This work was supported by NIH/NIEHS R03ES021581 (awarded to M.D.R.) and by the University of Rochester Environmental Health Center (NIH/NIEHS P30 ES001247).

1. Preparation of Fly Cultures, Solutions, and Embryo Handling Devices
<ol>
	<li>Prepare a cage culture of Drosophila. Place 500+ mating flies of the desired strain in a population cage fitted with a 10 cm grape-agar plate and a spot of yeast paste. Maintain culture in a 25 &#176;C humidity controlled incubator. NOTE:&#160;Cage cultures require a day or two of conditioning to obtain consistent embryo laying patterns. Grape plates with yeast paste are changed once in the morning and once in the evening during conditioning.</li>
	<li>Prepare EPS. Warm solutions of surfactants (cocamide DEA and ethoxylated alcohol) at 37 &#176;C. Pipette 18 ml of d-limonene to a glass scintillation vial equipped with a small stir bar. Pipette 1 ml each of the two surfactants (5% final concentration of each) to the d-limonene. Mix thoroughly and remove stir bar. NOTE: This stock solution of EPS is good for approximately 2 months at room temperature. The solution should be warmed at 37 &#176;C and swirled to fully solubilize surfactants prior to use. EPS and d-limonene should be stored in a glass container as they will dissolve some plastics over time.</li>
	<li>Prepare embryo dechorionation solution, incubation mediums, dye and drug solutions. Mix 25 ml of bleach with 25 ml H2O and place in shallow dish. Prepare modified basic incubation medium (MBIM) and MBIM-T according to prior recipe1,12. Prepare Shields and Sang M3 cell culture medium and PBS according to manufacturer&#8217;s protocol (see Materials List). Prepare stock solution of permeabilization dye at 10 mM concentration in DMSO.</li>
	<li>Assemble embryo-handling devices and supplies:
	<ol>
		<li>Prepare dechorionation and EPS treatment basket (see Figure 1A). Cut off a 3 cm section of a disposable 50 ml polypropylene centrifuge/culture tube with a fine-toothed saw. Make the welding surface flush by rubbing the tube section on sandpaper adhered to a benchtop surface. Weld the tube section to Nitex nylon mesh by melting the rim of the tube section over a flame and pressing on to the mesh on a glass plate. Let cool and trim off extra mesh. NOTE: The sheer sides and bottom allow for rapid and complete rinsing off of residual bleach and EPS at the respective steps. The welding steps should be carried out under a fume hood.</li>
		<li>Prepare a development basket. Cut off top portion of a 50 ml centrifuge/culture tube flush with the rim of the cap. Remove and modify the cap by cutting out a central opening and notches along the rim as shown in Figure 1B. Screw the cap over the mesh and on the threads of the cut off tube section and trim extra mesh. NOTE: The cap contains notches in the rim that permit diffusion to the bulk medium when positioned in the 60 mm reservoir dish (Figure 1B&#8217;).</li>
		<li>Prepare components of a slide chamber. Cut a square of DO membrane that is larger than the slide chamber opening. Apply a very thin layer of vacuum grease to the inside lip of the opening. Affix the DO membrane into the opening sealing it against the grease with the retainer ring. Trim excess DO membrane by cutting it back close to the retainer ring. NOTE: Specifications for the slide chamber can be found in Kiehart et al13.This chamber is not commercially available and requires custom fabrication by a machine shop.</li>
	</ol>
	</li>
</ol>
2. Staging, Dechorionation, and EPS Treatment of Embryos
<ol>
	<li>Staging embryos by timed collection. Set a fresh grape/yeast plate to the fly culture cage in the AM. Allow embryos to be laid for 1 hr at 25 &#176;C. Discard this plate and replace with a fresh grape/yeast plate for a subsequent embryo laying of 2 hr at 25 &#176;C. Collect this plate and place in 18 &#176;C incubator for further developmental staging. NOTE: 1 hr of development at 25 &#176;C is equal to 2 hr of development at 18 &#176;C. The effect of aging at 18 &#176;C versus 25 &#176;C on maintaining EPS permeabilization in late stage embryos can be seen in Figure 2.</li>
	<li>Dechorionation. Gently rinse embryos off of grape plate into mesh basket using 25 &#176;C tap water and a paintbrush. Rinse excess yeast away from the embryos in the basket under a gentle stream of tap water. Immerse the basket in 50% bleach for 2 min. Wash embryos thoroughly under a stream of tap water. NOTE: Gently squirt bleach solution on the embryos intermittently using a plastic pipette. While 2 min is usually sufficient for complete dechorionation, this incubation time should be checked by direct examination and adjusted accordingly. Maintain dechorionated embryos in the basket immersed in tap water and proceed immediately to the EPS step.</li>
	<li>EPS Treatment
	<ol>
		<li>Prepare six 60 mm dishes with approximately 10 ml of PBS in each. Prepare EPS dilution by dissolving 75 &#181;l EPS in 2.925 ml MBIM (1:40) in a 50 ml glass beaker with swirling. Note: A&#160;white emulsion forms from this mixture.</li>
		<li>Blot excess water from the bottom of the mesh basket with a lab wipe. Immerse basket in dilute EPS in beaker and immediately swirl to disperse embryos in the EPS solution in the bottom of the basket. Continue swirling motion for 30 sec. NOTE: EPS dilutions and exposure times can be varied to control permeability. It is recommended that optimal EPS dilutions and treatment times be established empirically with the strains of flies being used. Increasing exposure time to 60-90 sec is favorable for stage 12 and older embryos.</li>
		<li>Remove basket, blot away excess EPS with a lab wipe. Proceed with six sequential washes in 10 ml of PBS in the 60 mm dishes. Use a plastic pipette to gently squirt embryos with PBS in each of the six washes. Proceed to dye and drug treatment steps. NOTE: EPS can be disposed of down the sink.</li>
	</ol>
	</li>
</ol>
3. Dye and Drug Treatment of Permeabilized Embryos
<ol>
	<li>Dye Treatment
	<ol>
		<li>Add 5 &#181;l of 10 mM CY5 carboxylic acid dye* to 1 ml of MBIM-T (50 &#181;M final concentration) in a 1.5 ml microfuge tube and vortex to mix. NOTE: *Choice of dye depends on downstream analysis. CY5 carboxylic acid is effective for subsequent analyses using fixation and immunostaining. Rhodamine B is useful for analysis of living embryos. The red emission of Rhodamine B, and the green emission of its metabolites1, can present some complications with downstream applications using fluorescence. Drug or toxin can be added to the dye solution to initiate treatment at this stage, or to limit drug/toxin treatment to a pulse at this stage. Care should be taken to handle and dispose of drugs and toxins according to MSDS and environmental safety standards.</li>
		<li>Transfer embryos from the mesh basket to the dye solution using a paintbrush. Cap the tube and invert repeatedly to ensure the embryos float freely in suspension in the dye solution. Place tube on a nutating rocker for 15 min at room temperature.</li>
		<li>Remove tube from nutator and let embryos settle. Remove dye solution with a fine pipette and replace with 1 ml of MBIM-T to wash. Invert tube to re-suspend embryos completely. Let embryos settle and repeat with three more MBIM-T washes. Remove all MBIM-T from final rinse and proceed to incubation step.</li>
	</ol>
	</li>
	<li>Incubation for Embryo Development
	<ol>
		<li>Transfer embryos to one of two chambers: The development basket or the slide chamber. NOTE: The development basket is preferred for longer developmental periods, and is required if subsequent fixation and immunostaining steps are to be executed. The slide chamber is optimal for higher resolution time-lapse imaging and is most effective for short developmental periods (e.g., the early embryonic events). Drug or toxin can be added to the medium at various concentrations and embryo development can be monitored in real-time with live embryos or at an endpoint using standard fixation and immunostaining protocols.</li>
		<li>Development in Baskets
		<ol>
			<li>Clean a development basket by squirting with 70% ethanol, rinse thoroughly with de-ionized water and blot dry with lab wipe. Prepare 6 ml of incubation medium with desired concentration of drug or toxin. Place the development basket in medium in 60 mm dish taking care to not trap air bubbles under the mesh base. NOTE: Two media are commonly used: MBIM alone, or MBIM/M3 in a 50:50 mixture, the latter being more effective for longer development periods.</li>
			<li>Transfer permeabilized embryos to mesh surface in base of basket with a paintbrush. Gently squirt the embryos with media from the surrounding reservoir. Disperse the embryos with the paintbrush so that they are in a monolayer on the mesh. NOTE: Proceed to microscopy imaging and evaluate permeabilization and viability characteristics of the preparation (see step 4).</li>
		</ol>
		</li>
		<li>Development in the Slide Chamber
		<ol>
			<li>Invert slide chamber and apply a small bead of vacuum grease on the perimeter of the opening. Place 150 &#181;l of medium with desired amount of drug or toxin on surface of DO membrane within the opening. NOTE: Two media are commonly used: MBIM alone, or MBIM/M3 in a 50:50 mixture, the latter being more effective for longer development periods.</li>
			<li>Transfer permeabilized embryos to the drop of media with a paintbrush. Disperse the embryos making them settle on the membrane within the drop.</li>
			<li>Carefully apply a 25 mm circular coverslip over the opening thereby flattening the medium. Press down gently along the perimeter of the coverslip to form a seal with the grease. NOTE: Proceed to microscopy imaging to evaluate permeabilization and viability characteristics of the preparation (see step 4).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Identification of Permeabilized Viable Embryos
<ol>
	<li>Identify permeabilized embryos. Observe embryos under epifluorescence with a microscope equipped with a digital camera. Acquire images (in the blue wavelength to determine the profile yolk autofluorescence) of several fields using fixed microscope and camera settings.</li>
	<li>Determine permeabilization of embryos based on relative fluorescence intensity. Use a stereomicroscope with a large working distance to accommodate the basket and to allow manipulations of the embryos. The microscope should be equipped with a programmable X-Y-Z stage, epifluorescence illumination and a digital camera. Image the embryos taking care to record exposure and stage position parameters of each embryo image to enable re-evaluation of the same embryos at a later time point (see representative result in Figure 3). NOTE: Patterns of dye uptake will vary depending on the dye used, length of exposure and embryo age. A wide range of dye uptake across a single preparation is typical and reflects the variation in the degree of permeabilization.</li>
	<li>Identify viable embryos. After marking position of permeabilized embryos, continue with embryo development at room temperature or 25 &#176;C. Return basket or slide chamber to microscope. Observe under blue channel fluorescence and acquire image of yolk autofluorescence in previously identified permeabilized embryos. Assess viability according to normal progression of yolk distribution (see Rand&#160;et al.1 and representative result in Figure 3). NOTE: Other morphological features of the embryo observed with brightfield microscopy can be used to assess viability. It is recommended that establishing the level of permeability that is compatible with viability be determined empirically with each dye and strain of fly used.</li>
	<li>Evaluate drug or toxin effects in permeabilized viable embryos.</li>
	<li>Embryos processed with the above protocol are poised for a number of conventional analyses subsequent to drug or toxin exposure. Several of the various types of analyses are considered in the Discussion below and include direct observation of morphogenesis in live embryos as well as post- fixation immunostaining analyses. Both of these approaches are enhanced by use of vital dyes (e.g., GFP) that reveal gene expression patterns and cell lineage and morphology profiles.</li>
</ol>

<ol>
	<li>Rand, M. D., Kearney, A. L., Dao, J., Clason, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permeabilization+of+Drosophila+embryos+for+introduction+of+small+molecules.">Permeabilization of Drosophila embryos for introduction of small molecules.</a> Insect biochemistry and molecular biology. 40, 792-804 (2010).</li><li>Jaeger, J., Manu, J., Reinitz, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+blastoderm+patterning.">Drosophila blastoderm patterning.</a> Curr Opin Genet Dev. 22, 533-541 (2012).</li><li>Kopczynski, C. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+high+throughput+screen+to+identify+secreted+and+transmembrane+proteins+involved+in+Drosophila+embryogenesis.">A high throughput screen to identify secreted and transmembrane proteins involved in Drosophila embryogenesis.</a> Proceedings of the National Academy of Sciences of the United States of America. 95, 9973-9978 (1998).</li><li>Bejsovec, A., Chao, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T.+crinkled+reveals+a+new+role+for+Wingless+signaling+in+Drosophila+denticle+formation.">T. crinkled reveals a new role for Wingless signaling in Drosophila denticle formation.</a> Development. 139, 690-698 (2012).</li><li>Zappe, S., Fish, M., Scott, M. P., Solgaard, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+MEMS-based+Drosophila+embryo+injection+system+for+high-throughput+RNAi+screens.">Automated MEMS-based Drosophila embryo injection system for high-throughput RNAi screens.</a> Lab on a chip. 6, 1012-1019 (2006).</li><li>Delubac, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+system+with+integrated+microinjector+for+automated+Drosophila+embryo+injection.">Microfluidic system with integrated microinjector for automated Drosophila embryo injection.</a> Lab on a chip. 12, 4911-4919 (2012).</li><li>Mahowald, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fixation+problems+for+electron+microscopy+of+Drosophila+embryos.">Fixation problems for electron microscopy of Drosophila embryos.</a> Drosophila Info Serv. 36, 130-131 (1962).</li><li>Margaritis, L. H., Kafatos, F. C., Petri, W. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+eggshell+of+Drosophila+melanogaster.+I.+Fine+structure+of+the+layers+and+regions+of+the+wild-type+eggshell.">The eggshell of Drosophila melanogaster. I. Fine structure of the layers and regions of the wild-type eggshell.</a> Journal of cell science. 43, 1-35 (1980).</li><li>Mazur, P., Cole, K. W., Mahowald, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Critical+factors+affecting+the+permeabilization+of+Drosophila+embryos+by+alkanes.">Critical factors affecting the permeabilization of Drosophila embryos by alkanes.</a> Cryobiology. 29, 210-239 (1992).</li><li>Arking, R., Parente, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+RNA+inhibitors+on+the+development+of+Drosophila+embryos+permeabilized+by+a+new+technique.">Effects of RNA inhibitors on the development of Drosophila embryos permeabilized by a new technique.</a> The Journal of experimental zoology. 212, 183-194 (1980).</li><li>Li, J. S., Li, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Major+chorion+proteins+and+their+crosslinking+during+chorion+hardening+in+Aedes+aegypti+mosquitoes.">Major chorion proteins and their crosslinking during chorion hardening in Aedes aegypti mosquitoes.</a> Insect biochemistry and molecular biology. 36, 954-964 (2006).</li><li>Strecker, T. R., McGhee, S., Shih, S., Ham, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permeabilization+staining+and+culture+of+living+Drosophila+embryos.">Permeabilization staining and culture of living Drosophila embryos.</a> Biotech Histochem. 69, 25-30 (1994).</li><li>Kiehart, D. P., Montague, R. A., Rickoll, W. L., Foard, D., Thomas, G. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+microscopic+methods+for+the+analysis+of+cellular+movements+in+Drosophila+embryos.">High-resolution microscopic methods for the analysis of cellular movements in Drosophila embryos.</a> Methods in Cell Biology. Drosophila melanogaster: Practical uses in cell and molecular biology. 44, 518 (1994).</li><li>Patel, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+melanogaster:+Practical+uses+in+cell+and+molecular+biology.">Drosophila melanogaster: Practical uses in cell and molecular biology.</a> Methods in Cell Biology. 44, 445-487 (1994).</li><li>Engel, G. L., Delwig, A., Rand, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+methylmercury+on+Notch+signaling+during+embryonic+neural+development+in+Drosophila+melanogaster.">The effects of methylmercury on Notch signaling during embryonic neural development in Drosophila melanogaster.</a> Toxicol In Vitro. 26, 485-492 (2012).</li><li>Limbourg, B., Zalokar, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permeabilization+of+Drosophila+eggs.">Permeabilization of Drosophila eggs.</a> Developmental biology. 35, 382-387 (1973).</li><li>Schulman, V. K., Folker, E. S., Baylies, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+reversible+drug+delivery+to+internal+tissues+of+Drosophila+embryos.">A method for reversible drug delivery to internal tissues of Drosophila embryos.</a> Fly (Austin). 7, (2013).</li></ol>

The authors have nothing to disclose.

The above method outlines a means to obtaining viable Drosophila embryos that are accessible to small molecule treatments across a wide developmental range. This method introduces the novel and simple finding that aging embryos at 18 &#176;C enables permeabilization of late stage embryos with the same efficacy as previously seen only in early stage embryos. In addition, use of the far-red dye CY5 carboxylic acid as a permeability indicator has proven effective in post-fix applications and does not interfere with conventional red and green fluorescent markers that can be used to reveal developmental phenotypes. These findings significantly advance the efficacy and utility of the EPS method.
This method is amenable to analyses of both living and fixed embryo preparations. Using brightfield microscopy and the slide chamber set up, typical features to score in the first half of embryo development are morphogenetic movements such cellularization of the blastoderm, cephalic furrow formation, germband elongation and germband retraction1. With GFP or RFP reporters more specific endpoints can be discerned, e.g. early segmentation patterns and formation of neural structures in later development1. GFP and RFP reports also enable observation of developmental events in living permeabilized embryos in both the slide chamber and developmental basket preparations. A simple method to determine gross toxicity of an applied drug or chemical is to monitor the pattern of yolk protein auto-fluorescence in the blue channel to determine a delay or cessation of development1.
The EPS methodology also has great potential for broadening the investigative tools for non-model insects, in particular the mosquito, which shares a similar architecture of the eggshell. Application of small molecules to embryos of other insect species would open up an avenue of investigation where standard genetic approaches to functional studies are currently lacking. Embryos developed in baskets can be processed for formaldehyde fixation, therefore opening up analyses to the wide array of immunostaining reagents available for Drosophila studies. However, this step requires that a post-fix determination of permeabilization be feasible to correlate phenotypes with embryos that had been made accessible to the drug. Application of CY5 carboxylic acid dye has proven highly effective for this approach. CY5 carboxylic acid is efficiently taken up in permeabilized embryos (Figure 3A). During development CY5 is concentrated in the yolk, which is ultimately sequestered in the lumen of the forming gut at stage 14 and later (Figure 3B). After fixation, CY5 fluorescence is markedly decreased, yet reliably detectable in the gut and serves as a marker of those embryos that were effectively permeabilized at the outset (see representative result Figure 4). It should be noted that CY5 detection at this stage requires longer camera exposures (e.g., 1-4 seconds) for detection. Thus, scoring of phenotypes with immunostaining patterns can proceed using the CY5 signal to confirm similarly permeabilized embryos together with tissue specific markers (e.g., neural specific antibodies seen in Figure 4).
The biggest challenge in this method is the sensitivity of embryo viability to the permeabilization process, something that has long troubled prior attempts to develop this method9,10,16. Viability subsequent to permeabilization is starkly age-dependent, and increases dramatically the older the embryo is upon permeabilization9. Yet, as we have shown previously, permeability becomes increasingly difficult with age1. A recent report now demonstrates the ability to permeabilize late stage embryos (stage 14) by re-invoking heptane as a solvent in conjunction with d-limonene17. The broad utility of this latter method is not clear as only one drug effect was characterized (nocodazole) and application to earlier stage embryos was not described17. Furthermore, application of the 18 &#176;C development step outlined above yields late stage permeability and further avoids use of toxic organic solvents. The investigator who is new to the EPS protocol will experience variability in permeabilization and viability outcomes upon initial attempts. The steps outlined here give the investigator the tools to systematically vary conditions of permeabilization treatments and subsequent incubation steps to optimize conditions for specific strains of Drosophila they are working in their own laboratory setting.
An additional challenge with the method is the variability in chemical uptake seen from embryo to embryo. This variability is reflected in the heterogeneity of CY5 dye uptake seen in embryos immediately after dye treatment (Figure 3A). In contrast, Rhodamine B dye is more rapidly and evenly dispersed across the embryonic tissues than CY5 dye (Figure 2B&#8221;). Thus, some embryo-to-embryo variability may be harbored in the distribution properties of the chemical, drug or toxin of interest and is inherent to the method. Where quantification of dose is critical, it is recommended that uptake of the drug or toxin of interest is characterized through an alternative analytical method. Nonetheless, the ease of the above protocol, and the ability to screen hundreds of embryos, allows the investigator to evaluate dose responses and score characteristic phenotypes with little investment of resources, making for a powerful first approach to characterizing drugs or toxins in this highly developed model system.

The Drosophila embryo continues to be a premier model for investigation of fundamental mechanisms of development2. This powerful model is supported by a wide array of molecular genetic tools that permit manipulations of essentially any gene at any time point and within any developing organ. The small size, rapid development, and extensive characterization of morphogenesis of the Drosophila embryo make it a model of choice for genetic screens, many of which have uncovered fundamental developmental pathways3,4. Numerous phenotypes in the Drosophila embryo have been characterized and are easily interpretable, often providing a means to identify underlying molecular genetic mechanisms responsible for an abnormal trait.
Historically, a shortcoming of the fly embryo model has been the difficulty of introducing small molecules to embryonic tissues. This obstacle has posed limitations on: 1) using known bioactive small molecules as probes to interrogate developmental mechanisms and 2) using this established model to evaluate teratogenic or pharmacologic activity of uncharacterized small molecules. As a consequence, the screening potential of the fly embryo has been underutilized in characterization of small molecule activity.
Delivery of small molecules to the fly embryo can be achieved with two methods: 1) permeabilization of the eggshell and 2) microinjection. This article presents advances to the method of permeabilization that are easy to execute in the setting of a conventional Drosophila laboratory. It should be noted that recent advances in microinjection methods with microfluidics technology is also contributing to methods of introducing compounds to the embryo5,6. Introducing molecules to the embryo is prevented by a waxy layer of the eggshell7. The Drosophila eggshell consists of five layers. From the inside out they are: the vitelline membrane, the waxy layer, the inner chorionic layer, the endochorion and the exochorion8. The three outer chorionic layers can be removed by brief emersion of the embryo in dilute bleach, a step referred to as dechorionation. The exposed waxy layer can then be compromised by exposure to organic solvents, such as heptane and octane7,9, rendering the dechorionated embryo permeable, while it remains encased in the underlying vitelline membrane. However, use of these solvents introduces complications due to their toxicity and the difficulty in regulating their strong permeabilizing action, both of which have stark negative effects on embryo viability9,10.
A method of permeabilization using a composition termed embryo permeabilization solvent (EPS) has been previously described1. This solvent consists of d-limonene and plant-derived surfactants that enable the solvent to be miscible with aqueous buffers. The low toxicity of d-limonene and the ability to dilute the solvent to desired concentrations has yielded an effective method to generate permeable embryos with high viability1. However, two endogenous factors have continued to bring limitations to the application. First, embryos demonstrate heterogeneity in permeability after EPS treatment, even when care is taken to maintain close developmental staging. Second, embryos older than approximately eight hours have proven difficult to permeabilize, consistent with a hardening of the eggshell that occurs after egg laying11.
Described here are advances in the EPS method that: 1) assist in identifying and analyzing near-identically permeabilized embryos, even after fixation and immunostaining steps have been executed and 2) enable permeabilization of embryos at late developmental time points (&#62;8 hr, stage 12 and older). Specifically, application of a far-red dye, CY5 carboxylic acid, is described that serves as a permeability indicator, which persists in the embryo during development and after formaldehyde fixation. In addition, it is shown that rearing embryos at 18 &#176;C maintains the eggshell in an EPS sensitive state, enabling permeabilization of late stage embryos (stages 12-16).
These advances overcome the previously mentioned limitations to the EPS methodology. This application will therefore provide investigators with a means to introduce small molecules of interest to the embryo at distinct developmental time points while maintaining viability.

<table border="0" cellpadding="0" cellspacing="0" width="740">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Fly Cage</td>
			<td style="width:201px;">Flystuff.com</td>
			<td style="width:119px;">59-101</td>
			<td style="width:204px;">http://flystuff.com/general.php</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Cocamide DEA [Ninol 11-CM]&#160;</td>
			<td style="width:201px;">Stepan Chemical</td>
			<td style="width:119px;">call for special order</td>
			<td>http://www.stepan.com/</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Ethoxylated alcohol [Bio-soft 1-7]&#160;</td>
			<td style="width:201px;">Stepan Chemical</td>
			<td style="width:119px;">call for special order</td>
			<td>http://www.stepan.com/</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">d-limonene (Ultra high purity grade)</td>
			<td style="width:201px;">Florida Chemical Co.&#160;</td>
			<td style="width:119px;">call for special order</td>
			<td>http://www.floridachemical.com/</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium hypochlorite&#160;</td>
			<td style="width:201px;">Fisher</td>
			<td>SS290-4</td>
			<td>http://www.fishersci.com/</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tween-20&#160;</td>
			<td style="width:201px;">Fisher</td>
			<td>BP337</td>
			<td>http://www.fishersci.com/</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PBS powder</td>
			<td style="width:201px;">Sigma</td>
			<td>56064C</td>
			<td>http://www.sigmaaldrich.com/</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rhodamine B</td>
			<td style="width:201px;">Sigma</td>
			<td>R6626</td>
			<td>http://www.sigmaaldrich.com/</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CY5 carboxylic acid</td>
			<td style="width:201px;">Lumiprobe</td>
			<td>#23090</td>
			<td>http://www.lumiprobe.com/p/cy5-carboxylic-acid</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">DMSO</td>
			<td style="width:201px;">Sigma</td>
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a method of permeabilization of drosophila embryos for assays of small molecule activity

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of genetic manipulations with this model has supplanted pharmacological approaches that are commonplace in other animal models and cell-based assays. Here we describe recent advances in a protocol that enables application of small molecules to the developing fruit fly embryo. The method details steps to overcome the impermeability of the eggshell while maintaining embryo viability. Eggshell permeabilization across a broad range of developmental stages is achieved by application of a previously described d-limonene embryo permeabilization solvent (EPS1) and by aging embryos at reduced temperature (18 &#176;C) prior to treatments. In addition, use of a far-red dye (CY5) as a permeabilization indicator is described, which is compatible with downstream applications involving standard red and green fluorescent dyes in live and fixed preparations. This protocol is applicable to studies using bioactive compounds to probe developmental mechanisms as well as for studies aimed at evaluating teratogenic or pharmacologic activity of uncharacterized small molecules.

Embryo handling devices are pictured in Figure 1 to assist in visualizing the &#8220;home-made&#8221; devices for manipulation in the above Protocols. Results seen in Figure 2 illustrate the robust effect of rearing embryos at 18 &#176;C on their ability to be permeabilized by EPS at late stages of development. This condition is applied in the protocol step 2.1. Efficacy of the CY5 carboxylic acid dye to reveal the various levels of permeability typically seen in EPS treated embryos is seen in Figure 3. The developmental dynamics of the dye distribution in the yolk is also seen in Figure 3, revealing a criterion used to assess viability, as described in Protocol step 4.2. The utility of the CY5 dye in determining embryo permeabilization subsequent to toxin treatment, formaldehyde fixation and immunostaining is illustrated by the result in Figure 4.
<img alt="Figure 1" src="/files/ftp_upload/51634/51634fig1highres.jpg" /> 
Figure 1.&#160;Embryo handling devices for the EPS method. The flat-bottomed basket is used in dechorination and EPS exposure steps (A,A&#39;). The development basket is used for longer developmental exposures of permeabilized embryo (B,B&#39;). The slide chamber is used for shorter developmental exposures and higher resolution imaging of live embryos (C,C&#39;. See text for further description).
<img alt="Figure 2" src="/files/ftp_upload/51634/51634fig2highres.jpg" /> 
Figure 2.&#160;Effect of aging at 18 &#176;C on EPS efficacy in late stage embryos. Embryos were collected for two hours followed by aging at 18 &#176;C for 20 hr (Panels A-A&#8221;, B-B&#8221;) or at 25 &#176;C for 10 hr (Panel C-C&#8221;). Embryos at 18 &#176;C were then dechorionated and divided into two samples. The first sample was treated directly with 1 mM Rhodamine B dye in MBIM-T for 5 min, washed and visualized under brightfield and blue and red fluorescence channels (Panel A-A&#8221;). The second sample was treated with EPS (1:10 in MBIM for 1 min), washed and then treated with 1 mM Rhodamine B for 5 min and washed before visualization (Panel B-B&#8221;). Embryos raised at 25 &#176;C were dechorionated and treated directly with EPS (1:10 in MBIM for 1 min), washed and then treated with 1 mM Rhodamine B for 5 min before visualization (Panel C-C&#8221;). Embryos were determined to be at stage 14 by the folds in the gut revealed by yolk autofluorescence in the blue channel (Panel A&#8217;, B&#8217;, C&#8217;). Embryos raised at 18 &#176;C are impermeable prior to EPS treatment as seen by absence of Rhodamine B uptake (Panel A&#8221;). EPS treatment of 18 &#176;C embryos yields a high degree of permeability as seen by Rhodamine B uptake (Panel B&#8221;). Embryos raised at 25 &#176;C remain impermeable even with EPS treatment as seen by exclusion of Rhodamine B (Panel C&#8221;).
<img alt="Figure 3" src="/files/ftp_upload/51634/51634fig3highres.jpg" /> 
Figure 3.&#160;Incorporation of CY5 in permeable and viable embryos. Embryos were collected at 25 &#176;C for 2 hr and aged for 14 hr at 18 &#176;C (equivalent to 7-9 hr embryos at 25 &#176;C, stage 12). After dechorination, EPS treatment was done (1:40 in MBIM for 1 min) followed by incubation in CY5 dye (50 &#181;M in MBIM-T for 15 min). Embryos were washed three times in MBIM-T and transferred to development basket with MBIM in the reservoir. Development was allowed to proceed for 8 hr at room temperature. Uptake of CY5 (Red) is imaged in the far-red channel immediately after dye treatment and washing (Panel A) and after 8 hr development (Panel B). Distribution of the yolk is seen by autofluorescence in the blue channel. Dye uptake, hence permeability, is seen to vary from embryo to embryo. CY5 dye (red) is seen to localize to the yolk (blue), which becomes concentrated to the lumen of the gut at stage 16 (purple, Panel B).
<img alt="Figure 4" src="/files/ftp_upload/51634/51634fig4highres.jpg" /> 
Figure 4.&#160;Determination of permeabilization and methylmercury effects in fixed and immunostained embryos. Embryos were collected at 25 &#176;C for 2 hr and aged for 14 hr at 18 &#176;C (equivalent to 7-9 hr embryos at 25 &#176;C, stage 12). After dechorination, EPS treatment was done (1:40 in MBIM for 1 min) followed by incubation in CY5 dye (50 &#181;M in MBIM-T for 15 min) together with methylmercury (50 &#181;M MeHg, Panel B) or DMSO solvent control (0.1% final concentration, Panel A). Embryos were washed with MBIM-T and placed in a development basket with MBIM:M3 medium in the reservoir and aged for an additional 8 hr at room temperature. Embryos were then fixed in a two-phase 4% paraformaldehyde-heptane preparation by a standard protocol14. Staining was performed with anti-Fasciclin II (green in A,B and white in A&#8217;,B&#8217;) to label motor neurons and anti-elav antibodies (red in A, B) to label all neuron cell bodies. CY5 dye is revealed by direct fluorescence, which requires extended exposure due to diminished fluorescence intensity due to fixation (CY5 is pseudo-colored blue in all panels). The effects of MeHg are seen in the irregular patterning and clustering of the lateral chordotonal neuron cell bodies (elav-positive, labeled in red and denoted with white arrows in B versus A). In addition, a characteristic branching of the segmental (SN) (solid green arrows in A&#8217;) is seen to be highly variable with MeHg exposure (solid green arrows in B&#8221;) consistent with previously reported effects of MeHg on the embryo15. Projection of the intersegmental and segmental nerves at their roots are seen to be displaced posteriorly with MeHg exposure (open green arrow in B&#8217;). Note: Methylmercury is a potent neurotoxin. Care should be taken to wear gloves and eye protection when handling. Disposal should be done through an institutional environmental safety facility and service.

Watch this Scientific Journal Video about A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity at JoVE.com

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

Introduction of small molecules to the developing Drosophila embryo offers great potential for characterizing biological activity of novel compounds, drugs, and toxins as well as for probing fundamental developmental pathways. Methods described herein outline steps that overcome natural barriers to this approach, expanding the utility of the Drosophila embryo model.

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of ...

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12.4K Views.  University of Rochester School of Dentistry and Medicine. Introduction of small molecules to the developing Drosophila embryo offers great potential for characterizing biological activity of novel compounds, drugs, and toxins as well as for probing fundamental developmental pathways. Methods described herein outline steps that overcome natural barriers to this approach, expanding the utility of the Drosophila embryo model.

Video: A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

Neurocircuit Assays for Seizures in Epilepsy Mutants of Drosophila

Drosophila melanogaster is a useful tool for studying seizure like activity. A variety of mutants in which seizures can be induced through either physical shock or electrical stimulation is available for study of various aspects of seizure activity and behavior. All flies, including wild-type, will undergo seizure-like activity if stimulated at a high enough voltage. Seizure like activity is an all-or-nothing response and each genotype has a specific seizure threshold. The seizure threshold of a specific genotype of fly can be altered either by treatment with a drug or by genetic suppression or enhancement. The threshold is easily measured by electrophysiology. Seizure-like activity can be induced via high frequency electrical stimulation delivered directly to the brain and recorded through the dorsal longitudinal muscles (DLMs) in the thorax. The DLMs are innervated by part of the giant fiber system. Starting with low voltage, high frequency stimulation, and subsequently raising the voltage in small increments, the seizure threshold for a single fly can be measured.

Neurocircuit Assays for Seizures in Epilepsy Mutants of Drosophila

Using high frequency electrical stimulation, seizure-like activity can be induced in Drosophila. This activity is easily recorded from the giant fiber system.

Drosophila melanogaster is a useful tool for studying seizure like activity. A variety of mutants in which seizures can be induced through either physical ...

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 ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Two Types of Assays for Detecting Frog Sperm Chemoattraction

Sperm chemoattraction in invertebrates can be sufficiently robust that one can place a pipette containing the attractive peptide into a sperm suspension and microscopically visualize sperm accumulation around the pipette1. Sperm chemoattraction in vertebrates such as frogs, rodents and humans is more difficult to detect and requires quantitative assays. Such assays are of two major types - assays that quantitate sperm movement to a source of chemoattractant, so-called sperm accumulation assays, and assays that actually track the swimming trajectories of individual sperm.
Sperm accumulation assays are relatively rapid allowing tens or hundreds of assays to be done in a single day, thereby allowing dose response curves and time courses to be carried out relatively rapidly. These types of assays have been used extensively to characterize many well established chemoattraction systems - for example, neutrophil chemotaxis to bacterial peptides and sperm chemotaxis to follicular fluid. Sperm tracking assays can be more labor intensive but offer additional data on how chemoattractancts actually alter the swimming paths that sperm take. This type of assay is needed to demonstrate the orientation of sperm movement relative to the chemoattrractant gradient axis and to visualize characteristic turns or changes in orientation that bring the sperm closer to the egg.
Here we describe methods used for each of these two types of assays. The sperm accumulation assay utilized is called a "two-chamber" assay. Amphibian sperm are placed in a tissue culture plate insert with a polycarbonate filter floor having 12 &mu;m diameter pores. Inserts with sperm are placed into tissue culture plate wells containing buffer and a chemoatttractant carefully pipetted into the bottom well where the floor meets the wall (see Fig. 1). After incubation, the top insert containing the sperm reservoir is carefully removed, and sperm in the bottom chamber that have passed through the membrane are removed, pelleted and then counted by hemocytometer or flow cytometer.
The sperm tracking assay utilizes a Zigmond chamber originally developed for observing neutrophil chemotaxis and modified for observation of sperm by Giojalas and coworkers2,3. The chamber consists of a thick glass slide into which two vertical troughs have been machined. These are separated by a 1 mm wide observation platform. After application of a cover glass, sperm are loaded into one trough, the chemoattractant agent into the other and movement of individual sperm visualized by video microscopy. Video footage is then analyzed using software to identify two-dimensional cell movements in the x-y plane as a function of time (xyt data sets) that form the trajectory of each sperm.

Eggs and the extracellular coatings around eggs frequently release peptides, proteins and small molecules that communicate with sperm to guide them to the egg thereby promoting fertilization. Using frog sperm we describe and compare two classes of assays used to detect sperm chemoattraction &#x2013; sperm accumulation assays and sperm tracking assays.

Sperm chemoattraction in invertebrates can be sufficiently robust that one can place a pipette containing the attractive peptide into a sperm suspension ...

Small-scale Nuclear Extracts for Functional Assays of Gene-expression Machineries

A great deal of progress in understanding gene expression has been made using in vitro systems. For most studies, functional assays are carried out using extracts that are prepared in bulk from 10-50 or more liters of cells grown in suspension. However, these large-scale preparations are not amenable to rapidly testing in vitro effects that result from a variety of in vivo cellular treatments or conditions. This journal video article shows a method for preparing functional small-scale nuclear extracts, using HeLa cells as an example. This method is carried out using as few as three 150 mm plates of cells grown as adherent monolayers. To illustrate the efficiency of the small-scale extracts, we show that they are as active as bulk nuclear extracts for coupled RNA Polymerase II transcription/splicing reactions. To demonstrate the utility of the extract protocol, we show that splicing is abolished in extracts prepared from HeLa cells treated with the splicing inhibitor drug E7107. The small-scale protocol should be generally applicable to any process or cell type that can be investigated in vitro using cellular extracts. These include patient cells that are only available in limited quantities or cells exposed to numerous agents such as drugs, DNA damaging agents, RNAi, or transfection, which require the use of small cell populations. In addition, small amounts of freshly grown cells are convenient and/or required for some applications.

A protocol for preparation of robust, small-scale HeLa nuclear extracts is described. This protocol is valuable for assays that require use of small populations of cells, such as cells treated with drugs or RNAi. The method should be applicable to a wide variety of gene expression assays and other cell types, including patient cells.

A great deal of progress in understanding gene expression has been made using in vitro systems.  For most studies, functional assays are carried out using ...

High-throughput Screening for Small-molecule Modulators of Inward Rectifier Potassium Channels

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, atrial fibrillation, and pain1,2. For the most part, however, progress toward understanding their therapeutic potential or even basic physiological functions has been slowed by the lack of good pharmacological tools. Indeed, the molecular pharmacology of the inward rectifier family has lagged far behind that of the S4 superfamily of voltage-gated potassium (Kv) channels, for which a number of nanomolar-affinity and highly selective peptide toxin modulators have been discovered3. The bee venom toxin tertiapin and its derivatives are potent inhibitors of Kir1.1 and Kir3 channels4,5, but peptides are of limited use therapeutically as well as experimentally due to their antigenic properties and poor bioavailability, metabolic stability and tissue penetrance. The development of potent and selective small-molecule probes with improved pharmacological properties will be a key to fully understanding the physiology and therapeutic potential of Kir channels.
The Molecular Libraries Probes Production Center Network (MLPCN) supported by the National Institutes of Health (NIH) Common Fund has created opportunities for academic scientists to initiate probe discovery campaigns for molecular targets and signaling pathways in need of better pharmacology6. The MLPCN provides researchers access to industry-scale screening centers and medicinal chemistry and informatics support to develop small-molecule probes to elucidate the function of genes and gene networks. The critical step in gaining entry to the MLPCN is the development of a robust target- or pathway-specific assay that is amenable for high-throughput screening (HTS).
Here, we describe how to develop a fluorescence-based thallium (Tl+) flux assay of Kir channel function for high-throughput compound screening7,8,9,10.The assay is based on the permeability of the K+ channel pore to the K+ congener Tl+. A commercially available fluorescent Tl+ reporter dye is used to detect transmembrane flux of Tl+ through the pore. There are at least three commercially available dyes that are suitable for Tl+ flux assays: BTC, FluoZin-2, and FluxOR7,8. This protocol describes assay development using FluoZin-2. Although originally developed and marketed as a zinc indicator, FluoZin-2 exhibits a robust and dose-dependent increase in fluorescence emission upon Tl+ binding. We began working with FluoZin-2 before FluxOR was available7,8 and have continued to do so9,10. However, the steps in assay development are essentially identical for all three dyes, and users should determine which dye is most appropriate for their specific needs. We also discuss the assay's performance benchmarks that must be reached to be considered for entry to the MLPCN. Since Tl+ readily permeates most K+ channels, the assay should be adaptable to most K+ channel targets.

Methods for developing and validating a quantitative fluorescence assay for measuring the activity of inward rectifier potassium (Kir) channels for high-throughput compound screening is presented.

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, ...

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

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Chapters in this video

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

Method for Culture of Early Chick Embryos ex vivo (New Culture)

Double Whole Mount in situ Hybridization of Early Chick Embryos

Neurocircuit Assays for Seizures in Epilepsy Mutants of Drosophila

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

In-vivo Centrifugation of Drosophila Embryos

Upright Imaging of Drosophila Embryos

Two Types of Assays for Detecting Frog Sperm Chemoattraction

Small-scale Nuclear Extracts for Functional Assays of Gene-expression Machineries

High-throughput Screening for Small-molecule Modulators of Inward Rectifier Potassium Channels