Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts

Summary

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.

Abstract

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).

Introduction

The heart is the central organ of the cardiovascular system. The heart'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 cells^1,2. However, cardiac cells are electrically, metabolically and mechanically coupled^3,4. 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 pressure^9,10, electrocardiograms⁹, electrical action potentials (APs), ionic current recordings and flash photolysis of caged compounds^4,11. In addition, these measurements can be obtained by pacing the intact heart at higher frequencies closer to physiological rates. Although several articles^{9,11,12,13,14} 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 µm 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 microscopy^3,12 (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 µs. 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 (± 15°) 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.

Protocol

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

1. 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 µm filter.
2. Load the Tyrode solution into the 60 mL syringes and all the tubing of the horizontal Langendorff set-up^11,12 illustrated in Figure 4. Make sure to eliminate all air bubbles.
3. Equilibrate Tyrode solution with 100% O₂ using a submerged plastic "air stone" as illustrated in Figure 4.
   1. Connect plastic tubing to an O₂ tank.
   2. Add a plastic Tee adapter to lower a 5" tube into the Tyrode solution in the 60 mL syringe.
   3. Attach a plastic air stone to the end of the 5" tube so O₂ will bubble out into the Tyrode solution.
4. 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's aorta will be cannulated into the needle.

2. Animal Preparation and Heart Dissection

1. Weigh and inject the mouse with heparin (ex. Mouse of weight 20 g, inject with 20 units or 200 µL) 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.
   ​NOTE: Use 8 weeks old mice or 20 g parakeets.
2. Remove the heart from the thoracic cavity after euthanization. Extract parakeet hearts in exactly the same way as described for mice.
   1. Clean the mouse chest with ethanol.
   2. Using dissection scissors, make an incision in the lower abdomen and then cut up the sides toward the neck.
   3. Pull back the cut tissue and pin it down.
   4. Cut the diaphragm. Be careful when cutting the diaphragm to avoid damaging the heart.
   5. Remove the lungs and surrounding tissue.
   6. Use tweezers to scoop the heart without squeezing it. Cut the aorta as long as possible.
3. Transport the heart on a small weigh boat with approximately 1 mL Tyrode solution.
4. 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.
5. 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.

3. Cytosolic Ca^​2+ Measurements: Preparing Dye Rhod-2AM

1. Add 20 µL of the 20% pluronic (a non-ionic surfactant) in DMSO to a special packaged plastic vial provided by the dye manufacturer containing 50 µg of the dye.
2. Mix by pipetting up and down, avoiding bubbles.
3. 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.
4. Sonicate for 15-20 min in a bath sonicator.
5. Perfuse the dye using peristaltic pumps for 30 min at room temperature.
   1. Place the dye in the dye chamber.
   2. 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.
   3. Turn on the perfusion peristaltic pump to begin circulating the dye. Immediately close the 3-way valve below the 60 mL syringe.
   4. 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.

4. Intra-SR Ca^​2+ Measurements: Preparing Dye Mag-Fluo4AM

1. Prepare Mag-Fluo4AM in the same manner as Rhod-2AM. Refer to section 3 for step-wise instructions.
2. 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.
3. Add Tyrode solution to the horizontal chamber and warm up to 37 °C.
4. Retro-perfuse with Tyrode solution for 45 min to remove the cytosolic dye.

5. Membrane Potential Measurements: Preparing Dye Di-8-ANEPPS

1. Add 5 mL of 99% ethanol to the dye vial that contains 5 mg of the dye.
2. Aliquot 10 µL into 500 individual 1 mL plastic vials using a repeater pipette.
3. Desiccate in a speed vacuum and store at -20 °C.
4. Add 20 µL of 20% pluronic in DMSO to a plastic vial with 10 µg of the desiccated dye.
5. Mix by pipetting up and down, avoiding bubbles.
6. 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.
7. Sonicate for 20-25 min in a bath sonicator.
8. Perfuse into the heart for 30 min using peristaltic pumps. Refer to stepwise instructions in Section 3.5.

6 . Recording Epicardial Signals

1. 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.
2. Fill the horizontal chamber with Tyrode solution and turn on the peltier unit to bring the bath temperature to 37 °C.
3. Position the fiber optic on the surface of the heart.
   1. Place the fiber optic inside a 2 mL pipette and then attach the pipette to a micromanipulator.
   2. Use the micromanipulator to slightly press the fiber optic against the surface of the LV.
4. Externally pace the heart with a stimulator controlled by a wave generator.
   1. Program the wave generator to provide a square pulse with a width of 1 ms.
   2. Set the stimulator to be externally synchronized and connect the external input to the wave generator.
   3. To each output of the stimulator, connect a wire with an acupuncture needle soldered at the end.
   4. Place both acupuncture needles in the apex of the heart approximately 3 mm apart from each other.
   5. Only after the needles are placed in the tissue, turn on the output of the stimulator to prevent electric shock.
5. In the acquisition software, adjust acquisition frequency to 10 kHz.

7 . Recording Endocardial Signals

1. Refer to step 6.1 and 6.2 to stabilize the heart after loading the dye.
2. 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.
3. Place an intravitreal surgery sclerotomy adaptor to aid in positioning the first fiber optic into the endocardium using a micromanipulator.
4. Externally pace the heart. Refer to step 6.4.
5. In acquisition software, adjust acquisition frequency to 10 kHz.

Results

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...

Discussion

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^​2+ dynamics (Rhod 2 cytosolic Ca²⁺, ...

Materials

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