We thank the members of the Gelfand lab who contributed to the development of these protocols. Research reported in this publication was supported by the National Institute of General Medical Science of the National Institutes of Health under award number R01GM052111 to VIG and by Basque Government Department of Education, Universities and Research under award number BFI-2011-295 to UdC.

1. Drosophila S2 cells
<ol>
	<li>Preparation of ConA coated coverslips
	<ol>
		<li>Place coverslips in a ceramic rack and wash them by chromic acid immersion for 1 hr. [CAUTION: chromic acid is corrosive and can cause irritation of eyes, nose, throat, and skin].</li>
		<li>Rinse coverslips thoroughly with constantly running dH2O&#160;for 30 min until the acid is completely washed out.</li>
		<li>Let coverslips air dry and coat them with ConA (0.5 mg/ml solution in dH2O) for 30 min.</li>
		<li>Rinse coverslips with dH2O&#160;for 15 min and let them air dry. ConA coated coverslips can be stored up to 1 month.</li>
	</ol>
	</li>
	<li>Plating the cells
	<ol>
		<li>Allow S2 cells to exponentially grow in T25 or T75 cm2 flasks depending on the number of cells required for the experiment.</li>
		<li>Count the cell density using a hemocytometer. Add 1 ml growth medium (for example Insect-Xpress) into a 35 mm tissue culture dish with ConA-coated coverslip, and gently transfer ~1 x 105&#160;cells to the dish. This cell density is optimal for imaging because processes from different cells do not overlap.</li>
		<li>In order to induce formation of processes, immediately after plating add 1ml medium with 5 &#181;M CytoD to the dish (to the final concentration of 2.5 &#181;M CytoD). Allow full development of processes by incubating the cells at least for 2 hr at 25 &#186;C before imaging.</li>
	</ol>
	</li>
</ol>
2. Drosophila&#160;Primary Neuron Culture
<ol>
	<li>Preparation for neuron culture
	<ol>
		<li>Prepare Drosophila neuronal growth medium (Schneider&#39;s medium supplemented with: 20% fetal bovine serum, heat inactivated at 55 &#186;C for 30 min; 5 &#181;g/ml insulin; 100 &#181;g/ml penicillin, 100 &#181;g/ml streptomycin; 10 &#181;g/ml tetracycline). This supplemented Schneider&#39;s medium can be stored at 4 &#186;C for 6 months.</li>
		<li>Prepare apple juice agar vials for collecting Drosophila embryos (22.5 g agar, 12.0 g sucrose and 750 ml H2O in a 2-liter flask; autoclave for 20-25 min; add 250 ml apple juice and pour ~10 ml into each Drosophila vial; after solidification, plug vials with cotton balls, and wrap vials with plastic food wrap to avoid over drying of the food). The apple juice in agar vials can be stored at 4 &#186;C in well-sealed plastic bags for 4 weeks.</li>
		<li>Coat acid-washed 25 mm circle coverslips with ConA (see details in the Protocol for Drosophila S2 cells) and sterilize them under UV light for 10-15 min. These coverslips can be stored for a month.</li>
	</ol>
	</li>
	<li>Drosophila embryonic neuron culture
	<ol>
		<li>Keep young adult flies in apple juice agar vials supplemented with dry yeast for 1-2 days before collecting embryos for neuron preparation.</li>
		<li>On the collecting day, in order to synchronize the collected embryos, transfer adult flies to a fresh apple juice agar vial for 1 hr precollection in light and discard these embryos.</li>
		<li>Transfer flies to a new apple juice agar vial, collect embryos for 1 hr, and transfer the flies to another vial for another hour, both in dark to increase the embryo yield.</li>
		<li>Remove the adult flies from the apple juice agar vial and let the embryos develop another 4 hr&#160;to stages 9-11 at room temperature (4-6 hr-old embryos are between stage 9-11).</li>
		<li>Remove the embryos by squirting water into vials and loosening the embryos from the apple juice agar with a small paintbrush (at least 10 embryos are required for one good neuron prep).</li>
		<li>Transfer the embryos to a 100 &#181;m nylon mesh cell strainer with a transfer pipette to drain water (prerinse the transfer pipette in Drosophila embryo wash buffer, 0.4% NaCl, 0.03% Triton X-100).</li>
		<li>Collect the embryos from the strainer using a small paintbrush, and transfer them into a 1.5 ml microtube filled with 1 ml Drosophila embryo wash (Note: clean any agar or cotton fibers using the paintbrush or pipette tips). In this protocol, the 1.5 ml microtube comes with a matching disposable pellet pestle.</li>
		<li>Wash embryos in Drosophila embryo wash 3x.</li>
		<li>Dechorionate the embryos in a fresh 1:1 solution of&#160; commercial bleach and 95% ethanol for 10 min [CAUTION: bleach can cause irritation to eyes, nose, throat, and skin].</li>
		<li>Move to the tissue culture hood, and wash embryos 2x in supplemented Schneider&#39;s medium.</li>
		<li>Leave 200-300 &#181;l medium in the tube with the embryos, and gently mechanically dissociate dechorionated embryos using Kimble Chase disposable pellet pestles that have been sterilized by 70% ethanol (grind 10-15x).</li>
		<li>Centrifuge the homogenized mixture at 20 x&#160;g for 2 min, and transfer the supernatant to a fresh tube. [This step is optional; it helps to remove embryo debris]</li>
		<li>Centrifuge the supernatant at 550 x&#160;g for 2 min to pellet cells.</li>
		<li>Resuspend the pellet in 500 &#181;l supplemented Schneider&#39;s medium and spin cells down again at 550 x&#160;g for 2 min; repeat the resuspension-spinning cycle once.</li>
		<li>Finally resuspend the pellet in ~100 &#181;l supplemented Schneider&#39;s medium and plate the cells onto a 25 mm ConA-coated coverslip in a sterile 35 mm Petri dish. Incubate for 10 min to allow cell attachment.</li>
		<li>Add 2 ml of supplemented Schneider&#39;s medium, with 5 &#181;M CytoD to submerge the coverslip with attached cells.</li>
		<li>Incubate the dish in a humidified 25 &#186;C incubator overnight before imaging.</li>
	</ol>
	</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+Drosophila+S2+cells+for+light+microscopy.">Preparation of Drosophila S2 cells for light microscopy.</a> J. Vis. Exp. (40), e1982 (2010).</li><li>Ling, S. C., Fahrner, P. S., Greenough, W. T., Gelfand, V. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transport+of+Drosophila+fragile+X+mental+retardation+protein-containing+ribonucleoprotein+granules+by+kinesin-1+and+cytoplasmic+dynein.">Transport of Drosophila fragile X mental retardation protein-containing ribonucleoprotein granules by kinesin-1 and cytoplasmic dynein.</a> Proc. Natl. Acad. Sci. 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I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Initial+Neurite+Outgrowth+in+Drosophila+Neurons+Is+Driven+by+Kinesin-Powered+Microtubule+Sliding.">Initial Neurite Outgrowth in Drosophila Neurons Is Driven by Kinesin-Powered Microtubule Sliding.</a> Curr. Biol. 23, 1018-1023 (2013).</li><li>Pathak, D., Sepp, K. J., Hollenbeck, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+that+myosin+activity+opposes+microtubule-based+axonal+transport+of+mitochondria.">Evidence that myosin activity opposes microtubule-based axonal transport of mitochondria.</a> J. Neurosci. 30, 8984-8992 (2010).</li><li>Clark, I. E., 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=Drosophila+pink1+is+required+for+mitochondrial+function+and+interacts+genetically+with+parkin.">Drosophila pink1 is required for mitochondrial function and interacts genetically with parkin.</a> Nature. 441, 1162-1166 (2006).</li><li>Gunawardena, S., 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=Disruption+of+axonal+transport+by+loss+of+huntingtin+or+expression+of+pathogenic+polyQ+proteins+in+Drosophila.">Disruption of axonal transport by loss of huntingtin or expression of pathogenic polyQ proteins in Drosophila.</a> Neuron. 40, 25-40 (2003).</li><li>Koo, E. H., 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=Precursor+of+amyloid+protein+in+Alzheimer+disease+undergoes+fast+anterograde+axonal+transport.">Precursor of amyloid protein in Alzheimer disease undergoes fast anterograde axonal transport.</a> Proc. Natl. Acad. Sci. U.S.A. 87, 1561-1565 (1990).</li><li>Stokin, G. B., Goldstein, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axonal+transport+and+Alzheimer's+disease.">Axonal transport and Alzheimer's disease.</a> Annu. Rev. Biochem. 75, 607-627 (2006).</li><li>Bilsland, L. G., 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=Deficits+in+axonal+transport+precede+ALS+symptoms+in+vivo.">Deficits in axonal transport precede ALS symptoms in vivo.</a> Proc. Natl. Acad. Sci. U.S.A. 107, 20523-20528 (2010).</li><li>Lloyd, T. E., 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=The+p150(Glued)+CAP-Gly+domain+regulates+initiation+of+retrograde+transport+at+synaptic+termini.">The p150(Glued) CAP-Gly domain regulates initiation of retrograde transport at synaptic termini.</a> Neuron. 74, 344-360 (2012).</li><li>Moughamian, A. J., Holzbaur, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynactin+is+required+for+transport+initiation+from+the+distal+axon.">Dynactin is required for transport initiation from the distal axon.</a> Neuron. 74, 331-343 (2012).</li><li>Lorenzo, D. N., 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=Spectrin+mutations+that+cause+spinocerebellar+ataxia+type+5+impair+axonal+transport+and+induce+neurodegeneration+in+Drosophila.">Spectrin mutations that cause spinocerebellar ataxia type 5 impair axonal transport and induce neurodegeneration in Drosophila.</a> J. Cell. 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The authors declare that they have no competing financial interests.

Here we present detailed protocols for studying microtubule-dependent cargo transport in Drosophila S2 cells and primary neuron culture. Both S2 processes and neurites are structures filled by bundled microtubules arrays that serve as tracks for organelle transport.
Two strategies are typically used to label organelles: transfection of vectors encoding a fluorescent protein with an organelle-targeting sequence or staining with fluorescent dyes added directly to cultures. Common examples of cargo labels are listed in Table 1. Transient transfection or RNAi treatments of Drosophila S2 cells if necessary is performed in suspension4,6,7,9,10 before cells are plated on coverslips for imaging.
In order to get reliable results in S2 cell experiments, it is important to keep the following points in mind:
<ol>
	<li>Plate cells at optimal density for imaging. High density will result in overlapping of processes from different cells, while at low density cells have poor viability and are unable to form processes.</li>
	<li>Allow S2 cells to completely develop processes. We routinely incubate cells for 2 hr after plating but incubation up to 24 hr&#160;can be performed if necessary.</li>
	<li>S2 cells must be treated with CytoD before they attach to ConA coated coverslips. Adding the drug after cells are attached will not induce processes formation.</li>
	<li>When using dyes, change samples for imaging every 20-25 min; long-term incubation of cells with either Mitotracker or Lysotracker is toxic to the cells.</li>
	<li>We usually track cargo transport in S2 cells by imaging for 1 min at 1 frame/sec. Under this condition cells are not photo-damaged.</li>
	<li>Only analyze organelle movement in cell processes. Organelle density in the cell body is too high for reliable tracking and polarity of the movement could not be easily determined.</li>
</ol>
The Drosophila primary neuron culture preparation procedure was modified from the protocols by the Mahowald lab23 and Lieu lab24. This protocol allows you to obtain primary neuron culture within a day and it is ready for imaging and tracking the next day.
In this protocol, you may need to pay attentions to several key steps listed below:
<ol>
	<li>Stage embryos correctly. The collected embryos need to be synchronized and staged to stages 9-11, when neuroblast delamination from the ectoderm occurs. Staging embryos too early or too late may result in fewer neuroblasts in each embryo and thus failure of harvesting neurons out of the dissociated embryos.</li>
	<li>Dechorionate embryos to the right extent (all embryos change color from completely white to more transparent). Under-dechorionation could lead to difficulty in &#160;embryo&#160;dissociation, while over-dechorionation could damage the embryos. Dechorionated embryos become very sticky and soft, thus try to avoid pipetting or touching the embryos with pipette tips during the washing steps before dissociation.</li>
	<li>Pay special attention to embryos dissociation. We find that plastic disposable pellet pestles and matching microtubes give better dissociation than a glass dounce homogenizer, especially for small number of embryos. Under-dissociation will leave large cell clusters, while over-dissociation will lead to cells breakdown and massive accumulation of cell debris. Adjust the dissociation level according to the embryo numbers.</li>
	<li>Plating neuron on the glass coverslip to a reasonable density. Crowded neurons usually develop very short neurites, while sparse neurons often fail to survive.</li>
</ol>
Additionally, we noticed that primary cell cultures often contain muscle cells, and blood cells. Overnight treatment with CytoD helps to reduce the number of muscle cell and blood cell, and promotes neurite outgrowth.

Drosophila S2 cells have gained increasing popularity for studying many cellular processes. These cells were originally obtained from trypsinized late stage embryos of OregonR Drosophila melanogaster and maintain macrophage-like features1. Drosophila S2 cells offer many advantages over other cell lines. They are cultured at 25 &#176;C without CO2, and can be imaged for many hours at room temperature without the need of heating or gas exchange. More importantly, Drosophila S2 cells are very sensitive to RNAi treatment allowing high efficiency knockdown of proteins of interest. S2 cells are highly transfectable and stable cell lines can be selected in less than four weeks to allow stable ectopic expression of proteins of interest. S2 cells normally grow either loosely attached but not spread on a flask or in suspension. They can be induced to spread on coverslips precoated with a plant lectin concanavalin A (ConA). The periphery of the spread cells is especially thin and therefore is ideal for live cell microscopy. Detailed protocols for S2 cells culture, RNAi treatment, live light imaging and immunostaining can be found in the literature2,3. Our lab has adopted S2 cells as a model system to study microtubule-based cargo transport. Drosophila S2 cells treated with any drug that depolymerizes or severs F-actin, such as cytochalasin D (CytoD), grow long processes filled with uniformly polarized microtubules4-10, which makes it an ideal system to study cargo movement along microtubule arrays. Different parameters of transport in these processes (i.e. trajectories lengths, velocities, directionality etc.) can be easily measured using automated particle-tracking software, DiaTrack (Semasopht: http://www.microscopy.info/Organization/Details/4423; Dr. Pascal Vallotton: http://www.csiro.au/en/Organisation-Structure/Divisions/Mathematics-Informatics-and-Statistics/PascalVallotton.aspx#a4). These properties make S2 cells an appealing system for studying cargo transport along microtubules in tissue culture cells.
Pilot experiments in S2 cells can be extended to a physiologically important model of cargo transport studies in Drosophila primary neurons. Similar to S2 cells, neurons cultured from dissociated embryos are permeable to small molecules such as dyes and inhibitors, and high-resolution live imaging of organelle transport can be performed in neurons at room temperature. Using Drosophila with different genetic backgrounds, we can examine gene function in primary neurons by knockout (genetic loss-of-function mutations), knockdown (dsRNA injection or expression) or ectopic expression (transgenic flies). Specifically, fragmentation of actin filaments using CytoD treatment does not prevent neurite outgrowth; instead they grow faster, and thus we can study microtubule-dependent organelle transport in CytoD-treated neurons without the influence of actin filaments11. Therefore, primary neuronal cultures combine the advantages of tissue culture cells and fly genetics, which makes it a great system to study cargo transport in a physiological relevant system10,12. Since several familial neurodegenerative diseases could be caused by or associated with cargo transport defects (e.g.&#160;Parkinson&#39;s disease13, Huntington&#39;s disease14, Alzheimer&#39;s Disease15,16, Amyotrophic Lateral Sclerosis/ALS17, HMN7B and Perry syndrome18,19, and Spinocerebellar ataxia type 5/SCA520), primary neuronal cultures can be useful in analysis of the effects of specific mutations causing these diseases on microtubule-dependent transport.
Finally, important properties of microtubule-dependent transport are well conserved between mammals and flies, but Drosophila genetic and cell culture is less expensive and time-consuming, justifying the use of cultured Drosophila cells for studying essential aspects of organelle transport in normal and pathological conditions.

<table border="0" cellpadding="0" cellspacing="0" width="435">
	<colgroup>
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="61">
			<td height="61" style="height:61px;width:124px;">Powder Insect cell medium (Schneider&#8217;s)</td>
			<td style="width:133px;">Sigma&#160;</td>
			<td style="width:179px;">S9895</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Insect &#8211;Xpress medium</td>
			<td style="width:133px;">Fisher</td>
			<td style="width:179px;">BW12730Q</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:124px;">Insulin</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:179px;">I6634</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:124px;">Penicillin</td>
			<td style="width:133px;">Research Products International</td>
			<td style="width:179px;">P93000</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:124px;">Streptomycin</td>
			<td style="width:133px;">Research Products International</td>
			<td style="width:179px;">S62000</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Tetracycline hydrochloride</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:179px;">T7660-5G</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Fetal bovine serum</td>
			<td style="width:133px;">Atlanta Biologicals</td>
			<td style="width:179px;">S11150</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:124px;">Concanavalin A</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:179px;">C2010</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:124px;">Cytochalasin D</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">IC15077105</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">LysoTracker Red DND-99</td>
			<td style="width:133px;">Molecular Probes</td>
			<td style="width:179px;">L7528</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">100 &#181;m Cell strainer</td>
			<td style="width:133px;">Fisher Scientific</td>
			<td style="width:179px;">22363549</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">25 mm Circle cover glass</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">48380 080</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Drosophila Vials</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">89092-730</td>
		</tr>
		<tr height="81">
			<td height="81" style="height:81px;width:124px;">Disposable pellet pestles with matching microtubes</td>
			<td style="width:133px;">Kimble Chase</td>
			<td style="width:179px;">749520-0000</td>
		</tr>
	</tbody>
</table>

organelle transport in cultured drosophila cells: s2 cell line and primary neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.

Drosophila S2 cells attached on the ConA-coated coverslip spread into a flat round &#8220;pancake&#8221; shape (Figures 1A and 1C). However, when S2 cells plated on coverslips in the presence of CytoD and allowed to spread for 2-3 hr, they form long thin processes filled by parallel microtubules (Figures 1B and 1D). These microtubules have uniform polarity with plus-ends out7. Time-lapse fluorescence imaging of Drosophila S2 stably expressing mCherry-tubulin shows that processes are very dynamic, especially during the first 60 min after attachment (Figure 2). After 1 hr incubation, their growth slows down and 2-3 hr incubation ensures that majority of cells have fully grown processes. Here we show two representative examples of organelle transport along microtubules in S2 cells, lysosomes (labeled with LysoTracker, 200 nM; Figures 3A-B) and peroxisomes (labeled with peroxisome-targeted GFP, GFP-SKL)7 (stable transfection with pAC-GFP-SKL; Figures 3D-E). Time-lapse images of the cells were taken every 1 sec for 61 frames using a Nikon TU-2000 inverted microscope equipped with a Perfect Focus system (Nikon) and Coolsnap CCD camera (Roper Scientific) driven by Metamorph software. A 100-W halogen light source was used for fluorescence excitation to minimize photobleaching and phototoxicity. Each of the 61 frames in sequence was color coded according to the bar at upper right and images were superimposed to generate color-coded tracks. Thus moving particles generated rainbow tracks, while stationary particles appeared white10. ImageJ plug-in Temporal Color-Code (<a href="http://fiji.sc/Temporal-Color_Code">http://fiji.sc/Temporal-Color_Code</a>) was used for this processing. For quantitative analysis of organelle movement we used DiaTrack software (Semasopht Inc.), we only select organelles that are localized in processes because they can be easily tracked and the polarity of microtubule tracks is known. Figures 3C and 3F show typical distributions of retrograde and anterograde velocities obtained by lysosomes and peroxisomes tracking, respectively.
Similar techniques are used in our lab for analysis of organelle transport in primary cultured neurons. Neurons are a predominant cell type in mixed cultures obtained from stage 9-11 embryos. After overnight culturing they are easy to recognize by characteristic cell shape with neurites that are over 50 &#181;m long (Figure 4A). These cells are also positive for the pan-neuronal marker, Elav21 (Figures 4B and 4B&#39;). Processes in the neurons are filled with microtubules (Figures 4B and 4B&#34;). We can track organelle transport along the neurites using the techniques described above for S2 cells. Mitochondria can be labeled with mitochondrial GFP (Mito-GFP) under control of a motor neuron-specific promoter D4222 (Figures 5A and 5A&#39;), and peroxisomes can be marked by injection of DNA encoding a peroxisome-targeted mCherry7, into syncytial blastoderm stage embryos&#160;(Figures 5B and 5B&#39;).
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/50838/50838fig1.jpg" width="600px" /> 
Figure 1. Drosophila S2 process formation induced by CytoD.&#160;(A, C) Drosophila S2 cells attach and spread into flatten and round shapes when plated in ConA-precoated coverslips. (B, D) S2 cells plated in the presence of 2.5 &#181;m CytoD form long processes filled with microtubules after 3 hr incubation. A-B and C-D are transmitted light and fluorescent mCherry-tagged tubulin images, respectively. Scale bar, 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50838/50838fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/50838/50838fig2.jpg" width="600px" /> 
Figure 2. Time lapse of processes growth in a Drosophila S2 cell. Fluorescence time-lapse of a Drosophila S2 cell expressing mCherry-tubulin in the presence of 2.5 &#181;M CytoD. The numbers indicate the time after the cell attach to the coverslip. Scale bar, 5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50838/50838fig2highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/50838/50838fig3.jpg" width="600px" /> 
Figure 3. Organelle transport in Drosophila S2 cells. (A-C) Lysosome movement in Drosophila S2 cell. A representative 1 min-lysosome transport (labeled with 200 nM Lysotracker) in one cell (A, DIC image) represented by the artificial Temporal-Code (created by FIJI) (B). Velocities obtained from the analysis of lysosome movement tracked in processes (DiaTrack; 663 particles in 23 cells) (C). (D-F) Peroxisome movement in Drosophila S2 cell. A representative 1 min-peroxisome transport (labeled with pMT-GFP-SKL) in one cell (D, DIC image) represented by the artificial Temporal-Code (created by FIJI) (E). Velocities obtained from the analysis of peroxisome movement tracked in processes (DiaTrack; 228 particles in 20 cells) (F). Note that lysosomes move with longer and faster trajectories than peroxisomes. Scale bar, 5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50838/50838fig3highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/50838/50838fig4.jpg" width="600px" /> 
Figure 4. Primary Drosophila neurons from overnight culture. (A) An overnight cultured Drosophila neuron has characteristic neuronal morphology with long neurites. (B) The neuron expressed a pan-neuronal marker, Elav (red in B and white in B&#39;, DSHB, 7E8A10, 1:100) and with long neurites filled with microtubules (green in B and white in B&#39;&#39;, DM1&#945;, 1:1000). Scale Bar, 5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50838/50838fig4highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/50838/50838fig5.jpg" width="600px" /> 
Figure 5. Organelle transport in cultured Drosophila neurons. (A) Mitochondria in a cultured Drosophila neuron marked by Mito-GFP under control of a motor neuron-specific promoter, D42. (A&#39;) 5 min-mitochondrial movement in (A) is represented by the artificial Temporal-Color code (created by FIJI). (B) Peroxisomes in a cultured Drosophila neuron marked by pAC-SKL-mCherry, injected into early syncytial blastoderm stage embryos. (B&#39;) 2 min-peroxisome movement in (B) is represented by the artificial Temporal-Color code (created by FIJI). Scale bar, 5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50838/50838fig5highres.jpg" target="_blank">Click here to view larger image</a>.
<table border="0" cellpadding="0" cellspacing="0" width="955">
	<colgroup>
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr>
			<td>Organelle</td>
			<td>Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:207px;">Peroxisomes</td>
			<td>Peroxisomal targeting signal 1 tripeptide (SKL) tagged with a FP25</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:207px;">Mitochondria</td>
			<td>Mitochondrial targeting sequence of cytochrome c oxidase subunit VIII tagged with a FP26</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:207px;">Mitochondria</td>
			<td>Mitotracker (Dye which accumulates in active mitochondria)27</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:207px;">Lysosomes</td>
			<td>Lysotracker (Dye which accumulates in cellular compartments with low internal pH)28</td>
		</tr>
	</tbody>
</table>
Table 1. Most common organelle labeling strategies.
This table summarizes the most common organelle labeling strategies in tissue culture cells, including labeling of peroxisomes, mitochondria, and lysosomes.

Watch this Scientific Journal Video about Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons. at JoVE.com

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

organelle-transport-cultured-drosophila-cells-s2-cell-line-primary

Nanyang Technological University

Research

JoVE Journal

Biology

13.6K Views.  Feinberg School of Medicine, Northwestern University. Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Video: Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

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 ...

Isolation and Primary Culture of Rat Hepatic Cells

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology and other related subjects. The method based on two-step collagenase perfusion for isolation of intact hepatocytes was first introduced by Berry and Friend in 1969 1 and, since then, has undergone many modifications. The most commonly used technique was described by Seglenin 1976 2. Essentially, hepatocytes are dissociated from anesthetized adult rats by a non-recirculating collagenase perfusion through the portal vein. The isolated cells are then filtered through a 100 &mu;m pore size mesh nylon filter, and cultured onto plates. After 4-hour culture, the medium is replaced with serum-containing or serum-free medium, e.g. HepatoZYME-SFM, for additional time to culture. These procedures require surgical and sterile culture steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol, which allow consistently in the generation of viable hepatocytes in large numbers.

Primary hepatocytes provide a valuable tool to evaluate biochemical, molecular, and metabolic functions in a physiologically relevant experimental system. We describe a reliable protocol for rat in situ liver perfusion, which consistently generates viable hepatocytes up to 1.0 &times; 108 cells per preparation with cell viability between 88 ~ 96%.

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Generating a &#34;Humanized&#34; Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper distribution of the genetic material requires that sister kinetochores on every chromosome become bioriented by attaching to microtubules from opposite spindle poles before progressing into anaphase. However, erroneous, non-bioriented attachment states are common and cellular pathways exist to both detect and correct such attachments during cell division. The process by which improper kinetochore-microtubule interactions are destabilized is referred to as error correction. To study error correction in living cells, incorrect attachments are purposely generated via chemical inhibition of kinesin-5 motor, which leads to monopolar spindle assembly, and the transition from mal-orientation to biorientation is observed following drug washout. The large number of chromosomes in many model tissue culture cell types poses a challenge in observing individual error correction events. Drosophila S2 cells are better subjects for such studies as they possess as few as 4 pairs of chromosomes. However, small molecule kinesin-5 inhibitors are ineffective against Drosophila kinesin-5 (Klp61F). Here we describe how to build a Drosophila cell line that effectively replaces Klp61F with human kinesin-5, which renders the cells sensitive to pharmacological inhibition of the motor and suitable for use in the cell-based error correction assay.

Generating a "Humanized" Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

This protocol describes how to generate a Drosophila S2 cell line that is sensitive to small molecule inhibitors of kinesin-5. The use of these cells in a cell-based error correction assay is also outlined.

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper ...

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex ...

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory cells, EC cells sense luminal stimuli and convert them into serotonin (5-hyroxytryptamine, 5-HT) release events. However, the electrophysiology of these cells is poorly understood because they are difficult to culture and to identify. The method presented in this paper outlines primary EC cell cultures optimized for single cell electrophysiology. This protocol utilizes a transgenic cyan fluorescent protein (CFP) reporter to identify mouse EC cells in mixed primary cultures, advancing the approach to obtaining high-quality recordings of whole cell electrophysiology in voltage- and current-clamp modes.

Enterochromaffin (EC) cells comprise a small subset of gastrointestinal epithelial cells. EC cells are electrically excitable and release serotonin, yet difficulties in culturing and identifying EC cells have limited physiological studies. The method presented here establishes a primary culture model amenable to examination of single EC cells by electrophysiology.

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory ...