This protocol is currently used in our laboratory and has been refined over the years by many people including Drs David Sharp, Mijung Kwon, Patrizia Sommi, Dhanya Cheerambathur. We thank Dr Bill Sullivan (UCSC) who provided us with excellent advice on the manipulation and microinjection of early Drosophila embryos when our work in this system was being initiated (ref. 13). We thank all members of the Scholey laboratory. Our work on mitosis in Drosophila is supported by NIH grant GM55507.

Recipes:
Grape Juice plates:
<ul>
<li>5.5g bacto agar</li>
<li>14.5 g dextrose or glucose</li>
<li>7.15 g sucrose</li>
<li>45 ml grape juice concentrate (100% juice).</li>
<li>204.5 ml H20</li>
<li>625 &mu;l 10N NaOH</li>
</ul>
Mix all ingredients and microwave until boiling. Add 2.8 ml acid mix (acid mix: 20.9 ml propionic acid, 2.1 ml phosphoric acid, 27 ml H20)
Mix and pour onto 35 mm Petri dishes. Let solidify at room temperature for one or two days. If plates are not going to be used soon, seal with parafilm and keep at 4&deg;C (allow to equilibrate to RT before using).
Yeast paste:
Dissolve about one teaspoon of yeast (Sigma YSC2, yeast from Saccharomyces cerevisiae type II) in water to form a thick paste. Place a small amount of this on each grape juice plate just prior to use.
G-PEM buffer:
<ul>
<li>80 mM Na-PIPES, pH 6.9</li>
<li>1 mM MgCl2</li>
<li>1 mM EGTA</li>
<li>1 mM GTP</li>
</ul>
Injection buffer:
<ul>
<li>150 mM K-Aspartate</li>
<li>10 mM K-phosphate</li>
<li>20 mM imidazole, pH 7.2</li>
</ul>
Heptane glue:
Unroll double sticky tape and place in a 100ml bottle, add about 50ml heptane, seal the bottle, and rock for several days. When using, add heptane if the glue is too thick.
Dehydration chamber:
Take a 100 mm Petri dish, put one part of a 35 mm dish inside to make a "table" and add Drierite (anhydrous calcium sulfate) around it so that the height of Drierite is no higher than the "table" and cover. The coverslip with embryos will be placed on this "table" for dehydration before injection. It is a good idea to use at least some indicating Drierite and change it when it has changed color.
Protocol:
This protocol can be used for injection of virtually any soluble reagent into the Drosophila syncytial embryo: For example, either a fluorescent protein for observation or a target protein inhibitor or both.
Getting everything ready:
1.- 2 to 3 days before experiment, make grape juice plates and set up lay cage: Take a plastic bottle (6 oz, round bottom from Applied Scientific Drosophila), cut two small holes (about 1cm2) on opposite sides of the bottle and fill with a cotton piece (this will allow air to flow), tap the flies into the bottle and cover with a grape juice plate. Keep at 25&deg;C (or room temperature), flies should lay embryos for about 10 days.
2.- At least one day before the experiment pull needles for injection, we use a Kopf Needle/Pipette puller model 720 and thin walled glass filaments (World Precision Instruments tw100f). For injection of fluorescent tubulin, keep needles at 4&deg;C to prevent temperature-induced polymerization in the needle.
Tubulin preparation:
Note: All work with tubulin should be done at 4&deg;C to prevent polymerization.
<ol>
<li>Take labeled tubulin out of the freezer and put it on ice for 5 to 15 minutes.</li>
<li>Dilute with G-PEM buffer (4 to 10 fold depending on the intensity required).</li>
<li>Centrifuge at 13k rpm at 4&deg;C for 10-15 minutes.</li>
<li>Load 1 - 2 &mu;l into needle.</li>
</ol>
Antibody or chemical inhibitor:
Prepare needles as for tubulin, using an appropriate concentration (this may need to be tested). The antibody can be in injection buffer, G-PEM buffer or PBS. If another buffer is needed, this buffer will need to be injected as a control to make sure this buffer causes no damage on its own.
Note: Inhibitor and fluorescent tubulin can be mixed and injected together if the concentrations allow it. Otherwise, inject the fluorescent tubulin and wait 5 to 10 minutes before injecting the inhibitor. When performing a double injection, start with more embryos, since fewer embryos will recover normally.
Embryo collection:
<ol>
<li>Put a new grape juice plate on the lay cage.</li>
<li>Remove this plate after one hour, this is the first collection of the day and often is not very good. It can be discarded.</li>
<li>Change the plate every hour to keep collecting embryos for one hour each.</li>
</ol>
Coverslip preparation:
<ol>
<li>Put a 50 x 22mm coverslip on one side of a microscope slide. Tape the four corners to the slide so it doesn't move.</li>
<li>Using a cotton tipped applicator, put one layer of heptane glue in one line on the coverslip (the glue should not be viscous and should dry in a few seconds, otherwise add more heptane).</li>
<li>Put a piece of double sticky tape on the slide toward the side of the coverslip.</li>
</ol>
Embryo preparation:
Note: Embryos should be imaged about 2 hours after start of collection, so begin the following steps allowing sufficient time to finish them within this time-frame.
<ol>
<li>With a moistened brush carefully pick up the embryos from the grape juice plate and place on double sticky tape on slide.</li>
<li>Using the outer part of the tweezers, roll the embryos over the double sticky tape until the chorion (the outer membrane) breaks open.</li>
<li>Pick up the embryo by gently rolling it over the chorion so it sticks to the tweezer and place it on the heptane glue on the coverslip with the long side of the embryo parallel to the long side of the coverslip. Set up 10 to 20 embryos in one row.</li>
<li>Remove the coverslip and place it in the dehydration chamber for 3 to 8 minutes (this depends on the humidity of the room and the amount to be injected).</li>
<li>Place on a metal chamber with vacuum grease.</li>
<li>Cover the embryos with halocarbon oil 700 to avoid further dehydration. Embryos are now ready for injection.</li>
</ol>
Embryo injection:
<ol>
<li>Find the embryos under a 16x objective.</li>
<li>Move the embryos away, and find the needle without moving the focal plane.</li>
<li>Center the needle and move it up without moving it in x or y direction.</li>
<li>If the needle is not open, put the edge of the coverslip in the field of view, but not where the needle will be, and lower the needle to the same focal plane. Very carefully move the coverslip until it hits the needle and gently breaks it open. If the needle is open, go to step 5. (Needles can be opened with hydrofluoric acid before filling).</li>
<li>Put the embryos in view (but not where the needle will be), lower the needle into the oil and make sure you obtain nice liquid drops from the needle.</li>
<li>Carefully but steadily move the embryo into the needle and inject a drop into the embryo and move the embryo away. (Or follow the instructions of your injection apparatus).</li>
<li>After all the embryos have been injected, they are ready for observation on a confocal microscope.</li>
</ol>
Imaging:
Image the embryos on a confocal microscope with a 60 or 100X objective: Notes:
<ol>
<li>The time interval on a time lapse series should be at most a few seconds, since mitosis in the embryo is very fast, depending on the exact process to be studied.</li>
<li>A 3D series or a single plane can be acquired depending on the process of interest.</li>
<li>For fast-acting inhibitors, the transfer from injection microscope to imaging microscope has to be fast.</li>
</ol>
<img src="/files/ftp_upload/1382/1382fig1.jpg" alt="figure 1" /> Figure 1: Study of mitosis in the Drosophila syncytial embryo. Nuclei in the early embryo undergo rapid divisions without cytokinesis, during cycles 10 thru 13 nuclei form a monolayer at the cortex. A. Schematic of early embryo with nuclei at cortex. Embryo can express GFP and/or RFP labeled proteins and can be microinjected with other labeled proteins and/or inhibitors. B. Time lapse imaging of embryo expressing GFP-tubulin and RFP-histone. C. Plot of pole-pole separation for cycles 11, 12 and 13 in a wild type embryo. Spindle length exhibits periods of isometric length and periods of elongation, which are very consistent and reproducible. D. Dynamics of spindle microtubules. Injection of low concentration of rhodamine tubulin leads to speckles formed of a few tubulin subunits, which can be used to study microtubule dynamics.

<ol>
	<li>Brust-Mascher, I., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitotic+spindle+dynamics+in+Drosophila.">Mitotic spindle dynamics in Drosophila.</a> Int Rev Cytol. 259, 139-172 (2007).</li><li>Duffy, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GAL4+system+in+Drosophila:+a+fly+geneticist's+Swiss+army+knife.">GAL4 system in Drosophila: a fly geneticist's Swiss army knife.</a> Genesis. 34, 1-15 (2002).</li><li>Morris, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+methods+for+analyzing+the+functions+of+kinesins+in+early+embryos.">Microinjection methods for analyzing the functions of kinesins in early embryos.</a> Methods Mol Biol. 164, 163-172 (2001).</li><li>Brust-Mascher, I., Sommi, P., Cheerambathur, D. K., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinesin-5-dependent+poleward+flux+and+spindle+length+control+in+Drosophila+embryo+mitosis.">Kinesin-5-dependent poleward flux and spindle length control in Drosophila embryo mitosis.</a> Mol Biol Cell. 20, 1749-1762 (2009).</li><li>Brust-Mascher, I., Civelekoglu-Scholey, G., Kwon, M., Mogilner, A., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+for+anaphase+B:+role+of+three+mitotic+motors+in+a+switch+from+poleward+flux+to+spindle+elongation.">Model for anaphase B: role of three mitotic motors in a switch from poleward flux to spindle elongation.</a> Proc Natl Acad Sci U S A. 101, 15938-15943 (2004).</li><li>Brust-Mascher, I., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubule+flux+and+sliding+in+mitotic+spindles+of+Drosophila+embryos.">Microtubule flux and sliding in mitotic spindles of Drosophila embryos.</a> Mol Biol Cell. 13, 3967-3975 (2002).</li><li>Cheerambathur, D. K., Brust-Mascher, I., Civelekoglu-Scholey, G., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+partitioning+of+mitotic+kinesin-5+crosslinkers+between+microtubule-bound+and+freely+diffusing+states.">Dynamic partitioning of mitotic kinesin-5 crosslinkers between microtubule-bound and freely diffusing states.</a> J Cell Biol. 182, 421-428 (2008).</li><li>Cheerambathur, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+analysis+of+an+anaphase+B+switch:+predicted+role+for+a+microtubule+catastrophe+gradient.">Quantitative analysis of an anaphase B switch: predicted role for a microtubule catastrophe gradient.</a> J Cell Biol. 177, 995-1004 (2007).</li><li>Kwon, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chromokinesin,+KLP3A,+dives+mitotic+spindle+pole+separation+during+prometaphase+and+anaphase+and+facilitates+chromatid+motility.">The chromokinesin, KLP3A, dives mitotic spindle pole separation during prometaphase and anaphase and facilitates chromatid motility.</a> Mol Biol Cell. 15, 219-233 (2004).</li><li>Sharp, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+coordination+of+three+mitotic+motors+in+Drosophila+embryos.">Functional coordination of three mitotic motors in Drosophila embryos.</a> Mol Biol Cell. 11, 241-253 (2000).</li><li>Sharp, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+bipolar+kinesin,+KLP61F,+cross-links+microtubules+within+interpolar+microtubule+bundles+of+Drosophila+embryonic+mitotic+spindles.">The bipolar kinesin, KLP61F, cross-links microtubules within interpolar microtubule bundles of Drosophila embryonic mitotic spindles.</a> J Cell Biol. 144, 125-138 (1999).</li><li>Sharp, D. J., Rogers, G. C., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoplasmic+dynein+is+required+for+poleward+chromosome+movement+during+mitosis+in+Drosophila+embryos.">Cytoplasmic dynein is required for poleward chromosome movement during mitosis in Drosophila embryos.</a> Nat Cell Biol. 2, 922-930 (2000).</li><li>Sharp, D. J., Yu, K. R., Sisson, J. C., Sullivan, W., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antagonistic+microtubule-sliding+motors+position+mitotic+centrosomes+in+Drosophila+early+embryos.">Antagonistic microtubule-sliding motors position mitotic centrosomes in Drosophila early embryos.</a> Nat Cell Biol. 1, 51-54 (1999).</li><li>Waterman-Storer, C. M., Desai, A., Bulinski, J. C., Salmon, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+speckle+microscopy,+a+method+to+visualize+the+dynamics+of+protein+assemblies+in+living+cells.">Fluorescent speckle microscopy, a method to visualize the dynamics of protein assemblies in living cells.</a> Curr Biol. 8, 1227-1230 (1998).</li></ol>

This protocol is relatively straightforward, however, each step requires practice to make sure the embryos are not damaged. Careful control experiments always need to be done to ensure that reliable results are being obtained. One good way to begin this is by microinjecting buffer or rhodamine tubulin into control embryos expressing GFP-tubulin to make sure that mitosis proceeds normally through all cycles at the embryo surface (cycles 10 through 13). In control embryos you should not observe any physical connections bet...

microinjection techniques for studying mitosis in the drosophila melanogaster syncytial embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Watch this Scientific Journal Video about Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo at JoVE.com

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

microinjection-techniques-for-studying-mitosis-drosophila

Research

JoVE Journal

Biology

14.1K Views.  University of California, Davis. This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis. 

Video: Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

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 ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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  ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

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 ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular biology has been greatly fueled by the discovery of noncoding RNAs as major players in post-transcriptional regulation. Such a revolutionary perspective has been accompanied and triggered by the development of powerful technologies for profiling short RNAs expression, both at the high-throughput level (genome-wide identification) or as single-candidate analysis (steady state accumulation of specific species). Although several state-of-art strategies are currently available for dosing or visualizing such fleeing molecules, Northern Blot assay remains the eligible approach in molecular biology for immediate and accurate evaluation of RNA expression. It represents a first step toward the application of more sophisticated, costly technologies and, in many cases, remains a preferential method to easily gain insights into RNA biology. Here we overview an efficient protocol (Enhanced Northern Blot) for detecting weakly expressed microRNAs (or other small regulatory RNA species) from Drosophila melanogaster whole embryos, manually dissected larval/adult tissues or in vitro cultured cells. A very limited amount of RNA is required and the use of material from flow cytometry-isolated cells can be also envisaged.

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The aim of this publication is to visualize and discuss the operative steps of an Enhanced Northern Blot protocol on RNA extracted from Drosophila melanogaster embryos, cells, and tissues. This protocol is particularly useful for the efficient detection of small RNA species.

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular ...

Generation of Transgenic Hydra by Embryo Microinjection

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among multicellular animals. Hydra is used as a model for the study of regeneration, pattern formation, and stem cells. However, research efforts have been hampered by lack of a reliable method for gene perturbations to study molecular function. The development of transgenic methods has revitalized the study of Hydra biology1. Transgenic Hydra allow for the tracking of live cells, sorting to yield pure cell populations for biochemical analysis, manipulation of gene function by knockdown and over-expression, and analysis of promoter function. Plasmid DNA injected into early stage embryos randomly integrates into the genome early in development. This results in hatchlings that express transgenes in patches of tissue in one or more of the three lineages (ectodermal epithelial, endodermal epithelial, or interstitial). The success rate of obtaining a hatchling with transgenic tissue is between 10% and 20%. Asexual propagation of the transgenic hatchling is used to establish a uniformly transgenic line in a particular lineage. Generating transgenic Hydra is surprisingly simple and robust, and here we describe a protocol that can be easily implemented at low cost.

Generation of Transgenic Hydra by Embryo Microinjection

Stably transgenic Hydra are made by microinjection of plasmid DNA into embryos followed by random genomic integration and asexual propagation to establish a uniform line. Transgenic Hydra are used to track cell movements, overexpress genes, study promoter function, or knock down gene expression using RNAi.

Methods Article

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo