The authors have no Acknowledgements.

1. Cell Preparation
<ol>
	<li>Growing cells
	<ol>
		<li>Grow D. discoideum cells in AX medium at 23 &#176;C under light in tissue culture plates. Avoid keeping cells at high densities above 75% confluency. 
		NOTE: For an optimal response to heat stress, cells need to be in the exponential growth phase and should not have reached stationary phase.</li>
		<li>The day before imaging, split cells into low fluorescence medium (LFM) to 1 x 104 cells/ml. 
		NOTE: This step is necessary to reduce background fluorescence caused by the AX medium. The incubation time can be reduced to a minimum of 2 hr. However, vesicles taken up in AX medium can still generate some background signal after 2 hr.</li>
	</ol>
	</li>
	<li>Preparing cells for microscopy
	<ol>
		<li>Before imaging, harvest cells from the tissue plate in LFM and transfer an adequate number of cells into glass bottom dishes. 
		NOTE: Usually, 1 x 105 cells/ml is sufficient; however, if the experimental setup requires long imaging periods (longer than 8 hr), cell numbers need to be carefully adjusted, as D. discoideum cells will transition to the developmental cycle and start to stream and aggregate if cell numbers are high while nutrients are low.</li>
		<li>Allow the cells to settle for 20 min. Remove non-settled cells by carefully replacing the used medium with fresh medium.</li>
	</ol>
	</li>
</ol>
2. Imaging
NOTE: The protocol can be used on wide-field systems as well as in confocal microscopy setups. The usage of a temperature-controlled incubation box covering the objectives and the microscope stage is helpful, yet not essential for temperature control. The temperature of the incubation box is set to the highest temperature used in the experiment, e.g., to 40 &#176;C if heat stress is performed. Otherwise, the temperature is set to 25 &#176;C. Temperature changes should be made in advance to allow the system to adapt, as changes in temperature during imaging can affect the focus. 
NOTE: During imaging, the temperature is controlled using a cooling chamber. The default temperature for imaging non-stressed cells is 23 &#176;C. For heat stress conditions, the temperature is increased accordingly to the desired level. The piezo element in the cooling chamber can respond very quickly to the temperature shift. However, the focus of the images will change slightly (it can deviate to up to 20 &#181;m depending on the temperature shift). Thus manual refocusing is required within the first minutes of image acquisition, unless the used microscope is equipped with a hardware auto-focus.
<ol>
	<li>Image acquisition under non-stressed conditions
	<ol>
		<li>Image cells under non-stressed conditions at 23 &#176;C, acquire a minimum of 6 fields of view (FOVs). 
		NOTE: Non-stress condition can be visualized either by recording a short time-lapse (5-30 min) or acquisition of representative stills. For each condition or cell line acquire a minimum of 6 fields of view (FOVs). 
		NOTE: In case individual cells are of interest, position the respective cells in the middle of the FOV and record a 3 x 3 tile series with the FOV containing the cells of interest in the middle. D. discoideum cells are motile and might migrate out of the FOV. However, the 3 x 3 tile series increases the area potentially explored by the cell of interest and it remains traceable for time-lapse periods of ~6 hr.</li>
		<li>Acquire z-stacks of maximum 7 &#181;m in 1-0.25 &#181;m steps to capture the whole volume of the cell.</li>
		<li>Acquire a reference bright field image to assess the total number of cells.</li>
	</ol>
	</li>
	<li>Image acquisition for heat stress
	<ol>
		<li>Adjust the temperature of the cooling chamber to 30 &#176;C according to the manufacturer&#39;s instructions.</li>
		<li>After the temperature display has reached the desired value, refocus manually in the bright field channel and check all recorded FOVs before starting the time-lapse.</li>
		<li>Start a time-lapse for 4 hr of heat stress with a time interval of 5-10 min between time points. Check for correct focusing during the acquisition of the first two time points.</li>
	</ol>
	</li>
	<li>Image acquisition during recovery
	<ol>
		<li>After heat stress, reduce the temperature back to 23 &#176;C, wait for the temperature display to adjust and refocus manually.</li>
		<li>Record time-lapse movies in the recovery phase for 8-10 hr in 10-20 min time intervals. Check for correct focusing during the acquisition of the first time point.</li>
	</ol>
	</li>
</ol>
3. Image Analysis
NOTE: To quantify the number of cells harboring cytosolic foci, assess the total number of cells and the number of cells with cytosolic foci in each time frame. Due to the difficulty of segmenting cells from bright field images, the quantification needs to be done manually. In the following, data analysis is described for the free image processing package ((http://fiji.sc/Fiji). It requires the plugin &#8220;Cell Counter&#8221; (http://fiji.sc/Cell_Counter).
<ol>
	<li>Assessing total number of cells
	<ol>
		<li>Load the bright field image stack (XYT) by clicking &#34;File&#34; and &#34;Open&#34;.</li>
		<li>Open the &#34;Cell Counter&#34; plugin (&#34;Plugins&#34; -&#34;Cell Counter&#34;). Load the image stack by clicking &#34;Initialize&#34;.</li>
		<li>Choose the appropriate type of counters by ticking the appropriate box. 
		NOTE: use type 1 as counter for total number of cells and type 2 as counter for cells with cytosolic foci. Count all cells.</li>
		<li>Save the markers by clicking &#34;Save Markers&#34;.</li>
	</ol>
	</li>
	<li>Assessing the number of cells with cytosolic foci
	<ol>
		<li>Load the fluorescence image hyperstack (XZYT) (&#34;File&#34; - &#34;Open&#34;) and make a maximum intensity projection using the &#34;Z-Projector&#34; (Image/Stacks).</li>
		<li>Open the &#34;Cell Counter&#34; plugin. Load the resulting image stack (XYT) by clicking &#34;Initialize&#34;.</li>
		<li>Load the markers by clicking &#34;Load Markers&#34;. 
		CAUTION: The file names of the reference image and the fluorescence stack must be identical. If necessary, rename accordingly before initializing the image stack.</li>
		<li>Choose the appropriate marker type (see 3.1.3) and count cells with cytosolic foci. The existing markers can be used as guidance for the position of individual cells.</li>
		<li>Retrieve the number of counts per slice by clicking on &#34;Results&#34; and save the counts elsewhere for further calculations.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Bonner, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+for+the+Formation+of+Cell+Aggregates+by+Chemotaxis+in+the+Development+of+the+Slime+Mold+Dictyostelium+Discoideum.">Evidence for the Formation of Cell Aggregates by Chemotaxis in the Development of the Slime Mold Dictyostelium Discoideum.</a> Journal of Experimental Zoology. 106 (1), 1-26 (1947).</li><li>Loomis, W. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+signaling+during+development+of+Dictyostelium.">Cell signaling during development of Dictyostelium.</a> Developmental Biology. 391 (1), 1-16 (2014).</li><li>Nichols, J. M. E., Veltman, D., 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=Chemotaxis+of+a+model+organism:+progress+with+Dictyostelium.">Chemotaxis of a model organism: progress with Dictyostelium.</a> Current Opinion in Cell Biology. 36, 7-12 (2015).</li><li>Williams, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dictyostelium+finds+new+roles+to+model.">Dictyostelium finds new roles to model.</a> Genetics. 185 (3), 717-726 (2010).</li><li>M&#252;ller-Taubenberger, A., Kortholt, A., Eichinger, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simple+system+-+substantial+share:+The+use+of+Dictyostelium+in+cell+biology+and+molecular+medicine.">Simple system - substantial share: The use of Dictyostelium in cell biology and molecular medicine.</a> European Journal of Cell Biology. 92 (2), 45-53 (2013).</li><li>Malinovska, L., Palm, S., Gibson, K., Verbavatz, J. M., Alberti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dictyostelium+discoideum+has+a+highly+Q/N-rich+proteome+and+shows+an+unusual+resilience+to+protein+aggregation.">Dictyostelium discoideum has a highly Q/N-rich proteome and shows an unusual resilience to protein aggregation.</a> Proc Natl Acad Sci U S A. 112 (20), E2620-E2629 (2015).</li><li>Malinovska, L., Alberti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+misfolding+in+Dictyostelium:+Using+a+freak+of+nature+to+gain+insight+into+a+universal+problem.">Protein misfolding in Dictyostelium: Using a freak of nature to gain insight into a universal problem.</a> Prion. 9 (5), 339-346 (2015).</li><li>Schneider, K., Bertolotti, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surviving+protein+quality+control+catastrophes+-+from+cells+to+organisms.">Surviving protein quality control catastrophes - from cells to organisms.</a> Journal of Cell Science. 128 (21), 3861-3869 (2015).</li><li>Dobson, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+folding+and+misfolding.">Protein folding and misfolding.</a> Nature. 426 (6968), 884-890 (2003).</li><li>Fukui, Y., Yumura, S., Yumura, T. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Agar-overlay+immunofluorescence:+high-resolution+studies+of+cytoskeletal+components+and+their+changes+during+chemotaxis.">Agar-overlay immunofluorescence: high-resolution studies of cytoskeletal components and their changes during chemotaxis.</a> Methods in cell biology. 28, 347-356 (1987).</li><li>Graf, R., Rietdorf, J., Zimmermann, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+cell+spinning+disk+microscopy.">Live cell spinning disk microscopy.</a> Microscopy Techniques. 95, 57-75 (2005).</li><li>Samereier, M., Meyer, I., Koonce, M. P., Graef, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+Cell-Imaging+Techniques+for+Analyses+of+Microtubules+in+Dictyostelium.">Live Cell-Imaging Techniques for Analyses of Microtubules in Dictyostelium.</a> Methods in cell biology. 97, 341-357 (2010).</li><li>Herman, P. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stationary+phase+in+yeast.">Stationary phase in yeast.</a> Current opinion in microbiology. 5 (6), 602-607 (2002).</li><li>Loomis, W. F., Wheeler, S., Wheeler, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heat+shock+response+of+Dictyostelium.">Heat shock response of Dictyostelium.</a> Developmental Biology. 79 (2), 399-408 (1980).</li></ol>

The authors have nothing to disclose.

The protocol described here can be used to study the behavior of a particular protein of interest in response to heat-induced stress. Temperature increase to 30 &#176;C has been reported to trigger heat stress response and under these conditions the viability of D. discoideum is markedly reduced.
Modifications 
The protocol can be modified to compare the behavior of different proteins under the same stress conditions. For this, cells expressing different proteins with the same fluorescent tag are transferred into multi-well dishes, such as four chamber dishes (section 1.3.1). The protocol can also be applied on cells co-expressing proteins with different markers, such as GFP or RFP. This can be for example used to monitor the behavior of different components of the protein quality control (PQC) system. Cells expressing GFP-tagged aggregation marker and different RFP-tagged PCQ components can be observed using multi-well dished for imaging. This ensures the same stress conditions (speed of temperature increase/decrease, duration of the temperature increase/decrease) and allows for comparative studies.
Moreover, the protocol can be used to study the influence of the PQC system on the heat stress response. The activity of the components can be modulated by changing expression levels using genetic tools such as knock-out or overexpression or by using commercially available specific inhibitors6. The proteasome can be inhibited by adding MG132 (100 &#181;M) or lactacystin (10 &#181;M) to the growth medium. The chaperone Hsp90 can be inhibited using geldanamycin (6 &#181;M) or radicicol (10 &#181;M). The chaperone Hsp70 can be inhibited with VER-155088, although we could not confirm the efficacy of the inhibition in our experimental settings so far. Inhibitors should be added one day before imaging and the cells are incubated for 14 hr.
Critical Steps within the Protocol 
The critical step for the assessment of heat-induced perturbation is the state of the cells prior to the imaging experiment. Studies in yeast showed that cells acquire resistance to a variety of environmental stresses during stationary phase13. We also observed minimal responsiveness to applied heat stress if D. discoideum cells had reached stationary phase prior to imaging. Therefore, maintaining a constant cell number &#60;5 x 105 cells/ml is critical.
Moreover, high cell numbers can induce transition from the vegetative cycle of D. discoideum to the developmental cycle, thus triggering starvation pathways, which might lead to a different response to heat stress. If high cell numbers are reached in the recovery phase after heat stress, streaming and aggregating cells can interfere with data analysis as the cells move out of focus (see Figure 4).
Limitation of the Technique 
The loss of focus is the bottleneck of the protocol. During temperature changes, the focal plane can shift drastically, which requires manual refocusing in a time period immediately after changing the temperature. The jump in the focus can sometimes be too extreme to be overcome by software autofocus, especially if the time between the time points is high. However, if the used microscope is equipped with a hardware autofocus, this bottleneck can be circumvented.
Significance of the Technique with Respect to Existing Methods 
In comparison with existing setups such as the use of air-conditioned rooms, temperature-controlled incubation boxes covering the objectives and the microscope stage or temperature-controlled stages and objective collars, the use of a cooling chamber provides a more precise control of the ambient temperature. The Peltier-element in the cooling chamber can maintain a constant and uniform temperature throughout the experimental setup. In classical setups, local temperature differences might lead to different observation outcomes. Moreover, it can respond quickly and rapidly to induced changes in the temperature, while classical setups adapt very slowly to induced temperature changes, especially lowering the temperature. The Peltier-element in the cooling chamber can also cover a wider temperature range (15-40 &#176;C) than classical setups, with which reaching temperatures such as 15 &#176;C or 40 &#176;C is very difficult.
Dictyostelium discoideum is particularly sensitive to photo-toxicity. Previous studies used ascorbate as scavenger to reduce photo-toxicity. However, long imaging periods require additional supplementation. Moreover, the use of ascorbate is limited to studies where the mechanism of interest is not affected by the antioxidant supplement. We propose that temperature-controlled imaging can be used as an alternative approach to minimize photo-toxicity and could be combined with addition of ascorbate to further decrease toxicity.
Phototoxic effects can be minimized by providing a constant and uniform temperature of 23 &#176;C using a cooling chamber. Cells imaged using temperature control show fewer signs of photo-toxicity, such cell rounding, for a longer time period. This also allows imaging with higher intensities, smaller time intervals or more fields of view (FOVs).
General Application 
Heat stress at higher temperatures has been shown to trigger a different heat stress response14. Therefore, increasing the temperature to 34 &#176;C or 37 &#176;C might induce a different response. In addition to the applied temperature stress, the duration of a particular stress situation can be modified to study the immediate response or long-term adaptation to heat stress.
In general, the described protocol can be expanded to a wide set of applications. Because the precise temperature control considerably reduces photo-toxic effects, the protocol can be used in settings which require high exposure times and/or higher excitation light intensities, e.g., for visualizing proteins with a low expression level, or in combination with short time intervals between imaging, e.g., during time-lapse imaging. It can also be advantageous for imaging objects spanning the whole cell, e.g., microtubules, as these settings require a high number of optical z-sections.

Dictyostelium discoideum are solitary soil living amoebae that feed on bacteria and other microorganisms, which are taken up by phagocytosis. It has a unique and remarkable life cycle that has been a major area of research since its discovery1. The early interest in multicellular development2 and the molecular basis of chemotaxis3 was soon complemented by studies focusing on cell motility, cell polarity, innate immunity. In addition, D. discoideum was introduced as a model system for biomedical research4,5.
Recently, we established D. discoideum as a new system to study the protein quality control (PQC) system6,7. Its proteome is enriched in aggregation-prone prion-like proteins, which poses a challenge to protein quality control8. To investigate whether D. discoideum has developed special molecular mechanisms to control its highly aggregation-prone proteome, we studied the behavior of aggregation-prone marker proteins both under normal growth conditions and during stress. Stress conditions, such as heat stress, can be used to increase the rate of protein misfolding9. Therefore, we sought a system where we could induce temperature changes and simultaneously follow the behavior of marker proteins. For this purpose, we combined live-cell imaging with Peltier-controlled heating using a thermal stage insert (cooling chamber). This method allowed us to maintain a constant and uniform temperature as well as to induce a rapid, yet precise temperature change.
Live-cell imaging is used to study a variety of biological processes in D. discoideum. However, this approach faces two major limitations. First, the cells display a high motility and tend to migrate out of the field of view, thus tracking of individual cells often requires imaging of a large area. The cell mobility can be reduced by Agar overlay10, however, these conditions are not suitable for long term imaging due to a decline in viability. Second, D. discoideum cells show a particularly high sensitivity to photo-toxicity, which results in cell rounding and mitotic arrest11. Previous protocols addressed this issue by addition of ascorbate as a radical scavenger and reduction of exposure times12. The latter can be critical if the protein of interest is expressed at low levels and shows a weak fluorescence signal. The authors also suggest to provide a constant temperature of 21 &#176;C either by imaging in an air-conditioned room or by using temperature-controlled incubation boxes, which cover the objective and microscope stage12.
Here, we describe a method with improved temperature control by using a cooling chamber set at 23 &#176;C. During imaging our set-up significantly enhances the resistance to photo-toxicity. It allows the usage of higher exposure times and higher excitation light intensities. This is of particular importance during time-lapse imaging, as the time intervals need to be carefully balanced against the number of positions imaged and the exposure time used. The possibility to increase the numbers of imaged positions also allows the coverage of a wider imaging area and facilitates tracking of individual cells over a longer period of time.

<table border="0" cellpadding="0" cellspacing="0" width="587">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">AX Medium</td>
			<td style="width:85px;">ForMedium</td>
			<td style="width:119px;">AXM0102</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">LoFlo medium</td>
			<td>ForMedium</td>
			<td>LF1001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">MG132</td>
			<td style="width:85px;">Sigma</td>
			<td>C2211</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Lactacystin</td>
			<td style="width:85px;">Sigma</td>
			<td style="width:119px;">L6785</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Geldanamycin</td>
			<td style="width:85px;">Santa Cruz</td>
			<td style="width:119px;">sc-200617</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Radicicol</td>
			<td style="width:85px;">Santa Cruz</td>
			<td style="width:119px;">sc-200620</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">MatTek disch 35 mm</td>
			<td style="width:85px;">MatTek corporation</td>
			<td style="width:119px;">P35G-1.5-14-C</td>
			<td>glass bottom imaging dish</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CellViell cell culture dish</td>
			<td style="width:85px;">Greiner</td>
			<td align="right" style="width:119px;">627870</td>
			<td>4-compartments glass bottom imaging dish</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Thermal Stage Heater/Cooler Insert</td>
			<td style="width:85px;">Warner Istruments</td>
			<td style="width:119px;">TB-3/CCD</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Bipolar temperature controller</td>
			<td style="width:85px;">Warner Istruments</td>
			<td style="width:119px;">CL-100</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Liquid cooling System</td>
			<td style="width:85px;">Warner Instruments</td>
			<td style="width:119px;">LCS-1</td>
			<td></td>
		</tr>
	</tbody>
</table>

studying the protein quality control system of d. discoideum using temperature-controlled live cell imaging

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we showed that D. discoideum is remarkably resilient to protein aggregation and can be used to gain insights into the cellular protein quality control system. However, the use of D. discoideum as a model system poses several challenges to microscopy-based experimental approaches, such as the high motility of the cells and their susceptibility to photo-toxicity. The latter proves to be especially challenging when studying protein homeostasis, as the phototoxic effects can induce a cellular stress response and thus alter to behavior of the protein quality control system. 
Temperature increase is a commonly used way to induce cellular stress. Here, we describe a temperature-controllable imaging protocol, which allows observing temperature-induced perturbations in D. discoideum. Moreover, when applied at normal growth temperature, this imaging protocol can also noticeably reduce photo-toxicity, thus allowing imaging with higher intensities. This can be particularly useful when imaging proteins with very low expression levels. Moreover, the high mobility of the cells often requires the acquisition of multiple fields of view to follow individual cells, and the number of fields needs to be balanced against the desired time interval and exposure time.

The described imaging protocol can be used to analyze the behavior of aggregation-prone prion-like proteins in the amoeba D. discoideum during heat stress. Figure 1 shows the distribution of GFP-tagged polyglutamine protein Q103 in cells under normal growth conditions (Figure 1A), after 2 hours of heat stress at 30 &#176;C (Figure 1B) and after 6 hours of stress recovery and growth at normal growth conditions (Figure 1C). Upon temperature increase from 23 &#176;C to 30 &#176;C, the Q103-GFP marker coalesces from a diffuse distribution to punctate structures. Upon stress release and growth at normal temperature, these structures dissolve. The redistribution of Q103-GFP and formation of heat stress-induced cytosolic foci can be quantified using image analysis. Figure 2A shows the analysis of individual FOVs using Fiji and the plugin &#34;Cell counter&#34;. Figure 2B shows the quantification of the heat stress response. This shows that the cytosolic Q103-GFP foci increase in number with continuing heat stress. During heat stress, the protein quality control (PQC) system is overwhelmed, thus aggregation-prone proteins such as the prion-like protein Q103 cannot be maintained in a soluble state and increasingly aggregate in cytosolic foci. The cells also show a characteristic rounding, as depicted in Figure 1B. The reduction of cytosolic foci after stress removal suggests that the aggregated proteins are dissolved.
The imaging quality is sensitive to the initial cell density. Figure 3 shows an example of cells after stress recovery, comparing appropriate initial cell density (Figure 3A) and high initial cell density (Figure 3B). If the initial cell density was too high, cells started to form streams as depicted in Figure 3b. This which imposes difficulties on the subsequent quantification as the cells have moved out of the focal plane.
The described method can be used to monitor temperature-induced changes as described above. It can also be used to reduce photo-toxicity when imaging at physiological growth temperature. Figure 4 shows a comparison of cell imaged at 23 &#176;C in a cooling chamber (Figure 4A) and cells imaged without temperature control in an air conditioned room, set to 25 &#176;C (Figure 4B). Upon constant exposure to the excitation light, cells round up when not imaged under temperature-controlled conditions. This change in cell shape is indicating stress.
<img alt="Figure 1" src="/files/ftp_upload/54730/54730fig1.jpg" /> 
Figure 1: Aggregation-prone prion-like proteins change their cellular distribution during heat stress. GFP-tagged polyglutamine-rich huntingtin exon 1 (Q103) is constitutively expressed in AX2-214 D. discoideum cells (GFP signal depicted in inverted gray scale lookup table). (A) Under normal growth conditions at 23 &#176;C, Q103-GFP is diffusely distributed in the cytoplasm. (B) After 2 hr of heat stress at 30 &#176;C, Q103-GFP coalesces into cytosolic foci (red arrows). The cells show a characteristic shape change and round up. (C) Upon stress release and growth at 23 &#176;C, the cytosolic foci are dissolved. After 6 hours, no cytosolic foci can be observed. Scale bar = 10 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54730/54730fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54730/54730fig2.jpg" /> 
Figure 2: Quantification of heat-induced cytosolic foci formation. (A) Analysis of cells using the free image-processing package Fiji and the plugin &#34;Cell Counter&#34;. The total amount of cells (type 1, blue counter markers) is determined in the bright field image. The markers are loaded in the GFP-channel and the number of cells with cytosolic foci (type 2, red counter markers) is determined. (B) Quantification of foci after 3 hr of heat stress at 30 &#176;C and 3 hours of stress recovery at 23 &#176;C (n = 4 fields of view, error bar = SD). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54730/54730fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54730/54730fig3.jpg" /> 
Figure 3: Effect of cell density on long-term time-lapse imaging. (A) Cells imaged at initial cell densities &#60;105 cells/ml. After 3 hr of heat stress at 30 &#176;C and 6 hr of stress recovery at 23 &#176;C, cells still remain in the vegetative cycle and divide. Only few cells exit the focal plane. (B) Cells imaged at initial cell densities &#62;106 cells/ml. After 9 hr of imaging (settings as in A), cells have entered the developmental cycle and started streaming and aggregating. Cells in the aggregates cannot be clearly distinguished. A large portion of cells exits the focal plane and cannot be analyzed. Scale bar = 10 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54730/54730fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54730/54730fig4.jpg" /> 
Figure 4: Effect of temperature control on phototoxic effects. (A) Cells imaged for 10 min with temperature control at 23 &#176;C: 6 fields of view, z-stack (spacing 0.5 &#181;M, sample thickness 8 &#181;M), time lapse (total time 10 min, lapse 1 min), GFP channel (0.07 msec exposure time, 10% laser intensity), RFP channel (0.1 msec exposure time, 10% laser intensity). Cells show no signs of the characteristic morphological changes associated with phototoxic stress. (B) Cells imaged for 10 min without temperature control (same conditions as in A). Cells exhibit signs of phototoxic induced rounding, indicating stress. Scale bar = 10 &#181;m. The cells were imaged on a system based on a microscope with a 100x/1.4 numerical aperture (N.A.) objective. The images were collected with a CCD camera as 1,024 x 1,024 pixel files using 1 x 1 binning. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54730/54730fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging at JoVE.com

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The social amoebae Dictyostelium discoideum has recently been established as a system to study protein misfolding and proteostasis. Here, we describe a new imaging-based methodology to study temperature-induced protein aggregation and the cellular stress response in D. discoideum.

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we ...

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Nanyang Technological University

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JoVE Journal

Biology

6.1K Views.  Max-Planck Institute for Molecular Cell Biology and Genetics. The social amoebae Dictyostelium discoideum has recently been established as a system to study protein misfolding and proteostasis. Here, we describe a new imaging-based methodology to study temperature-induced protein aggregation and the cellular stress response in D. discoideum.

Video: Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based fractionation. The system is comprised of single-use, 8-sample capacity cartridges and a benchtop GELFREE Fractionation Instrument. During separation, a constant voltage is applied between the anode and cathode reservoirs, and each protein mixture is electrophoretically driven from a loading chamber into a specially designed gel column gel. Proteins are concentrated into a tight band in a stacking gel, and separated based on their respective electrophoretic mobilities in a resolving gel. As proteins elute from the column, they are trapped and concentrated in liquid phase in the collection chamber, free of the gel. The instrument is then paused at specific time intervals, and fractions are collected using a pipette. This process is repeated until all desired fractions have been collected. If fewer than 8 samples are run on a cartridge, any unused chambers can be used in subsequent separations.
This novel technology facilitates the quick and simple separation of up to 8 complex protein mixtures simultaneously, and offers several advantages when compared to previously available fractionation methods. This system is capable of fractionating up to 1mg of total protein per channel, for a total of 8mg per cartridge. Intact proteins over a broad mass range are separated on the basis of molecular weight, retaining important physiochemical properties of the analyte. The liquid phase entrapment provides for high recovery while eliminating the need for band or spot cutting, making the fractionation process highly reproducible1.

The accompanying video describes the use of the GELFREE 8100 Fractionation System, which partitions complex protein samples on the basis of molecular weight and recovers the fractions in liquid phase. The video describes how the technology works, how it is used, and provides resultant data, with polyacrylamide gel electrophoresis analysis of fractionated bovine liver homogenate.

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based ...

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

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 ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

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  ...

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

Mouse Fetal Whole Intestine Culture System for Ex Vivo Manipulation of Signaling Pathways and Three-dimensional Live Imaging of Villus Development

Most morphogenetic processes in the fetal intestine have been inferred from thin sections of fixed tissues, providing snapshots of changes over developmental stages. Three-dimensional information from thin serial sections can be challenging to interpret because of the difficulty of reconstructing serial sections perfectly and maintaining proper orientation of the tissue over serial sections. Recent findings by Grosse et al., 2011 highlight the importance of three- dimensional information in understanding morphogenesis of the developing villi of the intestine1. Three-dimensional reconstruction of singly labeled intestinal cells demonstrated that the majority of the intestinal epithelial cells contact both the apical and basal surfaces. Furthermore, three-dimensional reconstruction of the actin cytoskeleton at the apical surface of the epithelium demonstrated that the intestinal lumen is continuous and that secondary lumens are an artifact of sectioning. Those two points, along with the demonstration of interkinetic nuclear migration in the intestinal epithelium, defined the developing intestinal epithelium as a pseudostratified epithelium and not stratified as previously thought1. The ability to observe the epithelium three-dimensionally was seminal to demonstrating this point and redefining epithelial morphogenesis in the fetal intestine. With the evolution of multi-photon imaging technology and three-dimensional reconstruction software, the ability to visualize intact, developing organs is rapidly improving. Two-photon excitation allows less damaging penetration deeper into tissues with high resolution. Two-photon imaging and 3D reconstruction of the whole fetal mouse intestines in Walton et al., 2012 helped to define the pattern of villus outgrowth2. Here we describe a whole organ culture system that allows ex vivo development of villi and extensions of that culture system to allow the intestines to be three-dimensionally imaged during their development.

Mouse Fetal Whole Intestine Culture System for Ex Vivo Manipulation of Signaling Pathways and Three-dimensional Live Imaging of Villus Development

Improved imaging technology is allowing three-dimensional imaging of organs during development. Here we describe a whole organ culture system that allows live imaging of the developing villi in the fetal mouse intestine.

Most morphogenetic processes in the fetal intestine have been inferred from thin sections of fixed tissues, providing snapshots of changes over ...

Live-cell Imaging of Platelet Degranulation and Secretion Under Flow

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the formation of thrombi that occlude the vasculature and injure organs, with life-threatening consequences. This motivates scientific research on platelet function and the development of methods to track cell-biological processes as they occur under flow conditions.
A variety of flow models are available for the study of platelet adhesion and aggregation, two key phenomena in platelet biology. This work describes a method to study real-time platelet degranulation under flow during activation. The method makes use of a flow chamber coupled to a syringe-pump setup that is placed under a wide-field, inverted, LED-based fluorescence microscope. The setup described here allows for the simultaneous excitation of multiple fluorophores that are delivered by fluorescently labeled antibodies or fluorescent dyes. After live-cell imaging experiments, the cover glasses can be further processed and analyzed using static microscopy (i.e., confocal microscopy or scanning electron microscopy).

This work describes a fluorescence microscopy-based method for the study of platelet adhesion, spreading, and secretion under flow. This versatile platform enables the investigation of platelet function for mechanistic research on thrombosis and hemostasis.

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the ...

LDL Cholesterol Uptake Assay Using Live Cell Imaging Analysis with Cell Health Monitoring

The regulation of LDL cholesterol uptake through LDLR-mediated endocytosis is an important area of study in various major pathologies including metabolic disorder, cardiovascular disease, and kidney disease. Currently, there is no available method to assess LDL uptake while simultaneously monitoring for health of the cells. The current study presents a protocol, using a live cell imaging analysis system, to acquire serial measurements of LDL influx with concurrent monitoring for cell health. This novel technique is tested in three human cell lines (hepatic, renal tubular epithelial, and coronary artery endothelial cells) over a four-hour time course. Moreover, the sensitivity of this technique is validated with well-known LDL uptake inhibitors, Dynasore and recombinant PCSK9 protein, as well as by an LDL uptake promoter, Simvastatin. Taken together, this method provides a medium-to-high throughput platform for simultaneously screening pharmacological activity as well as monitoring of cell morphology, hence cytotoxicity of compounds regulating LDL influx. The analysis can be used with different imaging systems and analytical software.

This protocol provides an efficient approach to measuring LDL cholesterol uptake with real time influx rates using a live cell imaging system in various cell types. This technique provides a platform to screen the pharmacological activity of compounds affecting LDL influx while monitoring for cell morphology and hence potential cytotoxicity.

The regulation of LDL cholesterol uptake through LDLR-mediated endocytosis is an important area of study in various major pathologies including metabolic ...