We are grateful to Professor Richard Borgens, Center for Paralysis Research, Purdue University, for help in assembling the vibrating probe system. This study was supported by NEI grant NIH 1R01EY019101 to MZ and BR, and in part by grants from the California Institute of Regenerative Medicine RB1-01417, NSF MCB-0951199, and by an Unrestricted Grant from Research to Prevent Blindness, UC Davis Ophthalmology.

1. Probe Manufacture
Blank probes are purchased from World Precision Instruments (elgiloy/stainless parylene-coated microelectrodes) (see 'Table of specific reagents and equipment' below). The probe is cut 25-30 mm behind the tip and about 5 mm of parylene insulation at the cut end scraped away with a scalpel (#11 blade) to ensure a good connection. The probe is mounted in a gold R30 connector using electrically-conductive silver-loaded epoxy (e.g. Rite-Lok SL65) [*see note below]. The probe is stored overnight at room temperature to allow the epoxy to harden. Next, the probe tip is plated with gold and then platinum, using a nano-amp power supply. The probe tip is rinsed in acetone and connected to the negative output of a nano-amp power source. The probe tip is viewed under a dissecting microscope (x40) and placed first in gold plating solution (potassium dicyanoaurate (0.2% w/v KAu(CN)2 in distilled water (dH2O)). A reference wire connected to the positive output and placed in solution completes the circuit. A current of 5 nA is applied for 5 min, then increased to 20 nA until the tip is about half the desired final size (about 10-15 &mu;m). The probe tip is rinsed in dH2O and then placed in platinizing solution (chloroplatinic acid hydrate; 1% w/v H2PtCl6 * 6H2O) plus lead(II) acetate trihydrate (0.1% w/v Pb(CH3CO2)2 * 3H2O in dH2O). A current of 250 nA is applied for 5 min, then increased to 500 nA until the tip is about 80% of the desired final size. The current is increased to 1 &mu;A and applied in 1 sec bursts until the final tip diameter is obtained (about 30-35 &mu;m). Finally, the tip is rinsed in dH2O. Probes can be store at room temperature indefinitely. If probes are damaged, the gold R30 connectors can be re-used. 
[* Note: Some probe systems have the probe mounting connected directly to the amplifier to carry the signal. Other systems have a separate mounting point and signal connector. In the latter case, a short (2-3 cm) wire with an R30 connector at one end is soldered to the first R30 connector prior to mounting the probe (see Figure 2A).]
2. Probe System
The probe is attached to a piezo-electric bender mounted on a 3-dimensional micro-positioner (Figure 1). Probe vibration is controlled by the vibrating probe power supply which also allows adjustment of vibration amplitude and frequency. The probe power supply sends a reference signal to the lock-in amplifier, which also displays the vibration frequency and phase angle. It is also useful to connect an oscilloscope so one has a quick visual reference of probe vibration. The signal from the probe goes to the lock-in amplifier. The probe and sample to be measured can be viewed with a dissecting microscope (magnification x6 to x40) with fiber-optic illumination. During calibration and sample measurements, a reference and a ground (earth) wire must be in the solution (see Figure 2B).The amplifier is connected to a computer via an analog-digital (i/o) interface. Data is recorded using Strathclyde Electrophysiology Software's Whole Cell Program (WinWCP).
Lock-in amplifier settings: Sensitivity [200 &mu;V], Dynamic Resolution [normal], Offset [on], Expand [x1], Time Constant: pre [10 s], post [0.1 s]. If a faster response is required, the pre Time Constant can be reduced to 3 s. The offset control is used to bring the probe trace to near the center of the screen. If large responses are expected, the trace can be moved up or down.
WCP software settings: Record Duration [204.8 s], Samples Per Channel [1024], Sampling Interval [0.2 s], Voltage Range [+/- 0.2 V]. If large currents are anticipated, the Voltage Range can be increased to 1 V or 5 V.
3. Probe Settings
New probes have to be tested and their unique frequency and phase angle determined. The new probe is placed in the calibration chamber containing physiological saline (Figure 2B). The power supply is turned on and the frequency turned up until the maximum vibration is observed. This is the resonant frequency of the probe. Using the probe at this frequency can cause instability and produce noise in the recording, so the probe is 'de-tuned' by subtracting 10 Hz to give the probe's working frequency (typically 150-200 Hz). The vibration amplitude is adjusted so that the probe vibration distance is the same as the tip diameter, so that when the probe is vibrated a 'double image' of the probe tip is seen (see Figure 2B). To determine the phase angle, the probe is placed in saline in the calibration chamber and a current of 1.5 &mu;A/cm2 applied repeatedly. The phase angle on the lock-in amplifier is adjusted until there is no response. Adding or subtracting 90&deg; to this angle gives the maximum response, and this angle is the probe's working phase angle. The frequency and phase angle for each probe are noted for future use. During an experiment, it is important that these settings of frequency, amplitude and phase angle are not changed, as this will alter the probe's response. For convenience, when the current is flowing 'South-to-North', this should produce an upward deflection (termed here 'peak') in the recording trace, and current flowing 'North-to-South' should show a downward deflection (see Figure 3A). If this is the wrong way, then adding 180&deg; to the phase angle will fix it by flipping the responses round. See Reid et al.1 for detailed information on the theory behind probe function, calibration, etc.
Calibration: The response of the probe to a 'standardized' current of exactly 1.5 &mu;A/cm2, applied to the probe in a calibration chamber, is used to calculate the current in the sample (see Figure 2B, 3A). Prior to sample measurement, the probe is calibrated in appropriate solution, e.g. BSS+ Artificial Tear Solution for cornea. Current is applied in two directions: South-North and North-South, producing an upward and a downward deflection, respectively, equivalent to outward and inward currents, depending on the orientation of the sample. The probe trace should have a stable baseline and low noise (compare Figs. 3A and 3B). The probe is calibrated at the end of the experiment in used solution to compensate for osmolality change due to evaporation. When analyzing data, measurements from the first half of the experiment can be calculated using the starting calibration values, and measurements from the second half calculated using the end calibration.
Sample Measurement: A chamber may have to be designed to hold and immobilize the sample. For example, Figure 3C shows Petri dishes with wire loops to hold eyes for cornea measurements. The dish containing physiological saline is placed under the dissecting microscope and the area of interest on the sample placed in focus in the field of view. The probe is then placed in solution and orientated parallel to the sample surface and also in focus so it is on the same level as the point on the sample to be measured. The probe is moved as far as is convenient (e.g. 1-2 cm) away from the sample, the vibration turned on (and the computer software set to record) to establish a stable (horizontal) baseline (see Figure 4A). The probe is then moved into measuring position, about 50 &mu;m from the surface. When the new ('peak') value is stable, the probe is moved back to reference position and the trace returns to baseline. This can be repeated at regular time points to produce timelapse data, or the sample moved/rotated slightly and repeated at different positions to yield spatial current mapping data (see Figure 4B, 5B).
Data Analysis: Data are analyzed using WinWCP (see Figure 5A). The horizontal red 'zero' line is moved up or down so it is parallel to the trace baseline prior to the measuring peak. The vertical green measuring line is moved across to the trace baseline. The output reading in green at the bottom of the green line should be close to zero (e.g. 0.00012). This number shows the differnce between the point where the red and green lines cross, and the blue trace. The red line is then moved up until it is parallel with the top of the peak. The green output reading is the size of the peak in mV. The data for all measurement peaks, and the calibration data, are put into a Microsoft Excel spreadsheet template (see Table 1). Relevent information such as the date, probe number, voltage range (VR), solution used, measuring positions, time-points, etc. can also be put in the spreadsheet. The current direction (into or out of the sample: 'i/o') is noted, and inward currents are also given a negative value in the 'peak' column. The current in the right-hand column is calculated using the formula: current=peak*(1.5/calibration) where 1.5 is the current in &mu;A/cm2 applied at calibration. Thus: 'unw1'=12.45*(1.5/56.12)= 0.33276907 &mu;A/cm2.
4. Secrets to Success
As in all electrophysiology, proper grounding (earthing) of vital equipment helps to eliminate noise. Thus, at least the micro-positioner and the chassis of the microscope should be earthed, and maybe also the light source (see Figure 1). The user can also be a source of static electricity, so earthing via a wrist-band can prevent instability of the probe in some cases, but is not always necessary. A Faraday cage is not necessary, as the lock-in amplifier filters out all frequencies (e.g. 60Hz from power grid) apart from the frequency the probe is vibrating at. A vibration isolation table is useful but not essential. A solid, stable bench or table works just as well. Apart from the basic information in the Excel spreadsheet (see above) it is useful to record in one's lab-book additional useful information such as changes in temperature, drug additions, etc. If photographs are taken down the microscope, note the magnification. If relevant, it is also useful to draw a sketch of the sample(s) showing position and/or orientation of probe measurements (see Figure 6).
Challenging steps
<ol class="lalpha">
	<li>Probe making: there must be a good electrical connection between the probe and the R30 connector. If nothing happens at the electroplating stage, then this is probably the cause.</li>
	<li>Calibration: use appropriate saline or culture medium for the sample you will be measuring. Don't over- or under-fill the calibration chamber as this can alter the response. The surface of the liquid should be flat across the top of the chamber.</li>
	<li>Sample measurements: Plan in advance, e.g. do you need to make a special chamber to hold/mount the sample (see Figure 3C)? When measuring from a sample, the probe should be orientated with the long axis parallel to the sample surface, so that the probe's direction of vibration (and thus direction of current being measured) is perpendicular to the sample surface (for example, see Figure 4B). The sample can be moved and/or rotated for measurements at different positions. It is important to maintain a consistent distance between the probe and the sample surface when measuring. The current measured is proportional to the distance from the sample surface; as the probe moves away from the sample surface, the current drops by the inverse square law. That is, when measuring a current generated at the sample surface, the current detected is inversely proportional to the square of the distance from the surface. An eyepiece graticule can be used to judge the distance between the probe and the sample surface.</li>
</ol>
Troubleshooting
<ul>
	<li>Problem: no response at calibration. Solution: check saline is contacting both electrodes. Check battery in constant current calibrator.</li>
	<li>Problem: small response. Solution: clean probe in dH2O and/or acetone. Check phase angle.</li>
	<li>Problem: noise or unstable baseline (see Figure 3B). Solution: Check earthing wires.</li>
	<li>Problem: trace jumps off screen. Solution: do not allow probe tip to touch sample.</li>
</ul>
5. Representative Results
Figure 3A shows a good example of a calibration trace. Note the stable (horizontal) baseline, low noise and large response. As a comparison, Figure 3B shows a noisy trace with unstable baseline. Current measurements at different positions at a mouse cornea wound are shown in Figure 4B. The top panel shows the probe positions, the middle panel shows the probe traces as recorded on the computer and the lower panel is a graph of the currents at the different positions, showing a profile of wound current. Figure 5B shows measurements at a mouse skin wound made at regular time intervals to produce data on the wound current time-course.
<img src="/files/ftp_upload/2358/2358fig1.jpg" alt="Figure 1" /> Figure 1. Vibrating probe system. See text for detailed description. Blue text describes the function of the connecting wires. Green symbols show earthing points.
<img src="/files/ftp_upload/2358/2358fig2.jpg" alt="Figure 2" /> Figure 2. A. Probe mounting. The probe is glued into a gold R30 connector with silver-loaded epoxy, prior to electroporating. In some systems, a second connector with a short wire to carry the signal is soldered on. Scale bar 3 mm. The platinum ball is not to scale. B. Probe calibration. The constant current calibrator (left) applies a current of 1.5 &mu;A/cm2 to the probe in the calibration chamber (right). Lower left: close-up of a probe. When the probe is vibrated (lower right), the amplitude is adjusted so a double-image of the tip is seen. Scale bar 100 &mu;m.
<img src="/files/ftp_upload/2358/2358fig3.jpg" alt="Figure 3" /> Figure 3. A. Probe calibration trace. Example of a good probe calibration trace, with stable baseline, low noise and large response. Current flowing South-to-North gives an upward deflection, and current flowing North-to-South produces a downward deflection. B. An unstable, noisy probe trace. C. Chambers made for mounting mouse (left) or rat (right) eyes for cornea measurements. Scale bar 5mm.
<img src="/files/ftp_upload/2358/2358fig4.jpg" alt="Figure 4" /> Figure 4. Prove measurement examples. A. A stable baseline is established with the probe in reference position 1-2 from the sample. When the probe is moved to measuring position near the sample, it detects a current and the trace deflects upwards (outward current). B. Measurements at different positions across a mouse cornea would. Probe is orientated parallel to the surface, so vibration is perpendicular. Upward peaks show outward currents. Maximum current is seen at the wound edges (positions B &amp; F). The schematic of the probe is shown at measuring position F (right wound edge). Scale bar 300 &mu;m.
<img src="/files/ftp_upload/2358/2358fig5.jpg" alt="Figure 5" /> Figure 5. A. Analyzing probe traces. Upper panel: the red zero line is moved up or down so it is parallel with the trace baseline, then the green measuring line is moved across to the baseline so the output reading in green is close to zero (0.00012). Lower panel: the red line is then moved up until it is parallel with the top of the trace peak, and the output reading gives the size of the peak in mV (12.45). Current is calculated from this using the calibration data (see Table 1). B. Mouse skin wound timelapse data. Current before wounding is shown at time zero (red symbol). Measurements were made at the same position at a mouse skin wound at regular timepoints after wounding. After initial transient inward currents (below zero), the current reversed and the outward currents (positive) rose slowly and plateaued.
<img src="/files/ftp_upload/2358/2358fig6.jpg" alt="Figure 6" /> Figure 6. Lab-book sketches showing probe measuring positions. A. Rat brain; red dots show measuring positions and probe symbols show probe orientation. B. Rat cornea wound; red dots show measuring positions and arrows show direction of current measured.
<img src="/files/ftp_upload/2358/2358tab1.jpg" alt="Table 1" /> Table 1. Example of Excel spreadsheet for storing and quantifying probe data. pN2_cal1 = starting calibration; cal+ and cal- = calibration values in mV; VR = voltage range, peak = sample measurements in mV, i/o = flow of current (into or out of sample).

<ol>
	<li>Reid, B., Nuccitelli, R., Zhao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-invasive+measurement+of+bioelectric+currents+with+a+vibrating+probe.">Non-invasive measurement of bioelectric currents with a vibrating probe.</a> Nat. Protoc. 2, 661-9282 (2007).</li><li>Reid, B., Song, B., McCaig, C. D., Zhao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wound+healing+in+rat+cornea:+the+role+of+electric+currents.">Wound healing in rat cornea: the role of electric currents.</a> FASEB J. 19, 379-386 (2005).</li><li>Lois, N., Reid, B., Song, B., Zhao, M., Forrester, J. V., McCaig, C. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electric+currents+and+lens+regeneration+in+the+rat.">Electric currents and lens regeneration in the rat.</a> Exp. Eye Res. 90, 316-323 (2010).</li><li>Wang, E., Reid, B., Lois, N., Forrester, J. V., McCaig, C. D., Zhao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrical+inhibition+of+lens+epithelial+cell+proliferation:+an+additional+factor+in+secondary+cataract.">Electrical inhibition of lens epithelial cell proliferation: an additional factor in secondary cataract.</a> FASEB J. 19, 842-844 (2005).</li><li>Guo, A., Song, B., Reid, B., Gu, Y., Forrester, J. V., Jahoda, C., Zhao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+physiological+electric+fields+on+migration+of+human+dermal+fibroblasts.">Effects of physiological electric fields on migration of human dermal fibroblasts.</a> J. Invest. Derm. , (2010).</li><li>Reid, B., Song, B., Zhao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electric+currents+in+Xenopus+tadpole+tail+regeneration.">Electric currents in Xenopus tadpole tail regeneration.</a> Dev. Biol. 335, 198-207 (2009).</li><li>Zhao, M., Song, B., Pu, J., Wada, T., Reid, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrical+signals+control+wound+healing+through+phosphatidylinositol-3-OH+kinase-&#947;.">Electrical signals control wound healing through phosphatidylinositol-3-OH kinase-&#947;.</a> 442, 457-460 (2006).</li><li>Reid, B., EO, G. r. a. u. e. -. H. e. r. n. a. n. d. e. z., Mannis, M. J., Zhao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulating+endogenous+electric+currents+in+human+corneal+wounds+-+a+novel+approach+of+bioelectric+stimulation+without+electrodes.">Modulating endogenous electric currents in human corneal wounds - a novel approach of bioelectric stimulation without electrodes.</a> Cornea. , (2010).</li><li>Nuccitelli, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+ultrasensitive+vibrating+probe+for+measuring+steady+extracellular+currents.">An ultrasensitive vibrating probe for measuring steady extracellular currents.</a> J. Cell Biol. 63, 614-628 (1974).</li><li>Hotary, K. B., Nuccitelli, R., Robinson, K. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+computerized+2-dimensional+vibrating+probe+for+mapping+extracellular+current+patterns.">A computerized 2-dimensional vibrating probe for mapping extracellular current patterns.</a> J. Neurosci. Meth. 43, 55-67 (1992).</li><li>Nuccitelli, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endogenous+ion+currents+and+DC+electric+fields+in+multicellular+animal+tissues.">Endogenous ion currents and DC electric fields in multicellular animal tissues.</a> Bioelectromagnetics Supplement. 1, 147-157 (1992).</li><li>Levin, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioelectric+mechanisms+in+regeneration:+Unique+aspects+and+future+perspectives.">Bioelectric mechanisms in regeneration: Unique aspects and future perspectives.</a> Seminars in Cell Dev. Biol. 20, 543-556 (2009).</li><li>Zhao, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electric+fields+in+wound+healing+-+An+overriding+signal+that+directs+cell+migration.">Electric fields in wound healing - An overriding signal that directs cell migration.</a> Seminars in Cell Dev. Biol. 20, 674-682 (2009).</li></ol>

No conflicts of interest declared.

We describe a low cost, basic, but highly sensitive vibrating probe system for measuring non-invasively electric current in a variety of biological systems.
Possible Modifications
<ol>
	<li>If platinum/iridium electrodes (World Precision Instruments; cat # PTM23B20) are used instead of stainless steel, then the gold plating stage can be eliminated.</li>
</ol>
Applications
We have used the vibrating probe to measure electric current in: ra...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Eligoy-Stainless Electrode</td>
<td>&nbsp;</td>
<td>World Precision Instruments</td>
<td>SSM33A70</td>
<td>76 mm, 7 M&Omega;, 1-2&mu;m tip</td>
</tr>
<tr>
<td>Gold R30 connector</td>
<td>&nbsp;</td>
<td>www.vectorelect.com</td>
<td>R30</td>
<td>Re-usable</td>
</tr>
<tr>
<td>Silver-loaded epoxy</td>
<td>&nbsp;</td>
<td>3M</td>
<td>SL65</td>
<td>Mix 1-part Resin with 1-part Hardener</td>
</tr>
<tr>
<td>Dissecting microscope</td>
<td>&nbsp;</td>
<td>Olympus</td>
<td>SZ40</td>
<td>Magnification x6 to x40</td>
</tr>
<tr>
<td>Potassium dicyanoaurate (KAu(CN)2)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>379867</td>
<td>CAUTION: Toxic</td>
</tr>
<tr>
<td>Chloroplatinic acid hydrate (H2PtCl6 x 6H2O)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>520896</td>
<td>CAUTION: Toxic</td>
</tr>
<tr>
<td>Lead(II) acetate trihydrate (Pb(CH3CO2)2 x 3H2O)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>185191</td>
<td>CAUTION: Toxic</td>
</tr>
<tr>
<td>Nano-Amp power source</td>
<td>&nbsp;</td>
<td>Made in-house</td>
<td>-</td>
<td>Powered by six 1.5 V (AAA) batteries</td>
</tr>
<tr>
<td>3-dimensional micro-positioner</td>
<td>&nbsp;</td>
<td>Line Tool Co.</td>
<td>Model H</td>
<td>&nbsp;</td>
<tr>
<td>Lock-in amplifier</td>
<td>&nbsp;</td>
<td>Stanford Research Systems</td>
<td>SR530</td>
<td>&nbsp;</td>
<tr>
<td>Digital I/O interface</td>
<td>&nbsp;</td>
<td>National instruments</td>
<td>PCI-6220</td>
<td>&nbsp;</td>
<tr>
<td>Shielded Connector Block with BNC connections</td>
<td>&nbsp;</td>
<td>National instruments</td>
<td>BNC-2110</td>
<td>&nbsp;</td>
<tr>
<td>Strathclyde Electrophysiology Software</td>
<td>&nbsp;</td>
<td>University of Strathclyde Institute of Pharmacy and Biomedical Sciences, UK</td>
<td>WinWCP V4.1.5</td>
<td>Free download from: <a href="http://spider.science.strath.ac.uk/sipbs/software_ses.htm">http://spider.science.strath.ac.uk/sipbs/software_ses.htm</a></td>
</tr>
<tr>
<td>Calibration Chamber</td>
<td>&nbsp;</td>
<td>Made in-house</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Constant Current Calibrator</td>
<td>&nbsp;</td>
<td>Vibrating Probe Company</td>
<td>&nbsp;</td>
<td>Powered by one 9 V (PP3) battery</td>
</tr>		</tbody>
		</table>
		</div>

measurement of bioelectric current with a vibrating probe

Electric fields, generated by active transport of ions, are present in many biological systems and often serve important functions in tissues and organs. For example, they play an important role in directing cell migration during wound healing. Here we describe the manufacture and use of ultrasensitive vibrating probes for measuring extracellular electric currents. The probe is an insulated, sharpened metal wire with a small platinum-black tip (30-35 &mu;m), which can detect ionic currents in the &mu;A/cm2 range in physiological saline. The probe is vibrated at about 200 Hz by a piezoelectric bender. In the presence of an ionic current, the probe detects a voltage difference between the extremes of its movement. A lock-in amplifier filters out extraneous noise by locking on to the probe's frequency of vibration. Data are recorded onto computer. The probe is calibrated at the start and end of experiments in appropriate saline, using a chamber which applies a current of exactly 1.5 &mu;A/cm2. We describe how to make the probes, set up the system and calibrate. We also demonstrate the technique of cornea measurement, and show some representative results from different specimens (cornea, skin, brain).

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Measurement of Bioelectric Current with a Vibrating Probe

The manufacture, calibration and use of non-invasive vibrating probes to measure bioelectric current in various biological systems is described.

Electric fields, generated by active transport of ions, are present in many biological systems and often serve important functions in tissues and organs. ...

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14.1K Views.  University of California, Davis. The manufacture, calibration and use of non-invasive vibrating probes to measure bioelectric current in various biological systems is described.

Video: Measurement of Bioelectric Current with a Vibrating Probe

Measurement of Aggregate Cohesion by Tissue Surface Tensiometry

Rigorous measurement of intercellular binding energy can only be made using methods grounded in thermodynamic principles in systems at equilibrium. We have developed tissue surface tensiometry (TST) specifically to measure the surface free energy of interaction between cells. The biophysical concepts underlying TST have been previously described in detail1,2. The method is based on the observation that mutually cohesive cells, if maintained in shaking culture, will spontaneously assemble into clusters. Over time, these clusters will round up to form spheres. This rounding-up behavior mimics the behavior characteristic of liquid systems. Intercellular binding energy is measured by compressing spherical aggregates between parallel plates in a custom-designed tissue surface tensiometer. The same mathematical equation used to measure the surface tension of a liquid droplet is used to measure surface tension of 3D tissue-like spherical aggregates. The cellular equivalent of liquid surface tension is intercellular binding energy, or more generally, tissue cohesivity. Previous studies from our laboratory have shown that tissue surface tension (1) predicts how two groups of embryonic cells will interact with one another1-5, (2) can strongly influence the ability of tissues to interact with biomaterials6, (3) can be altered not only through direct manipulation of cadherin-based intercellular cohesion7, but also by manipulation of key ECM molecules such as FN8-11 and 4) correlates with invasive potential of lung cancer12, fibrosarcoma13, brain tumor14 and prostate tumor cell lines15. In this article we will describe the apparatus, detail the steps required to generate spheroids, to load the spheroids into the tensiometer chamber, to initiate aggregate compression, and to analyze and validate the tissue surface tension measurements generated.

We describe a method of measuring binding energy, expressible as tissue surface tension, between cells within 3D tissue-like aggregates. Differences in tissue surface tension have been demonstrated to correlate with invasiveness of lung, muscle, and brain tumors, and are fundamental determinants of establishing spatial relationships between different cell types.

Rigorous measurement of intercellular binding energy can only be made using methods grounded in thermodynamic principles in systems at equilibrium.  We ...

Fluorescent Nanoparticles for the Measurement of Ion Concentration in Biological Systems

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion fluxes at the cellular level are required for initiating action potentials in excitable cells.2 Sodium regulation plays an important role in both of these cases; however, no method currently exists for continuously monitoring sodium levels in vivo 3 and intracellular sodium probes 4 do not provide similar detailed results as calcium probes. In an effort to fill both of these voids, fluorescent nanosensors have been developed that can monitor sodium concentrations in vitro and in vivo.5,6 These sensors are based on ion-selective optode technology and consist of plasticized polymeric particles in which sodium specific recognition elements, pH-sensitive fluorophores, and additives are embedded.7-9 Mechanistically, the sodium recognition element extracts sodium into the sensor. 10 This extraction causes the pH-sensitive fluorophore to release a hydrogen ion to maintain charge neutrality within the sensor which causes a change in fluorescence. The sodium sensors are reversible and selective for sodium over potassium even at high intracellular concentrations.6 They are approximately 120 nm in diameter and are coated with polyethylene glycol to impart biocompatibility. Using microinjection techniques, the sensors can be delivered into the cytoplasm of cells where they have been shown to monitor the temporal and spatial sodium dynamics of beating cardiac myocytes.11 Additionally, they have also tracked real-time changes in sodium concentrations in vivo when injected subcutaneously into mice.3 Herein, we explain in detail and demonstrate the methodology for fabricating fluorescent sodium nanosensors and briefly demonstrate the biological applications our lab uses the nanosensors for: the microinjection of the sensors into cells; and the subcutaneous injection of the sensors into mice.

Fluorescent nanoparticles produced in our lab are used for imaging ion concentrations and ion fluxes in biological systems such as cells during signaling and interstitial fluid during physiological homeostasis.

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion ...

Microfluidic Fabrication of Polymeric and Biohybrid Fibers with Predesigned Size and Shape

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and used to dictate the shape as well as the diameter of a core stream.&#160;Grooves in the top and bottom of a microfluidic channel were designed to direct the sheath fluid and shape the core fluid.&#160;By matching the viscosity and hydrophilicity of the sheath and core fluids, the interfacial effects are minimized and complex fluid shapes can be formed.&#160;Controlling the relative flow rates of the sheath and core fluids determines the cross-sectional area of the core fluid.&#160;Fibers have been produced with sizes ranging from 300 nm to ~1 mm, and fiber cross-sections can be round, flat, square, or complex as in the case with double anchor fibers. Polymerization of the core fluid downstream from the shaping region solidifies the fibers.&#160;Photoinitiated click chemistries are well suited for rapid polymerization of the core fluid by irradiation with ultraviolet light.&#160;Fibers with a wide variety of shapes have been produced from a list of polymers including liquid crystals, poly(methylmethacrylate), thiol-ene and thiol-yne resins, polyethylene glycol, and hydrogel derivatives. Minimal shear during the shaping process and mild polymerization conditions also makes the fabrication process well suited for encapsulation of cells and other biological components.

Two adjacent fluids passing through a grooved microfluidic channel can be directed to form a sheath around a prepolymer core; thereby&#160;determining both shape and cross-section. Photoinitiated polymerization, such as thiol click chemistry, is well suited for rapidly solidifying the core fluid into a microfiber with predetermined size and shape.

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and ...

Quantitative Optical Microscopy: Measurement of Cellular Biophysical Features with a Standard Optical Microscope

We describe the use of a standard optical microscope to perform quantitative measurements of mass, volume, and density on cellular specimens through a combination of bright field and differential interference contrast imagery. Two primary approaches are presented: noninterferometric quantitative phase microscopy (NIQPM), to perform measurements of total cell mass and subcellular density distribution, and Hilbert transform differential interference contrast microscopy (HTDIC)&#160;to determine volume. NIQPM is based on a simplified model of wave propagation, termed the paraxial approximation, with three underlying assumptions: low numerical aperture (NA) illumination, weak scattering, and weak absorption of light by the specimen. Fortunately, unstained cellular specimens satisfy these assumptions and low NA illumination is easily achieved on commercial microscopes. HTDIC is used to obtain volumetric information from through-focus DIC imagery under high NA illumination conditions. High NA illumination enables enhanced sectioning of the specimen along the optical axis. Hilbert transform processing on the DIC image stacks greatly enhances edge detection algorithms for localization of the specimen borders in three dimensions by separating the gray values of the specimen intensity from those of the background. The primary advantages of NIQPM and HTDIC lay in their technological accessibility using &#8220;off-the-shelf&#8221; microscopes. There are two basic limitations of these methods: slow z-stack acquisition time on commercial scopes currently abrogates the investigation of phenomena faster than 1 frame/minute, and secondly, diffraction effects restrict the utility of NIQPM and HTDIC to objects from 0.2 up to 10 (NIQPM) and 20 (HTDIC) &#956;m in diameter, respectively. Hence, the specimen and its associated time dynamics of interest must meet certain size and temporal constraints to enable the use of these methods. Excitingly, most fixed cellular specimens are readily investigated with these methods.

We describe the use of a standard optical microscope to perform quantitative measurements of cellular mass, volume, and density through a combination of bright field and differential interference contrast imagery.

We describe the use of a standard optical microscope to perform quantitative measurements of mass, volume, and density on cellular specimens through a ...

A Microfluidic Technique to Probe Cell Deformability

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

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

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

Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. Kelvin probe force microscopy (KPFM) is a module of atomic force microscopy (AFM) that measures the contact potential difference between surfaces at the nano-scale. The combination of KPFM with AFM allows for the simultaneous generation of surface potential and topographical maps of biological samples such as bacterial cells. Here, we employ KPFM to examine the effects of surface potential on microbial adhesion to medically relevant surfaces such as stainless steel and gold. Surface potential maps revealed differences in surface potential for microbial membranes on different material substrates. A step-height graph was generated to show the difference in surface potential at a boundary area between the substrate surface and microorganisms. Changes in cellular membrane surface potential have been linked with changes in cellular metabolism and motility. Therefore, KPFM represents a powerful tool that can be utilized to examine the changes of microbial membrane surface potential upon adhesion to various substrate surfaces. In this study, we demonstrate the procedure to characterize the surface potential of individual methicillin-resistant Staphylococcus aureus USA100 cells on stainless steel and gold using KPFM.

Here, we present a protocol explaining the use of Kelvin probe force microscopy as a tool for generating high resolution nano-scale surface potential maps. This tool was applied to assess the role of surface potential on the binding capacity of microorganisms to substrate surfaces.

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. ...

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 ...

Fabrication of a Dipole-assisted Solid Phase Extraction Microchip for Trace Metal Analysis in Water Samples

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water samples. A brief overview of the evolution of chip-based SPE techniques is provided. This is followed by an introduction to specific polymeric materials and their role in SPE. To develop an innovative dipole-assisted SPE technique, a chlorine (Cl)-containing SPE functionality was implanted into a poly(methyl methacrylate) (PMMA) microchip. Herein, diverse analytical techniques including contact angle analysis, Raman spectroscopic analysis, and laser ablation-inductively coupled plasma-mass spectrometry (LA-ICP-MS) analysis were employed to validate the utility of the implantation protocol of the C-Cl moieties on the PMMA. The analytical results of the X-ray absorption near-edge structure (XANES) analysis also demonstrated the feasibility of the Cl-containing PMMA used as an extraction medium by virtue of the dipole-ion interactions between the highly electronegative C-Cl moieties and the positively charged metal ions.

The fabrication protocol of a dipole-assisted solid phase extraction microchip for the trace metal analysis is presented.

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water ...

Rapid Subtractive Patterning of Live Cell Layers with a Microfluidic Probe

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one particular implementation of the MFP, the hierarchical hydrodynamic flow confinement (hHFC), multiple liquids are simultaneously brought in contact with a substrate. Local chemical action and liquid shaping using the hHFC, is exploited to create cell patterns by locally lysing and removing cells. By utilizing the scanning ability of the MFP, user-defined patterns of cell monolayers are created. This protocol enables rapid, real-time and spatially controlled cell patterning, which can allow selective cell-cell and cell-matrix interaction studies.

We present a protocol to perform subtractive patterning of live cell monolayers on a surface. This is achieved by local and selective lysis of adherent cells using a microfluidic probe (MFP). The cell lysate retrieved from local regions can be used for downstream analysis, enabling molecular profiling studies.

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one ...

Optimized Setup and Protocol for Magnetic Domain Imaging with In Situ Hysteresis Measurement

This paper elaborates the sample preparation protocols required to obtain optimal domain patterns using the Bitter method, focusing on the extra steps compared to standard metallographic sample preparation procedures. The paper proposes a novel bespoke rig for dynamic domain imaging with in situ BH (magnetic hysteresis) measurements and elaborates the protocols for the sensor preparation and the use of the rig to ensure accurate BH measurement. The protocols for static and ordinary dynamic domain imaging (without in situ BH measurements) are also presented. The reported method takes advantage of the convenience and high sensitivity of the traditional Bitter method and enables in situ BH measurement without interrupting or interfering with the domain wall movement processes. This facilitates establishing a direct and quantitative link between the domain wall movement processes&#8211;microstructural feature interactions in ferritic steels with their BH loops. This method is anticipated to become a useful tool for the fundamental study of microstructure&#8211;magnetic property relationships in steels and to help interpret the electromagnetic sensor signals for non-destructive evaluation of steel microstructures.

Optimized Setup and Protocol for Magnetic Domain Imaging with In Situ Hysteresis Measurement

This paper elaborates the sample and sensor preparation procedures and the protocols for using the test rig particularly for dynamic domain imaging with in situ BH measurements in order to achieve optimal domain pattern quality and accurate BH measurements.

This paper elaborates the sample preparation protocols required to obtain optimal domain patterns using the Bitter method, focusing on the extra steps ...