We thank Dr. Alicia Mattiazzi for critical discussion of the presented work. This work was supported by a grant from NIH (R01 HL-084487) to ALE.

This protocol and all mice handling was approved by the UC Merced Institutional Animal Care and Use Committee (No. 2008-201). Experiments with parakeets were conducted in 1999 according to general policies for animal use established by the scientific commission of the Venezuelan Institute for Scientific Research (IVIC).
1. Langendorff Set Up Preparation
<ol>
	<li>Prepare Tyrode solution containing the following solute concentrations in mM: 140 NaCl, 5.4 KCl, 2 CaCl2, 1 MgCl2, 0.33 Na2HPO4, 10 glucose, 10 HEPES. Adjust the pH of the Tyrode solution to 7.4 with NaOH and filter the solution through a 0.22 &#181;m filter.</li>
	<li>Load the Tyrode solution into the 60 mL syringes and all the tubing of the horizontal Langendorff set-up11,12 illustrated in Figure 4. Make sure to eliminate all air bubbles.</li>
	<li>Equilibrate Tyrode solution with 100% O2 using a submerged plastic &#34;air stone&#34; as illustrated in Figure 4.
	<ol>
		<li>Connect plastic tubing to an O2 tank.</li>
		<li>Add a plastic Tee adapter to lower a 5&#34; tube into the Tyrode solution in the 60 mL syringe.</li>
		<li>Attach a plastic air stone to the end of the 5&#34; tube so O2 will bubble out into the Tyrode solution.</li>
	</ol>
	</li>
	<li>Place a non-absorbable surgical suture around a needle used as a cannula. The needle is coupled to the manifold (see Figure 4) that allows retro-perfusion with different solutions. Finally, the heart&#39;s aorta will be cannulated into the needle.</li>
</ol>
2. Animal Preparation and Heart Dissection
<ol>
	<li>Weigh and inject the mouse with heparin (ex. Mouse of weight 20 g, inject with 20 units or 200 &#181;L) 15 min before euthanizing by cervical dislocation. Anesthetize parakeets according to the animal use guides of your IACUC protocol and then proceed with cervical dislocation. 
	&#8203;NOTE: Use 8 weeks old mice or 20 g parakeets.</li>
	<li>Remove the heart from the thoracic cavity after euthanization. Extract parakeet hearts in exactly the same way as described for mice.
	<ol>
		<li>Clean the mouse chest with ethanol.</li>
		<li>Using dissection scissors, make an incision in the lower abdomen and then cut up the sides toward the neck.</li>
		<li>Pull back the cut tissue and pin it down.</li>
		<li>Cut the diaphragm. Be careful when cutting the diaphragm to avoid damaging the heart.</li>
		<li>Remove the lungs and surrounding tissue.</li>
		<li>Use tweezers to scoop the heart without squeezing it. Cut the aorta as long as possible.</li>
	</ol>
	</li>
	<li>Transport the heart on a small weigh boat with approximately 1 mL Tyrode solution.</li>
	<li>Using a non-absorbable surgical suture, tie the aorta onto the horizontal Langendorff apparatus via a needle. Tie the heart aorta with the aid of two fine tweezers. Begin retro-perfusion by opening the valve located in series with the 60 mL syringes containing Tyrode solution.</li>
	<li>Allow the heart to stabilize for 10 min. Use this time to clean blood and the fatty tissue surrounding the heart before loading the dye. Pull the fatty tissue near the base of the heart with tweezers and cut using small dissection scissors. Be sure to do this under the view of a dissecting microscope.</li>
</ol>
3. Cytosolic Ca&#8203;2+ Measurements: Preparing Dye Rhod-2AM
<ol>
	<li>Add 20 &#181;L of the 20% pluronic (a non-ionic surfactant) in DMSO to a special packaged plastic vial provided by the dye manufacturer containing 50 &#181;g of the dye.</li>
	<li>Mix by pipetting up and down, avoiding bubbles.</li>
	<li>Transfer the mixed DMSO with the non-ionic surfactant and the dye from the special packaged plastic vial into a clear glass vial. Add 1 mL of Tyrode solution to the clear glass vial.</li>
	<li>Sonicate for 15-20 min in a bath sonicator.</li>
	<li>Perfuse the dye using peristaltic pumps for 30 min at room temperature.
	<ol>
		<li>Place the dye in the dye chamber.</li>
		<li>Use a mechanical clamp to compress all the other tubing lines connected to the manifold. The clamp is placed above the manifold. This will prevent any backflow into the tubing connected to the 60 mL syringes.</li>
		<li>Turn on the perfusion peristaltic pump to begin circulating the dye. Immediately close the 3-way valve below the 60 mL syringe.</li>
		<li>Position a small tube that is connected to the suction peristaltic pump next to the heart to recirculate the dye that has been perfused into the heart.</li>
	</ol>
	</li>
</ol>
4. Intra-SR Ca&#8203;2+ Measurements: Preparing Dye Mag-Fluo4AM
<ol>
	<li>Prepare Mag-Fluo4AM in the same manner as Rhod-2AM. Refer to section 3 for step-wise instructions.</li>
	<li>After loading the dye, open the valve located in series with the 60 mL syringes containing Tyrode solution to begin retro-perfusion. Be sure to remove the clamp above the manifold.</li>
	<li>Add Tyrode solution to the horizontal chamber and warm up to 37 &#176;C.</li>
	<li>Retro-perfuse with Tyrode solution for 45 min to remove the cytosolic dye.</li>
</ol>
5. Membrane Potential Measurements: Preparing Dye Di-8-ANEPPS
<ol>
	<li>Add 5 mL of 99% ethanol to the dye vial that contains 5 mg of the dye.</li>
	<li>Aliquot 10 &#181;L into 500 individual 1 mL plastic vials using a repeater pipette.</li>
	<li>Desiccate in a speed vacuum and store at -20 &#176;C.</li>
	<li>Add 20 &#181;L of 20% pluronic in DMSO to a plastic vial with 10 &#181;g of the desiccated dye.</li>
	<li>Mix by pipetting up and down, avoiding bubbles.</li>
	<li>Transfer the mix containing DMSO with pluronic and dye from the plastic vial to a graduated cylinder (10 mL). Add Tyrode solution to a final volume of 5 mL.</li>
	<li>Sonicate for 20-25 min in a bath sonicator.</li>
	<li>Perfuse into the heart for 30 min using peristaltic pumps. Refer to stepwise instructions in Section 3.5.</li>
</ol>
6 . Recording Epicardial Signals
<ol>
	<li>After loading the dye, retro-perfuse the heart with Tyrode solution by removing the clamp above the manifold. Open the valve located in series with the 60 mL syringe containing Tyrode solution. Retro-perfuse Tyrode solution for 10 min to stabilize the heart.</li>
	<li>Fill the horizontal chamber with Tyrode solution and turn on the peltier unit to bring the bath temperature to 37 &#176;C.</li>
	<li>Position the fiber optic on the surface of the heart.
	<ol>
		<li>Place the fiber optic inside a 2 mL pipette and then attach the pipette to a micromanipulator.</li>
		<li>Use the micromanipulator to slightly press the fiber optic against the surface of the LV.</li>
	</ol>
	</li>
	<li>Externally pace the heart with a stimulator controlled by a wave generator.
	<ol>
		<li>Program the wave generator to provide a square pulse with a width of 1 ms.</li>
		<li>Set the stimulator to be externally synchronized and connect the external input to the wave generator.</li>
		<li>To each output of the stimulator, connect a wire with an acupuncture needle soldered at the end.</li>
		<li>Place both acupuncture needles in the apex of the heart approximately 3 mm apart from each other.</li>
		<li>Only after the needles are placed in the tissue, turn on the output of the stimulator to prevent electric shock.</li>
	</ol>
	</li>
	<li>In the acquisition software, adjust acquisition frequency to 10 kHz.</li>
</ol>
7 . Recording Endocardial Signals
<ol>
	<li>Refer to step 6.1 and 6.2 to stabilize the heart after loading the dye.</li>
	<li>Using a sharp 23Ga point about the size of the fiber optic, make a small hole in the surface of the heart in the LV near the septum.</li>
	<li>Place an intravitreal surgery sclerotomy adaptor to aid in positioning the first fiber optic into the endocardium using a micromanipulator.</li>
	<li>Externally pace the heart. Refer to step 6.4.</li>
	<li>In acquisition software, adjust acquisition frequency to 10 kHz.</li>
</ol>

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The authors have nothing to disclose.

This paper is centered in describing local field fluorescence techniques to evaluate the function of cardiac myocytes ex vivo. The study of these cells in a coupled environment is not only more physiological, but is also highly appropriate for assessing organ-level pathologies. The cellular events underlying excitation-contraction coupling (ECC) can be evaluated at the whole organ level with the use of molecular probes that monitor intracellular Ca&#8203;2+ dynamics (Rhod 2 cytosolic Ca2+, ...

The heart is the central organ of the cardiovascular system. The heart&#39;s contraction is initiated by an increase in intracellular [Ca2+]. The relationship between electrical excitability and changes in intracellular Ca2+ release has been historically studied in enzymatically dissociated cells1,2. However, cardiac cells are electrically, metabolically and mechanically coupled3,4. When isolated, the myocytes are not only physically uncoupled, but myocytes from different layers are mixed during the dissociation5. Furthermore, despite the enormous advantages that have emerged from the study of isolated cells under voltage clamp conditions,6,7,8 the intrinsic nature of the heart as an electrical syncytium always poses the question of how functionally different are dissociated cells from those present in the tissue3.
In this manuscript, we describe the advances obtained in the knowledge of cardiac physiology by the use of local field fluorescence microscopy (LFFM) techniques in the intact heart. LFFM uses fluorescent indicators to measure multiple physiological variables such as cytosolic Ca2+, intra sarcoplasmic reticulum (SR) Ca2+ and membrane potential. These measurements can be obtained simultaneously and in conjunction with ventricular pressure9,10, electrocardiograms9, electrical action potentials (APs), ionic current recordings and flash photolysis of caged compounds4,11. In addition, these measurements can be obtained by pacing the intact heart at higher frequencies closer to physiological rates. Although several articles9,11,12,13,14 have been published by our group using LFFM techniques, the presumption of technical complexities associated with this technique has prevented its massive use in studying ex vivo physiological phenomena in the heart and other organs.
The LFFM technique (Figure 1) is based on epifluorescence measurements obtained using a multimode optical fiber in contact with the tissue. Like any contact fluorescence imaging technique, the optical resolution depends on the diameter and the numerical aperture (NA) of the fiber. A higher NA and smaller diameter of the fiber will increase the spatial resolution of the measurements. NAs and fiber diameters can range from 0.22 to 0.66 and from 50 &#181;m to 1 mm, respectively. Increasing the NA will improve the signal to noise ratio (S/N) by accepting photons arriving from a larger solid angle. In order to act as an epifluorescence device, the light beam is focused into the optical fiber with an aspheric lens or an epifluorescence objective where the NA of the lens and the fiber match. This matching maximizes the energy transfer for excitation and for collecting back the photons emitted by the fluorophore.
In order to excite the exogenous fluorescent indicators loaded in the tissue, different light sources and illumination modes can be utilized. Our pioneering studies using the pulsed local field fluorescence microscopy3,12 (PLFFM) employed a low-cost picosecond laser (Figure 1a, PLFFM). This type of light source has the huge advantage of exciting a big fraction of fluorophore molecules under the illumination area without substantially bleaching the dye due to the short pulse durations12. Additionally, the use of ultrashort pulses allowed the assessment of the fluorescence lifetime of the dye12. The fluorescence lifetime is a property that can be used to quantify the fraction of dye molecules bound to Ca2+. Unfortunately, the temporal jittering of the pulses and variations in amplitude from pulse-to-pulse limit the application of this experimental strategy to cases where the change in fluorescence produced by the ligand binding to the dye is large.
Continuous Wave (CW) lasers are usually used as the main illumination source in LFFM (Figure 1b, CLFFM). The laser beam can continuously illuminate the tissue or can be ferroelectrically modulated. The ferroelectric modulation of the beam allows the generation of microsecond pulses of light. This modulation can be controlled by external hardware. This procedure not only dramatically reduces the temporal jittering of light pulses but also allows mixing beams of different wavelengths. The mixing of beams is done by multiplexing rays from different lasers. As a consequence, multiple dyes having different spectral properties can be excited to perform measurements of a variety of physiological variables, for example, Rhod-2 for cytosolic Ca2+, MagFluo4 for intra-SR Ca2+ and di-8-ANEPPS for membrane potential.
Although lasers present various advantages as the light source in LFFM, other types of light sources can be used including light-emitting diodes (LEDs). In this case, the excitation light source consisted of an InGaN LED (Figure 1c, PLEDFM). In LEDs, photons are spontaneously emitted when electrons from the conduction band recombine with holes in the valence band. The difference with solid-state lasers is that the emission is not stimulated by other photons. This results in a non-coherent beam and a wider spectral emission for LEDs.
Different types of high power LEDs can be used. For AP recordings using Di-8-ANEPPS and for Ca2+ transients recorded using Fluo-4 or Mag-Fluo-4, we used an LED that has a typical peak emission at 485 nm (blue) and a half width of 20 nm (Figure 1d). For Ca2+ transients recorded with Rhod-2, the LED had a typical peak emission at 540 nm (green) and a half width of 35 nm (Figure 1d). LEDs emit in a band wavelength and therefore require filters to narrow their spectral emission. In addition, pulsed light can be generated at a rate of 1.6 kHz with duration of 20 &#181;s. The LEDS were pulsed with a fast power MOSFET field effect transistor. Simultaneous recordings with different indicators can be performed by time-multiplexing the LEDs. Unfortunately, light emitted by LEDs is more difficult to focus onto a fiber optic compared to a laser beam. Thus, the main drawback of using LEDs is that their emission profiles have angular displacements (&#177; 15&#176;) from the main axis, and an auxiliary optic must be used to correct it.
In all of the optical configurations previously described, the excitation light is reflected with the aid of a dichroic mirror. The beam is subsequently focused by an aspheric lens and a microscope objective onto a multimode fiber optic that is positioned on the tissue. As in any epifluorescence arrangement, the dichroic mirror also serves to separate the excitation from the emitted light. The emitted light spectrum travels back through a barrier filter to remove any reflected excitation. Finally, the emitted light is focused with an objective onto a photodetector (Figure 1).
The transduction from light to electrical current is performed by silicon avalanche photodiodes. These diodes have a fast response and a high sensitivity allowing low light detection. The photocurrent produced by the avalanche photodiodes can be amplified in two ways: a transimpedance amplifier having a resistive feedback element (Figure 1e) or by an integrator to convert the current into a voltage (Figure 1f). Using the first approach, the output voltage is proportional to the photocurrent and the feedback resistor. A typical example of the resistive detection of picosecond laser pulses is shown in Figures 2a, 2b and 2c. Panel 2a illustrates the output of the transimpedance amplifier and panel 2b shows a time expansion of the interval indicated with an asterisk (*). A peak tracking algorithm was implemented to detect the peak (red) and the base (green) for the fluorescent responses12. The measurement of the base fluorescence provides information of both the dark current of the avalanche photodiode and the interferences introduced by ambient light and electromagnetic coupling. A representation of peaks and bases is shown in Figure 2c. This figure illustrates the fluorescence emitted by the dye (Rhod-2) bound to Ca2+ during the cardiac cycle of a beating parakeet heart.
In the second method, the output voltage of the integrator is a function of the current and capacitive feedback (Figures 2d, 2e, and 2f). Figure 2f shows two consecutive integration cycles: the first with no external illumination and the second with applied light pulses from a pulsed LED. A detailed description is presented in Figures 2g and 2h. This approach, although more laborious, provides a larger S/N due to the absence of thermal noise in the feedback capacitor. The instrument includes a timing stage that generates all the control and multiplexing of the excitation light and commands the headstage integration and reset periods. This is usually performed with a digital signal processing circuit that also performs a digital differentiation of the integrated output signal by computing an on-line regression of the data. In the case of using a resistive feedback, any A/D acquisition board can be used.
Finally, our LFFM technique is highly versatile and can be adapted to record from more than one region. Adding a beam splitter in the light path allows us to split the light into two optical fibers. Each optical fiber can then be placed on different regions of a tissue to, independently, excite and record emission from the exogenous fluorescent probes. This modification permits us to assess how anatomical regional differences influence physiological variables. Figure 3 shows a beam splitter being employed to split the CW excitation light such that two optical fibers are used to measure transmural electrical or intracellular [Ca2+] levels with minor-invasiveness. Transmural signals can be recorded by placing one fiber on the endocardium and the other on the epicardium layer of the ventricular wall. Therefore, the LFFM technique has the ability to measure the time course of cellular signals in different regions and can be used to test if regional changes occur under pathological situations.

<table><tbody><tr><td>Sodium chloride</td><td>Sigma</td><td>7647-14-5</td><td></td></tr><tr><td>D-(+)-glucose</td><td>Sigma</td><td>50-99-7</td><td></td></tr><tr><td>Potassium chloride</td><td>Sigma</td><td>7447-40-7</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>7365-45-9</td><td></td></tr><tr><td>Sodium phosphate</td><td>Sigma</td><td>10049-21-5</td><td></td></tr><tr><td>Calcium chloride solution</td><td>Sigma</td><td>10043-52-4</td><td></td></tr><tr><td>Magnesium chloride solution</td><td>Sigma</td><td>7786-30-3</td><td></td></tr><tr><td>Sodium hyrdoxide</td><td>Sigma</td><td>S-8045</td><td></td></tr><tr><td>0.2 &amp;mu;m nylon membrane filter</td><td>Whatman</td><td>7402-004</td><td></td></tr><tr><td>Manifold MPP</td><td>Warner</td><td>64-0216&amp;nbsp;</td><td></td></tr><tr><td>21G1.5 Precision glide needle</td><td>B-D</td><td>305167</td><td></td></tr><tr><td>Black Braided silk string (non-absorbable surgical suture)</td><td>AllMech Tech</td><td>LOOK-SP105</td><td></td></tr><tr><td>Heparin sodium injection 1000USP/mL</td><td>AllMech Tech</td><td>NDC63323-540-11</td><td></td></tr><tr><td>DMSO D8779</td><td>Sigma</td><td>67-68-5</td><td></td></tr><tr><td>Blebbistatin</td><td>Sigma</td><td>856925-71-8</td><td></td></tr><tr><td>Pluronic F-127 20% solution</td><td>Biotium</td><td>59004</td><td></td></tr><tr><td>Materflex C/L peristaltic pump</td><td>Cole-Parmer</td><td>77122-26</td><td></td></tr><tr><td>Isostim stimulator</td><td>World Precision Instruments</td><td>A320RC</td><td></td></tr><tr><td>Waveform generator</td><td>Teledyne Lecroy</td><td>WaveStation 2012</td><td></td></tr><tr><td>Ultrasonic cleaner FS20</td><td>Fisher Scientific</td><td>1533530</td><td></td></tr><tr><td>Tygon tubing ID:1/32&quot; OD:3/32&quot; Wall 1/32&quot;</td><td>Component Supply</td><td>TET-031A</td><td></td></tr><tr><td>Tygon tubing ID:3/32&quot; OD:5/32&quot; Wall 1/32&quot;</td><td>Component Supply</td><td>TET-094A</td><td></td></tr><tr><td>Adapter luer lock to 3-way valve</td><td>Cole-Parmer</td><td>EW-31200-80</td><td></td></tr><tr><td>Tee adapters and plastic fittings</td><td>Cole-Parmer</td><td>6365-90</td><td></td></tr><tr><td>Plastic clamp</td><td>WaterZoo</td><td>2465</td><td></td></tr><tr><td>Peltier</td><td>TE Technology</td><td>TE-127-2.0-2.5</td><td></td></tr><tr><td>Rhod-2AM</td><td>ThermoFisher Scientific</td><td>R1245MP</td><td></td></tr><tr><td>Di-8-ANEPPS</td><td>ThermoFisher Scientific</td><td>D3167</td><td></td></tr><tr><td>Mag-Fluo-4AM</td><td>ThermoFisher Scientific</td><td>M14206</td><td></td></tr><tr><td>Acupunture needles</td><td>LHASA</td><td>TC1.20x13</td><td></td></tr><tr><td>60 mL BD syringe with luer-lok</td><td>Fisher Scientific</td><td>14-820-11</td><td></td></tr><tr><td>LabView</td><td>National Instruments</td><td></td><td></td></tr><tr><td>Speed vacuum Eppendorf Vagufuge</td><td>Fisher Scientific</td><td>07-748-13</td><td></td></tr><tr><td>Digital signal processing circuit DSP TMS 320</td><td>Texas Instrument</td><td></td><td></td></tr><tr><td>Longpass Dichroic mirror 567 nm</td><td>ThorLabs</td><td>DMLP567L</td><td></td></tr><tr><td>Objective 10X NA 0.25 DIN AchromaticFinite Intl Standard Objective</td><td>Edmund Optics</td><td>Stock# 33-437</td><td></td></tr><tr><td>Objective 20X NA 0.40 DIN Achromatic Finite Intl Standard Objective</td><td>Edmund Optics</td><td>Stock# 33-438</td><td></td></tr><tr><td>Longpass colored glass filter 590 nm&amp;nbsp;</td><td>ThorLabs</td><td>FGL590</td><td></td></tr><tr><td>Green Nd-YAG laser 532 nm, 500 mW</td><td></td><td></td><td></td></tr><tr><td>Micromanipulator for laser</td><td>Siskiyou</td><td>MX130R</td><td></td></tr><tr><td>Multimode fiber optic 200 &amp;micro;m NA 0.39</td><td>ThorLabs</td><td>FT200UMT</td><td></td></tr><tr><td>LEDs blue</td><td>Lumileds</td><td>L135-B475003500000</td><td></td></tr><tr><td>LEDs green</td><td>Lumileds</td><td>L135-G525003500000</td><td></td></tr><tr><td>Cube beam splitter, non-polarizing</td><td>ThorLabs</td><td>BS007</td><td></td></tr><tr><td>36&quot; Length, Dovetail Optical Rail</td><td>Edmund Optics</td><td>54-402</td><td></td></tr><tr><td>2.5&quot; Width, Dovetail Carrier</td><td>Edmund Optics</td><td>54-404</td><td></td></tr><tr><td>0.75&quot; Travel, micrometer stage</td><td>Edmund Optics</td><td>37-983</td><td></td></tr><tr><td>Dovetail optical rail 3&quot;</td><td>ThorLabs</td><td>RLA075/M</td><td></td></tr><tr><td>Dovetail rail carrier 1&quot;</td><td>ThorLabs</td><td>RC1</td><td></td></tr><tr><td>Avalanche photodiode Helix 902</td><td>Digi-Key</td><td>HELIX-902-200</td><td></td></tr><tr><td>Objective holder XY Translator&amp;nbsp;</td><td>ThorLabs</td><td>ST1XY-S</td><td></td></tr><tr><td>Aluminum breadboard 6&quot; x 6&quot;</td><td>ThorLabs</td><td>MB6</td><td></td></tr><tr><td>Nexus otpical table 4&#039; x 6&#039;</td><td>ThorLabs</td><td>T46HK</td><td></td></tr><tr><td>Stainless steel cap screws</td><td>ThorLabs</td><td>HW-KIT5/M</td><td></td></tr><tr><td>Acrylic&amp;nbsp; sheet</td><td>Home Depot/Lowes</td><td></td><td></td></tr><tr><td>Sylgard Silicone elastomer kit</td><td>Dow Corning</td><td>Sylgard 184</td><td></td></tr><tr><td>Epoxy gel&amp;nbsp;</td><td>Walmart/Home Depot</td><td>2-part 5 min clr 1oz</td><td></td></tr><tr><td>21G1.5 Precision glide needle</td><td>B-D</td><td>305167</td><td></td></tr><tr><td>23G Precision glide needle</td><td>B-D</td><td>305145</td><td></td></tr><tr><td>Mounting base, 25 mm x 75 mm x 10 mm</td><td>ThorLabs</td><td>BA1/M</td><td></td></tr><tr><td>Wire shelve posts 36&quot;</td><td>Alera</td><td>AALESW59PO36SR</td><td></td></tr><tr><td>Wire shelves</td><td>Alera</td><td>ALESW582424SR</td><td></td></tr><tr><td>Post and angle clamp</td><td>ThorLabs</td><td>SWC/M-P5</td><td></td></tr><tr><td>Glass syringe for dye chamber</td><td>Wheaton</td><td>W851020</td><td></td></tr><tr><td>Rubber stopper</td><td>Home Science Tools</td><td>CE-STOP01C</td><td></td></tr></tbody></table>

local field fluorescence microscopy: imaging cellular signals in intact hearts

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and arrhythmias can only be fully understood at the whole organ level. Here, we present the spectroscopic technique of local field fluorescence microscopy (LFFM) that allows the measurement of cellular signals in the intact heart. The technique is based on a combination of a Langendorff perfused heart and optical fibers to record fluorescent signals. LFFM has various applications in the field of cardiovascular physiology to study the heart under normal and pathological conditions. Multiple cardiac variables can be monitored using different fluorescent indicators. These include cytosolic [Ca2+], intra-sarcoplasmic reticulum [Ca2+] and membrane potentials. The exogenous fluorescent probes are excited and the emitted fluorescence detected with three different arrangements of LFFM epifluorescence techniques presented in this paper. The central differences among these techniques are the type of light source used for excitation and on the way the excitation light is modulated. The pulsed LFFM (PLFFM) uses laser light pulses while continuous wave LFFM (CLFFM) uses continuous laser light for excitation. Finally, light-emitting diodes (LEDs) were used as a third light source. This non-coherent arrangement is called pulsed LED fluorescence microscopy (PLEDFM).

AP and Ca2+ transients in endocardium and epicardium
In order to compare signals across the ventricular wall, one fiber optic is positioned in the endocardium and the other in the epicardium. Comparing the morphology of an AP recorded from the endocardium with one from the epicardium is the best way to assess the transmural function. The ventricular wall is highly hetero...

Watch this Scientific Journal Video about Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts at JoVE.com

Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts

In the heart, molecular events coordinate the electrical and contractile function of the organ. A set of local field fluorescence microscopy techniques presented here enables the recording of cellular variables in intact hearts. Identifying mechanisms defining the cardiac function is critical in understanding how the heart works under pathological situations.

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

8.2K Views.  University of California, Merced. In the heart, molecular events coordinate the electrical and contractile function of the organ. A set of local field fluorescence microscopy techniques presented here enables the recording of cellular variables in intact hearts. Identifying mechanisms defining the cardiac function is critical in understanding how the heart works under pathological situations. 

Video: Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts

Imaging Cleared Embryonic and Postnatal Hearts at Single-cell Resolution

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, and allow for the study of the cellular and molecular basis of normal and abnormal cardiac morphogenesis. Recent emerging technologies of retrospective single clonal analyses make the study of cardiac morphogenesis at single cell resolution feasible. However, tissue opacity and light scattering of the heart as imaging depth is increased hinder whole-heart imaging at single cell resolution. To overcome these obstacles, a whole-embryo clearing system that can render the heart highly transparent for both illumination and detection must be developed. Fortunately, in the last several years, many methodologies for whole-organism clearing systems such as CLARITY, Scale, SeeDB, ClearT, 3DISCO, CUBIC, DBE, BABB and PACT have been reported. This lab is interested in the cellular and molecular mechanisms of cardiac morphogenesis. Recently, we established single cell lineage tracing via the ROSA26-CreERT2; ROSA26-Confetti system to sparsely label cells during cardiac development. We adapted several whole embryo-clearing methodologies including Scale and CUBIC (clear, unobstructed brain imaging cocktails and computational analysis) to clear the embryo in combination with whole mount staining to image single clones inside the heart. The heart was successfully imaged at single cell resolution. We found that Scale can clear the embryonic heart, but cannot effectively clear the postnatal heart, while CUBIC can clear the postnatal heart, but damages the embryonic heart by dissolving the tissue. The methods described here will permit the study of gene function at a single clone resolution during cardiac morphogenesis, which, in turn, can reveal the cellular and molecular basis of congenital heart defects.

We describe a protocol to volumetrically image fluorescent protein labeled cells deep inside intact embryonic and postnatal hearts. Utilizing tissue-clearing methods in combination with whole mount staining, single fluorescent protein-labeled cells inside an embryonic or postnatal heart can be imaged clearly and accurately.

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, ...

Developmental Biology

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

Capturing the Cardiac Injury Response of Targeted Cell Populations via Cleared Heart Three-Dimensional Imaging

Cardiovascular disease outranks all other causes of death and is responsible for a staggering 31% of mortalities worldwide. This disease manifests in cardiac injury, primarily in the form of an acute myocardial infarction. With little resilience following injury, the once healthy cardiac tissue will be replaced by fibrous, non-contractile scar tissue and often be a prelude to heart failure. To identify novel treatment options in regenerative medicine, research has focused on vertebrates with innate regenerative capabilities. One such model organism is the neonatal mouse, which responds to cardiac injury with robust myocardial regeneration. In order to induce an injury in the neonatal mouse that is clinically relevant, we have developed a surgery to occlude the left anterior descending artery (LAD), mirroring a myocardial infarction triggered by atherosclerosis in the human heart. When matched with the technology to track changes both within cardiomyocytes and non-myocyte populations, this model provides us with a platform to identify the mechanisms that guide heart regeneration. Gaining insight into changes in cardiac cell populations following injury once relied heavily on methods such as tissue sectioning and histological examination, which are limited to two-dimensional analysis and often damage the tissue in the process. Moreover, these methods lack the ability to trace changes in cell lineages, instead providing merely a snapshot of the injury response. Here, we describe how technologically advanced methods in lineage tracing models, whole organ clearing, and three-dimensional (3D) whole-mount microscopy can be used to elucidate mechanisms of cardiac repair. With our protocol for neonatal mouse myocardial infarction surgery, tissue clearing, and 3D whole organ imaging, the complex pathways that induce cardiomyocyte proliferation can be unraveled, revealing novel therapeutic targets for cardiac regeneration.

Cardiomyocyte proliferation following injury is a dynamic process that requires a symphony of extracellular cues from non-myocyte cell populations. Utilizing lineage tracing, passive CLARITY, and three-dimensional whole-mount confocal microscopy techniques, we can analyze the influence of a variety of cell types on cardiac repair and regeneration.

Cardiovascular disease outranks all other causes of death and is responsible for a staggering 31% of mortalities worldwide. This disease manifests in ...

Light-sheet Fluorescence Microscopy for the Study of the Murine Heart

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. Traditional light-sheet techniques involve the use of a single illumination beam directed orthogonally at sample tissue. Images of large samples that are produced using a single illumination beam contain stripes or artifacts and suffer from a reduced resolution due to the scattering and absorption of light by the tissue. This study uses a dual-sided illumination beam and a simplified CLARITY optical clearing technique for the murine heart. These techniques allow for deeper imaging by removing lipids from the heart and produce a large field of imaging, greater than 10 x 10 x 10 mm3. As a result, this strategy enables us to quantify the ventricular dimensions, track the cardiac lineage, and localize the spatial distribution of cardiac-specific proteins and ion-channels from the post-natal to adult mouse hearts with sufficient contrast and resolution.

This study uses a dual-sided illumination light-sheet fluorescence microscopy (LSFM) technique combined with optical clearing to study the murine heart.

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. ...

Bioengineering

Optical Imaging of Isolated Murine Ventricular Myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically ...

Simultaneous Brightfield, Fluorescence, and Optical Coherence Tomographic Imaging of Contracting Cardiac Trabeculae Ex Vivo

In cardiac muscle, intracellular Ca2+ transients activate contractile myofilaments, causing contraction, macroscopic shortening, and geometric&#160;deformation. Our understanding of the internal relationships between these events has been limited because we can neither &#39;see&#39; inside the muscle nor precisely track the spatio-temporal nature of excitation-contraction dynamics. To resolve these problems, we have constructed a device that combines a suite of imaging modalities. Specifically, it integrates a brightfield microscope to measure local changes of sarcomere length and tissue strain, a fluorescence microscope to visualize the Ca2+ transient, and an optical coherence tomograph to capture the tissue&#39;s geometric&#160;changes throughout the time-course of a cardiac cycle. We present here the imaging infrastructure and associated data collection framework. Data are collected from isolated rod-like tissue structures known as trabeculae carneae. In our instrument, a pair of position-controlled platinum hooks hold each end of an ex vivo muscle sample while it is continuously superfused with nutrient-rich saline solution. The hooks are under independent control, permitting real-time control of muscle length and force. Lengthwise translation enables the piecewise scanning of the sample, overcoming limitations associated with the relative size of the microscope imaging window (540 &#181;m by 540 &#181;m) and the length of a typical trabecula (&#62;2000 &#181;m). Platinum electrodes at either end of the muscle chamber stimulate the trabecula at a user-defined rate. We exploit the stimulation signal as a trigger for synchronizing the data from each imaging window to reconstruct the entire sample twitching under steady-state conditions. Applying image-processing techniques to these brightfield imaging data provides tissue displacement and sarcomere length maps. Such a collection of data, when incorporated into an experiment-modeling pipeline, will provide a deeper understanding of muscle contractile homogeneity and heterogeneity in physiology and pathophysiology.

Simultaneous Brightfield, Fluorescence, and Optical Coherence Tomographic Imaging of Contracting Cardiac Trabeculae Ex Vivo

This protocol presents a collection of sarcomere, calcium, and macroscopic geometric&#160;data from an actively contracting cardiac trabecula ex vivo. These simultaneous measurements are made possible by the integration of three imaging modalities.

In cardiac muscle, intracellular Ca2+ transients activate contractile myofilaments, causing contraction, macroscopic shortening, and ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a significant role in certain types of arrhythmia. Because arrhythmias are spatially distinct, emergent phenomena, they must be investigated at the tissue level. However, methods for directly probing SR Ca2+ in the intact heart remain limited. This article describes the protocol for dual optical mapping of transmembrane potential (Vm) and free intra-SR [Ca2+] ([Ca2+]SR) in the Langendorff-perfused rabbit heart. This approach takes advantage of the low-affinity Ca2+ indicator Fluo-5N, which has minimal fluorescence in the cytosol where intracellular [Ca2+] ([Ca2+]i) is relatively low but exhibits significant fluorescence in the SR lumen where [Ca2+]SR is in the millimolar range. In addition to revealing SR Ca2+ characteristics spatially across the epicardial surface of the heart, this approach has the distinct advantage of simultaneous monitoring of Vm, allowing for investigations into the bidirectional relationship between Vm and SR Ca2+ and the role of SR Ca2+ in arrhythmogenic phenomena.

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

This article describes the detailed protocol and equipment necessary for dual optical mapping of transmembrane potential (Vm) and free intra-sarcoplasmic reticulum (SR) Ca2+ in the Langendorff-perfused rabbit heart. This method allows for direct observation and quantification of Vm and SR Ca2+ dynamics in the intact heart.

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a ...

Mesoscopic Optical Imaging of Whole Mouse Heart

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition (fibrosis) and cellular misalignment, can affect electrical conduction, introduce electromechanical dysfunctions and, eventually, lead to arrhythmia. Current predictive models of these functional alterations are based on non-integrated and low-resolution structural information. Placing this framework on a different order of magnitude is challenging due to the inefficacy of standard imaging methods in performing high-resolution imaging in massive tissue. In this work, we describe a methodological framework that allows imaging of whole mouse hearts with micrometric resolution. The achievement of this goal has required a technological effort where advances in tissue transformation and imaging methods have been combined. First, we describe an optimized CLARITY protocol capable of transforming an intact heart into a nanoporous, hydrogel-hybridized, lipid-free form that allows high transparency and deep staining. Then, a fluorescence light-sheet microscope able to rapidly acquire images of a mesoscopic field of view (mm-scale) with the micron-scale resolution is described. Following the mesoSPIM project, the conceived microscope allows the reconstruction of the whole mouse heart with micrometric resolution in a single tomographic scan. We believe that this methodological framework will allow clarifying the involvement of the cytoarchitecture disarray in the electrical dysfunctions and pave the way for a comprehensive model that considers both the functional and structural data, thus enabling a unified investigation of the structural causes that lead to electrical and mechanical alterations after the tissue remodeling.

We report a method for mesoscopic reconstruction of the whole mouse heart by combining new advancements in tissue transformation and staining with the development of an axially scanned light-sheet microscope.

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition ...

Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts

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Rozdziały w tym wideo

Imaging Cleared Embryonic and Postnatal Hearts at Single-cell Resolution

Optical Mapping of Langendorff-perfused Rat Hearts

Capturing the Cardiac Injury Response of Targeted Cell Populations via Cleared Heart Three-Dimensional Imaging

Light-sheet Fluorescence Microscopy for the Study of the Murine Heart

Optical Imaging of Isolated Murine Ventricular Myocytes

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Simultaneous Brightfield, Fluorescence, and Optical Coherence Tomographic Imaging of Contracting Cardiac Trabeculae Ex Vivo

Imaging Local Ca²⁺ Signals in Cultured Mammalian Cells

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca²⁺ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

Mesoscopic Optical Imaging of Whole Mouse Heart