This work was supported by the Lehigh University Start-Up Fund, the NIH (R00EB010071 to C.Z., R15EB019704 to C.Z. and A.L., R03AR063271 to A.L., and R01AG014713 and R01MH060009 to R.E.T.), the NSF (1455613 to C.Z. and A.L.), the Cure Alzheimer's Fund (to R.E.T.), and the Massachusetts General Hospital (Executive Committee on Research Award to A.L.). M.C. and Y.M. was supported by the National Key Basic Research Program of China (973 Program) under Grant No. 2014CB340404.

1. Preparation of OCM System for Optical Imaging of Drosophila16
<ol>
	<li>Select a spectrometer and a high-speed line scan camera that provides a frame rate of at least 80 frame/sec so the OCM system will be able to resolve the heartbeat of Drosophila.</li>
	<li>Use a broadband light source to ensure the axial resolution of 2 &#181;m to identify the heart structure of Drosophila.</li>
	<li>Use a 10X objective to obtain a high transverse resolution.</li>
	<li>Use a 45&#176; rod mirror to reflect the reference arm light beam and to generate an annular sample arm light beam to extend the depth of focus in the specimens.</li>
	<li>Develop a custom computer program to control the OCM system and perform measurements.</li>
</ol>
2. Drosophila Culture
<ol>
	<li>&#160;Standard Fly Food Preparation
	<ol>
		<li>Put ~ 5 ml instant Drosophila formula into a polystyrene vial tube with the assistance of a paper chute.</li>
		<li>Pour ~ 8 ml water into the formula to properly saturate the food.</li>
		<li>Add different supplements to the standard fly food for different experiments. When preparing for all-trans-retinal (ATR) food for the optogenetic pacing experiment35, use a pipette to extract 100 mM ATR and dissolve into ~ 8 ml water to get the ATR concentration of 1 mM in food. After mixing the solution uniformly, pour the solution into the formula and stir it sufficiently.</li>
		<li>To prepare a high-fat-diet for studying obesity related heart dysfunctions in Drosophila,10,15 mix ~10 ml formula with 15 ml water in a cup and heat for 30 sec in a microwave oven. Put some organic extra virgin coconut oil in another cup and heat it for 90 sec in the microwave oven.</li>
		<li>Extract 7.5 ml coconut oil and mix with the prepared formula sufficiently to make the weight/volume ratio of coconut oil to food ~ 30/100, and then extract ~ 2 ml mixed food and put it to the bottom of a tube.</li>
		<li>Wait for 1 min until the medium is thoroughly saturated. Compact the food carefully with a flat surface to optimize the living conditions for the Drosophila. Add 6 - 8 grains of yeast to the prepared formula, and plug the tube with a cluster of cotton.</li>
	</ol>
	</li>
	<li>Fruit Fly Crosses and Culture
	<ol>
		<li>Take a tube with prepared standard fly food, and remove the plugged cotton. Carefully transfer the adult flies (male and female) to the tube, and plug the tube with cotton immediately. Check the cotton to make sure that there is no gap between the cotton and the tube wall to prevent flies from escaping from the tube.</li>
		<li>Keep the fruit flies in the incubator at 25 &#176;C for cross breeding. Most of the genes are active and the cellular proteins are synthesized at 25 &#176;C36-39.</li>
		<li>Take the tube out from the incubator after 8 hand transfer the adult flies out of the tube to obtain the eggs at similar age for experimental control.</li>
		<li>Continue culturing the eggs in the incubator at 25 &#176;C, which is the standard temperature for Drosophila development with the development period of 8.5 days40,41. 
		NOTE: Temperature influences the developmental period (egg to adult) and expression level of various genes.</li>
	</ol>
	</li>
</ol>
3. Performing Optical Imaging with OCM
<ol>
	<li>Mount Fly Larva for Optical Imaging 
	NOTE: The egg of Drosophila hatches in 22 - 24 hr at 25 &#176;C to the first instar larva (L1). The second instar larva emerges after another 24 hr. The largest larval form is the third instar larva, which molts after about 24 hr. Structural characteristics in larva can be used to distinguish their different developmental stages. The size of the mouthparts between the first instar and the second instar is different. The mouth hooks of the first instar larva are very small and look like two pairs of tiny black spots, while the mouth hooks of the second instar larva are larger and the structure is clearer. The spiracles are usually used to identify the second instar and the third instar. The second instar larva has clubbed anterior spiracles, while, for the third instar, the anterior spiracles are branched. A dark orange ring will begin to appear at the tip of the posterior spiracles in the third instar larva.
	<ol>
		<li>Apply a piece of double sided tape to a clean microscope glass slide. Expel the air bubbles under the tape to avoid the reflections caused by air bubbles during imaging.</li>
		<li>Take one of the tubes with the cultured flies out from the incubator at the larval stage.</li>
		<li>Identify the larva in the media, remove it from the media with a soft brush and place on a clean tissue. Remove any food stuck to the larva with a wet soft brush and dry it on the tissue.</li>
		<li>Move the cleaned fly to a tissue under the objective lens of a wide field microscope.</li>
		<li>Adjust the focus of the microscope to find a clear view of the fly. Identify the right developmental stage of the larva by its structural characteristics with the microscope.</li>
		<li>Position the fly using the soft brush. Ensure the body is straight with the dorsal side facing upward to prepare for mounting on the glass slide by the dorsal side. Perform this step under the microscope.</li>
		<li>Ensure the larva is completely dry before mounting on the tape. Otherwise, the larva will not adhere to the tape.</li>
		<li>Stick the dorsal side of the positioned fly to the double side tape on the glass slide with moderate pressure. Note that too much pressure may kill the fly and too little force will lead to fly movement during imaging.</li>
	</ol>
	</li>
	<li>Optical Imaging of Drosophila at Larval Stages (L2 and L3) with OCM 
	NOTE: A broad lumen of the heart tube can be found located in the segments between A5 to A8 at the larval stages (Figure 1). The transverse OCM M-mode images (2D + time) were acquired at the A7 segment of the heart tube for each larva to facilitate the systolic and diastolic analysis.
	<ol>
		<li>Place the mounted larva on the adjustable sample stage of the OCM system along the y-transverse direction with the dorsal side facing upward below the objective lens. A small hole in the sample stage is necessary for placing the larva to avoid its contact with the stage plane.</li>
		<li>Adjust the sample stage to move the heart tube of the fruit fly to the focal plane of the imaging beam. To easily find the A7 segment, find the posterior region of the heart tube with the real time cross sectional OCM images in the image acquisition software. Then move the stage forward until the A7 segment is visible.</li>
		<li>Set parameters of the image acquisition software to 100 A-scans per B-scan (frame), 100 B-scans, and the scanner voltage to cover ~ 0.28 mm in the x-transverse direction, and 0 V in the y-transverse direction. Click the &#34;start&#34; button in the software to acquire the background noise data for background subtraction by blocking the sample beam path with a dark cloth. 
		NOTE: 3 of the 100 frames can be used for the background subtraction.</li>
		<li>Set parameters of the data acquisition software to 128 A-scans per B-scan, 4096 B-scans, and the scanner voltage to cover ~ 0.28 mm in the x-transverse direction, and 0 V in the y-transverse direction. Click the &#34;start&#34; button in the software to acquire the transverse M-mode images across the A7 segment of the fly heart tube over a region covering 0.28 x 0.57 mm2&#160;for about 30 sec.</li>
		<li>Block the imaging beam using a dark cloth during the data saving process to avoid lengthy exposure of the fly heart to the imaging light.</li>
		<li>Repeat the measurement for 5 times to get reliable measurement of the heart function.</li>
		<li>Set parameters of the image acquisition software to 400 A-scans per B-scan, 800 B-scans, and the scanner voltage to cover ~ 1.7 mm in the x-transverse direction, and ~ 4 mm in the y-transverse direction. Move the stage in both directions to ensure the entire fruit fly can be imaged. Click the &#34;start&#34; button in the software to acquire one dataset to obtain images of the fruit fly in 3 dimensions. Note: The 3D fly structure can be rendered using Amira 3D software</li>
		<li>Use a wet soft brush to moisten the measured fly and gently remove it from the glass slide. Move it into a separate tube for continuous development. Label the tube for longitudinal study through the next developmental stages.</li>
	</ol>
	</li>
	<li>Image Drosophila at Pupal Stages 
	NOTE: All the fruit flies were taken out for imaging from PD1 to PD5. As shown in the larva schematic in Figure 1b, a broad lumen remains in A5 to A8 segments of the heart tube until PD1. From PD2, a conical chamber starts to develop between A1 to A4 segments. To acquire consistent images and facilitate heart analysis, transverse M-mode images were obtained from the A7 segment at PD1, and from A1 segment after PD2, as marked in Figure 1b.
	<ol>
		<li>Image Drosophila at PD1 
		NOTE: Drosophila will have a white puparium for a short time window (0 - 1 hr) during PD1. This time window is ideal for performing optical imaging of early pupa because the high transparency leads to higher light penetration for the OCM imaging.
		<ol>
			<li>As the fruit flies are found on the tube wall when they become pupa, remove the pupa from individual tubes for imaging at PD1 with a wet soft brush, and clean the pupa with the brush if there is food stuck to the body.</li>
			<li>Mount the fruit fly on a small glass slide directly with the wet brush and keep the dorsal side facing upward (Figure 1a). Make sure the glass slide is small enough to fit back into the tube once imaging at this stage is complete.</li>
			<li>Remove excessive water from the side of the fly body.</li>
			<li>Put the glass slide on the sample stage of the OCM system, keeping the fruit fly on top. Find clear real-time image of the A7 segment of the fly heart utilizing the same strategy described in the larva measurement.</li>
			<li>Set the same parameters of the data acquisition software as in section 3.2, and image the heartbeats at the A7 segment to acquire transverse M-mode and 3D images.</li>
			<li>After imaging, use a tweezer to place the glass slide with pupa back into the tube for continuous culture.</li>
		</ol>
		</li>
		<li>Image Drosophila at PD2 to PD5 Stages 
		NOTE: Since the specimen becomes more and more opaque during the pupal stages, the penetration depth of the imaging system will be reduced.
		<ol>
			<li>Use a tweezer to carefully remove the glass slide mounted with the fly at PD2 from the tube for imaging. At PD2, the specimen shell becomes yellowish and the body becomes less transparent compared to PD1 (Figure 1).</li>
			<li>Put the slide on the sample stage of the OCM system.</li>
			<li>Adjust the sample stage to move the fly into the focal plane of imaging beam of the OCM system. Find the anterior end of the heart tube with real-time cross-sectional OCM image. Move ~ 50 &#181;m back in the posterior direction to find the A1 segment of the heart tube. 
			NOTE: At this point of heart development (PD2), the conical chamber will be very small and may not be beating.</li>
			<li>Collect transverse M-mode datasets from the A1 segment as well as 3D data using the same method as previous developmental stages.</li>
			<li>Put the slide back to the tube carefully for continuous culture. 
			NOTE: At PD3, the color of the specimen in the shell is darker than that at PD2 stage. At PD4 stage, black stripes can be observed inside the shell of the specimens. Some flies will develop into adult from this stage in the following day, while others will evolve into PD5. At PD5 stage, black stripes are even more obviously seen in the fruit flies. These flies will become adults in the following day.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Image Drosophila at the Adult Stage 
	NOTE: At the adult stage, female and male flies can be distinguished by the size of the body and the color of the lower abdomen. Female adults have larger size, while males are smaller and dark-colored in the lower abdomen.
	<ol>
		<li>Take the tube out from the incubator when the fruit fly develops into an adult, and transfer the adult fly to a ~ 45 ml empty vial.</li>
		<li>Dip the absorbent end (~ 1 cm length, ~ 3 mm diameter) of a wand into the anesthesia, put the wand into the vial, and plug the tube with a cluster of cotton to keep the anesthetic end just below the plugged cotton and to anesthetize the fly for 3 min. The duration of anesthesia depends on the size of the fly, and may vary between 2.5 to 3.5 min (for example: male for 2.5 min, female for 3 or 3.5 min).</li>
		<li>Prepare a glass slide with a piece of double sided tape.</li>
		<li>Move the anesthetized fly onto the glass slide with dorsal side facing upward using the soft brush.</li>
		<li>Separate the wings using a tweezer and stick the wings on the tape under a microscope to fix the fly and expose the heart region for imaging.</li>
		<li>Image the fly from the A1 segment of the fly heart (Figure 1). At the end of the experiment, the fly may be sacrificed.</li>
	</ol>
	</li>
</ol>
4. Imaging Analysis16
<ol>
	<li>Develop Matlab programs to convert the 2D and 3D binary files collected with the image acquisition software to image files.</li>
	<li>Use ImageJ to identify the heart tube region in the transverse M-mode images and a magic wand algorithm to create a mask of the heart region for each transverse M-mode image. Segment the masked region and use a peak-finding algorithm to identify the systolic and diastolic locations. Calculate the time dependent heart diameter changes from the transverse M-mode images.</li>
	<li>Based on the acquired time dependent heart diameters, calculate the cardiac parameters such as HR, cardiac activity period (CAP),end diastole diameter (EDD), end systole diameter (ESD), end diastole area (EDA), and end systole area (ESA). Calculate the fractional shortening (FS) with <img alt="Equation 1" src="/files/ftp_upload/55002/55002eq1.jpg" /></li>
	<li>Use ImageJ to analyze the 3D OCM images to visualize the structural development of the fly heart.</li>
</ol>

The authors declare no conflicts of interests related to the current study.

The rapid heartbeat of Drosophila, with a maximum HR around 400 bpm at larval and adult stages, requires high imaging speed to resolve the heart diastoles and systoles (no less than 80 frames/sec based on experiences). Due to the small heart chamber size and micron scale heart wall thickness (5 - 10 &#181;m), a high spatial resolution (better than 2 &#181;m) is required for resolving the heart tube structures. In this study, a high resolution and ultrahigh speed OCM system was developed, where a spectrometer wit...

Longitudinal study of the heart in small animals contributes to understanding a variety of human related cardiovascular diseases, such as gene related congenital heart defects1,2. In the past decades, various animal models, such as mouse3,4, Xenopus5,6, zebrafish7,8, avian9, and Drosophila10-16, have been used to conduct the human heart-development related research. The mouse model has been widely used to study normal and abnormal cardiac development and cardiac defect phenotypes due to its similarities with the human heart3,4. The Xenopus embryo is especially useful in the study of heart development due to its easy handling and partial transparency5,6. The transparency of the embryo and early larva of the zebrafish model allows for easy optical observation of cardiac development7,8. The avian model is a common subject of developmental heart studies because the heart can be easily accessed after removing the eggshells and the morphological similarity of avian hearts to humans9. The Drosophila model has some unique features which make it ideal for performing longitudinal studies of the heart. First, the heart tube of Drosophila is ~ 200 &#181;m below the dorsal surface, which provides convenience for optical access and observation of the heart. Additionally, many molecular mechanisms and genetic pathways are conserved between Drosophila and vertebrates. The orthologs of over 75% of human disease genes were found in Drosophila, which have made it widely used in transgenic studies11,13. Furthermore, it has a short life cycle and low maintenance costs, and has been commonly used as a specimen model for developmental biology research14-16.
Previous reports described the protocols for monitoring Drosophila cardiac functions such as the heartbeat. However, dissection procedures were required17,18. Optical imaging provides an effective way to visualize cardiac development in animals due to its non-invasive nature. Different optical imaging modalities have been applied in performing animal cardiac study, such as two-photon microscopy19, confocal microscopy20,21, light sheet microscopy22, and optical coherence tomography (OCT)16,23-26. Comparatively, OCT is capable of providing great imaging depth in small animal hearts without using contrast agents, while keeping a high resolution and an ultrahigh imaging speed, which are important for imaging live animals. Additionally, the low cost of developing an OCT system has popularized this technique for optical imaging of specimens. OCT has been successfully used for the longitudinal study of Drosophila. Using OCT, cardiac morphological and functional imaging has been performed to study the heart structures, the functional roles of genes, and the mechanisms of cardiovascular defects in mutant models during cardiac development. For example, age-dependent cardiac function decline was confirmed with down-regulated angiotensin-converting enzyme-related (ACER) gene in Drosophila with OCT27. Phenotyping of gene related cardiomyopathy was demonstrated in Drosophila using OCT28-33. Research using OCT also revealed the functional role of the human SOX5 gene in the heart of Drosophila34. Compared with OCT, OCM uses an objective with a higher numerical aperture to provide better transverse resolution. In the past, the heart dysfunction caused by silencing an ortholog human circadian gene dCry/dClock has been studied using a custom OCM system15,16, as well as the effect of high-fat-diet on cardiomyopathies in Drosophila to understand obesity induced human cardiac diseases.15
Here, the experimental protocol is summarized for longitudinal study of the cardiac morphological and functional changes in Drosophila at second instar (L2), third instar (L3), pupa day 1 (PD1), pupa day 2 (PD2), pupa day 3 (PD3), pupa day 4 (PD4), pupa day 5 (PD5), and adult (Figure 1) using OCM to facilitate study of human-related congenital cardiac diseases. Cardiac functional parameters, such as HR and CAP were quantitatively analyzed at different developmental stages to reveal the cardiac development features.

<table><tbody><tr><td>Custom OCM imaging system</td><td>Developed in our lab</td><td></td><td></td></tr><tr><td>my Temp Mini Digital Incubator</td><td>Benchmark</td><td>H2200-HC</td><td></td></tr><tr><td>Cover glass</td><td>AmScope</td><td>200PCS</td><td></td></tr><tr><td>Cotton Ball</td><td>RITE AID</td><td></td><td></td></tr><tr><td>Instant Drosophila Formula</td><td>CAROLINA</td><td>formula 4-24</td><td></td></tr><tr><td>Yeast</td><td>ActiveDry</td><td></td><td></td></tr><tr><td>Microscope</td><td>SONY</td><td>WILD M420</td><td></td></tr><tr><td>Brush</td><td>Loew-Cornell</td><td>245B</td><td>being used to move specimens</td></tr><tr><td>Labview software</td><td>National Instruments</td><td></td><td></td></tr><tr><td>ImageJ</td><td>National Institutes of Health</td><td></td><td></td></tr><tr><td>Matlab</td><td>Mathworks</td><td></td><td></td></tr><tr><td>Tweezer</td><td>Wiha</td><td>AA SA</td><td>to fix the fruit fly wings</td></tr><tr><td>FlyNap</td><td>Carolina Biological Supply Company</td><td>4,224,898</td><td></td></tr><tr><td>Scotch Permanent Double Sided Tape, 3 M</td><td>Scotch</td><td></td><td></td></tr><tr><td>Pipette</td><td>Fisherbrand</td><td>MU18837</td><td></td></tr><tr><td>Organic Extra Coconut Oil</td><td>Spring Valley</td><td>13183</td><td></td></tr><tr><td>Microscope Slide</td><td>CapitolBrand</td><td>M3504-E</td><td></td></tr><tr><td>&lt;i&gt;Drosophila&lt;/i&gt; Vials</td><td>SEOH</td><td>8401SS</td><td></td></tr><tr><td>All-trans-retinal</td><td>Sigma-Aldrich Co.</td><td>R2500</td><td></td></tr></tbody></table>

drosophila preparation and longitudinal imaging of heart function in vivo using optical coherence microscopy (ocm)

Longitudinal study of the heartbeat in small animals contributes to understanding structural and functional changes during heart development. Optical coherence microscopy (OCM) has been demonstrated to be capable of imaging small animal hearts with high spatial resolution and ultrahigh imaging speed. The high image contrast and noninvasive properties make OCM ideal for performing longitudinal studies without requiring tissue dissections or staining. Drosophila has been widely used as a model organism in cardiac developmental studies due to its high number of orthologous human disease genes, its similarity of molecular mechanisms and genetic pathways with vertebrates, its short life cycle, and its low culture cost. Here, the experimental protocols are described for the preparation of Drosophila and optical imaging of the heartbeat with a custom OCM system throughout the life cycle of the specimen. By following the steps provided in this report, transverse M-mode and 3D OCM images can be acquired to conduct longitudinal studies of the Drosophila cardiac morphology and function. The en face and axial sectional OCM images and the heart rate (HR) and cardiac activity period (CAP) histograms, were also shown to analyze the heart structural changes and to quantify the heart dynamics during Drosophila metamorphosis, combined with the videos constructed with M-mode images to trace cardiac activity intuitively. Due to the genetic similarity between Drosophila and vertebrates, longitudinal study of heart morphology and dynamics in fruit flies could help reveal the origins of human heart diseases. The protocol here would provide an effective method to perform a wide range of studies to understand the mechanisms of cardiac diseases in humans.

The longitudinal cardiac imaging was conducted using the fruit flies with the 24B-GAL4/+ strain at room temperature with OCM. Measurements were performed at L2, L3, and at 8 hr intervals from PD1 to PD4, and adult day 1 (AD1) to track the metamorphosis process (Table 1). Larva, early pupa, late pupa and adult flies were mounted on the glass slides as seen in Figure 1A. The segment features of the heart for larval and adult flies were shown in the schematic representations in Figu...

Watch this Scientific Journal Video about Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy OCM at JoVE.com

Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy OCM

Here, the experimental protocols are described for preparing Drosophila at different developmental stages and performing longitudinal optical imaging of Drosophila heartbeats using a custom optical coherence microscopy (OCM) system. The cardiac morphological and dynamical changes can be quantitatively characterized by analyzing the heart structural and functional parameters from OCM images.

Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy (OCM)

Longitudinal study of the heartbeat in small animals contributes to understanding structural and functional changes during heart development. Optical ...

drosophila-preparation-longitudinal-imaging-heart-function-vivo-using

Research

JoVE Journal

Biology

10.8K Views.  Lehigh University. Here, the experimental protocols are described for preparing Drosophila at different developmental stages and performing longitudinal optical imaging of Drosophila heartbeats using a custom optical coherence microscopy (OCM) system. The cardiac morphological and dynamical changes can be quantitatively characterized by analyzing the heart structural and functional parameters from OCM images.

Video: Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy OCM

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

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

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

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

Bioengineering

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Neuroscience

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

Developmental Biology

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Semi-automated Optical Heartbeat Analysis of Small Hearts

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our Semi-automatic Optical Heartbeat Analysis (SOHA) uses a novel movement detection algorithm that is able to detect cardiac movements associated with individual contractile and relaxation events. The program provides a host of physiologically relevant readouts including systolic and diastolic intervals, heart rate, as well as qualitative and quantitative measures of heartbeat arrhythmicity. The program also calculates heart diameter measurements during both diastole and systole from which fractional shortening and fractional area changes are calculated. Output is provided as a digital file compatible with most spreadsheet programs. Measurements are made for every heartbeat in a record increasing the statistical power of the output. We demonstrate each of the steps where user input is required and show the application of our methodology to the analysis of heart function in all three genetically tractable heart models.

We have developed a Semi-automated Optical Heartbeat Analysis method (SOHA) for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts. We demonstrate the application of our methodology to the analysis of heart function in fruit fly and embryonic mouse hearts.

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Novel And Efficient Method for Drosophila Heart Fluorescence Staining with Cryosectioning

Drosophila heart models are widely employed in studying cardiac aging and modeling&#160;human cardiac diseases. However, the dissection of Drosophila hearts before imaging is a meticulous time-intensive process that requires advanced training and motor skills. To address these challenges, we present an innovative protocol that utilizes cryosectioning for the fluorescence imaging of Drosophila heart tissue. The protocol has been demonstrated in imaging the adult Drosophila heart but could be adapted for developmental stages. The method enhances both the efficiency and accessibility of fluorescence staining while preserving the integrity of the tissue. This protocol simplifies the process without compromising the quality of imaging, thereby reducing the dependency on technicians with highly developed training and motor skills. Specifically, we replace complex techniques, such as capillary vacuum suction, with more straightforward methods like tissue embedding. This approach allows for the visualization of cardiac structures with greater ease and reproducibility. We demonstrate the utility of this protocol by effectively detecting key cardiac markers and&#160;achieving high-resolution fluorescence and immunostaining imaging that unveils intricate details of heart morphology and cellular organization. This method provides a robust and accessible tool for researchers exploring Drosophila cardiac biology, facilitating detailed analyses of heart development, function, and disease models.

The Drosophila heart sectioning and fluorescence imaging protocol simplifies studying heart structure and pathologies. This approach involves straightforward sectioning, staining, and imaging, bypassing the technical expertise needed for traditional dissection. It enhances accessibility, making Drosophila a more widely usable model for cardiac-related research within the broader scientific community.

Drosophila heart models are widely employed in studying cardiac aging and modeling&#160;human cardiac diseases. However, the dissection of Drosophila ...

Genetics

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Developing Drosophila melanogaster Models for Imaging and Optogenetic Control of Cardiac Function

Using Drosophila melanogaster (fruit fly) as a model organism has ensured significant progress in many areas of biological science, from cellular organization and genomic investigations to behavioral studies. Due to the accumulated scientific knowledge, in recent years, Drosophila was brought to the field of modeling human diseases, including heart disorders. The presented work describes the experimental system for monitoring and manipulating the heart function in the context of a whole live organism using red light (617 nm) and without invasive procedures. Control over the heart was achieved using optogenetic tools. Optogenetics combines the expression of light-sensitive transgenic opsins and their optical activation to regulate the biological tissue of interest. In this work, a custom integrated optical coherence tomography (OCT) imaging and optogenetic stimulation system was used to visualize and modulate the functioning D. melanogaster heart at the 3rd instar larval and early pupal developmental stages. The UAS/GAL4 dual genetic system was employed to express halorhodopsin (eNpHR2.0) and red-shifted channelrhodopsin (ReaChR), specifically in the fly heart. Details on preparing D. melanogaster for live OCT imaging and optogenetic pacing are provided. A lab-developed integration software processed the imaging data to create visual presentations and quantitative characteristics of Drosophila heart function. The results demonstrate the feasibility of initiating cardiac arrest and bradycardia caused by eNpHR2.0 activation and performing heart pacing upon ReaChR activation.

Developing Drosophila melanogaster Models for Imaging and Optogenetic Control of Cardiac Function

The present protocol describes the generation of Drosophila melanogaster expressing eNpHR2.0 or ReaChR opsins in the heart for OCT imaging and optogenetic heart pacing. Detailed instructions for Drosophila OCT imaging and heart beating modulation, including the simulation of restorable heart arrest, bradycardia, and tachycardia in live animals at different developmental stages, are reported.

Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy (OCM)

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