Name: local field fluorescence microscopy: imaging cellular signals in intact hearts
Uploaded: 2017-03-08T13:00:00
Description: Watch this Scientific Journal Video about Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts at JoVE.com

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<...

<ol>
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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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		<tr>
			<td>Acupunture needles</td>
			<td>LHASA</td>
			<td>TC1.20x13</td>
			<td></td>
		</tr>
		<tr>
			<td>60mL 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 567nm</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&#160;</td>
			<td>ThorLabs</td>
			<td>FGL590</td>
			<td></td>
		</tr>
		<tr>
			<td>Green Nd-YAG laser 532nm, 500mW</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&#181;m NA 0.39</td>
			<td>ThorLabs</td>
			<td>FT200UMT</td>
			<td></td>
		</tr>
		<tr>
			<td>LEDs blue</td>
			<td>Lumileds</td>
			<td colspan="2">L135-B475003500000</td>
		</tr>
		<tr>
			<td>LEDs green</td>
			<td>Lumileds</td>
			<td colspan="2">L135-G525003500000</td>
		</tr>
		<tr>
			<td>Cube beam splitter, non-polarizing</td>
			<td>ThorLabs</td>
			<td>BS007</td>
			<td></td>
		</tr>
		<tr>
			<td>36&#34; Length, Dovetail Optical Rail</td>
			<td>Edmund Optics</td>
			<td>54-402</td>
			<td></td>
		</tr>
		<tr>
			<td>2.5&#34; Width, Dovetail Carrier</td>
			<td>Edmund Optics</td>
			<td>54-404</td>
			<td></td>
		</tr>
		<tr>
			<td>0.75&#34; Travel, micrometer stage</td>
			<td>Edmund Optics</td>
			<td>37-983</td>
			<td></td>
		</tr>
		<tr>
			<td>Dovetail optical rail 3&#34;</td>
			<td>ThorLabs</td>
			<td>RLA075/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Dovetail rail carrier 1&#34;</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&#160;</td>
			<td>ThorLabs</td>
			<td>ST1XY-S</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminum breadboard 6&#34;x6&#34;</td>
			<td>ThorLabs</td>
			<td>MB6</td>
			<td></td>
		</tr>
		<tr>
			<td>Nexus otpical table 4&#39;x6&#39;</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&#160; 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&#160;</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, 25mmx75mmx10mm</td>
			<td>ThorLabs</td>
			<td>BA1/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Wire shelve posts 36&#34;</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 ...

The overall goal of this local field fluorescence microscopy imaging technique is to use molecular probes to measure intracellular calcium levels, or membrane potentials from heart regions difficult to access at the whole heart level. This method can help us to answer key cardiovascular questions related to cardiac arrest and ischemia. The main advantage of this technique is that cardiac properties get measured from cellular events in the intact organ where the cells continue to be electrically, mechanically, and metabolically coupled.First, prepare the horizontal Langendorff apparatus. Load freshly prepared Tyrode solution into the 60 milliliter syringes, and add all the associated tubing of the setup. Once filled, eliminate all of the air bubbles.Then, equilibrate the loaded solution with pure oxygen gas delivered from a submerged plastic airstone. Next, place a non-absorbable surgical suture around a needle. This will serve as a cannula to retroperfuse the heart when attached to the apparatus.Next, collect the heart after injecting heparin and euthanizing the animal. First, clean the chest with ethanol. The demonstration is with a mouse, but the same procedure works for parakeet heart harvesting.Next, using dissection scissors, make an incision in the lower abdomen and then cut up the sides toward the neck. Then, carefully cut open the diaphragm, avoiding the heart. Now, pull back the cut tissue and pin it down.Next, remove the lungs and surrounding tissues to expose the heart. Once exposed, use tweezers to scoop out the heart without squeezing it. Detach it from the animal by cutting the aorta, and leave as much aorta attached to the heart as possible.Then, load the heart into a small weigh boat with one milliliter of Tyrode solution. Under a dissecting microscope, quickly remove the blood and the fatty tissue surrounding the heart using tweezers and scissors. Using a non-absorbable surgical suture and a needle, attach the aorta to the horizontal connection of the Langendorff apparatus.Secure the knot using two fine tweezers. Then, begin the retroperfusion by opening the valve to release the Tyrode solution. Clean the surrounding tissue and suction away debris to maintain the horizontal chamber clean while the heart stabilizes for 10 minutes.Make the dye fresh. For Rhod-2 AM, add 20 microliters of 20%Pluronic and DMSO to the manufacturer's 50 microgram package of dye. Mix by pipetting, avoiding bubbles.Then, transfer the liquified dye into a clear glass vial, and add one milliliter of Tyrode solution. Thoroughly dissolve the dye in this mixture using 15 to 20 minutes of sonication. Next, perfuse the prepared dye using a peristaltic pump at room temperature.First, load the dye in the dye chamber, and then, clamp all the tubing lines connected to the manifold, and keep the clamps above the manifold to prevent backflow. Next, switch on the pump and immediately close the three way valve below the 60 milliliter syringe. Position the small tube attached to the suction end of the pump next to the heart to recirculate the dye after it has been perfused.Take a small gauge needle and pinch it through the apex to relieve pressure. Then, let the heart perfuse with dye for 30 minutes. During this time, it's very important to evaluate the level of the dye in order for bubbles to not get into the tissue, which would cause ischemia and tissue damage.After perfusing the dye, retroperfuse the heart with Tyrode solution. Remove the clamp above the manifold. Open the valve, and then allow the heart to stabilize for 10 minutes.During the retroperfusion, fill the horizontal chamber with Tyrode solution and turn on the Peltier unit to bring the bath temperature to 37 degrees Celsius. To prepare the fiber optic light for the experiment, first place the fiber optic inside a two milliliter pipette. And then attach the pipette to a micromanipulator.Use the micromanipulator to slightly press the fiber optic against the surface of the lateral ventricle. Next, program a wave generator to provide a square pulse with a width of one millisecond. And set a heart stimulator to be externally synchronized and connected to the wave generator.At each of the two stimulator outputs, attach a wire with an acupuncture needle soldered at the end. Attach two acupuncture needles to the heart's apex, about three millimeters apart. Only once the needles are placed, start the stimulator, thus avoiding accidental shocks.Now, to record the epicardial signal, adjust the acquisition frequency in the software to 10 kilohertz. To collect the endocardial signal, follow the same procedure, but use a 23 gauge needle to poke a hole in the lateral ventricle near the septum. And then, using a micromanipulator, position the second fiber optic into the endocardium.To access transmural function, the morphology of action potentials recorded from the endocardium and the epicardium were compared. Fluorescence recordings were made after treating the heart with di-8-ANEPPS using the CLFFM technique. The action potential was slower in the endocardium, particularly in phase one.The duration of the action potential was described as the time it takes for the action potential to repolarize to a certain degree. When the durations were normalized and shifted to overlap at the zero phase, they could be compared. There was indeed a significant difference during phase one.Next, Rhod-2 AM was used to monitor intracellular calcium. In order to compare regional differences in the intracellular calcium, the kinetics of the calcium transience were assessed. By comparison, the time to peak of the transience was about two milliseconds longer in the endocardium.Overall, the calcium transience from the endocardium had significantly slower kinetics than the epicardium. Once mastered this technique can be used to record intracellular calcium transience and action potentials in regions difficult to access in the heart. The Langendorff portion of the setup makes it very practical to perfuse different chemicals into the heart, and assess the effects on the cardiac properties.

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Research

JoVE Journal

Biology

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

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, ...

Cellular Redox Profiling Using High-content Microscopy

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, excessive ROS levels induce a state of oxidative stress, which is accompanied by irreversible oxidative damage to DNA, lipids and proteins. Thus, quantification of ROS provides a direct proxy for cellular health condition. Since mitochondria are among the major cellular sources and targets of ROS, joint analysis of mitochondrial function and ROS production in the same cells is crucial for better understanding the interconnection in pathophysiological conditions. Therefore, a high-content microscopy-based strategy was developed for simultaneous quantification of intracellular ROS levels, mitochondrial membrane potential (&#916;&#936;m) and mitochondrial morphology. It is based on automated widefield fluorescence microscopy and image analysis of living adherent cells, grown in multi-well plates, and stained with the cell-permeable fluorescent reporter molecules CM-H2DCFDA (ROS) and TMRM (&#916;&#936;m and mitochondrial morphology). In contrast with fluorimetry or flow-cytometry, this strategy allows quantification of subcellular parameters at the level of the individual cell with high spatiotemporal resolution, both before and after experimental stimulation. Importantly, the image-based nature of the method allows extracting morphological parameters in addition to signal intensities. The combined feature set is used for explorative and statistical multivariate data analysis to detect differences between subpopulations, cell types and/or treatments. Here, a detailed description of the assay is provided, along with an example experiment that proves its potential for unambiguous discrimination between cellular states after chemical perturbation.

This paper presents a high-content microscopy workflow for simultaneous quantification of intracellular ROS levels, as well as mitochondrial membrane potential and morphology &#8211; jointly referred to as mitochondrial morphofunction &#8211; in living adherent cells using the cell-permeant fluorescent reporter molecules 5-(and-6)-chloromethyl-2&#39;,7&#39;-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) and tetramethylrhodamine methylester (TMRM).

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, ...

Oligomerization Dynamics of Cell Surface Receptors in Living Cells by Total Internal Reflection Fluorescence Microscopy Combined with Number and Brightness Analysis

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of clustering. Here, we describe an imaging approach to determine the average oligomeric state of mEGFP-tagged-receptor homocomplexes in the membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis. N&#38;B is a method similar to fluorescence-correlation spectroscopy (FCS) and photon counting histogram (PCH), which are based on the statistical analysis of the fluctuations of the fluorescence intensity of fluorophores diffusing in and out of an illumination volume during an observation time. In particular, N&#38;B is a simplification of PCH to obtain information on the average number of proteins in oligomeric mixtures. The intensity fluctuation amplitudes are described by the molecular brightness of the fluorophore and the average number of fluorophores within the illumination volume. Thus, N&#38;B considers only the first and second moments of the amplitude distribution, namely, the mean intensity and the variance. This is, at the same time, the strength and the weakness of the method. Because only two moments are considered, N&#38;B cannot determine the molar fraction of unknown oligomers in a mixture, but it only estimates the average oligomerization state of the mixture. Nevertheless, it can be applied to relatively small time series (compared to other moment methods) of images of live cells on a pixel-by-pixel basis, simply by monitoring the time fluctuations of the fluorescence intensity. It reduces the effective time-per-pixel to a few microseconds, allowing acquisition in the time range of seconds to milliseconds, which is necessary for fast oligomerization kinetics. Finally, large cell areas as well as sub-cellular compartments can be explored.

We describe an imaging approach for the determination of the average oligomeric state of mEGFP-tagged-receptor oligomers induced by ligand binding in the plasma membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis.

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of ...