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><tbody><tr><td>AX Medium</td><td>ForMedium</td><td>AXM0102</td><td></td></tr><tr><td>LoFlo medium</td><td>ForMedium</td><td>LF1001</td><td></td></tr><tr><td>MG132</td><td>Sigma</td><td>C2211</td><td></td></tr><tr><td>Lactacystin</td><td>Sigma</td><td>L6785</td><td></td></tr><tr><td>Geldanamycin</td><td>Santa Cruz</td><td>sc-200617</td><td></td></tr><tr><td>Radicicol</td><td>Santa Cruz</td><td>sc-200620</td><td></td></tr><tr><td>MatTek disch 35 mm</td><td>MatTek corporation</td><td>P35G-1.5-14-C</td><td>glass bottom imaging dish</td></tr><tr><td>CellViell cell culture dish</td><td>Greiner</td><td>627870</td><td>4-compartment glass-bottom imaging dish</td></tr><tr><td>Thermal Stage Heater/Cooler Insert</td><td>Warner Istruments</td><td>TB-3/CCD</td><td></td></tr><tr><td>Bipolar temperature controller</td><td>Warner Istruments</td><td>CL-100</td><td></td></tr><tr><td>Liquid cooling System</td><td>Warner Instruments</td><td>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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/files/ftp_upload/54730/54730fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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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6.2K 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

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

Developmental Biology

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One challenge is that embryogenesis dynamically unfolds over a period of about 13 h; this half day-long timescale has constrained the scope of experiments by limiting the number of embryos that can be imaged. Here, we describe a semi-high-throughput protocol that allows for the simultaneous 3D time-lapse imaging of development in 80&#8211;100 embryos at moderate time resolution, from up to 14 different conditions, in a single overnight run. The protocol is straightforward and can be implemented by any laboratory with access to a microscope with point visiting capacity. The utility of this protocol is demonstrated by using it to image two custom-built strains expressing fluorescent markers optimized to visualize key aspects of germ-layer specification and morphogenesis. To analyze the data, a custom program that crops individual embryos out of a broader field of view in all channels, z-steps, and timepoints and saves the sequences for each embryo into a separate tiff stack was built. The program, which includes a user-friendly graphical user interface (GUI), streamlines data processing by isolating, pre-processing, and uniformly orienting individual embryos in preparation for visualization or automated analysis. Also supplied is an ImageJ macro that compiles individual embryo data into a multi-panel file that displays maximum intensity fluorescence projection and brightfield images for each embryo at each time point. The protocols and tools described herein were validated by using them to characterize embryonic development following knock-down of 40 previously described developmental genes; this analysis visualized previously annotated developmental phenotypes and revealed new ones. In summary, this work details a semi-high-throughput imaging method coupled with a cropping program and ImageJ visualization tool that, when combined with strains expressing informative fluorescent markers, greatly accelerates experiments to analyze embryonic development.

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

This work describes a semi-high-throughput protocol that allows simultaneous 3D time-lapse imaging of embryogenesis in 80&#8211;100 C. elegans embryos in a single overnight run. Additionally, image processing and visualization tools are included to streamline data analysis. The combination of these methods with custom reporter strains enables detailed monitoring of embryogenesis.

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One ...

Monitoring Protein Aggregation Kinetics In Vivo using Automated Inclusion Counting in Caenorhabditis elegans

Protein aggregation into insoluble inclusions is a hallmark of a variety of human diseases, many of which are age-related. The nematode Caenorhabditis elegans is a well-established model organism that has been widely used in the field to study protein aggregation and toxicity. Its optical transparency enables the direct visualization of protein aggregation by fluorescence microscopy. Moreover, the fast reproductive cycle and short lifespan make the nematode a suitable model to screen for genes and molecules that modulate this process.
However, the quantification of aggregate load in living animals is poorly standardized, typically performed by manual inclusion counting under a fluorescence dissection microscope at a single time point. This approach can result in high variability between observers and limits the understanding of the aggregation process. In contrast, amyloid-like protein aggregation in vitro is routinely monitored by thioflavin T fluorescence in a highly quantitative and time-resolved fashion.
Here, an analogous method is presented for the unbiased analysis of aggregation kinetics in living C. elegans, using a high-throughput confocal microscope combined with custom-made image analysis and data fitting. The applicability of this method is demonstrated by monitoring inclusion formation of a fluorescently labeled polyglutamine (polyQ) protein in the body wall muscle cells. The image analysis workflow allows the determination of the numbers of inclusions at different timepoints, which are fitted to a mathematical model based on independent nucleation events in individual muscle cells. The method described here may prove useful to assess the effects of proteostasis factors and potential therapeutics for protein aggregation diseases in a living animal in a robust and quantitative manner.

Monitoring Protein Aggregation Kinetics In Vivo using Automated Inclusion Counting in Caenorhabditis elegans

Here, a method is presented for the analysis of protein aggregation kinetics in the nematode Caenorhabditis elegans. Animals from an age-synchronized population are imaged at different time points, followed by semiautomated inclusion counting in CellProfiler and fitting to a mathematical model in AmyloFit.

Protein aggregation into insoluble inclusions is a hallmark of a variety of human diseases, many of which are age-related. The nematode Caenorhabditis ...

Biochemistry

Real-Time Imaging of Germline Stem Cells in Drosophila Ex Vivo Germaria Culture

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to explore the dynamics of a myriad of developmental processes, such as cell division, stemness, differentiation, migration, apoptosis, autophagy, cellular adhesion, etc., over time. Recently, we have implemented an extended ex vivo culture and live imaging of the female Drosophila GSC niche. Using a Drosophila line harboring a GFP::Par-1 transgene as an example, this method allows the visualization of the GSCs&#39; asymmetric division within their niche and the description of the changes in the spectrosome morphology along the cell cycle. Here, we present a detailed protocol for the ex vivo culture of Drosophila germaria, enabling prolonged visualization of the female GSC niche. Importantly, this protocol is broadly applicable to live imaging GSCs with multiple fluorescently tagged proteins of interest that are available in stock centers and/or in the Drosophila research community.

Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein Par-1.

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

Neuroscience

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Few naturally-occurring plasmids are maintained in mammalian cells. Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus (EBV) and Kaposi's Sarcoma-associated herpesvirus (KSHV), which cause multiple human malignancies 1-3. These two genomes are replicated in a licensed manner, each using a single viral protein and cellular replication machinery, and are passed to daughter cells during cell division despite their lacking traditional centromeres 4-8.
Much work has been done to characterize the replications of these plasmid genomes using methods such as Southern blotting and fluorescence in situ hybridization (FISH). These methods are limited, though. Quantitative PCR and Southern blots provide information about the average number of plasmids per cell in a population of cells. FISH is a single-cell assay that reveals both the average number and the distribution of plasmids per cell in the population of cells but is static, allowing no information about the parent or progeny of the examined cell.
Here, we describe a method for visualizing plasmids in live cells. This method is based on the binding of a fluorescently tagged lactose repressor protein to multiple sites in the plasmid of interest 9. The DNA of interest is engineered to include approximately 250 tandem repeats of the lactose operator (LacO) sequence. LacO is specifically bound by the lactose repressor protein (LacI), which can be fused to a fluorescent protein. The fusion protein can either be expressed from the engineered plasmid or introduced by a retroviral vector. In this way, the DNA molecules are fluorescently tagged and therefore become visible via fluorescence microscopy. The fusion protein is blocked from binding the plasmid DNA by culturing cells in the presence of IPTG until the plasmids are ready to be viewed.
This system allows the plasmids to be monitored in living cells through several generations, revealing properties of their synthesis and partitioning to daughter cells. Ideal cells are adherent, easily transfected, and have large nuclei. This technique has been used to determine that 84% of EBV-derived plasmids are synthesized each generation and 88% of the newly synthesized plasmids partition faithfully to daughter cells in HeLa cells. Pairs of these EBV plasmids were seen to be tethered to or associated with sister chromatids after their synthesis in S-phase until they were seen to separate as the sister chromatids separated in Anaphase10. The method is currently being used to study replication of KSHV genomes in HeLa cells and SLK cells. HeLa cells are immortalized human epithelial cells, and SLK cells are immortalized human endothelial cells. Though SLK cells were originally derived from a KSHV lesion, neither the HeLa nor SLK cell line naturally harbors KSHV genomes11. In addition to studying viral replication, this visualization technique can be used to investigate the effects of the addition, removal, or mutation of various DNA sequence elements on synthesis, localization, and partitioning of other recombinant plasmid DNAs.

A method of observing individual DNA molecules in live cells is described. The technique is based on the binding of a fluorescently tagged lac repressor protein to binding sites engineered into the DNA of interest. This method can be adapted to follow many recombinant DNAs in live cells over time.

Few naturally-occurring plasmids are maintained in mammalian cells.  Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus  (EBV) ...

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage (CoTrAC)

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method makes it possible to count the numbers of protein molecules produced in one cell during sequential, five-minute time windows. It requires a fluorescence microscope with laser excitation power density of ~0.5 to 1 kW/cm2, which is sufficiently sensitive to detect single fluorescent protein molecules in live cells. The fluorescent reporter used in this method consists of three parts: a membrane targeting sequence, a fast-maturing, yellow fluorescent protein and a protease recognition sequence. The reporter is translationally fused to the N-terminus of a protein of interest. Cells are grown on a temperature-controlled microscope stage. Every five minutes, fluorescent molecules within cells are imaged (and later counted by analyzing fluorescence images) and subsequently photobleached so that only newly translated proteins are counted in the next measurement.
Fluorescence images resulting from this method can be analyzed by detecting fluorescent spots in each image, assigning them to individual cells and then assigning cells to cell lineages. The number of proteins produced within a time window in a given cell is calculated by dividing the integrated fluorescence intensity of spots by the average intensity of single fluorescent molecules. We used this method to measure expression levels in the range of 0-45 molecules in single 5 min time windows. This method enabled us to measure noise in the expression of the &lambda; repressor CI, and has many other potential applications in systems biology.

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage CoTrAC

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC), to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method has been used to follow the stochastic expression dynamics of a transcription factor, the &lambda; repressor CI 1.

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells ...

A Flexible Chamber for Time-Lapse Live-Cell Imaging with Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy is a label-free chemical imaging technology. Live-cell imaging with SRS has been demonstrated for many biological and biomedical applications. However, long-term time-lapse SRS imaging of live cells has not been widely adopted. SRS microscopy often uses a high numerical aperture (NA) water-immersion objective and a high NA oil-immersion condenser to achieve high-resolution imaging. In this case, the gap between the objective and the condenser is only a few millimeters. Therefore, most commercial stage-top environmental chambers cannot be used for SRS imaging because of their large thickness with a rigid glass cover. This paper describes the design and fabrication of a flexible chamber that can be used for time-lapse live-cell imaging with transmitted SRS signal detection on an upright microscope frame. The flexibility of the chamber is achieved by using a soft material - a thin natural rubber film. The new enclosure and chamber design can be easily added to an existing SRS imaging setup. The testing and preliminary results demonstrate that the flexible chamber system enables stable, long-term, time-lapse SRS imaging of live cells, which can be used for various bioimaging applications in the future.

We report a stage-top, flexible environmental chamber for time-lapse imaging of live cells using upright stimulated Raman scattering microscopy with transmitted signal detection. Lipid droplets were imaged in SKOV3 cells treated with oleic acid for up to 24 h with a 3 min time interval.

Stimulated Raman scattering (SRS) microscopy is a label-free chemical imaging technology. Live-cell imaging with SRS has been demonstrated for many ...

Bioengineering

Detecting, Visualizing and Quantitating the Generation of Reactive Oxygen Species in an Amoeba Model System

Reactive oxygen species (ROS) comprise a range of reactive and short-lived, oxygen-containing molecules, which are dynamically interconverted or eliminated either catalytically or spontaneously. Due to the short life spans of most ROS and the diversity of their sources and subcellular localizations, a complete picture can be obtained only by careful measurements using a combination of protocols. Here, we present a set of three different protocols using OxyBurst Green (OBG)-coated beads, or dihydroethidium (DHE) and Amplex UltraRed (AUR), to monitor qualitatively and quantitatively various ROS in professional phagocytes such as Dictyostelium. We optimised the beads coating procedures and used OBG-coated beads and live microscopy to dynamically visualize intraphagosomal ROS generation at the single cell level. We identified lipopolysaccharide (LPS) from E. coli as a potent stimulator for ROS generation in Dictyostelium. In addition, we developed real time, medium-throughput assays using DHE and AUR to quantitatively measure intracellular superoxide and extracellular H2O2 production, respectively.

We adapted a set of protocols for the measurement of reactive oxygen species (ROS) that can be applied in various amoeba and mammalian cellular models for qualitative and quantitative studies.

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

Summary

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Chapters in this video

Long-term Live Imaging of Drosophila Eye Disc

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

Monitoring Protein Aggregation Kinetics In Vivo using Automated Inclusion Counting in Caenorhabditis elegans

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage (CoTrAC)

A Flexible Chamber for Time-Lapse Live-Cell Imaging with Stimulated Raman Scattering Microscopy

Detecting, Visualizing and Quantitating the Generation of Reactive Oxygen Species in an Amoeba Model System