We thank Dr. Olu Ajijola (UCLA Cardiac Arrhythmia Center) for expert support for the in vivo experiments. This work was supported by NIH U01 EB025138 (JLA, CS).

All animal experiments were approved by the University of California, Los Angeles Animal Research Committee and performed following the guidelines set forth by the National Institutes of Health Guide for the Care and Use of Laboratory Animals (8th edition, 2011). Adult male Yorkshire pigs of approximately 75 kg were used for in vivo studies10.
1. Capacitive immunoprobe fabrication and functionalization
<ol>
	<li>Cut a 25 cm length of perfluoroalkoxy (PFA)-coated platinum wire (see Table of Materials) and strip approximately 5 mm of PFA coating from one end using a scalpel, being careful not to cut into the platinum wire.</li>
	<li>Insert the stripped end of the platinum wire into a gold-plated 1 mm male connector pin and crimp the connector pin teeth around the stripped end of the platinum wire using needle-nose pliers (see Table of Materials).</li>
	<li>Solder the platinum wire to the gold-plated connector pin. Be careful not to use an excessive amount of solder.</li>
	<li>Prepare dopamine solution by dissolving 50 mg of dopamine HCl in 50 mL of 10 mM phosphate-buffered saline (PBS, pH 6.0) by stirring.</li>
	<li>Once the dopamine is completely dissolved, place the tip of the platinum wire in the vessel containing the freshly made dopamine-supplemented PBS. Plug a gold connector pin into a channel of the headstage (see Table of Materials).</li>
	<li>Connect the AgCl disc electrode (ground electrode, see Table of Materials) to the ground channel in the headstage. Place the AgCl disc in the vessel containing dopamine-supplemented PBS and platinum wire; be careful only to submerge the disc electrode and not any length of the wire or solder. Connect a wire shunt into the reference channels of the headstage prior to proceeding.</li>
	<li>Open the interactive data acquisition software (see Table of Materials). Prepare a sawtooth electrodeposition command potential protocol with the following parameters: start potential = &#8722;0.6 V; end potential = +0.65 V; scan rate = 0.04 V&#8729;s-1; duration of deposition = 420 s. Begin the polydopamine deposition protocol, ensuring that all the wires are connected properly.</li>
	<li>After completing the polydopamine deposition, remove the AgCl ground pellet and the tip of the platinum wire from the vessel, being careful not to disturb the tip of the platinum wire electrode. Place the tip of the wire into a microtube containing PBS (pH 7.4) for 2-5 min as the antibody solution is prepared; ensure the wire tip does not contact the sides or bottom of the microtube. 
	NOTE: Antibody solution can be made during polydopamine deposition; however, the transfer of the platinum wire from the dopamine-containing vessel to the microtube of PBS following polydopamine deposition should not be skipped.</li>
	<li>Prepare the antibody solution. Combine the antibody of interest with PBS (pH 7.4) in a 1:20 ratio in a vessel of an appropriate size (e.g., a microtube). 
	NOTE: The anti-NPY monoclonal antibody (see Table of Materials) used here was aliquoted at 1 mg/mL; an example antibody preparation here would be 4 &#181;L of antibody to 76 &#181;L of PBS.</li>
	<li>Soak the polydopamine-deposited tip of the platinum electrode in antibody solution for a minimum of 2 h at room temperature, again ensuring the platinum wire tip is suspended in solution and not resting on the interior surface of the microtube. 
	NOTE: Recent implementation of this technique has favored using the platinum wire electrode immediately following this step instead of wet or dry storage for later use.</li>
	<li>After soaking in the antibody solution, briefly rinse the newly-functionalized capacitive immunoprobe (CI probe) tip in PBS (pH 7.4). The probe is now ready for use.</li>
</ol>
2. Experimental setup for in vitro detection and measurement of peptide
<ol>
	<li>Place the functional tip of the CI probe into the flow chamber, taking care not to disturb the tip of the electrode in any way, as doing so may damage the sensory tip of the probe. 
	NOTE: The flow chamber was created by pouring silicone elastomer (see Table of Materials) into a 35 mm culture dish with an elongated ovoid space-filler in the center of the dish. After hardening, the ovoid shape is removed from the elastomer. The chamber is then superfused with Tris-buffered saline (TBS) and allowed a flow rate of 3 mL/min. Ensure the inflow and outflow maintain the fluid level in the chamber such that no tidal action of the superfusate is observed. The flow must remain in place for as long as the CI probe is in use.</li>
	<li>Prior to the first experimental test, perform a TBS standard run to condition the CI probe. Set up the following command voltage protocol: positive step potential = +100 mV; negative step potential = &#8722;5 mV; step duration = 20 ms; duration of acquisition = 600 s. 
	NOTE: It is important to allow for the equilibration of the probe during the initial phase of cycling command potential prior to data acquisition.</li>
	<li>Create a solution of the peptide of interest using the same TBS to maintain the superfusate&#39;s composition. Set up a manifold system where the superfusate can be switched between TBS and peptide-supplemented TBS without introducing bubbles into the tubing system or flow chamber. 
	NOTE: Synthetic porcine NPY peptide (see Table of Materials) was used in the present study.</li>
	<li>Set up the peptide-sensing data acquisition protocol using TBS standard parameters (see step 2.2.). 
	&#8203;NOTE: In this implementation, the duration of each experimental test was 360 s (120 s TBS, 120 s peptide-supplemented TBS, 120 s TBS).</li>
</ol>
3. Adaptation of the CI probe for in vivo use
<ol>
	<li>Prior to polydopamine deposition (step 1.7.), thread the exposed tip of the platinum wire electrode through a 22 G hypodermic needle, leaving approximately 2 mm beyond the tip of the needle. Using forceps, gently bend back the tip of the platinum wire electrode, creating a &#34;barb&#34; that hangs from the end of the hypodermic needle.
	<ol>
		<li>Gently withdraw the needle from the barbed tip, leaving enough wire to place in the vessel without the needle contacting the fluid. Proceed with steps 1.4.-1.11. 
		NOTE: Depending on the in vivo setup, it may be necessary to cut a length of platinum wire longer than 25 cm.</li>
	</ol>
	</li>
	<li>Before connecting the CI probe to the headstage, ensure the entire electrical setup is properly grounded. Failure to do so may introduce unwanted electrical interference during the experimental recordings.</li>
	<li>Anesthetize the animals following a previously published report10.</li>
	<li>Perform surgery to expose the region of interest. 
	NOTE: A median sternotomy was performed in the present study to expose the heart. Please see Kluge et al.10 for details on animal surgery.</li>
	<li>Gently remove the functionalized tip from the PBS rinse (step 1.11.), forward the hydodermic needle to the barbed C.I. probe and gently implant it in the region of interest prior to plugging the gold connector pin into the headstage. Once implanted, withdraw the hypodermic needle, leaving the electrode in place. 
	NOTE: For the present study, the probe was placed in the mid-lateral wall of the left ventricular myocardium10.</li>
	<li>After ensuring proper electrical setup, proceed with standard and experimental testing protocols (step 2.2. and step 2.4.).</li>
	<li>Upon completion of the experiments, euthanize the animal following institutionally approved techniques. 
	NOTE: In the present study, the animals are euthanized under deep anesthesia via induction of ventricular fibrillation10.</li>
</ol>

<ol>
	<li>Chow, S. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+Biomarkers+for+the+prevention,+assessment,+and+management+of+heart+failure:+A+scientific+statement+from+the+American+Heart+Association.">Role of Biomarkers for the prevention, assessment, and management of heart failure: A scientific statement from the American Heart Association.</a> Circulation. 135 (22), 1054-1091 (2017).</li><li>Goldstein, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adrenal+responses+to+stress.">Adrenal responses to stress.</a> Cellular and Molecular Neurobiology. 30 (8), 1433-1440 (2010).</li><li>Ullman, B., Hulting, J., Lundberg, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prognostic+value+of+plasma+neuropeptide-Y+in+coronary+care+unit+patients+with+and+without+acute+myocardial+infarction.">Prognostic value of plasma neuropeptide-Y in coronary care unit patients with and without acute myocardial infarction.</a> European Heart Journal. 15 (4), 454-461 (1994).</li><li>Farrell, D. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Angiotensin+II+modulates+catecholamine+release+into+interstitial+fluid+of+canine+myocardium+in+vivo.">Angiotensin II modulates catecholamine release into interstitial fluid of canine myocardium in vivo.</a> American Journal of Physiology-Heart and Circulatory Physiology. 281 (2), 813-822 (2001).</li><li>Ardell, J. L., Foreman, R. D., Armour, J. A., Shivkumar, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+sympathectomy+and+spinal+cord+stimulation+attenuate+reflex-mediated+norepinephrine+release+during+ischemia+preventing+ventricular+fibrillation.">Cardiac sympathectomy and spinal cord stimulation attenuate reflex-mediated norepinephrine release during ischemia preventing ventricular fibrillation.</a> JCI Insight. 4 (23), 131648 (2019).</li><li>Franco-Cereceda, A., Lundberg, J. M., Dahlof, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuropeptide+Y+and+sympathetic+control+of+heart+contractility+and+coronary+vascular+tone.">Neuropeptide Y and sympathetic control of heart contractility and coronary vascular tone.</a> Acta Physiologica Scandinavica. 124 (3), 361-369 (1985).</li><li>Habecker, B. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+and+cellular+neurocardiology:+development,+and+cellular+and+molecular+adaptations+to+heart+disease.">Molecular and cellular neurocardiology: development, and cellular and molecular adaptations to heart disease.</a> The Journal of Physiology. 594 (14), 3853-3875 (2016).</li><li>Hoang, J. D., Salavatian, S., Yamaguchi, N., Swid, M. A., Vaseghi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+sympathetic+activation+circumvents+high-dose+beta+blocker+therapy+in+part+through+release+of+neuropeptide+Y.">Cardiac sympathetic activation circumvents high-dose beta blocker therapy in part through release of neuropeptide Y.</a> JCI Insight. 5 (11), 135519 (2020).</li><li>Rigel, D. F. <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+neuropeptides+on+heart+rate+in+dogs:+comparison+of+VIP,+PHI,+NPY,+CGRP,+and+NT.">Effects of neuropeptides on heart rate in dogs: comparison of VIP, PHI, NPY, CGRP, and NT.</a> American Journal of Physiology. 255, 311-317 (1988).</li><li>Kluge, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+measurement+of+cardiac+neuropeptide+dynamics+by+capacitive+immunoprobe+in+the+porcine+heart.">Rapid measurement of cardiac neuropeptide dynamics by capacitive immunoprobe in the porcine heart.</a> American Journal of Physiology-Heart and Circulatory Physiology. 320 (1), 66-76 (2021).</li><li>Berggren, C., Bjarnason, B., Johansson, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capacitive+biosensors.">Capacitive biosensors.</a> Electroanalysis. 13 (3), 173-180 (2001).</li><li>Prodromidis, M. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impedimetric+immunosensors-A+review.">Impedimetric immunosensors-A review.</a> Electrochimica Acta. 55 (14), 4227-4233 (2010).</li><li>Lee, H., Dellatore, S. M., Miller, W. M., Messersmith, P. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mussel-inspired+surface+chemistry+for+multifunctional+coatings.">Mussel-inspired surface chemistry for multifunctional coatings.</a> Science. 318 (5849), 426-430 (2007).</li><li>Leszczyszyn, D. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Secretion+of+catecholamines+from+individual+adrenal+medullary+chromaffin+cells.">Secretion of catecholamines from individual adrenal medullary chromaffin cells.</a> Journal of Neurochemistry. 56 (6), 1855-1863 (1991).</li><li>Pihel, K., Schroeder, T. J., Wightman, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+and+selective+cyclic+voltammetric+measurements+of+epinephrine+and+norepinephrine+as+a+method+to+measure+secretion+from+single+bovine+adrenal+medullary+cells.">Rapid and selective cyclic voltammetric measurements of epinephrine and norepinephrine as a method to measure secretion from single bovine adrenal medullary cells.</a> Analytical Chemistry. 66 (24), 4532-4537 (1994).</li><li>Jaffe, E. H., Marty, A., Schulte, A., Chow, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extrasynaptic+vesicular+transmitter+release+from+the+somata+of+substantia+nigra+neurons+in+rat+midbrain+slices.">Extrasynaptic vesicular transmitter release from the somata of substantia nigra neurons in rat midbrain slices.</a> The Journal of Neuroscience. 18 (10), 3548-3553 (1998).</li><li>Walsh, P. L., Petrovic, J., Wightman, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinguishing+splanchnic+nerve+and+chromaffin+cell+stimulation+in+mouse+adrenal+slices+with+fast-scan+cyclic+voltammetry.">Distinguishing splanchnic nerve and chromaffin cell stimulation in mouse adrenal slices with fast-scan cyclic voltammetry.</a> American Journal of Physiology-Cell Physiology. 300 (1), 49-57 (2011).</li><li>Wolfe, J. T., Wang, H., Perez-Reyes, E., Barrett, P. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stimulation+of+recombinant+Ca(v)3.2,+T-type,+Ca(2+)+channel+currents+by+CaMKIIgamma(C).">Stimulation of recombinant Ca(v)3.2, T-type, Ca(2+) channel currents by CaMKIIgamma(C).</a> The Journal of Physiology. 538, 343-355 (2002).</li><li>Chan, S. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+in+vivo+detection+of+myocardial+norepinephrine+levels+in+the+beating+porcine+heart.">Fast in vivo detection of myocardial norepinephrine levels in the beating porcine heart.</a> American Journal of Physiology-Heart and Circulatory Physiology. 318 (5), 1091-1099 (2020).</li><li>Chow, R. H., von R&#252;den, L., Sakmann, B., Neher, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Single-Channel Recording, Second Edition. , 245-275 (1995).</li><li>Ren, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Facile,+high+efficiency+immobilization+of+lipase+enzyme+on+magnetic+iron+oxide+nanoparticles+via+a+biomimetic+coating.">Facile, high efficiency immobilization of lipase enzyme on magnetic iron oxide nanoparticles via a biomimetic coating.</a> BMC Biotechnology. 11, 63 (2011).</li></ol>

The authors declare no conflicts of interest, financial or otherwise.

The present protocol describes the manufacture and testing of a capacitive immunoprobe (CI probe) capable of detecting and measuring biomarkers of interest in both in vitro and in vivo settings. Detection is achieved by trapping the biomarker at the electrode tip. The trapping event alters the capacitive junction between a platinum wire capacitive immunoprobe and the surrounding conductive fluid environment, measured as a change in charge current in response to a potential shift in the probe. A unique e...

Several chemical methods for detecting and quantifying biomarkers are routinely utilized in both protein chemistry and clinical diagnostics, particularly in cancer diagnoses and the assessment of cardiovascular disease progression. Currently, methods such as high-performance liquid chromatography (HPLC), enzyme-linked immunosorbent assay (ELISA), and mass spectrometry rely on sample collection from the vascular compartment1,2,3 by bulk fluid draw or the interstitial compartment by microdialysis. Microdialysis employs a semipermeable membrane tube of known length that is placed in a region of interest. Collection fluid is perfused through the tube over several minutes4 to collect the sample for analysis5, thus limiting temporal resolution. In this manner, samples collected only provide an averaged value over time of the local microenvironment and are limited by the perfusion rate and collection of sufficient sample volume. Moreover, these methods require the pooling of experimental data and signal averaging; therefore, they may fail to account for variability between subjects. Importantly, the time between sample collection and subsequent offline analysis precludes immediate clinical intervention and therapeutics.
In the present protocol, the use of a capacitive immunoprobe biosensor (CI probe) for the time-resolved electrical detection of specific bioactive peptides is outlined. Neuropeptide Y (NPY), released from post-ganglionic sympathetic neurons that innervate the vasculature, endocardium, cardiomyocytes, and intracardiac ganglia, is a major neuromodulatory peptide transmitter in the cardiovascular system6,7,8,9. The method presented here is designed to measure NPY, and the experimental feasibility is demonstrated in a porcine heart model. However, this approach applies to any bioactive peptide for which a selective antibody is available10. This method relies on the capacitive junction between a platinum wire probe and the conductive fluid at the functionalized tip11,12. In this application, the interaction was mediated through an antibody against the target neuropeptide (NPY), which was bound to the electrode tip, interfacing the conductive fluid environment. This functionalization was achieved through electrodeposition of reactive polydopamine onto the tip of the platinum wire probe10,13.
When the antibody-functionalized probe is placed in a region of interest in vivo, evoked endogenous NPY release leads to binding to the trapping antibodies on the probe tip, and the conductive fluid at the electrode surface is displaced by the NPY protein. Local alteration in the electrical environment results in the displacement of high-mobility, high-dielectric fluid with an immobile, statically charged molecule. This alters the electrode-fluid interface and, thus, its capacitance, which is measured as a change in charge current in response to a step-function command potential. A negative &#34;reset&#34; potential is employed immediately following each individual measurement cycle to repel bound NPY from the antibody through electrostatic interaction, thus clearing the antibody binding sites for subsequent rounds of measurement10. This effectively allows for the measurement of NPY in a time-resolved manner. The unique CI technique overcomes the limitations of the microdialysis-based immunochemical methods outlined above to measure dynamic biomarker levels from a single experiment without data pooling or signal averaging over several experiments9, providing data in nearly real time. Moreover, the ability to adapt this method to any biomarker of interest for which there exists an appropriate antibody on a time-resolved and localized scale provides a major technical advance in immunochemical measurement for the evaluation of disease progression and the guidance of therapeutic interventions.
The software for data acquisition and analysis was custom-written in IGOR Pro (a fully interactive software environment). An analog to digital converter (A/D) system issued a command voltage under computer control and acquired data from a custom amplifier. The amplifier possessed certain unique features. These included a feedback resistor (switchable) for each of the four acquisition channels, allowing for choosing 1 MOhm or 10 MOhm feedback voltage clamp circuits to integrate the variability of the electrode. A stage unit with a single head and mutual ground/reference circuit for all the four acquisition channels was also built to place the device close by the chest in a single physical module. A 1 MOhm feedback resistor setting was used to collect all the reported data.
The filter and gain settings were telegraphed from the amplifier and recorded within the data file. Data were filtered at 1 kHz via a 2-pole analog Bessel filter digitized at 10 kHz. The difference in potential between the probe and surrounding conductive solution creates a Helmholtz capacitive layer at the probe tip. Ligand binding to the antibody at the probe tip results in an altered local charge and, thus, a change in the Helmholtz capacitance. This change in the capacitive component of the circuit results in a shift in the magnitude of injected charge required to bring the probe to the potential in the step-function voltage protocol. Thus, the binding of a specific ligand to the functionalized probe results in an alteration in the electrode capacitance measurement as a change in peak capacitive current.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>AgCl disc electrode</td>
			<td>Warner Instruments (Holliston, MA)</td>
			<td>64-1307</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-NPY monoclonal antibody</td>
			<td>Abcam, (Cambridge, MA)</td>
			<td>ab112473</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom multichannel amplifier/ 1 M&#937; feedback resistor multichannel headstage</td>
			<td>NPI Electronic, (Tamm, Germany)</td>
			<td>NA</td>
			<td>Based on NPI VA-10M multichannel amplifier</td>
		</tr>
		<tr>
			<td>Dopamine HCl</td>
			<td>Sigma Aldrich (St. Louis, MO)</td>
			<td>H8502-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Gold-plated male connector pin</td>
			<td>AMP-TE Connectivity (Amplimite)</td>
			<td>6-66506-1</td>
			<td></td>
		</tr>
		<tr>
			<td>HEKA LIH 8+8 analog-to-digital/digital-to-analog device</td>
			<td>HEKA Elektronik, (Holliston, MA)</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Igor Pro data acquisition software, v. 7.08</td>
			<td>WaveMetrics, (Lake Oswego, OR)</td>
			<td></td>
			<td>Software driving command potential and data acquisition was custom written</td>
		</tr>
		<tr>
			<td>Masterflex L/S Standard Digital peristaltic pump</td>
			<td>Cole Palmer, (Vernon Hills, IL)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PFA-coated platinum wire</td>
			<td>A-M Systems, (Sequim, WA)</td>
			<td>773000</td>
			<td>0.005&#8221; bare diameter, 0.008&#8221; coated diameter</td>
		</tr>
		<tr>
			<td>Silicone elastomer</td>
			<td>World Precision Instruments (Sarasota, FL)</td>
			<td>SYLG184</td>
			<td></td>
		</tr>
		<tr>
			<td>Synthetic porcine NPY peptide</td>
			<td>Bachem (Torrance, CA)</td>
			<td>4011654</td>
			<td></td>
		</tr>
		<tr>
			<td>Synthetic porcine NPY peptide</td>
			<td>Bachem (Torrance, CA)</td>
			<td>4011654</td>
			<td></td>
		</tr>
	</tbody>
</table>

time-resolved in vivo measurement of neuropeptide dynamics by capacitive immunoprobe in porcine heart

The ability to measure biomarkers in vivo relevant to the assessment of disease progression is of great interest to the scientific and medical communities. The resolution of results obtained from current methods of measuring certain biomarkers can take several days or weeks to obtain, as they can be limited in resolution both spatially and temporally (e.g., fluid compartment microdialysis of interstitial fluid analyzed by enzyme-linked immunosorbent assay [ELISA], high-performance liquid chromatography [HPLC], or mass spectrometry); thus, their guidance of timely diagnosis and treatment is disrupted. In the present study, a unique technique for detecting and measuring peptide transmitters in vivo through the use of a capacitive immunoprobe biosensor (CI probe) is reported. The fabrication protocol and in vitro characterization of these probes are described. Measurements of sympathetic stimulation-evoked neuropeptide Y (NPY) release in vivo are provided. NPY release is correlated to the sympathetic release of norepinephrine for reference. The data demonstrate an approach for the fast and localized measurement of neuropeptides in vivo. Future applications include intraoperative real-time assessment of disease progression and minimally invasive catheter-based deployment of these probes.

Electrode fabrication and characterization 
A flexible capacitive immunoprobe (CI probes) was fabricated, and a representative image is depicted in Figure 1A. The electrode potential was set by a computer-controlled voltage clamp circuit (Figure 1B), and the electrode was immersed in a polydopamine solution made in PBS. Polydopamine was electrodeposited onto the conductive electrode tip13 for functionalization. The c...

Watch this Scientific Journal Video about Time-Resolved In Vivo Measurement of Neuropeptide Dynamics by Capacitive Immunoprobe in Porcine Heart at JoVE.com

Time-Resolved In Vivo Measurement of Neuropeptide Dynamics by Capacitive Immunoprobe in Porcine Heart

Established immunochemical methods to measure peptide transmitters in vivo rely on microdialysis or bulk fluid draw to obtain the sample for offline analysis. However, these suffer from spatiotemporal limitations. The present protocol describes the fabrication and application of a capacitive immunoprobe biosensor that overcomes the limitations of the existing techniques.

Time-Resolved In Vivo Measurement of Neuropeptide Dynamics by Capacitive Immunoprobe in Porcine Heart

The ability to measure biomarkers in vivo relevant to the assessment of disease progression is of great interest to the scientific and medical ...

time-resolved-vivo-measurement-neuropeptide-dynamics-capacitive

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

1.9K Views.  Case Western Reserve University. Established immunochemical methods to measure peptide transmitters in vivo rely on microdialysis or bulk fluid draw to obtain the sample for offline analysis. However, these suffer from spatiotemporal limitations. The present protocol describes the fabrication and application of a capacitive immunoprobe biosensor that overcomes the limitations of the existing techniques.

Video: Time-Resolved In Vivo Measurement of Neuropeptide Dynamics by Capacitive Immunoprobe in Porcine Heart

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not have been accomplished by other approaches1. With the use of the calcium- and voltage-sensitive dyes, optical mapping allows measurement of transmembrane action potentials and calcium transients with high spatial resolution without the physical contact with the tissue. This makes measurements of the cardiac electrical activity possible under many conditions where the use of electrodes is inconvenient or impossible1. For example, optical recordings provide accurate morphological changes of membrane potential during and immediately after stimulation and defibrillation, while conventional electrode techniques suffer from stimulus-induced artifacts during and after stimuli due to electrode polarization1. 
The Langendorff-perfused rabbit heart is one of the most studied models of human heart physiology and pathophysiology. Many types of arrhythmias observed clinically could be recapitulated in the rabbit heart model. It was shown that wave patterns in the rabbit heart during ventricular arrhythmias, determined by effective size of the heart and the wavelength of reentry, are very similar to that in the human heart2. It was also shown that critical aspects of excitation-contraction (EC) coupling in rabbit myocardium, such as the relative contribution of sarcoplasmic reticulum (SR), is very similar to human EC coupling3. Here we present the basic procedures of optical mapping experiments in Langendorff-perfused rabbit hearts, including the Langendorff perfusion system setup, the optical mapping systems setup, the isolation and cannulation of the heart, perfusion and dye-staining of the heart, excitation-contraction uncoupling, and collection of optical signals. These methods could be also applied to the heart from species other than rabbit with adjustments to flow rates, optics, solutions, etc.
Two optical mapping systems are described. The panoramic mapping system is used to map the entire epicardium of the rabbit heart4-7. This system provides a global view of the evolution of reentrant circuits during arrhythmogenesis and defibrillation, and has been used to study the mechanisms of arrhythmias and antiarrhythmia therapy8,9. The dual mapping system is used to map the action potential (AP) and calcium transient (CaT) simultaneously from the same field of view10-13. This approach has enhanced our understanding of the important role of calcium in the electrical alternans and the induction of arrhythmia14-16.

This article describes the basic procedures for conducting optical mapping experiments in the Langendorff-perfused rabbit heart using the panoramic imaging system, and the dual (voltage and calcium) imaging modality.

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not ...

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural biology 1. In recent years, this method has achieved great success in studying structures of proteins and macromolecular complexes 2, 3. Compared with electron cryomicroscopy (cryoEM), in which frozen hydrated protein samples are embedded in a thin layer of vitreous ice 4, negative staining is a simpler sample preparation method in which protein samples are embedded in a thin layer of dried heavy metal salt to increase specimen contrast 5. The enhanced contrast of negative stain EM allows examination of relatively small biological samples. In addition to determining three-dimensional (3D) structure of purified proteins or protein complexes 6, this method can be used for much broader purposes. For example, negative stain EM can be easily used to visualize purified protein samples, obtaining information such as homogeneity/heterogeneity of the sample, formation of protein complexes or large assemblies, or simply to evaluate the quality of a protein preparation.
In this video article, we present a complete protocol for using an EM to observe negatively stained protein sample, from preparing carbon coated grids for negative stain EM to acquiring images of negatively stained sample in an electron microscope operated at 120kV accelerating voltage. These protocols have been used in our laboratory routinely and can be easily followed by novice users.

Visualizing protein samples by negative stain electron microscopy (EM) has become a popular structural analysis method. It is useful for quantitative structural analysis, such as calculating a 3D reconstruction of the molecules being studied, and also for qualitative examination of the quality of protein preparations. In this article we present detailed protocols for preparing the EM grids, staining the sample and visualizing the sample in an electron microscope. Novice users can follow these protocols easily and to utilize negative stain EM as a routine assay, in addition to other biochemical assays, for evaluating their protein samples.

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural ...

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Procedure for Decellularization of Porcine Heart by Retrograde Coronary Perfusion

Perfusion-based whole organ decellularization has recently gained interest in the field of tissue engineering as a means to create site-specific extracellular matrix scaffolds, while largely preserving the native architecture of the scaffold. To date, this approach has been utilized in a variety of organ systems, including the heart, lung, and liver 1-5. Previous decellularization methods for tissues without an easily accessible vascular network have relied upon prolonged exposure of tissue to solutions of detergents, acids, or enzymatic treatments as a means to remove the cellular and nuclear components from the surrounding extracellular environment6-8. However, the effectiveness of these methods hinged upon the ability of the solutions to permeate the tissue via diffusion. In contrast, perfusion of organs through the natural vascular system effectively reduced the diffusion distance and facilitated transport of decellularization agents into the tissue and cellular components out of the tissue. Herein, we describe a method to fully decellularize an intact porcine heart through coronary retrograde perfusion. The protocol yielded a fully decellularized cardiac extracellular matrix (c-ECM) scaffold with the three-dimensional structure of the heart intact. Our method used a series of enzymes, detergents, and acids coupled with hypertonic and hypotonic rinses to aid in the lysis and removal of cells. The protocol used a Trypsin solution to detach cells from the matrix followed by Triton X-100 and sodium deoxycholate solutions to aid in removal of cellular material. The described protocol also uses perfusion speeds of greater than 2 L/min for extended periods of time. The high flow rate, coupled with solution changes allowed transport of agents to the tissue without contamination of cellular debris and ensured effective rinsing of the tissue. The described method removed all nuclear material from native porcine cardiac tissue, creating a site-specific cardiac ECM scaffold that can be used for a variety of applications.

A method to rapidly and completely remove cellular components from an intact porcine heart through retrograde perfusion is described. This method yields a site specific cardiac extracellular matrix scaffold which has the potential for use in multiple clinical applications.

Perfusion-based  whole organ decellularization has recently gained interest in the field of  tissue engineering as a means to create site-specific ...

Millifluidics for Chemical Synthesis and Time-resolved Mechanistic Studies

Procedures utilizing millifluidic devices for chemical synthesis and time-resolved mechanistic studies are described by taking three examples. In the first, synthesis of ultra-small copper nanoclusters is described. The second example provides their utility for investigating time resolved kinetics of chemical reactions by analyzing gold nanoparticle formation using in situ X-ray absorption spectroscopy. The final example demonstrates continuous flow catalysis of reactions inside millifluidic channel coated with nanostructured catalyst.

Millifluidic devices are utilized for controlled synthesis of nanomaterials, time-resolved analysis of reaction mechanisms and continuous flow catalysis.

Procedures utilizing millifluidic devices for chemical synthesis and time-resolved mechanistic studies are described by taking three examples. In the ...

Quantification of Global Diastolic Function by Kinematic Modeling-based Analysis of Transmitral Flow via the Parametrized Diastolic Filling Formalism

Quantitative cardiac function assessment remains a challenge for physiologists and clinicians.&#160;Although historically invasive methods have comprised the only means available, the development of noninvasive imaging modalities (echocardiography, MRI, CT) having high temporal and spatial resolution provide a new window for quantitative diastolic function assessment. Echocardiography is the agreed upon standard for diastolic function assessment, but indexes in current clinical use merely utilize selected features of chamber dimension (M-mode) or blood/tissue motion (Doppler) waveforms without incorporating the physiologic causal determinants of the motion itself.&#160;The recognition that all left ventricles (LV) initiate filling by serving as mechanical suction pumps allows global diastolic function to be assessed based on laws of motion that apply to all chambers. What differentiates one heart from another are the parameters of the equation of motion that governs filling. Accordingly, development of the Parametrized Diastolic Filling (PDF) formalism&#160;has shown that the entire range of clinically observed early transmitral flow (Doppler E-wave) patterns are extremely well fit by the laws of damped oscillatory motion.&#160;This permits analysis of individual E-waves in accordance with a causal mechanism (recoil-initiated suction) that yields three (numerically) unique lumped parameters whose physiologic analogues are chamber stiffness (k), viscoelasticity/relaxation (c), and load (xo).&#160;The recording of transmitral flow (Doppler E-waves) is standard practice in clinical cardiology and, therefore, the echocardiographic recording method is only briefly reviewed. Our focus is on determination of the PDF parameters from routinely recorded E-wave data.&#160;As the highlighted results indicate, once the PDF parameters have been obtained from a suitable number of load varying E-waves, the investigator is free to use the parameters or construct indexes from the parameters (such as stored energy 1/2kxo2, maximum A-V pressure gradient kxo, load independent index of diastolic function, etc.) and select the aspect of physiology or pathophysiology to be quantified.

Accurate, causality-based quantification of global diastolic function has been achieved by kinematic modeling-based analysis of transmitral flow via the Parametrized Diastolic Filling (PDF) formalism. PDF generates unique stiffness, relaxation,&#160;and load parameters and elucidates &#39;new&#39; physiology while providing sensitive and specific indexes of dysfunction.

Quantitative cardiac function assessment remains a challenge for physiologists and clinicians.&#160;Although historically invasive methods have comprised ...

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and trade one defect for another. Tissue engineering holds the promise to address this increasing demand. However, most tissue engineering strategies fail to generate stable and functional tissue substitutes because of poor vascularization. This paper focuses on an in vivo tissue engineering chamber model of intrinsic vascularization where a perfused artery and a vein either as an arteriovenous loop or a flow-through pedicle configuration is directed inside a protected hollow chamber. In this chamber-based system angiogenic sprouting occurs from the arteriovenous vessels and this system attracts ischemic and inflammatory driven endogenous cell migration which gradually fills the chamber space with fibro-vascular tissue. Exogenous cell/matrix implantation at the time of chamber construction enhances cell survival and determines specificity of the engineered tissues which develop. Our studies have shown that this chamber model can successfully generate different tissues such as fat, cardiac muscle, liver and others. However, modifications and refinements are required to ensure target tissue formation is consistent and reproducible. This article describes a standardized protocol for the fabrication of two different vascularized tissue engineering chamber models in vivo.

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

This is a guideline for constructing in vivo vascularized tissue using a microsurgical arteriovenous loop or a flow-through pedicle configuration inside a tissue engineering chamber. The vascularized tissues generated can be employed for organ regeneration and replacement of tissue defects, as well as for drug testing and disease modeling.

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in every tissue to be identified and characterized. Our laboratory is actively working to make technological advances in X-ray micro-computed tomography (micro-CT) that will bring the diagnostic power of histology to the study of full tissue volumes at cellular resolution (i.e., an X-ray Histo-tomography modality). Toward this end, we have made targeted improvements to the sample preparation pipeline. One key optimization, and the focus of the present work, is a straightforward method for rigid embedding of fixed and stained millimeter-scale samples. Many of the published methods for sample immobilization and correlative micro-CT imaging rely on placing the samples in paraffin wax, agarose, or liquids such as alcohol. Our approach extends this work with custom procedures and the design of a 3-dimensional printable apparatus to embed the samples in an acrylic resin directly into polyimide tubing, which is relatively transparent to X-rays. Herein, sample preparation procedures are described for the samples from 0.5 to 10 mm in diameter, which would be suitable for whole zebrafish larvae and juveniles, or other animals and tissue samples of similar dimensions. As proof of concept, we have embedded the specimens from Danio, Drosophila, Daphnia, and a mouse embryo; representative images from 3-dimensional scans for three of these samples are shown. Importantly, our methodology leads to multiple benefits including rigid immobilization, long-term preservation of laboriously-created resources, and the ability to re-interrogate samples.

We developed protocols and designed a custom apparatus to enable embedding of millimeter-scale specimens. We present sample preparation procedures with an emphasis on embedding in acrylic resin and polyimide tubing to achieve rigid immobilization and long-term storage of specimens for the interrogation of tissue architecture and cell morphology by micro-CT.

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Time-Resolved In Vivo Measurement of Neuropeptide Dynamics by Capacitive Immunoprobe in Porcine Heart

Summary

Explore More Videos

Chapters in this video

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

Procedure for Decellularization of Porcine Heart by Retrograde Coronary Perfusion

Millifluidics for Chemical Synthesis and Time-resolved Mechanistic Studies

Quantification of Global Diastolic Function by Kinematic Modeling-based Analysis of Transmitral Flow via the Parametrized Diastolic Filling Formalism

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques