We thank the Medical Research Council UK for core funding (U105115237) to RRK.

1. Preparation of Cells and Materials
<ol>
	<li>Cultivate cells either on SM agar plates in conjunction with bacteria such as Klebsiella aerogenes, or in nutrient medium such as HL5 as described23. 
	NOTE: Growing knockout mutants that are defective in macropinocytosis on bacteria helps prevent accumulation of secondary mutations that may increase the rate of macropinocytosis. Keep the passage number below 4 after plating cells from stocks for optimum results. Mutants should be stored as frozen stocks as soon as possible after isolation.</li>
	<li>To cultivate cells on SM agar plates, grow K. aerogenes to confluence in SM medium. Add 200&#8211;300 &#956;L of bacteria onto an SM agar plate and spread. Take a sterile loop of cells (when transferring from another bacterial plate) and spread at one edge of the plate. Incubate at 22 &#176;C for up to a week.</li>
	<li>Dissolve Tetra-methyl-rhodamine isothiocyanate (TRITC-) dextran to 50 mg/mL in water, filter using 0.22 &#956;m filters into 1 mL aliquots and store at -20 &#176;C. Aliquots can be kept indefinitely. 
	NOTE: 155 kDa is the typical size used with this method, as it can be bought cheaply in bulk, but smaller dextrans measure macropinocytosis just as effectively in Dictyostelium24. Different non-quenchable dextrans, such as cascade blue or Alexa-647, are alternatives if different fluorophores are required.</li>
</ol>
2. Converting Qualitative Measurements into Quantitative (Optional)
<ol>
	<li>Perform a fluid uptake assay using cells in suspension.
	<ol>
		<li>Pellet axenically growing cells at 300 x g for 3 min, discard the supernatant and resuspend in nutrient medium to 1x107 cells/mL. Add TRITC-dextran to 0.5 mg/mL and shake at 180 rpm, 22 &#176;C.</li>
		<li>Dilute 0.8 mL samples into 0.7 mL of ice-cold KK2 at 0, 30, 60, 90, 120 min. Wash once in 1.5 mL of ice-cold KK2 buffer23 in a benchtop centrifuge, resuspend to 1 mL in the same buffer and store on ice. 
		NOTE: washing immediately or once all samples have been collected yields equivalent results.</li>
	</ol>
	</li>
	<li>Determine the volume of fluid internalized by the cells.
	<ol>
		<li>Set the fluorimeter to measure fluorescence using an excitation wavelength of 544 nm and emission wavelength of 574 nm with a 10 nm slit. Set up a dilution series of TRITC-dextran and use it to measure the fluorescence for given volumes of dextran. Create a calibration curve.</li>
		<li>Measure the fluorescence internalized by the cells in section 2.1 using 0.9 mL of the cell suspension. Calculate the volume of fluid internalized per cell using the calibration curve24.</li>
	</ol>
	</li>
	<li>Dilute the remainder of the cells from each time point separately into 0.5 mL ice cold KK2. Filter through a 70 &#181;m cell strainer into 5 mL polystyrene flow cytometry tubes. Set the flow cytometer so that cells from each time point are distinct from each other and record their fluorescence (see sections 3.4 and 3.5).</li>
	<li>Measure the fluorescence of calibration beads in the flow cytometer using the settings above and record it. 
	NOTE: This is the reference for all future fluorescence measurements on the flow cytometer: before using again run the beads and adjust the settings so they have the same fluorescence. This will give a very narrow defined peak. Putting a gate around the fluorescence peak can make this easier.</li>
	<li>Repeat three times and plot the cell fluorescence data obtained in sections 2.2 and 2.3 against each other. Plot a line of best fit and use the equation to convert the qualitative data from the flow cytometry into quantitative units.</li>
</ol>
3. Measuring Fluid Uptake
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58434/58434fig1.jpg" /> 
Figure 1: Schematic of high-throughput measurement of macropinocytosis. (A) Grow Dictyostelium on an SM plate seeded with K. aerogenes bacteria (yellow). Harvest cells from the feeding front (orange), avoiding cells that are already developed (green), into 25 mL of KK2 buffer. Vortex to dissociate, pellet by 3 min centrifugation at 300 x g, then wash 3 times in 50 mL of KK2 buffer, discarding the supernatant each time. Resuspend to 1 x 105 cells/mL in HL5 growth medium and add 50 &#181;L into three wells per sample of a flat bottom 96-well plate. Incubate at 22 &#176;C for 24 h. (B) Dilute TRITC-dextran to 1 mg/mL in HL5 growth medium from a stock solution of 50 mg/mL. Add 50 &#181;L to each sample (excluding the 0 min uptake control wells), and incubate at 22 &#176;C for 1 h, after which add the dextran to the 0 min uptake control. (C) Immediately decant the media, pat the plate on a tissue to remove excess medium and submerge in a bath of ice-cold KK2 buffer, filling the wells to wash. Decant the buffer and pat dry again. Add 100 &#181;L of 5 mM sodium azide dissolved in KK2MC to detach the cells. Take to flow cytometer for measurement of internalized fluorescence. <a href="https://www.jove.com/files/ftp_upload/58434/58434fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Set up a flat-bottomed 96-well tissue-culture plate (Figure 1A). Use cells grown on bacteria and to transfer them into HL5 medium (containing 100 &#181;g/mL of dihydrostreptomycin, 100 &#181;g/mL ampicillin and 50 &#181;g/mL kanamycin) 24 h before the assay; this allows macropinocytosis to be upregulated from the low levels seen in cells grown on bacteria24. Alternatively, dilute cells directly from axenic culture, incubating for 24 h before the assay, to reduce errors due to the cells being diluted from different densities.
	<ol>
		<li>Harvest cells from the feeding front into 25 mL of KK2 in a 50 mL centrifuge tube. Dissociate cells by vortexing and pellet at 300 x g for 3 min. Discard the supernatant, resuspend the pellet and wash 3 times at 300 x g for 3 min in 50 mL of KK2, discarding the supernatant each time to remove the bacteria.</li>
		<li>Determine the cell density (using a hemacytometer or other cell counting system) and dilute into HL5 containing antibiotics to 1 x 105 cells/mL. Pipette 50 &#956;L into each well, using three wells for each condition. Incubate at 22 &#176;C for 24 h. Remember to set up a 0 min uptake control. 
		NOTE: An alternative medium, e.g. SIH, VL6 can be used instead.</li>
	</ol>
	</li>
	<li>Load cells with TRITC-dextran (Figure 1B).
	<ol>
		<li>Dilute the dextran to 1 mg/mL in the medium used (this can be increased to 2.5&#8211;5 mg/mL when assessing cells with very low uptake). Add 50 &#956;L to each well (giving a final concentration of 0.5 mg/mL TRITC-dextran) and return the plate to 22 &#176;C for 1 h, as this allows significant dextran accumulation but exocytosis of dextran has not yet begun. 
		NOTE: A repeater pipette allows this step to be done more rapidly, reducing error.</li>
	</ol>
	</li>
	<li>Prepare cells for flow cytometry (Figure 1C).
	<ol>
		<li>Immediately prior to washing, add 50 &#956;L dextran-containing media to the 0 min uptake controls.</li>
		<li>Decant the medium and pat dry on a tissue. Wash by submerging the plate in ice-cold KK2, then decant. 
		NOTE: some strains with adherence defects may become detached during this step, e.g. a knockout of both homologs of Talin (talA-/talB-)25. Use caution when working with cell lines that attach poorly and aspirate media if required.</li>
		<li>Add 100 &#956;L of ice-cold 5 mM sodium azide dissolved in KK2MC (KK2 + 2 mM MgSO4 and 100 &#956;M CaCl2). 
		CAUTION: Sodium azide is extremely toxic. Use a dust mask and safety glasses when working with the powder. Always wear gloves and lab coat. Avoid heat and do not mix with acid. 
		NOTE: Cells rapidly detach, and exocytosis is prevented24. A microscope can be used to check this.</li>
	</ol>
	</li>
	<li>Measure fluid uptake by flow cytometry
	<ol>
		<li>If planning to convert relative values to absolute ones, use beads to standardize flow cytometer settings (see section 2). Alternatively, use cells loaded with dextran as in section 2 to ensure that the forward scatter and side scatter are set appropriately to isolate cells (Figure 2A), ensuring the machine is not blocked, (Figure 2B) and adjust the parameters to measure the internalized fluorescence (Figure 2C).</li>
		<li>Attach high-throughput sampling system and add plate. Set up a protocol to measure the fluorescence (for TRITC-dextran use a yellow-green laser to excite and measure in the 582 nm channel or similar) of up to 65 &#956;L of the cell suspension and run.</li>
	</ol>
	</li>
	<li>Analyze flow cytometry results
	<ol>
		<li>Gate on the cells using forward scatter and side scatter (Figure 2A). Calculate the median fluorescence of the cells using the statistics option. Set these parameters using one sample and apply to all samples. 
		NOTE: The forward scatter/side scatter profiles can vary between samples when using different mutants or inhibitors.</li>
		<li>Determine the mean of the three wells for each condition and subtract the 0 min uptake control. Either convert to quantitative values using the equation determined in section 2 or normalize to the control.</li>
	</ol>
	</li>
</ol>
4. Performing Fluid Uptake Time-courses
<ol>
	<li>Set up cells as in section 3, with three wells for each time point and strain/condition.</li>
	<li>Add dextran labelled medium to the different wells sequentially. A typical time-course measures fluid uptake at 0, 30, 60, 90, 120 and 180 min. Add dextran-containing medium for the 180 min time point wells, followed 60 min later by adding it to the wells for the 120 min time point, etc. (Figure 3A).</li>
	<li>At 0 min, add 50 &#956;L of dextran-containing medium and immediately decant and wash the plate as in section 3.3. Analyze as described in section 3.5.</li>
</ol>
5. Dose Response Curves
<ol>
	<li>Set up cells as described in section 3, with three wells for each strain and condition.</li>
	<li>Dilute the compound of interest into medium containing 1 mg/mL dextran at double the desired maximum final concentration of the compound. Prepare a second tube containing medium with dextran with the same proportion of vehicle as the first. Vortex to mix.</li>
	<li>Create a dilution series of the compound of interest in 200 &#956;L dextran containing medium per sample (Figure 4A). Vortex to mix. Add 50 &#956;L of the medium to each sample well for 1 h.</li>
	<li>Wash and analyze as described in section 3.3 onwards.</li>
</ol>
6. Phagocytosis and Membrane Uptake
<ol>
	<li>To measure membrane uptake, prepare the cells as described in section 3. Add 50 &#956;L of medium containing 20 &#956;M FM 1-43 for a final concentration of 10 &#956;M. After loading, wash and prepare cells for flow cytometry as in section 3.3. Measure in the 585 nm channel or similar, using a blue laser to excite. 
	NOTE: As membrane trafficking is more rapid than fluid-phase, 10 min is a more appropriate time point to use than 1 h26. Alternative dyes that fluoresce only when incorporated into the membrane can be used instead.</li>
	<li>Use either fluorescently labelled beads or bacteria to measure phagocytosis. Fluorescent yeast cannot be used, as they are inseparable from Dictyostelium by forward and side scatter24. Set up cells as described in section 3 and analyze as described27.
	<ol>
		<li>To measure phagocytosis of beads add yellow-green labelled beads as described in section 3.2 to a final concentration of 5 x 107 beads/mL (1.75 and 2 &#956;m) or 1 x 108 beads/mL (1 and 1.5 &#956;m) for 1 h before washing and preparing for flow cytometry as in section 3.3. Measure in the 525 nm channel or similar, using a blue laser to excite. 
		Note: Higher concentrations will lead to cell detachment.</li>
		<li>To measure phagocytosis of bacteria, add Texas-Red labelled Escherichia coli bioparticles as described in section 3.2 to 1 x 108 bacteria/mL for 1 h before washing and preparing for flow cytometry as in section 3.3. Measure in the 610 nm channel or similar, using a yellow-green laser to excite.</li>
	</ol>
	</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dictyostelium+myosin+I+double+mutants+exhibit+conditional+defects+in+pinocytosis.">Dictyostelium myosin I double mutants exhibit conditional defects in pinocytosis.</a> Journal of Cell Biology. 131, 1205-1221 (1995).</li><li>Mercer, J., Helenius, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vaccinia+virus+uses+macropinocytosis+and+apoptotic+mimicry+to+enter+host+cells.">Vaccinia virus uses macropinocytosis and apoptotic mimicry to enter host cells.</a> Science. 320 (5875), 531-535 (2008).</li><li>Paschke, P., 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=Rapid+and+efficient+genetic+engineering+of+both+wild+type+and+axenic+strains+of+Dictyostelium+discoideum.">Rapid and efficient genetic engineering of both wild type and axenic strains of Dictyostelium discoideum.</a> PLoS One. 13 (5), e0196809 (2018).</li><li>Bloomfield, G., Tanaka, Y., Skelton, J., Ivens, A., Kay, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Widespread+duplications+in+the+genomes+of+laboratory+stocks+of+Dictyostelium+discoideum.">Widespread duplications in the genomes of laboratory stocks of Dictyostelium discoideum.</a> Genome Biology. 9 (4), R75 (2008).</li><li>Fey, P., Dodson, R. J., Basu, S., Chisholm, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=One+stop+shop+for+everything+Dictyostelium:+dictyBase+and+the+Dicty+Stock+Center+in+2012.">One stop shop for everything Dictyostelium: dictyBase and the Dicty Stock Center in 2012.</a> Methods in Molecular Biology. 983, 59-92 (2013).</li></ol>

The authors have no conflicts of interest to disclose.

Whereas other methods to assess fluid uptake are low throughput, washing the cells in situ and the use of sodium azide to detach cells are the critical steps in this method, which allow high-throughput measurement of macropinocytosis, membrane uptake, or phagocytosis by Dictyostelium. As the cells are attached to a surface and the medium is not, they can be left attached while the medium around them is first thrown off and then changed by immersion in buffer and thrown off again. Sodium azide, which depletes cellular ATP and depolarizes the membrane30, is then used to detach the cells, and also prevents exocytosis without affecting cell viability24.
While using flow cytometry to measure macropinocytosis by Dictyostelium gives a very accurate measurement of fluid uptake very quickly, to establish the reason why a particular strain or condition has altered fluid uptake, further investigation using microscopy is required24. It should also be noted that previously published results have, in some cases, shown a difference in fluid uptake by mutant strains grown either on a surface (as in this case), or in shaking suspension (as in the standard protocol)31. Using this method may mean that, in rare cases, apparent fluid uptake defects are missed. Additionally, when measuring phagocytosis, only low concentrations of particles can be used. The maximum rate of phagocytosis that can be determined with this technique is far below the real maximum, although it is still possible to measure relevant differences in phagocytosis between strains and conditions24. To determine the maximum rate of phagocytosis, uptake must be measured in shaking suspension by an alternative protocol27. Cells that have phagocytosed beads have increased side scatter, so this should be corrected for accordingly when setting up the flow cytometer.
Flow cytometry can be used to measure fluid uptake in mammalian cells32, however the higher proportion of fluid phase uptake by other endocytic pathways than seen in Dictyostelium is a concern. In addition, cells are typically detached using trypsin at 37 &#176;C, allowing further endocytic progression of internalized dextran. Ice-cold sodium azide does not cause macrophages to detach from a surface (Williams, unpublished observation), making this technique not applicable to mammalian cells without further optimization.
High throughput measurement of macropinocytosis has the potential to be used to screen quickly and cheaply for the effects of inhibitors, genetic mutation or gene knockdown on Dictyostelium cells. Mutants should always be compared to their direct parent only. If the reader has no prior preference for Dictyostelium strain, non-axenic strains such as DdB or NC4 are more &#34;wild-type&#34; than axenic ones and can be manipulated as effectively as axenic strains33. Otherwise, Ax2 strains are the axenic strains with the fewest genome duplications34, while many strains of Ax4 are Talin A knockouts and should be avoided if possible23. Most previously published strains can be ordered from the Dicty Stock Center35.
This technique allows greater investigative possibilities than was previously possible into the effects of different conditions, inhibitors and mutations on macropinocytosis by Dictyostelium.

Large-scale non-specific fluid uptake by macropinocytosis is important in several biological contexts1, including antigen sampling by immune cells2, pathogen entry into host cells3, cancer cell proliferation4 and the spreading of prion diseases5. In mammalian and Dictyostelium cells, actin6,7, PI(3,4,5)P38,9,10 (although the exact nature of the lipid differs between the two11), activated Ras12,13, and activated Rac14,15 are important for efficient fluid uptake by macropinocytosis, although there remain many unanswered questions about how the macropinocytic patch is formed, organized and eventually internalized. Discovering more proteins important for macropinocytosis, and subsequent determination of how they are important in the various biological contexts, will give a more comprehensive understanding of macropinocytosis and potentially allow development of targeted treatments for a range of conditions.
Dictyostelium is an ideal model system for studying macropinocytosis. The high level of constitutive macropinocytosis in standard laboratory strains means that fluid uptake is over 90% due to macropinocytosis6. This allows macropinocytosis to be measured solely by determining fluid uptake, unlike mammalian cells where the proportion of fluid uptake due to macropinocytosis is much lower. That macropinocytosis is so well defined and easily visualized12 in this system similarly offers distinct advantages for investigating core conserved components of the macropinosome over other systems where there may be multiple regulatory signals16,17.
The standard technique used to measure macropinocytosis by mammalian cells involves fixing cells after pulsing with dextran for a short period of time followed by microscopy to determine the area of a cell that is occupied by dextran-positive vesicles18. This technique does not however account for the possibility of macropinosomes shrinking upon entering the cell, which has been reported in Dictyostelium19, and only takes into account single planes of the cell, meaning the volume internalized is unclear. An alternative technique, of counting the number of macropinosomes internalized in a given time, has the same downsides20. Using Dictyostelium avoids these issues; however, existing techniques for measuring fluid uptake by Dictyostelium are relatively labor intensive, using a large amount of both cells and dextran21. Cells are shaken at high density in fluorescent dextran and samples removed at various time-points for determination of the internalized fluorescence using a fluorimeter. Cells prepared this way can be analyzed by flow cytometry to gain single cell, rather than population-level, resolution22, although this remains low-throughput.
Here, the standard technique to measure internalized fluid using fluorescent dextran as a label is adapted to a 96-well plate format, with the samples analyzed by flow cytometry using a high-throughput sampling (HTS) attachment. Cells are fed non-quenchable fluorescent dextran for a pre-determined length of time, washed by immersion in ice-cold buffer and detached using 5 mM sodium azide, which also stops exocytosis. Cells in each well are then analyzed by flow cytometry. This method was designed to overcome the limitations of the above methods and allow simultaneous comparison of the fluid uptake of large numbers of strains/conditions while using fewer resources and reducing the labor involved.

<table>
	<tbody>
		<tr>
			<td>LSR_II flow cytometer</td>
			<td>BD Biosciences</td>
			<td>-</td>
			<td>Other Flow cytometers can also do this role, e.g. the LSRFortessa by BD</td>
		</tr>
		<tr>
			<td>TRITC-dextran (155 kDa)</td>
			<td>Sigma-Aldrich</td>
			<td>T1287</td>
			<td>Other non-quenchable dextrans, and other sizes are also fine</td>
		</tr>
		<tr>
			<td>HL5 medium</td>
			<td>Formedium</td>
			<td>HLGCFG</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well tissue culture plate</td>
			<td>Corning</td>
			<td>3596</td>
			<td>Any flat-bottom tissue culture treated 96-well plate will work</td>
		</tr>
		<tr>
			<td>Dihydrostreptomycin sulfate</td>
			<td>Sigma-Aldrich</td>
			<td>PHR1517</td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin sodium</td>
			<td>Formdium</td>
			<td>AMP50</td>
			<td></td>
		</tr>
		<tr>
			<td>Kanamycin monosulfate</td>
			<td>Sigma-Aldrich</td>
			<td>60615</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>VWR</td>
			<td>103694M</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate hydrate</td>
			<td>VWR</td>
			<td>25169.295</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>VWR</td>
			<td>1.02382.0250</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium dihydrogen phopshate</td>
			<td>VWR</td>
			<td>1.04877.1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Di-potassium hydrogen phosphate</td>
			<td>VWR</td>
			<td>1.05104.1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorimeter</td>
			<td>Perkin-Elmer</td>
			<td>LS 50 B</td>
			<td></td>
		</tr>
		<tr>
			<td>FM1-43</td>
			<td>Thermofisher</td>
			<td>T35356</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoresbrite YG-carboxy microspheres 1.00 &#181;m</td>
			<td>Polysciences</td>
			<td>15702-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoresbrite YG-carboxy microspheres 1.50 &#181;m</td>
			<td>Polysciences</td>
			<td>09719-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoresbrite YG-carboxy microspheres 1.75 &#181;m</td>
			<td>Polysciences</td>
			<td>17687-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoresbrite YG-carboxy microspheres 2.00 &#181;m</td>
			<td>Polysciences</td>
			<td>09847-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Texas Red E. coli bioparticles</td>
			<td>Thermofisher</td>
			<td>E2863</td>
			<td></td>
		</tr>
		<tr>
			<td>Flow-set fluorospheres</td>
			<td>Beckman Coulter</td>
			<td>6607007</td>
			<td>Calibration Beads</td>
		</tr>
		<tr>
			<td>SM agar</td>
			<td>Formedium</td>
			<td>SMACFG</td>
			<td></td>
		</tr>
		<tr>
			<td>0.22 &#181;m syringe filter</td>
			<td>Elkay Laboratory Products</td>
			<td>E25-PS22-50S</td>
			<td></td>
		</tr>
		<tr>
			<td>10 mL Syringe</td>
			<td>Becton Dickinson</td>
			<td>302188</td>
			<td></td>
		</tr>
		<tr>
			<td>Round-bottom polystyrene tubes</td>
			<td>Corning</td>
			<td>352058</td>
			<td>Use a tube that will fit onto your flow cytometer.</td>
		</tr>
		<tr>
			<td>70 &#181;m cell strainer</td>
			<td>Falcon</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL centrifuge tube</td>
			<td>Sarstedt</td>
			<td>62.547.004</td>
			<td></td>
		</tr>
		<tr>
			<td>Repeating pipette</td>
			<td>Eppendorf</td>
			<td>M4-SK</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL repeating pipette tips</td>
			<td>Eppendorf</td>
			<td>30089650</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma-Aldrich</td>
			<td>D2650-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>LY294002</td>
			<td>Cayman Chemical Company</td>
			<td>70920</td>
			<td></td>
		</tr>
		<tr>
			<td>Nocodazole</td>
			<td>Sigma-Aldrich</td>
			<td>M1404-2MG</td>
			<td></td>
		</tr>
	</tbody>
</table>

high-throughput measurement of dictyostelium discoideum macropinocytosis by flow cytometry

Large-scale non-specific fluid uptake by macropinocytosis is important for the proliferation of certain cancer cells, antigen sampling, host cell invasion and the spread of neurodegenerative diseases. The commonly used laboratory strains of the amoeba Dictyostelium discoideum have extremely high fluid uptake rates when grown in nutrient medium, over 90% of which is due to macropinocytosis. In addition, many of the known core components of mammalian macropinocytosis are also present, making it an excellent model system for studying macropinocytosis. Here, the standard technique to measure internalized fluid using fluorescent dextran as a label is adapted to a 96-well plate format, with the samples analyzed by flow cytometry using a high-throughput sampling (HTS) attachment.
Cells are fed non-quenchable fluorescent dextran for a pre-determined length of time, washed by immersion in ice-cold buffer and detached using 5 mM sodium azide, which also stops exocytosis. Cells in each well are then analyzed by flow cytometry. The method can also be adapted to measure membrane uptake and phagocytosis of fluorescent beads or bacteria.
This method was designed to allow measurement of fluid uptake by Dictyostelium in a high-throughput, labor and resource efficient manner. It allows simultaneous comparison of multiple strains (e.g. knockout mutants of a gene) and conditions (e.g. cells in different media or treated with different concentrations of inhibitor) in parallel and simplifies time-courses.

Once the technique has been performed and cells are loaded with dextran and ready for analysis (Figure 1), ensure the flow cytometer is not blocked and adjust the forward scatter/side scatter profile to look like the cells shown in Figure 2A. If the machine is blocked, it will look more like that shown in Figure 2B and must be unblocked before continuing. Ensure the parameters show control axenic cells have high internalized dextran fluorescence at longer time points and low internalized fluorescence at shorter ones (Figure 2C).
When looking for differences between mutants, it is likely that there will be one of three phenotypes. The mutants could have normal fluid uptake, they could have a partial defect, or fluid uptake could be completely abolished. Figure 2D shows a strain with normal fluid uptake, in this case the standard laboratory strain Ax2, a mutant with a ~50% decrease in fluid uptake (Ax2 rasG-14) and one with abolished fluid uptake (Ax3 gefB-28). Obtain the average median fluid uptake (section 3.5) and use it to either calculate the volume of fluid internalized (as in Figure 3B) or compare the data to a control (as in Figure 4B and 4C).
When performing a fluid uptake time course, as in Figure 3A, the internalized fluorescence should increase for 60&#8211;90 min, after which the dextran begins to be exocytosed and a plateau is reached (Figure 3B). Using 60 min as the standard time point when comparing macropinocytosis in different mutants/conditions therefore allows a good signal to be achieved, and no signal is lost due to exocytosis. Mutants where exocytosis is severely obstructed may take longer to reach a plateau29.
When treating cells with inhibitors that are effective against macropinocytosis in Dictyostelium (set up as in Figure 4A), the dextran internalized in 1 h will go down to almost nothing at higher inhibitor concentrations in the majority of cases (Figure 4B). Some inhibitors may not be 100% effective, however, e.g. nocodazole only inhibits up to 50% of fluid uptake by macropinocytosis when added acutely (Figure 4C). If the inhibitors are not effective, the cells will internalize a similar amount of dextran as the control, and a decrease in fluorescence will not be seen. This technique allows a large range of different inhibitors and inhibitor concentrations to be screened for effects on fluid uptake by macropinocytosis very quickly, reducing the time spent optimizing the inhibitor treatment.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58434/58434fig2.jpg" /> 
Figure 2: Set up of flow cytometer and representative data. (A) The forward scatter (FSC) and side scatter (SSC) profiles of cells should be set so the cells can be easily distinguished. An example of how Ax2 cells should look is shown. (B) If the flow cytometer is blocked, as in this example, the cells have very low side scatter. The laser will not excite the fluorophores properly and the machine should be unblocked before continuing. Any data obtained while the machine was blocked should be discarded. (C) The fluorescence should be set so that a 0 min uptake sample has low fluorescence, which increases when cells have been incubated for longer in the fluorescent medium, as shown in this example taken from Williams &#38; Kay 201824. (D) Examples of cells that have been incubated with TRITC dextran for 1 h with normal macropinocytosis (Ax2, green), reduced macropinocytosis (Ax2 rasG-, HM172614, orange) and abolished macropinocytosis (Ax3 gefB-, HM177628, blue). <a href="https://www.jove.com/files/ftp_upload/58434/58434fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58434/58434fig3.jpg" /> 
Figure 3: Performing fluid uptake time-courses in 96-well plates. (A) Dextran should be added to each set of samples sequentially, with the same finish time. Then wash the wells, detach and measure the internalized fluorescence by flow cytometry. Example times to add the dextran are shown here. (B) Fluid uptake time-course of Ax2 cells performed in 96-well plates. Taken from Williams &#38; Kay 201824, error bars show the standard error of three independent experiments. <a href="https://www.jove.com/files/ftp_upload/58434/58434fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58434/58434fig4.jpg" /> 
Figure 4: Fluid uptake dose response curves. (A) Add the compound of interest, in this case the PI3K inhibitor LY294002, to HL5 containing 1 mg/mL TRITC-dextran at double the desired final maximum concentration. Mix with HL5 growth medium + 1 mg/mL dextran containing vehicle alone in various proportions to generate a dilution series of 200 &#181;L medium per condition. Add to wells as normal for 1 h before washing and measuring internalized fluorescence. (B) Fluid uptake dose response curve for Ax2 cells incubated with the LY294002-containing medium from A. Adapted from Williams &#38; Kay 201824. Fluid uptake is normalized to an untreated control. Error bars show the standard error of three independent experiments. (C) Fluid uptake dose response curve for Ax2 cells incubated with the nocodazole. Adapted from Williams &#38; Kay 201824. Fluid uptake is normalized to an untreated control. Error bars show the standard error of three independent experiments. <a href="https://www.jove.com/files/ftp_upload/58434/58434fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry at JoVE.com

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

Macropinocytosis, large-scale non-specific fluid uptake, is important in many areas of clinical biology including immunology, infection, cancer, and neurodegenerative diseases. Here, existing techniques have been adapted to allow high-throughput, single-cell resolution measurement of macropinocytosis in the macropinocytosis model organism Dictyostelium discoideum using flow cytometry.

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

Large-scale non-specific fluid uptake by macropinocytosis is important for the proliferation of certain cancer cells, antigen sampling, host cell invasion ...

high-throughput-measurement-dictyostelium-discoideum-macropinocytosis

Nanyang Technological University

Research

JoVE Journal

Biology

7.7K Views.  MRC-Laboratory of Molecular Biology, Cambridge, UK. Macropinocytosis, large-scale non-specific fluid uptake, is important in many areas of clinical biology including immunology, infection, cancer, and neurodegenerative diseases. Here, existing techniques have been adapted to allow high-throughput, single-cell resolution measurement of macropinocytosis in the macropinocytosis model organism Dictyostelium discoideum using flow cytometry.

Video: High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

IP-FCM:  Immunoprecipitation Detected by Flow Cytometry

Immunoprecipitation detected by flow cytometry (IP-FCM) is an efficient method for detecting and quantifying protein-protein interactions. The basic principle extends that of sandwich ELISA, wherein the captured primary analyte can be detected together with other molecules physically associated within multiprotein complexes. The procedure involves covalent coupling of polystyrene latex microbeads with immunoprecipitating monoclonal antibodies (mAb) specific for a protein of interest, incubating these beads with cell lysates, probing captured protein complexes with fluorochrome-conjugated probes, and analyzing bead-associated fluorescence by flow cytometry. IP-FCM is extremely sensitive, allows analysis of proteins in their native (non-denatured) state, and is amenable to either semi-quantitative or quantitative analysis. As additional advantages, IP-FCM requires no genetic engineering or specialized equipment, other than a flow cytometer, and it can be readily adapted for high-throughput applications.

The IP-FCM method is presented, which allows a sensitive, robust, biochemical assessment of native protein-protein interactions, without requiring genetic engineering or large sample sizes.

Immunoprecipitation detected by flow cytometry (IP-FCM) is an efficient method for detecting and quantifying protein-protein interactions. The basic ...

Quantitative Measurement of GLUT4 Translocation to the Plasma Membrane by Flow Cytometry

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces glucose uptake by insulin-sensitive tissues, most notably the brain, skeletal muscle, and adipocytes. Patients suffering from type-2 diabetes and/or obesity often develop insulin resistance and are unable to control their glucose homeostasis. New insights into the mechanisms of insulin resistance may provide new treatment strategies for type-2 diabetes.
The GLUT family of glucose transporters consists of thirteen members distributed on different tissues throughout the body1. Glucose transporter type 4 (GLUT4) is the major transporter that mediates glucose uptake by insulin sensitive tissues, such as the skeletal muscle. Upon binding of insulin to its receptor, vesicles containing GLUT4 translocate from the cytoplasm to the plasma membrane, inducing glucose uptake. Reduced GLUT4 translocation is one of the causes of insulin resistance in type-2 diabetes2,3.
The translocation of GLUT4 from the cytoplasm to the plasma membrane can be visualized by immunocytochemistry, using fluorophore-conjugated GLUT4-specific antibodies.
Here, we describe a technique to quantify total amounts of GLUT4 translocation to the plasma membrane of cells during a chosen duration, using flow cytometry. This protocol is rapid (less than 4 hours, including incubation with insulin) and allows the analysis of as few as 3,000 cells or as many as 1 million cells per condition in a single experiment. It relies on anti-GLUT4 antibodies directed to an external epitope of the transporter that bind to it as soon as it is exposed to the extracellular medium after translocation to the plasma membrane.

This protocol describes a rapid technique to quantify the translocation of GLUT4 from the cytoplasm to the plasma membrane of cells by flow cytometry.

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces ...

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

High-Throughput Measurement and Classification of Organic P in Environmental Samples

Many types of organic phosphorus (P) molecules exist in environmental samples1. Traditional P measurements do not detect these organic P compounds since they do not react with colorimetric reagents2,3. Enzymatic hydrolysis (EH) is an emerging method for accurately characterizing organic P forms in environmental samples4,5. This method is only trumped in accuracy by Phosphorus-31 Nuclear Magnetic Resonance Spectroscopy (31P-NMR) -a method that is expensive and requires specialized technical training6. We have adapted an enzymatic hydrolysis method capable of measuring three classes of phosphorus (monoester P, diester P and inorganic P) to a microplate reader system7. This method provides researchers with a fast, accurate, affordable and user-friendly means to measure P species in soils, sediments, manures and, if concentrated, aquatic samples. This is the only high-throughput method for measuring the forms and enzyme-lability of organic P that can be performed in a standard laboratory. The resulting data provides insight to scientists studying system nutrient content and eutrophication potential.

This protocol describes a high-throughput method of enzymatic hydrolysis that utilizes a microplate reader to measure and classify soil phosphorus as P monoesters, P diesters and inorganic P. Up to 96 samples can be measured at one time in a standard laboratory.

Many types of organic phosphorus (P) molecules exist in environmental samples1.  Traditional P measurements do not detect these organic P compounds since ...

Imaging G-protein Coupled Receptor (GPCR)-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

Many eukaryotic cells can detect gradients of chemical signals in their environments and migrate accordingly 1. This guided cell migration is referred as chemotaxis, which is essential for various cells to carry out their functions such as trafficking of immune cells and patterning of neuronal cells 2, 3. A large family of G-protein coupled receptors (GPCRs) detects variable small peptides, known as chemokines, to direct cell migration in vivo 4. The final goal of chemotaxis research is to understand how a GPCR machinery senses chemokine gradients and controls signaling events leading to chemotaxis. To this end, we use imaging techniques to monitor, in real time, spatiotemporal concentrations of chemoattractants, cell movement in a gradient of chemoattractant, GPCR mediated activation of heterotrimeric G-protein, and intracellular signaling events involved in chemotaxis of eukaryotic cells 5-8. The simple eukaryotic organism, Dictyostelium discoideum, displays chemotaxic behaviors that are similar to those of leukocytes, and D. discoideum is a key model system for studying eukaryotic chemotaxis. As free-living amoebae, D. discoideum cells divide in rich medium. Upon starvation, cells enter a developmental program in which they aggregate through cAMP-mediated chemotaxis to form multicullular structures. Many components involved in chemotaxis to cAMP have been identified in D. discoideum. The binding of cAMP to a GPCR (cAR1) induces dissociation of heterotrimeric G-proteins into G&gamma; and G&beta;&gamma; subunits 7, 9, 10. G&beta;&gamma; subunits activate Ras, which in turn activates PI3K, converting PIP2 into PIP3 on the cell membrane 11-13. PIP3 serve as binding sites for proteins with pleckstrin Homology (PH) domains, thus recruiting these proteins to the membrane 14, 15. Activation of cAR1 receptors also controls the membrane associations of PTEN, which dephosphorylates PIP3 to PIP2 16, 17. The molecular mechanisms are evolutionarily conserved in chemokine GPCR-mediated chemotaxis of human cells such as neutrophils 18. We present following methods for studying chemotaxis of D. discoideum cells. 1. Preparation of chemotactic component cells. 2. Imaging chemotaxis of cells in a cAMP gradient. 3. Monitoring a GPCR induced activation of heterotrimeric G-protein in single live cells. 4. Imaging chemoattractant-triggered dynamic PIP3 responses in single live cells in real time. Our developed imaging methods can be applied to study chemotaxis of human leukocytes.

Imaging G-protein Coupled Receptor GPCR-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

Here, we describe detailed live cell imaging methods for investigating chemotaxis. We present fluorescence microscopic methods to monitor spatiotemporal dynamics of signaling events in migrating cells. Measurement of signaling events permits us to further understand how a GPCR-signaling network achieves gradient sensing of chemoattractants and controls directional migration of eukaryotic cells.

Many eukaryotic cells can detect gradients of chemical signals in their environments and migrate accordingly 1.  This guided cell migration is referred as ...

Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of different cell types in vitro and in vivo.1-5 Although there are literally thousands of publications using such dyes, some of the most commonly encountered cell tracking applications include monitoring of:
<ol>
	<li>stem and progenitor cell quiescence, proliferation and/or differentiation6-8</li>
	<li>antigen-driven membrane transfer9 and/or precursor cell proliferation3,4,10-18 and</li>
	<li>immune regulatory and effector cell function1,18-21.</li>
</ol>
Commercially available cell tracking dyes vary widely in their chemistries and fluorescence properties but the great majority fall into one of two classes based on their mechanism of cell labeling. "Membrane dyes", typified by PKH26, are highly lipophilic dyes that partition stably but non-covalently into cell membranes1,2,11. "Protein dyes", typified by CFSE, are amino-reactive dyes that form stable covalent bonds with cell proteins4,16,18. Each class has its own advantages and limitations. The key to their successful use, particularly in multicolor studies where multiple dyes are used to track different cell types, is therefore to understand the critical issues enabling optimal use of each class2-4,16,18,24.
The protocols included here highlight three common causes of poor or variable results when using cell-tracking dyes. These are:
<ol>
	<li>Failure to achieve bright, uniform, reproducible labeling. This is a necessary starting point for any cell tracking study but requires attention to different variables when using membrane dyes than when using protein dyes or equilibrium binding reagents such as antibodies.</li>
	<li>Suboptimal fluorochrome combinations and/or failure to include critical compensation controls. Tracking dye fluorescence is typically 102 - 103 times brighter than antibody fluorescence. It is therefore essential to verify that the presence of tracking dye does not compromise the ability to detect other probes being used.</li>
	<li>Failure to obtain a good fit with peak modeling software. Such software allows quantitative comparison of proliferative responses across different populations or stimuli based on precursor frequency or other metrics. Obtaining a good fit, however, requires exclusion of dead/dying cells that can distort dye dilution profiles and matching of the assumptions underlying the model with characteristics of the observed dye dilution profile.</li>
</ol>
Examples given here illustrate how these variables can affect results when using membrane and/or protein dyes to monitor cell proliferation.

Successful use of cell tracking dyes to monitor immune cell function and proliferation involves several critical steps. We describe methods for: 1) obtaining bright, uniform, reproducible label-ing with membrane dyes; 2) selecting fluorochromes and data acquisition conditions; and 3) choosing a model to quantify cell proliferation based on dye dilution.

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of ...

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and pathophysiology. This protocol describes a technique where mitochondria isolated from rodent tissue are immunolabeled and analyzed by flow cytometry. Mitochondria are isolated from rodent spinal cords and subjected to a rapid enrichment step so as to remove myelin, a major contaminant of mitochondrial fractions prepared from nervous tissue. Isolated mitochondria are then labeled with an antibody of choice and a fluorescently conjugated secondary antibody. Analysis by flow cytometry verifies the relative purity of mitochondrial preparations by staining with a mitochondrial specific dye, followed by detection and quantification of immunolabeled protein. This technique is rapid, quantifiable and high-throughput, allowing for the analysis of hundreds of thousands of mitochondria per sample. It is applicable to assess novel proteins at the mitochondrial surface under normal physiological conditions as well as the proteins that may become mislocalized to this organelle during pathology. Importantly, this method can be coupled to fluorescent indicator dyes to report on certain activities of mitochondrial subpopulations and is feasible for mitochondria from the central nervous system (brain and spinal cord) as well as liver.

Described here is a method to detect and quantify mitochondrial outer membrane proteins by immunolabeling of mitochondria isolated from rodent tissue and analysis by flow cytometry. This method can be extended to assess functional aspects of mitochondrial subpopulations.

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and ...

High-throughput Screening for Chemical Modulators of Post-transcriptionally Regulated Genes

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays focus on transcriptional regulation, being essentially aimed at the identification of promoter-targeting molecules. As a result, post-transcriptional mechanisms are largely uncovered by gene expression targeted drug development. Here we describe a cell-based assay aimed at investigating the role of the 3&#39; untranslated region (3&#8217; UTR) in the modulation of the fate of its mRNA, and at identifying compounds able to modify it. The assay is based on the use of a luciferase reporter construct containing the 3&#8217; UTR of a gene of interest stably integrated into a disease-relevant cell line. The protocol is divided into two parts, with the initial focus on the primary screening aimed at the identification of molecules affecting luciferase activity after 24 hr of treatment. The second part of the protocol describes the counter-screening necessary to discriminate compounds modulating luciferase activity specifically through the 3&#8217; UTR. In addition to the detailed protocol and representative results, we provide important considerations about the assay development and the validation of the hit(s) on the endogenous target. The described cell-based reporter gene assay will allow scientists to identify molecules modulating protein levels via post-transcriptional mechanisms dependent on a 3&#8217; UTR.

Here we describe a cell-based reporter gene assay as a valuable tool to screen chemical libraries for compounds modulating post-transcriptional control mechanisms exerted through 3&#8217; UTR.

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays ...

Investigating the Function of Coronin A in the Early Starvation Response of Dictyostelium discoideum by Aggregation Assays

Dictyostelium discoideum amoeba are found in soil, feeding on bacteria. When food sources become scarce, they secrete factors to initiate a multicellular development program, during which single cells chemotax towards aggregation centers1-4. This process is dependent on the release of cyclic adenosine monophosphate (cAMP)5. cAMP is produced in waves through the concerted action of adenylate cyclase and phosphodiesterases, and binds to G protein-coupled cAMP receptors6,7. A widely used assay to analyze the mechanisms involved in the developmental cycle of the lower eukaryote Dictyostelium discoideum is based on the observation of cell aggregation in submerged conditions8,9. This protocol describes the analysis of the role of coronin A in the developmental cycle by starvation in tissue-culture plates submerged in balanced salt solution (BSS)10. Coronin A is a member of the widely conserved protein family of coronins that have been implicated in a wide variety of activities11,12. Dictyostelium cells lacking coronin A are unable to form multicellular aggregates, and this defect can be rescued by supplying pulses of cAMP, suggesting that coronin A acts upstream of the cAMP cascade10. The techniques described in these studies provide robust tools to investigate functions of proteins during the initial stages of the developmental cycle of Dictyostelium discoideum upstream of the cAMP cascade. Therefore, utilizing this aggregation assay may allow the further study of coronin A function and advance our understanding of coronin biology.

Investigating the Function of Coronin A in the Early Starvation Response of Dictyostelium discoideum by Aggregation Assays

The social amoeba Dictyostelium discoideum undergoes a developmental transition into a multicellular organism when starved. The evolutionary conserved protein coronin A plays a crucial role in the initiation of development. Using aggregation assays as our main method, we aim to elucidate the role of coronin A in early development.

Dictyostelium discoideum amoeba are found in soil, feeding on bacteria. When food sources become scarce, they secrete factors to initiate a multicellular ...

Assessment of Dictyostelium discoideum Response to Acute Mechanical Stimulation

Chemotaxis, or migration up a gradient of a chemoattractant, is the best understood mode of directed migration. Studies using social amoeba Dictyostelium discoideum revealed that a complex signal transduction network of parallel pathways amplifies the response to chemoattractants, and leads to biased actin polymerization and protrusion of a pseudopod in the direction of a gradient. In contrast, molecular mechanisms driving other types of directed migration, for example, due to exposure to shear flow or electric fields, are not known. Many regulators of chemotaxis exhibit localization at the leading or lagging edge of a migrating cell, as well as show transient changes in localization or activation following global stimulation with a chemoattractant. To understand the molecular mechanisms of other types of directed migration we developed a method that allows examination of cellular response to acute mechanical stimulation based on brief (2 - 5 s) exposure to shear flow. This stimulation can be delivered in a channel while imaging cells expressing fluorescently-labeled biosensors to examine individual cell behavior. Additionally, cell population can be stimulated in a plate, lysed, and immunoblotted using antibodies that recognize active versions of proteins of interest. By combining both assays, one can examine a wide array of molecules activated by changes in subcellular localization and/or phosphorylation. Using this method we determined that acute mechanical stimulation triggers activation of the chemotactic signal transduction and actin cytoskeleton networks. The ability to examine cellular responses to acute mechanical stimulation is important for understanding the initiating events necessary for shear flow-induced motility. This approach also provides a tool for studying the chemotactic signal transduction network without the confounding influence of the chemoattractant receptor.

Assessment of Dictyostelium discoideum Response to Acute Mechanical Stimulation

Here we describe methods for assessing cellular response to acute mechanical stimulation. In the microscopy-based assay, we examine localization of fluorescently-labeled biosensors following brief stimulation with shear flow. We also test activation of various proteins of interest in response to acute mechanical stimulation biochemically.

Chemotaxis, or migration up a gradient of a chemoattractant, is the best understood mode of directed migration. Studies using social amoeba Dictyostelium ...

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

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IP-FCM: Immunoprecipitation Detected by Flow Cytometry

Quantitative Measurement of GLUT4 Translocation to the Plasma Membrane by Flow Cytometry

Flow Cytometry Purification of Mouse Meiotic Cells

High-Throughput Measurement and Classification of Organic P in Environmental Samples

Imaging G-protein Coupled Receptor (GPCR)-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

High-throughput Screening for Chemical Modulators of Post-transcriptionally Regulated Genes

Investigating the Function of Coronin A in the Early Starvation Response of Dictyostelium discoideum by Aggregation Assays

Assessment of Dictyostelium discoideum Response to Acute Mechanical Stimulation