Name: trap-rc, translating ribosome affinity purification from rare cell populations of drosophila embryos
Uploaded: 2015-09-10T14:00:00
Duration: 10 min 26 s
Description: Watch this Scientific Journal Video about TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos at JoVE.com

The authors thank Nicolas Allegre, Maud Peyny and Jean-Philippe Da Ponte for their excellent technical assistance. This work was supported by the Agence Nationale de la Recherche (ANR-JC-CARDIAC-SPE), the INSERM starting grant, ANR ID-CELL-SPE grant, Equipe FRM and the AFM 15845 grant.

1. Fly Line Generation&#160;
<ol>
	<li>Generate two different constructs. In the first construct, RpL10A cDNA (ribosomal protein subunit L10a) is cloned downstream of GFP into pUASP-PL-GFP-Nter plasmid (modified version from32). The use of a germline promoter allows a more polyvalent usage of the transgenic line. 
	NOTE: Obtain the second construct by cloning the regulatory sequence driving tissue-specific expression of the slouch gene upstream of GAL4 (TF) using pPTGAL4 plasmid.</li>
	<li>Inject plasmids separately into w1118 flies and insert constructs into the fly genome by P-element insertion (according to standard procedure)33.</li>
	<li>Select transformants in order to recover fly lines with constructs on two different chromosomes. The final TRAP line is created by combining these two lines (genetic crosses) in order to get one double-transgenic stable line. F0: w-; Cyo, UAS EGFP::RpL10A; MKRS/TM6B x w-;Cyo/Scu;Slouch Gal4/TM6B. F1: w-; Cyo, UAS EGFP::RpL10A; Slouch Gal4/TM6B. Massively amplify this line and collect the desired-staged embryos.</li>
</ol>
2. Embryo Collection
<ol>
	<li>Prepare 8 large cylindrical population cages containing around 40 g of young flies per cage (around 180,000 flies/cage).</li>
	<li>Proceed with so called pre-lays that allow flies to clear developing embryos from their oviducts. For that, lay the flies for 1 hr on two 11 cm-diameter petri dishes containing a mixture of solidified agar and grape juice and cover &#8531; of the surface with freshly-made yeast paste. After 3 exchanges of grape juice plates collect the real embryos on similar freshly made plates. 
	NOTE: Incubation time will vary according to developmental time-window studied.</li>
	<li>Remove plates from cages and leave them at the same temperature for the time needed to obtain correct developmental stage (e.g., 3 hr of egg-laying + 10 hr of additional incubation gives embryos from stage 10-13 hr After Egg Laying (AEL)). Resuspend the plate content (embryos, yeast paste, dead flies) with 50 ml of water using a brush.</li>
	<li>Go through a series of sieves (700 &#181;m, 355 &#181;m, 112 &#181;m) to separate embryos from dead flies and remaining body parts and collect embryos retained on the smaller-diameter sieve, then wash briefly in deionized water. Do not use more than 18 plates per collection as it can clog the sieves.</li>
	<li>Dechorionate embryos in 4.5% bleach in deionized water for 2 min and rinse thoroughly with deionized water for 30-60 sec. Incubate embryos in PBS 0.01% Tween 20, 100 &#181;g/ml cycloheximide, for 5-10 min at RT under agitation.</li>
	<li>Dry embryos on cellulose absorbent sheets and transfer them to microcentrifuge tubes. Weigh the tubes and flash-freeze the embryos by immersion in liquid nitrogen. At this stage, the dried embryos can be stored for several months at &#8722;80 &#176;C.</li>
</ol>
3. Preparation of Magnetic Beads Coupled to GFP Antibody
<ol>
	<li>Thoroughly resuspend the Protein G magnetic beads in the original bottle by gentle agitation.</li>
	<li>Transfer 90 &#181;l of beads to an RNase-free tube and collect them on a magnet (30&#8211;60 sec). 90 &#181;l of beads is necessary to perform immunoprecipitation (IP) with 1 ml of lysate containing 60-80 mg/mL proteins. Respect ratios to prevent bead saturation. Use several tubes according to protein amount.</li>
	<li>Wash beads with 1&#215; PBS 0.01% Tween 20 (500 &#181;l) and collect beads on the magnet to discard supernatant. Add 30 &#181;g of GFP antibody in 350 &#181;l of PBS 0.01% Tween 20 to the beads and incubate with slow end over end mixing for 30 min at RT.</li>
	<li>Collect beads on the magnet (30-60 sec), discard supernatant and rinse in 500 &#181;l of polysome extraction buffer (Hepes 10 mM, KCl 150 mM, MgCl2 5 mM, DTT 0.5 mM, Triton 1%, Cycloheximide 100 &#181;g/ml, Protease inhibitor 1&#215;, RNase inhibitor 100 U).</li>
	<li>Collect beads on the magnet and add 500 &#181;l of blocking buffer (BSA 0.1 &#181;g/&#181;l, yeast tRNA 0.1 &#181;g/&#181;l, glycogen 0.1 &#181;g/&#181;l in polysome extraction buffer).</li>
	<li>Incubate for 30 min at RT and repeat this step once with fresh blocking buffer.</li>
	<li>Collect beads on the magnet and rinse in 500 &#181;l of polysome extraction buffer, then recollect beads and proceed immediately to the immunopurification step (section 6).</li>
</ol>
4. Lysate Preparation
<ol>
	<li>Before starting, set up the program on the multi-directional fast speed beads grinder: 2&#215;10 sec at 5,000 rpm, 15 sec pause. Transfer dried embryos (1.5 g) to 15 ml tubes prechilled on ice containing polysome extraction buffer, and homogenize immediately. Homogenize 1.5 g of embryos in 4 ml of polysome extraction buffer.</li>
	<li>Spread homogenized embryos in two prechilled 2 mL tubes and grind the embryos by running the homogenizer program. Transfer the lysate into fresh prechilled microcentrifuge tubes on ice and prepare a postnuclear supernatant by centrifugation at 4 &#176;C for 10 min at 2,000 g.</li>
	<li>Transfer supernatant to a fresh prechilled microcentrifuge tube on ice, add 1/100 sample volume of 10% Nonidet P-40 to the supernatant (final concentration = 0.1%), and mix it gently by inverting the tube.</li>
	<li>Pulse-centrifuge the sample in a minifuge to collect the liquid at the bottom of the tube, add 1/9 sample volume of 300 mM 1,2-Diheptanoyl-sn-Glycero-3-Phosphocholine (DHPC) (final concentration = 30 mM), mix it gently by inverting the tube, and incubate the mixture on ice for 5 min (mix by inverting several times during incubation).</li>
	<li>Prepare the post-mitochondrial supernatant by centrifugation at 4 &#176;C for 10 min at 20,000 g. Measure protein concentration by the Bradford method: concentration in lysate should be between 60-80 mg/ml. Take the supernatant to a fresh new prechilled microcentrifuge tube and proceed immediately to the pre-absorption step (section 5).</li>
</ol>
5. Pre-absorption
<ol>
	<li>Thoroughly resuspend the magnetic beads by gentle agitation and transfer 30 &#181;l of beads into a microcentrifuge tube at RT (30 &#181;l of beads/1 ml embryonic lysate).</li>
	<li>Collect beads on the magnet, pipette off the supernatant, and resuspend the beads in polysome extraction buffer (500 &#181;l). Collect beads on the magnet, add embryonic lysate, and incubate for 1 hr at 4 &#176;C on a rotator.
	<ol>
		<li>Remove beads and keep supernatant on ice for the immunopurification step. Before immunopurification, keep 100 &#181;l of supernatant (input) to compare global RNA sample with the RNA sample collected after immunopurification, by RT-qPCR for example.</li>
	</ol>
	</li>
</ol>
6. Immunopurification
<ol>
	<li>Add 1 ml (corresponding to 60-80 mg/ml protein) of pre-absorbed lysate to an RNase-free tube containing 90 &#181;l of blocked beads coupled to GFP antibody. Incubate the samples at 4 &#176;C for 2 hr with gentle end-over-end mixing in a tube rotator. Alternatively, perform the incubation O/N at 4 &#176;C.</li>
	<li>After incubation, collect beads on the magnet. Use a minifuge to spin down the beads, then resuspend them in 500 &#181;l of wash buffer (Hepes 10mM, KCl 350mM, MgCl2 5mM, Nonidet P-40 1%, DTT 0.5 mM, cycloheximide 100 &#181;g/ml, RNase inhibitor 100 U) and collect them on the magnet. Repeat this step two more times.</li>
	<li>Change the beads to a new prechilled RNase-free tube and wash twice. Collect beads on a magnet and proceed to RNA extraction by adding 1 ml TRiZol directly to the beads and follow the manufacturer&#8217;s instructions.</li>
</ol>
7. RNA Clean-up and Quality Assessment
<ol>
	<li>Use an RNeasy Micro Kit as per the manufacturer&#8217;s instructions. At completion, elute RNA with (x) &#181;l RNase-free water. Warm for 2 min at 60 &#176;C, and store snap-frozen samples at -80 &#176;C. Check RNA quality using bioanalyzer (follow the manufacturer&#8217;s instructions).</li>
	<li>To test the specificity of the TRAPed RNA, perform reverse transcription on 3 ng of purified RNA (input and IP) according to the manufacturer&#8217;s instructions (superscript III). Use the cDNA obtained for qPCR using gene-specific primer sets.</li>
</ol>

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The authors have nothing to disclose.

This paper describes a modified Translating Ribosome Affinity Purification protocol (dubbed &#8216;TRAP-rc&#8217;) dedicated to the study of rare cell populations in Drosophila embryos. Information is provided on the key steps for successful isolation of specific RNA with a yield suitable for microarray or RNA-seq analysis, i.e., 1) quantity of biological material required and optimized procedure for embryo lysis and polysome extraction; 2) beads/antibody-to-lysate ratios for optimal immunoprecipitation; 3) steps allowing reduction of background including wash buffer composition so as to run TRAP-rc experiments with high specificity and sensitivity.
This protocol yields reproducible data making it readily applied to any other cell types. This technique makes it possible to analyze differential gene expression at the level of actively-translating mRNA and thus pave the way to understanding protein expression in a specific cell type at a specific time window. Note however that protein abundance will depend on the rate of the translation and degradation processes. A major limitation of the TRAP method is its inability to measure protein content in an accurate quantitative manner or to detect post-translational modification. Improving the quantification by measuring ribosome density along the mRNA body and, in doing so, in a cell-type-specific manner can make TRAP combined with ribosome footprinting a very powerful tool. In a recent paper34, this combination was performed in human embryonic kidney 293 cells. The authors ran nuclease footprinting followed by affinity purification of an inducible biotinylated form of RpL10A using streptavidin beads. This is a proof of principle that it is possible to perform ribosome footprinting in a whole organism while targeting a specific cell type, the limitation being the amount of biological material required for the purpose.
Performing TRAP experiments in parallel with global transcriptomic analyses will make it possible to track proportions between newly-transcribed and actively- translating RNA. This will be deeply informative of the potential post-transcriptional mechanisms taking place in specific developmental contexts or specific cell populations&#8212;by microRNAs for example.
In conclusion, TRAP is a highly efficient, specific and sensitive method for identifying RNAs bound to actively-translating ribosomes in a cell-specific manner. This method can be used in a wide variety of organisms and tissue types. The new TRAP-rc protocol described here was optimized for rare cell populations (less than 1% of total cell number) and for a quantity of final material sufficient for subsequent analyses at whole-genome level.
A possible limitation of this method is the requirement of a strong driver to produce tagged ribosomes in sufficient amount to compete with endogenous untagged ones in the targeted cell type. In a recent study25, authors estimated to 10-30% the ratio of tagged versus untagged ribosomes.
To overcome this problem increasing UAS-RpL10A-EGFP copy number should considerably improve this balance. Alternatively, adding to the described genetic background a UAS-GAL4 transgene will amplify the production of GAL4 protein and indirectly increase expression of RpL10A-EGFP. This should favor a better occupancy of tagged ribosomes on mRNA.
Note that this already efficient approach can be made even more powerful by adapting complementary methods to bring a more quantitative aspect or to discover and unravel molecular mechanisms that are essential for gene expression control.

Understanding how cells acquire specific properties during development is crucial in order to appreciate the complexity of organs and their potential evolution towards pathological conditions. There is a big research push to decipher the gene regulatory networks that underly cell specification and differentiation by unrevealing global gene expression profiles, Pol II binding status, transcription-factor occupancy on regulatory sequences or post-translational modifications of histones.
To assess gene expression at a global level microarrays and RNA-seq approaches are used. Microarrays allow to detect mRNA abundance, whereas RNA-seq is better geared to informing on the different types of RNAs including coding and noncoding RNAs like microRNAs, lncRNAs or snRNAs. However, these techniques cannot differentiate actively-transcribed, steady-state mRNA, actively-translating mRNA, and mRNA entering the degradation process. Measuring steady-state mRNA levels is known to be a poor estimation of proteome composition1-3. In contrary, identifying actively-translating mRNA gives a more accurate picture of protein production.
However, transcriptomic studies have mainly been led at whole-organism level by comparing gain or loss of function of a specific factor to wild-type condition4-7 or on dissected tissue, making gene expression profiles difficult to interpret. Recent advances in cell targeting tools, affinity purification and sensitivity improvements now allow to perform genome-wide analyses on small cell populations or even single cells in a living organism.
In Drosophila, large collections of transgenic lines enable specific temporal and spatial targeting of different tissues and cell subpopulations via the GAL4/UAS system8-10 yielding more accurate quantification of the molecular mechanisms required for cell function.
Among newly developed approaches polysome profiling by fractionation was described as a way to capture actively-translating RNA11. This method based on sucrose gradient centrifugation allows the selection of polysome fraction and determination of mRNA translated genome-wide when analyzed with microarrays or deep sequencing. Polysome isolation can be coupled with nuclease digestion to identify ribosomal footprints corresponding to RNA fragments protected by the ribosomes during the translation process12. Ribosomal footprinting increases the accuracy of quantification of RNA translation and RNA sequence-level resolution and ribosome positioning, makes it possible to measure rate of translation. However, to date it has not been applied to cell-specific isolation of translatomes, mainly due to the strong decrease in RNA yield obtained at the end of the ribosome footprinting process. Given that the size selection of RNA may generate potential false-positives from different RNPs that could also be protecting RNA in a similar manner, further developments are needed to improve ribosomal footprinting and adapt it to cell-specific approaches.
One of such methods, called the Translating Ribosome Affinity Purification (TRAP) has been described in 2008 by Heiman et al. and applied to isolate the translatome from a subset of neurons in mice. By epitope-tagging a ribosomal protein in a cell-type-specific manner, polysomes can be selectively purified without the laborious step of cell dissociation and sorting. In this case, the 60S ribosomal protein L10a at the surface of the ribosome was tagged with eGFP 13. Since 2008, several studies have used this method in different species, from mice14-19 to X. laevis20,21, zebrafish22,23 and Arabidopsis thaliana 24. The TRAP method has also been adapted to the Drosophila model and used to purify cell-type-specific mRNAs from neuronal cells 25 and astrocytes 26. The versatile binary GAL4/UAS system was used to express GFP-tagged RpL10A in a tissue-specific manner. Heads of adult flies were dissected to perform polysome selection from neuronal cells (targeted by pan-neural Elav-Gal4 driver) and isolated RNAs were sequenced. The large number of up-regulated genes corresponded to those known to be expressed in the nervous system, indicating that the approach has good specificity and prompting us to adapt it to rare cell populations (less than 1% of cells) present in developing Drosophila embryos.
Within the Drosophila model, the embryonic stage is a model of choice for the study of developmental processes, as the molecular actors and morphogenetic movements are evolutionarily conserved. The only tissue-specific transcriptomic approaches conducted so far in this model have been performed using cell or nuclear sorting and have only been able to study steady-state transcriptome 27-31, creating a need for methods dedicated to tissue/cell-specific translatome profiling in the fly embryo. Here we report the first TRAP protocol dedicated to Drosophila embryos. With this method, engaged-in-translation mRNA in a very restricted cell population of around 100 muscle cells per embryo was successfully isolated with high quality and specificity. The binary GAL4/UAS system is used to drive the expression of GFP-tagged RpL10A in subpopulation of muscles without any toxicity, no apparent phenotypes or developmental delay as shown previously25. Drosophila embryos possess 30 muscles per hemisegment that will give rise to the musculature of the larva. By using a regulatory region discovered in the vicinity of the slouch identity gene, 6 out of the 30 multi-nucleated muscles are targeted. In this assay, ribosomes are immobilized on the mRNA using the translation elongation inhibitor cycloheximide. Cytosolic extracts are then used for affinity purification with GFP antibody-coated magnetic beads. Quality of purified RNA was validated using bioanalyzer. Quantitative reverse transcription PCR (RT-qPCR) was used to determine the specificity and sensitivity of mRNA isolation and demonstrated that our optimized protocol is highly efficient.

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			<td>UC DAVIS/NIH Neuromab Faculty Antibodies Incorporated</td>
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			<td>Applied Biosystem&#160;</td>
			<td>AM95306</td>
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			<td>Cellulose waddin unbleached&#160;</td>
			<td>VWR</td>
			<td>115-2597&#160;</td>
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			<td>Sieves 112, 355, 710 um&#160;</td>
			<td>Retsch</td>
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			<td>TRiZol&#160;</td>
			<td>Invitrogen</td>
			<td>15596-018</td>
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			<td>MagRack 6</td>
			<td>GE HealthCare&#160;</td>
			<td>28948964</td>
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			<td>Low binding filter tips&#160;</td>
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			<td>Rnase free tube 1,5 mL&#160;</td>
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trap-rc, translating ribosome affinity purification from rare cell populations of drosophila embryos

Measuring levels of mRNAs in the process of translation in individual cells provides information on the proteins involved in cellular functions at a given point in time. The protocol dubbed Translating Ribosome Affinity Purification (TRAP) is able to capture this mRNA translation process in a cell-type-specific manner. Based on the affinity purification of polysomes carrying a tagged ribosomal subunit, TRAP can be applied to translatome analyses in individual cells, making it possible to compare cell types during the course of developmental processes or to track disease development progress and the impact of potential therapies at molecular level. Here we report an optimized version of the TRAP protocol, called TRAP-rc (rare cells), dedicated to identifying engaged-in-translation RNAs from rare cell populations. TRAP-rc was validated using the Gal4/UAS targeting system in a restricted population of muscle cells in Drosophila embryos. This novel protocol allows the recovery of cell-type-specific RNA in sufficient quantities for global gene expression analytics such as microarrays or RNA-seq. The robustness of the protocol and the large collections of Gal4 drivers make TRAP-rc a highly versatile approach with potential applications in cell-specific genome-wide studies.

The Drosophila embryonic somatic muscle system is composed of 30 muscles per hemisegment, and each muscle has a specific set of properties: code of identity genes, position, number of fusion events, attachment sites and innervation. Identifying translated mRNA in a specific muscle subset will give information on the proteins required to form these specific muscle characteristics. Using the TRAP-based method, RNA from a subpopulation of muscles expressing the identity gene slouch was purified (Figure 1A-B). A major concern of this approach is the quality and specificity of the isolated RNA. The optimized protocol reported here enabled systematic recovery of high-quality RNA assessed with bioanalyzer analysis. The profile obtained shows no degradation of RNA (Figure 1C).
In other studies developed around the TRAP method, questions were raised over the specificity of the data. Here we improve the method to suppress background linked to non-specific binding of RNA to the beads or the tubes and by the optimization of the wash buffer. In order to assess the background level and efficiency of isolating slouch-expressing cells, we led RT-qPCR trials on 3 replicates of the same embryonic stage. Fold-enrichments of 4 different genes (mef2, slouch, prospero, soxNeuro) were calculated compared to input and normalized against the RpL32 gene (Figure 1D). These results showed 2.3-fold enrichment of pan-muscular gene mef2 and5.6-fold enrichment of the slouch gene. In contrast, the two genes expressed in the neural system were depleted compared to input. Very similar fold change values were observed on 3 biological replicates, demonstrating that the protocol is robust. With Slouch-Gal4 driver and starting with 1.5 g, around 1.5x10^6 embryos were obtained that contain 100 GFP cells/embryo (total of 150&#215;10^6 positive cells).
After running the TRAP experiment, the yield is around 25&#8211;45 ng of specific RNA depending on stage-of-interest. This material is enough to run microarray analysis or RNA-seq using an amplification protocol to build an RNA-seq library. Efficiency will depend strongly on the strength of the driver used to express RpL10A-EGFP.
<img alt="Figure 1" src="/files/ftp_upload/52985/52985fig1.jpg" /> 
Figure 1. Quality and specificity assessment of TRAP-isolated RNA from Slouch-GAL4&#62;UASRpL10A-EGFP embryos. 
(A-B) Confocal images of a stage-16 embryo showing RpL10A-EGFP expression specifically in the six Slouch muscle cells per hemisegment (A). Co-localization of RpL10A-EGFP with the general muscle marker Beta3tubulin is observed on merge picture (B). (C) Quality control of TRAPed RNA ran on bioanalyzer showing perfect integrity of rRNA 18S and 28S. (D) RT-qPCR analysis showing high specificity of TRAP-isolated mRNA experiments on 3 biological replicates of stage-16 embryos. Fold change is calculated compared to input and normalized against the RpL32 gene. Mef2 transcripts present in all muscle lineages are 2.3-fold-enriched compared to input whereas more restricted slouch transcripts are 5.6-fold-enriched. Neural cells expressing prospero and soxN genes are depleted 2.2-fold and 5.5-fold respectively. Error bars represent standard deviation (n=3). <a href="https://www.jove.com/files/ftp_upload/52985/52985fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos at JoVE.com

TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos

Translating Ribosome Affinity Purification (TRAP) is able to capture cell-type-specific translation of mRNA. Here we report the first TRAP protocol dedicated to isolation of mRNA in rare cell populations of Drosophila embryos.

TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos

Measuring levels of mRNAs in the process of translation in individual cells provides information on the proteins involved in cellular functions at a given ...

trap-rc-translating-ribosome-affinity-purification-from-rare-cell

Research

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Biology

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 ...

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 ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Robust Generation of Hepatocyte-like Cells from Human Embryonic Stem Cell Populations

Despite progress in modelling human drug toxicity, many compounds fail during clinical trials due to unpredicted side effects. The cost of clinical studies are substantial, therefore it is essential that more predictive toxicology screens are developed and deployed early on in drug development (Greenhough et al 2010). Human hepatocytes represent the current gold standard model for evaluating drug toxicity, but are a limited resource that exhibit variable function. Therefore, the use of immortalised cell lines and animal tissue models are routinely employed due to their abundance. While both sources are informative, they are limited by poor function, species variability and/or instability in culture (Dalgetty et al 2009). Pluripotent stem cells (PSCs) are an attractive alternative source of human hepatocyte like cells (HLCs) (Medine et al 2010). PSCs are capable of self renewal and differentiation to all somatic cell types found in the adult and thereby represent a potentially inexhaustible source of differentiated cells. We have developed a procedure that is simple, highly efficient, amenable to automation and yields functional human HLCs (Hay et al 2008 ; Fletcher et al 2008 ; Hannoun et al 2010 ; Payne et al 2011 and Hay et al 2011). We believe our technology will lead to the scalable production of HLCs for drug discovery, disease modeling, the construction of extra-corporeal devices and possibly cell based transplantation therapies.

This article will focus on the generation of human hepatic endoderm from human embryonic stem cell populations.

Despite progress in modelling human drug toxicity, many compounds fail during clinical trials due to unpredicted side effects. The cost of clinical ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Identification of Protein Interacting Partners Using Tandem Affinity Purification

A critical and often limiting step in understanding the function of host and viral proteins is the identification of interacting cellular or viral protein partners. There are many approaches that allow the identification of interacting partners, including the yeast two hybrid system, as well as pull down assays using recombinant proteins and immunoprecipitation of endogenous proteins followed by mass spectrometry identification1. Recent studies have highlighted the utility of double-affinity tag mediated purification, coupled with two specific elution steps in the identification of interacting proteins. This approach, termed Tandem Affinity Purification (TAP), was initially used in yeast2,3 but more recently has been adapted to use in mammalian cells4-8.
As proof-of-concept we have established a tandem affinity purification (TAP) method using the well-characterized eukaryotic translation initiation factor eIF4E9,10.The cellular translation factor eIF4E is a critical component of the cellular eIF4F complex involved in cap-dependent translation initiation10. The TAP tag used in the current study is composed of two Protein G units and a streptavidin binding peptide separated by a Tobacco Etch Virus (TEV) protease cleavage sequence. The TAP tag used in the current study is composed of two Protein G units and a streptavidin binding peptide separated by a Tobacco Etch Virus (TEV) protease cleavage sequence8. To forgo the need for the generation of clonal cell lines, we developed a rapid system that relies on the expression of the TAP-tagged bait protein from an episomally maintained plasmid based on pMEP4 (Invitrogen). Expression of tagged murine eIF4E from this plasmid was controlled using the cadmium chloride inducible metallothionein promoter.
Lysis of the expressing cells and subsequent affinity purification via binding to rabbit IgG agarose, TEV protease cleavage, binding to streptavidin linked agarose and subsequent biotin elution identified numerous proteins apparently specific to the eIF4E pull-down (when compared to control cell lines expressing the TAP tag alone). The identities of the proteins were obtained by excision of the bands from 1D SDS-PAGE and subsequent tandem mass spectrometry. The identified components included the known eIF4E binding proteins eIF4G and 4EBP-1. In addition, other components of the eIF4F complex, of which eIF4E is a component were identified, namely eIF4A and Poly-A binding protein. The ability to identify not only known direct binding partners as well as secondary interacting proteins, further highlights the utility of this approach in the characterization of proteins of unknown function.

Tandem affinity purification is a robust approach for the identification of protein binding partners. As proof of concept, this methodology was applied to the well-characterized translation initiation factor eIF4E to co-precipitate the host cell factors involved in translation initiation. This method is easily adapted to any cellular or viral protein.

A critical and often limiting step in understanding the function of host and viral proteins is the identification of interacting cellular or viral protein ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

Purification of Transcripts and Metabolites from Drosophila Heads

For the last decade, we have tried to understand the molecular and cellular mechanisms of neuronal degeneration using Drosophila as a model organism. Although fruit flies provide obvious experimental advantages, research on neurodegenerative diseases has mostly relied on traditional techniques, including genetic interaction, histology, immunofluorescence, and protein biochemistry. These techniques are effective for mechanistic, hypothesis-driven studies, which lead to a detailed understanding of the role of single genes in well-defined biological problems. However, neurodegenerative diseases are highly complex and affect multiple cellular organelles and processes over time. The advent of new technologies and the omics age provides a unique opportunity to understand the global cellular perturbations underlying complex diseases. Flexible model organisms such as Drosophila are ideal for adapting these new technologies because of their strong annotation and high tractability. One challenge with these small animals, though, is the purification of enough informational molecules (DNA, mRNA, protein, metabolites) from highly relevant tissues such as fly brains. Other challenges consist of collecting large numbers of flies for experimental replicates (critical for statistical robustness) and developing consistent procedures for the purification of high-quality biological material. Here, we describe the procedures for collecting thousands of fly heads and the extraction of transcripts and metabolites to understand how global changes in gene expression and metabolism contribute to neurodegenerative diseases. These procedures are easily scalable and can be applied to the study of proteomic and epigenomic contributions to disease.

Purification of Transcripts and Metabolites from Drosophila Heads

We describe here the procedures for the extraction and purification of mRNA and metabolites from Drosophila heads. We are applying these techniques to better understand the cellular perturbations underlying neuronal degeneration. These methodologies can be easily scaled and adapted for other "omic" projects.

For the last decade, we have tried to understand the molecular and cellular mechanisms of neuronal degeneration using Drosophila as a model organism. ...