Name: monitoring heart function in larval drosophila melanogaster for physiological studies
Uploaded: 2009-11-16T17:02:00
Duration: 9 min 37 s
Description: Watch this Scientific Journal Video about Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies at JoVE.com

Funding provided by the G. Ribble Fellowships in the Dept of Biology, the Kentucky Young Researchers and Undergraduate Education-eUreKa! Office (Univ. of KY).

INTRODUCTION
 The cellular mechanisms in the function of the heart for both insects and vertebrates are amazingly similar. Drosophila melanogaster are being used for investigating gene mutations that are responsible for disease states in other animals including humans (Bier and Bodmer, 2004; Peron et al., 2009). Now various diseases are being induced in this tractable genetic organism (Dowse et al., 1995; Johnson et al., 1998; Ganetzky, 2000; Bier and Bodmer, 2004; Ocorr et al., 2007a, b) and with the hope that in time gene therapy can be tested for regain of normal function. In addition Drosophila serve as a good model for physiological function at a cellular level.
Many of the past studies on Drosophila heart have been targeted toward developmental aspects in the embryo (Azpiazu and Frasch 1993, Bodmer 1993, Bodmer et al. 1990). A few studies of physiological function occurred in the pre-pupal, pupal or adult stages (Ashton et al., 2001; He and Adler, 2001; Molina and Cripps, 2001; Ponzielli et al., 2002; Sl&aacute;ma and Farka&#353;, 2005; Wessells and Bodmer, 2004). However the pupal stage is one of metamorphosis and changing hormones as well as fluctuating various biogenic amines. Also the heart is undergoing structural transformation which can prove to be difficult to control for variables when investigating physiology of the myocytes (Dulcis et al. 2005; Johnson et al. 2002; Miller, 1997; Papaefthimiou and Theophilidis, 2001). The heart is known to be myogenic in the larval stage and can readily be removed or left in situ while being bathed by a defined physiological saline to limit compounding variables such as circulating hormones or peptides (Dowse et al. 1995; Johnson et al. 1997; Dasari and Cooper, 2006; Feng et al. 2004). The myogenic nature of the larval heart is comparable to the mammalian heart in that there are pacemakers that drive the rest of the heart to pump fluid in a direction to be effective in bathing organs. The chronotropic and ionotropic nature of the Drosophila larval heart is of interest since it could serve as a rapid means to test fundamental principles and pathological effects for mammalian heart function as well as a helping to develop a comparative model for cellular physiology such as ionic regulation of pacemaker cells.
The larval heart is very susceptible to biogenic amines and peptides which vary in the hemolymph depending on food source or intrinsic state of the animal (Dasari and Cooper, 2006; Johnson et al. 1997, 2000; Nichols et al. 1999; Zornik et al. 1999). Addressing how endogenous or exogenous compounds influence the heart mechanistically is of interest. Possibly novel insecticides with fewer effects on other organisms can be developed if we gain a better understanding of insect physiology and pharmacology.
The cellular mechanism of action of the neurotransmitters and cardiac modulators in larva have not been described to date as there has not been sufficient understanding of the ionic currents and channel types present in the larval heart that contribute and regulate pacemaker activity. As far as we are aware, there are no reports documenting extracellular or intracellular recordings of myocytes to assess ionic currents to address the mechanistic effects of modulators in larval Drosophila.
In this report we present various methods to record heart rate and physiological properties of the larval heart. 
METHODS
Intact larvae:
<ol>
<li>A good means of visualizing a intact beating larval heart is directly with a microscope; however, the larva must remain still enough to count of the contractions. We development methods to monitor heart rate in freely moving larvae as well as restrained larvae. Depending on ones experimental question one approach might be more suitable than other.</li>
<li>These approaches could be used to follow an individual over extended periods of time if care is used to avoid dehydration. These methods can also be used to assess pharmacological agents introduced in the diet or to examine various times in development in mutational lines or with induction of heat shock genes.</li>
<li>The first unrestrained method we term the 'Ant farm'. This technique consists of two glass plates spaced apart by a thin layer of larvae food. The larvae are able to be visualized within one plane of focus. This technique was also used to monitor by electrical activity circadian movement in larva (Cooper and Cooper, 2004). This technique consists of two glass plates (microscope slides; 75 x 25 mm; J. Melvin Freed Brand) narrowly spaced (1 to 1.5 mm) apart by a thin layer of larvae food (e.g. moist corn meal- a modified version of Lewis, 1960) so that the larvae are able to be visualized within one plane of focus. Spacers commonly used for gel electrophoreses plates (mini gel Bio-Rad; Life Science Research, Hercules, CA 94547, USA) work very well since they can be purchased with varying thickness for use with 1st , 2nd or 3rd instar larvae ant farm procedures. Also an option is to use a solid plastic of a given thickness and cut out the region to use as the crawling space.</li>
<li>To prevent larvae from crawling out from the edges of the two plates of the glass a plastic of the desired thickness is used. Slightly tilting the platform at 20 to 45 degrees causes the larvae to remain, the majority of the time, with their head pointed downward and their tail containing the spiracles out above the food or within a air passage in the food layer.</li>
<li>This &quot;Ant Farm Technique&quot; also allows video imaging within a single plane with food of uniform thickness. In this configuration the larvae tend not to crawl throughout the food, but instead to eat and gradual move around in the 2D plane. White light is projected from the underside of the microscope stage with a mirror so that it can be moved accordingly for the best contrast of the heart or the two trachea which move while the heart contracts. A microscope (adjustable zoom 0.67 to 4.5; World Precision Instrument; Model 501379) is used. A 2X base objective and tube objective 0.5X is used to gain enough spatial resolution and magnification to cover a 1cm by 0.5 cm rectangle. A mounted camera through a trinocular mount is used (Mintron, MTV; World Precision Instrument). The ambient temperature is maintained at 20&deg;C.
 <img src="/files/ftp_upload/1596/1596fig1.jpg" alt="figure 1" /> Figure 1. Dorsal view of an intact 3rd instar larva. The movement of trachea due to pulling of the attachments from heart is used to observe the heart rate.</li>
<li>To monitor the freely moving animal one can place a few drops of a dilute food mixture over the animals head. This should be done in a glass dish so the transmitted light can pass through the larvae for visualizing the heart tube. The same microscopy set up as described above can be used to watch the heart beat and obtain counts. </li>
<li>Another approach is to restrain the larva to one location by using double stick tape on a glass slide and placing the ventral side of the larva to the tape (Baker et al., 1999). However this approach does not work well if the tape gets wet when feeding the larvae. To avoid the tape getting wet one can use Vaseline (injected out of a small needle around the base of the larvae and around the tape edge). Here one can feed larva over time without having to chase the larvae into the focus plane or while it is moving on a dish. If one wishes to free the larvae the tape can be moistened and it looses it's adhesiveness to the animal.</li>
</ol>
If one is not interested in freeing the larvae for experimentation one could use the permanent method of restraining the larva by gluing the animal to a glass slide. With use of super glue method, the animal can eat and even be covered in a moist solution while remaining adhered to the glass cover slip. The procedures are:
<ol>
<li>Take a clean slide and place a cover slip at one end of it.</li>
<li>Put a small dab of superglue at one corner of the cover slip.</li>
<li>Locate your Drosophila larva and remove it from the test tube.</li>
<li>Place the larva in a Petri dish and rinse it with a small amount of water to remove any excess food.</li>
<li>Soak up reaming food with the corner of a small tissue or paper towel.</li>
<li>Gently pick up larva with tweezers and place it on your slide on the opposite end from your cover slip.</li>
<li>Place the slide under the microscope and adjust your lends on the larva. The larva should be on its stomach with its back facing upwards. You can distinguish between the two sides of the larva because their backs feature two 'racing stripes' which are the trachea. The stomach has faint horizontal grooves running along it with very fine black hairs.</li>
<li>If the larva is facing the incorrect way, simply turn the right way by gently flipping it over with your tweezers.</li>
<li>With a new set of tweezers used specifically for glue take a small dab from the drop at the end of your cover slip and place it at the corner of the opposite end. You should use a fractionally amount of glue; just enough to cover the head of the tweezers. Also, make sure you wipe off the ends of your tweezers so that they do not become glued shut.</li>
<li>Under the microscope, double check to make sure the larva is still in the correct position. If it has turned over, see step eight.</li>
<li>Now, with the tweezers used to handle larva, pick up the larva and place it gently on the fresh patch of glue. Make sure the black mouth hooks are located near or at the edge of the cover slip and neither they or the brown spiracles come in contact with the glue.</li>
<li>Carefully press down on the larva to flatten it out.</li>
<li>Now that the larva is in place, you can administer the substances which you wish to test them.</li>
<li>This is best accomplished if you use a syringe to draw up the substance and place it in small doses by the larva's head for it to ingest.</li>
<li>Finally, the heart rate can be observed by counting the number of pulses of the spiracles in one minute.</li>
</ol>
In situ: Dissection and counting heart rate:
The preparations are slit along the mid-ventral longitudinal axis and pinned flat. The preparation dish consisted of a glass slide (VWR) with magnetic tape (Best Buys retail out-let) adhered to one side. A hole in the center of the magnetic strip allows the preparation to be viewed with transmitted light. Dissecting pins (Fine Scientific Tools, WA) are bent and glued to paper clips. The paper clips are easily maneuvered on the magnetic tape to hold the filleted preparation in place. This type of recording dish has been previously described for utilization of pinning ganglia isolated from the leech ventral nerve cord (Muller et al., 1981).
<ol>
 <li>Place intact 3rd instar larva on dissection plate. Rotate larva on its ventral side until the trachea are no longer visible through the cuticle.</li>
 <li>Place four pins in the intact larva. Two pins go on either side of the mouth hooks, and two pins go on either side of the spiracles. </li>
 <li>Add saline, and make a horizontal incision a small distance from the head pins, then continue with a horizontal incision down the length of the longitudinal axis. </li>
 <li>Stop cutting a short distance from the true heart. Cut around the true heart and along side the spiracles.</li>
 <li>Lift one dorsal pin and hook it underneath the cuticle. Use the pin to open the body cavity and spread it open. Repeat this with the remaining pins.</li>
 <li>Remove the guts, being careful not to damage either the trachea or heart. It should be noted that some structures are attached to the heart and can damage it if removed. These include several fat bodies, and the brain. If you can see the heart being stretched while removing the guts, immediately stop attempting to remove that structure.</li>
</ol>
The heart is now exposed and ready for exposure to experimental conditions.
<img src="/files/ftp_upload/1596/1596fig2.jpg" alt="figure 2" /> Figure 2. Ventral dissection of a 3rd instar to view the heart directly. Pinning of the animal on its back after dissection is used to directly apply the compounds on the heart with or without the CNS intact. Small arrow indicates where the heart and aorta are separated for transected dorsal vessel studies (see below). Tr, Trachea; Sp, Spiracles; H, Heart; AO, Aorta.
This dissection technique has been used to directly assess pharmacological agents on the heart of Drosophila larvae (Gu and Singh, 1995). The dissection time is 3-6 min. The preparation is allowed to relax while bathed in HL3 saline for 3-5 min after dissection. 
The HL3 saline was carefully controlled to be at pH 7.2 since HR slows at high pH and speeds up at lower pH (Badre et al., 2005). Gu and Singh (1995) used pH 7.0 for the pharmacological analysis of the heart and also showed maintained viability. During these trials the saline remained aerated by agitation of solution through repetitively injecting saline, through a 21-gauge needle, into a beaker. 
To quantify HR either direct observations can be used or the images can be recorded on to VHS tapes or a digital camera for playback on a screen. A photodiode method has previously been described (Dasari and Cooper, 2006).
Exposure to heat shock:
<ol>
 <li>Dissect larva.</li>
 <li>Place dissected preparation on cover plate.</li>
 <li>Heat water bath.</li>
 <li>Check temperature inside of bath by floating a temperature probe on a cover plate.</li>
 <li>When temperature is satisfactory, place preparation and cover plate in bath.</li>
 <li>Leave preparation in for a desired amount of time and remove when finished.</li>
</ol>
One can use what every heat pulse and temperature required for their experimentation.
Electrically drive the heart to pace:
<ol>
<li>Fill a focal electrode (approximately 20 microns in diameter) by using a syringe inserted and sealed on the open end and drawing the room temperature (21&deg; C) fly saline (HL3) into the electrode. Mount the electrode into the manipulator, checking that the wire is in the saline inside the electrode and the stimulator is turned off.</li>
<li>Place a Drosophila larva 3rd instar larval heart dissected preparation on the dissecting microscope and secure it in place. Place the grounding wire into the saline of the preparation, careful to avoid contact with the pins and sides of the preparation.</li>
<li>Use the manipulator to position the tip of the electrode onto the top of the heart, adjusting the focus of the microscope as needed. The best way to achieve this is to focus on the electrode while moving the tip downwards.</li>
<li>When the tip is perceived to be in position, the stimulator (model S9 Grass Instruments, Quincy, Massachusetts, USA) should be turned on with the frequency at 2 Hz, duration at 0.5 to 1 ms and voltage on the lowest setting. The delay setting should be between 12 and 14.</li>
<li>While observing the heart's rhythm, the frequency should be turned to approximately 3 Hz and the voltage slowly increased until the heart's rhythm seen corresponds to the rhythm produced by the stimulator. Normally the dissected fly heart beats at a frequency of approximately 2 Hz, so when the stimulated rhythm is higher than the intrinsic rate, the heart will be seen, the heart will be seen to beat at a faster rate than normal. Voltage should not surpass approximately 20 V as cell damage can occur at high voltages, making the preparation useless. Typically, one can stimulate the heart between 2 and 8 V, depending on distance and the seal created on the heart.</li>
<li>If maximum voltage is reached and no effect is seen, the polarity can be reversed. If this also does not work, the probe should be moved to better establish a connection on the heart.</li>
<li>Once a stimulation driven rhythm is established, the rhythm can be manipulated by varying the frequency and duration settings. It should be made known, however, that frequencies above 4 Hz can put the heart into tetany. Various alcohols and pharmacological agents can be added and their effect on gap junctions can be observed once the desired rhythm is established. Blocked gap junctions do not conduct a current and the propagated rhythm will cease.
 
<img src="/files/ftp_upload/1596/1596fig3.jpg" alt="figure 3"/> 
<img border="0" src="/files/ftp_upload/1596/1596fig4tmb.jpg" alt="figure 4"/>
Please <a href="https://www.jove.com/files/ftp_upload/1596/1596fig4.jpg">click here</a> to see a larger version of this figure.
</li>
</ol>
Measure field potentials:
<ol>
 <li>Glass electrodes are made from Kimax glass (outer diameter: 1.5 mm). The glass tube is pulled and fire-polished to produce patch tips with inside diameters ranging from 10 to 20 &mu;m.</li>
 <li>The lumen of the electrode is filled with the bathing medium. The amplifier and recording on line to a computer is the same as that used below for the intracellular recordings.</li>
 <li>The lumen of a 'macro-patch' recording electrode (St&uuml;hmer et al.,1983) is placed directly over a region of the heart. The spontaneous pacing of the muscle fibers produce a field potential that can be monitored with the focal electrode.</li>
 <li>Care needs to be given by gently lowering the lumen and raising it over each region of the heart to be monitored. The potentials are recorded through a macro-patch electrode.</li>
</ol>
Direct counting of heart rate and monitoring electrical events is possible. Altering the bathing media while recording these potentials is also feasible to asses the effects on the field potentials.
Measure intracellular potentials:
To monitor the transmembrane potentials of the myocytes a region of heart is impaled with sharp intracellular electrode (20 to 30 mOhm resistance) filled with 3 M KCl. A standard head stage and amplifier for intracellular recording can be used; however we used a model Axonclamp 2B (Molecular Devices, Sunnyvale, CA, USA) amplifier and 1 X LU head stage. It appears that the caudal end of the heart is more rigid and this is where the pacemaker cells reside since contraction initiates in this region the majority of the time (Badre et al., 2005; Dasari and Cooper 2006; Dasari et al., 2007).

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A., Magnabosco, A., Reggiani, C., Megighian, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+action+potential+to+contraction:+Neural+control+and+excitation-contraction+coupling+in+larval+muscles+of+Drosophila.">From action potential to contraction: Neural control and excitation-contraction coupling in larval muscles of Drosophila.</a> Comp. Biochem.Physiol. A: Molecular &#38; Integrative Physiol. 154, 173-183 (2009).</li><li>Ponzielli, R., Astier, M., Chartier, A., Gallet, A., Therond, P., Semeriva, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heart+tube+patterning+in+Drosophila+requires+integration+of+axial+and+segmental+information+provided+by+the+Bithorax+Complex+genes+and+hedgehog+signaling.">Heart tube patterning in Drosophila requires integration of axial and segmental information provided by the Bithorax Complex genes and hedgehog signaling.</a> Develop. 129, 4509-4521 (2002).</li><li>Sl ma, K., Farkas, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heartbeat+patterns+during+the+postembryonic+development+of+Drosophila+melanogaster.">Heartbeat patterns during the postembryonic development of Drosophila melanogaster.</a> J. Insect Physiol. 51, 489-503 (2005).</li><li>St hmer, W., Roberts, W. S., Almers, W., Sakmann, B., Neher, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Single channel recordings. The loose patch clamp, 123-132 (1983).</li><li>Wessells, R. 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<ol>
<li>Dissection tools: Fine #5 tweezers and fine scissors (all obtained from Fine Science Tools (USA), Inc., 373-G Vintage Park Drive, Foster City, CA 94404-1139)</li>
<li>Dissecting microscope with zoom function for dissection and counting heart rate. For focal stimulation of the heart this is needed as well.</li>
<li>For extracellular and intracellular recordings a compound microscope with upright objectives (4 x and 20X) is used. Standard intracellular amplifier and A/D board for on line recording to a computer. Electrical signals are recorded on line to a <a href="http://www.adinstruments.com/products/pl3504#overview" target="_blank">PowerLab/4s interface</a> (ADInstruments, Australia). We use standard <a href="http://www.adinstruments.com/products/software/research/LabChart-Software/" target="_blank">software from ADInstruments named Chart or Scope</a>.</li>
<li>Chemicals: All saline chemicals are obtained from Sigma chemical company (St. Louis, MO).</li>
<li>For intracellular recordings we use glass capillary tubing (catalogue # 30-31-0 from FHC, Brunswick, ME, 04011, USA) and for focal macropatch recording and stimulating electrodes we use Kimax-51, Kimble Products Art. No. 34502, ID 0.8-1.1mm, length 100mm. The intracellular electrode should have a resistance of 20 to 30 mOhm. The macropatch electrode is constructed by breaking off the tip of the glass after a fine tip was made from an electrode puller. The broken off tip needs to be a clean perpendicular break about 20&mu;M in diameter. The tip is then heat polished to about 10&mu;M inner diameter for the focal extracellular recording. For the stimulating electrode to drive the heart we use a final tip of about 20-50 &mu;M in diameter. The shaft of the electrode is run over a heating element to cause it to bend about 45 degrees with a gradual bend. This produces a flat or perpendicular electrode lumen over the heart tube as the angle with the micro-manipulator will produce about another 45 degrees to the preparation.</li>
</ol>

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.

Watch this Scientific Journal Video about Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies at JoVE.com

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.

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  ...

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Research

JoVE Journal

Biology

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Drosophila Larval NMJ Immunohistochemistry

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 31 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. Immunohistochemistry can be used to clearly image the components of the NMJ. In this article, we demonstrate how to use antibody staining to visualize the Drosophila larval NMJ.

Drosophila Larval NMJ Immunohistochemistry

This protocol demonstrates how to perform immunohistochemistry on dissected Drosophila larva.

Visually Mediated Odor Tracking During Flight in Drosophila

Flying insects use visual cues to stabilize their heading in a wind stream. Many animals additionally track odors carried in the wind. As such, visual stabilization of upwind tracking directly aids in odor tracking. But do olfactory signals directly influence visual tracking behavior independently from wind cues? Additionally, recent advances in olfactory molecular genetics and neurophysiology have motivated novel quantitative behavioral analyses to assess the behavioral influence of (e.g.) genetically inactivating specific olfactory activation circuits. We modified a magnetic tether system originally devised for vision experiments by equipping the arena with narrow laminar flow odor plumes. Here we focus on experiments that can be performed after a fly is tethered and is able to navigate in the magnetic arena. We show how to acquire video images optimized for measuring body angle, how to judge stable odor tracking, and we illustrate two experiments to examine the influence of visual cues on odor tracking.

Here we describe how to optimize the acquired video image for an olfactory magnetic-tether (OMT) apparatus. We also describe two sample experimental protocols for studying visuo-olfactory fusion.

Flying insects use visual cues to stabilize their heading in a wind stream. Many animals additionally track odors carried in the wind. As such, visual ...

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 ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work examining post-embryonic through adult cardiac development has been limited7-10. Examining the changing morphology of the maturing and aging heart are restricted by the lack of techniques available for staging and isolating juvenile and adult hearts. In order to analyze heart development over the fish's lifespan, we dissect zebrafish hearts at numerous stages and photograph them for further analysis11. The morphological features of the heart can easily be quantified and individual hearts can be further analyzed by a host of standard methods. Zebrafish grow at variable rates and maturation correlates better with fish size than age, thus, post-fixation, we photograph and measure fish length as a gauge of fish maturation. This protocol explains two distinct, size dependent dissection techniques for zebrafish, ranging from larvae 3.5mm standard length (SL) with hearts of 100&mu;m ventricle length (VL), to adults, with SL of 30mm and VL 1mm or larger. Larval and adult fish have quite distinct body and organ morphology. Larvae are not only significantly smaller, they have less pigment and each organ is visually very difficult to identify. For this reason, we use distinct dissection techniques.
We used pre-dissection fixation procedures, as we discovered that hearts dissected directly after euthanization have a more variable morphology, with very loose and balloon like atria compared with hearts removed following fixation. The fish fixed prior to dissection, retain in vivo morphology and chamber position (data not shown). In addition, for demonstration purposes, we take advantage of the heart (myocardial) specific GFP transgenic Tg(myl7:GFP)twu34 (12), which allows us to visualize the entire heart and is particularly useful at early stages in development when the cardiac morphology is less distinct from surrounding tissues. Dissection of the heart makes further analysis of the cell and molecular biology underlying heart development and maturation using in situ hybridization, immunohistochemistry, RNA extraction or other analytical methods easier in post-embryonic zebrafish. This protocol will provide a valuable technique for the study of cardiac development maturation and aging.

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

A clear, standardized method for dissection and isolation of the zebrafish heart at multiple developmental stages are described. Annotation and quantification techniques are also discussed.

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

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.

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.

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