.

We describe the detailed procedures and strategies to measure the mechanical properties and mechanical unfolding pathways of single protein molecules using an atomic force microscope. We also show representative results as a reference for selection and justification of good single protein molecule recordings.

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. ...

Biochemistry

Investigating Single Molecule Adhesion by Atomic Force Spectroscopy

Atomic force spectroscopy is an ideal tool to study molecules at surfaces and interfaces. An experimental protocol to couple a large variety of single molecules covalently onto an AFM tip is presented. At the same time the AFM tip is passivated to prevent unspecific interactions between the tip and the substrate, which is a prerequisite to study single molecules attached to the AFM tip. Analyses to determine the adhesion force, the adhesion length, and the free energy of these molecules on solid surfaces and bio-interfaces are shortly presented and external references for further reading are provided. Example molecules are the poly(amino acid) polytyrosine, the graft polymer PI-g-PS and the phospholipid POPE (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine). These molecules are desorbed from different surfaces like CH3-SAMs, hydrogen terminated diamond and supported lipid bilayers under various solvent conditions. Finally, the advantages of force spectroscopic single molecule experiments are discussed including means to decide if truly a single molecule has been studied in the experiment.

A protocol to couple a large variety of single molecules covalently onto an AFM tip is presented. Procedures and examples to determine the adhesion force and free energy of these molecules on solid supports and bio-interfaces are provided.

Atomic force spectroscopy is an ideal tool to study molecules at surfaces and interfaces. An experimental protocol to couple a large variety of single ...

Engineering

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

Insights into the Interactions of Amino Acids and Peptides with Inorganic Materials Using Single-Molecule Force Spectroscopy

The interactions between proteins or peptides and inorganic materials lead to several interesting processes. For example, combining proteins with minerals leads to the formation of composite materials with unique properties. In addition, the undesirable process of biofouling is initiated by the adsorption of biomolecules, mainly proteins, on surfaces. This organic layer is an adhesion layer for bacteria and allows them to interact with the surface. Understanding the fundamental forces that govern the interactions at the organic-inorganic interface is therefore important for many areas of research and could lead to the design of new materials for optical, mechanical and biomedical applications. This paper demonstrates a single-molecule force spectroscopy technique that utilizes an AFM to measure the adhesion force between either peptides or amino acids and well-defined inorganic surfaces. This technique involves a protocol for attaching the biomolecule to the AFM tip through a covalent flexible linker and single-molecule force spectroscopy measurements by atomic force microscope. In addition, an analysis of these measurements is included.

Here we present a protocol to measure the force of interactions between a well-defined inorganic surface and either peptides or amino acids by single-molecule force spectroscopy measurements using an atomic force microscope (AFM). The information obtained from the measurement is important to better understand the peptide-inorganic material interphase.

The interactions between proteins or peptides and inorganic materials lead to several interesting processes. For example, combining proteins with minerals ...

Chemistry

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions are likely affected by the mechanical environment in which binding and signaling take place. A recent study demonstrated that mechanical force can regulate antigen recognition by and triggering of the T-cell receptor (TCR). This was made possible by a new technology we developed and termed fluorescence biomembrane force probe (fBFP), which combines single-molecule force spectroscopy with fluorescence microscopy. Using an ultra-soft human red blood cell as the sensitive force sensor, a high-speed camera and real-time imaging tracking techniques, the fBFP is of ~1 pN (10-12 N), ~3 nm and ~0.5 msec in force, spatial and temporal resolution. With the fBFP, one can precisely measure single receptor-ligand binding kinetics under force regulation and simultaneously image binding-triggered intracellular calcium signaling on a single live cell. This new technology can be used to study other membrane receptor-ligand interaction and signaling in other cells under mechanical regulation.

We describe a technique for concurrently measuring force-regulated single receptor-ligand binding kinetics and real-time imaging of calcium signaling in a single T lymphocyte.

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions ...

Covalent Attachment of Single Molecules for AFM-based Force Spectroscopy

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and surfaces. For a true single molecule experiment, covalent attachment of the probe molecule is essential because only then can hundreds of force-extension traces with one and the same single molecule be obtained. Many traces are in turn necessary to prove that a single molecule alone is probed. Additionally, passivation is crucial for preventing unwanted interactions between the single probe molecule and the AFM cantilever tip as well as between the AFM cantilever tip and the underlying surface. The functionalization protocol presented here is reliable and can easily be applied to a variety of polymers. Characteristic single molecule events (i.e., stretches and plateaus) are detected in the force-extension traces. From these events, physical parameters such as stretching force, desorption force and desorption length can be obtained. This is particularly important for the precise investigation of stimuli-responsive systems at the single molecule level. As exemplary systems poly(ethylene glycol) (PEG), poly(N-isopropylacrylamide) (PNiPAM) and polystyrene (PS) are stretched and desorbed from SiOx (for PEG and PNiPAM) and from hydrophobic self-assembled monolayer surfaces (for PS) in aqueous environment.

Covalent attachment of probe molecules to atomic force microscopy (AFM) cantilever tips is an essential technique for the investigation of their physical properties. This allows us to determine the stretching force, desorption force and length of polymers via AFM-based single molecule force spectroscopy with high reproducibility.

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and ...

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral budding, among other vital cellular processes. For efficient biological activity, cell membranes should be rigid enough to maintain the integrity of the cell and its compartments yet fluid enough to allow membrane components, such as proteins and functional domains, to diffuse and interact. This delicate balance of elastic and fluid membrane properties, and their impact on biological function, necessitate a better understanding of collective membrane dynamics over mesoscopic length and time scales of key biological processes, e.g., membrane deformations and protein binding events. Among the techniques that can effectively probe this dynamic range is neutron spin echo (NSE) spectroscopy. Combined with deuterium labeling, NSE can be used to directly access bending and thickness fluctuations as well as mesoscopic dynamics of select membrane features. This paper provides a brief description of the NSE technique and outlines the procedures for performing NSE experiments on liposomal membranes, including details of sample preparation and deuteration schemes, along with instructions for data collection and reduction. The paper also introduces data analysis methods used to extract key membrane parameters, such as the bending rigidity modulus, area compressibility modulus, and in-plane viscosity. To illustrate the biological importance of NSE studies, select examples of membrane phenomena probed by NSE are discussed, namely, the effect of additives on membrane bending rigidity, the impact of domain formation on membrane fluctuations, and the dynamic signature of membrane-protein interactions.

This paper describes the protocols for sample preparation, data reduction, and data analysis in neutron spin echo (NSE) studies of lipid membranes. Judicious deuterium labeling of lipids enables access to different membrane dynamics on mesoscopic length and time scales, over which vital biological processes occur.

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral ...