The authors thank Rocco D&#39;Antuono for proofreading aspects of the manuscript, as well as all present and past members of the Molecular Cell Biology of Autophagy (MCBA) lab for the discussions that led to the refinement of these protocols.&#160;A. v.V., S.d.T., E.A., S.A.T, were supported by The Francis Crick Institute which receives its core funding from Cancer Research UK (CC2134), the UK Medical Research Council (CC2134). This research was funded in whole, or in part, by the Wellcome Trust (CC2134). For&#160;the purpose of open access, the author has applied a CC BY public copyright licence to any Author Accepted Manuscript version arising from this submission.

All the reagents used in this study are commercially available, except for the ATG9A DNA constructs and homemade STO-215 antibody (see Table of Materials), which are available upon request. The analysis tools described here are based on open-source software (FIJI/ImageJ)18.
1. Cell culture
<ol>
	<li>Maintain HEK293A cells in a T150 tissue culture treated flask to 80%-90% confluence in high-glucose DMEM (Dulbecco&#39;s Modified Eagle Medium) supplemented with 10% FBS (fetal bovine serum) and 4 mM L-glutamine (see Table of Materials). Incubate the cells at 37 &#176;C in a humidified tissue culture incubator at 10% CO2. 
	NOTE: HEK293A cells are used in the present study, as they have a robust canonical autophagy response upon amino acid starvation, which is, in particular, detected by an increase in lipidated, membrane-associated LC31,9,19, and are suitable for imaging.</li>
	<li>Passage the cells by aspirating the medium from the flask using a serological pipette. Wash the cells once with 15 mL of 1x PBS (phosphate-buffered saline) or a similar solution before adding 2 mL of Trypsin-EDTA (ethylenediaminetetraacetic acid) solution to detach the cells. Collect the detached cells with 8 mL of DMEM, and seed back a number of cells so that 80%-90% confluency is reached after 2 days for the experiments described here.</li>
</ol>
2. Endogenous staining of ATG9A
<ol>
	<li>Seed HEK293A cells (or cells of choice) on sterile No. 1.5 glass coverslips placed into a 24-well plate so that they are ~80% confluent the next day. This typically gives 7 x 104 cells/well in 500 &#956;L of DMEM. 
	NOTE: For HEK293A cells or other loosely adherent cell lines, the coverslips need to be coated with poly-D-Lysine at 0.1 mg/mL in deionized water. Add 500 &#956;L of the poly-D-lysine (see Table of Materials) on the top of the coverslips placed into the wells for 10 min at room temperature (RT), followed by three washes with deionized water and a last wash in DMEM. Proceed to seeding the cells after the coating, and take care that they are evenly spread out in the culture dish.</li>
	<li>Treat the cells according to the specific experimental conditions (i.e., starvation for 2 h in EBSS [Earle&#39;s Balanced Salt Solution]). 
	NOTE: The EBSS composition is 1 g/L D-glucose, 6.8 g/L NaCl, 0.4 g/L KCl, 0.151 g/L CaCl2.2H2O, 0.2 g/L mM MgSO4.7H2O, 0.124 g/L NaH2HPO4.2H2O, and 2.2 g/L NaHCO3 (see Table of Materials) dissolved in distilled water.</li>
	<li>Aspirate the medium, and carefully replace it with 500 &#956;L of 4% formaldehyde solution in PBS supplemented with 0.1 mM CaCl2 and 0.1 mM MgCl2 for 20 min at RT to fix the cells.</li>
	<li>Aspirate the 4% formaldehyde solution from each well, replacing it with 500 &#956;L of PBS. Repeat this wash step three times. Do not let the coverslips dry out or remain without liquid.</li>
	<li>Quench the free aldehyde groups by using 500 &#956;L of 50 mM NH4Cl solution in PBS for 10 min at RT.</li>
	<li>Aspirate the PBS, and replace it with 500 &#956;L of a 50 &#956;g/mL digitonin solution in PBS (stock solution 1 mg/mL in deionized water, see Table of Materials) for 5 min at RT to permeabilize the cells.</li>
	<li>Aspirate the digitonin solution from each well, and replace it with 500 &#956;L of PBS. Repeat this wash step three times.</li>
	<li>Aspirate the PBS from each well, and replace it with 500 &#956;L of blocking solution (5% BSA [bovine serum albumin] diluted in PBS) for 30 min at RT.</li>
	<li>Aspirate the blocking solution, and replace it with 500 &#956;L of PBS.</li>
	<li>Using tweezers, collect the coverslips from the well, carefully remove the excess PBS solution using thin tissue wipes or cellulose filter paper, and gently lay down each coverslip, cell side down, onto a 50 &#956;L drop of primary antibody solution (e.g., Armenian Hamster 14F2 diluted to 0.9 &#956;g/mL in 1% BSA/PBS solution, see Table of Materials). Incubate in a humidified chamber for 1 h at RT. 
	NOTE: For easy handling, the drops of antibody solution can be placed on a sheet of self-sealing thermoplastic film (see Table of Materials) within another container instead of directly onto a solid surface.</li>
	<li>Collect the coverslips using tweezers, and, after gently draining off the excess primary antibody solution using thin tissue wipes, replace the coverslips (cell side up) in the 24-well plate, and wash them three times with PBS.</li>
	<li>Repeat step 2.10. Using tweezers, collect the coverslips from each well, carefully draining off the excess PBS using thin tissue wipes, and gently lay down each coverslip (cell side down) onto a 50 &#956;L drop of secondary antibody solution diluted 1:1,000 in 1% BSA/PBS solution (e.g., Cy3 Goat Anti-Armenian Hamster IgG, which has minimal crossreactivity with bovine, human, mouse, rabbit, and rat serum proteins, see Table of Materials). Incubate in a humidified chamber for 1 h at RT. 
	NOTE: Optional: A cytoskeleton marker can be used in addition to the secondary antibody for the subsequent image analysis (i.e., Alexa Fluor 647 Phalloidin diluted 1:1,000, see Table of Materials). For easy handling, the drops of antibody solution can be placed on a sheet of sealing film within another container instead of directly onto a solid surface.</li>
	<li>Collect the coverslips using tweezers and, after draining off the excess secondary antibody solution, place the coverslips in the 24-well plate (cell side up) and wash them three times with 500 &#956;L PBS.</li>
	<li>Using tweezers, collect the coverslips, carefully draining off the excess PBS using thin tissue wipes, and gently lay down each coverslip (cell side down) onto a 50 &#956;L drop of 1:4,000 Hoechst solution (Hoechst 33342) in PBS. Incubate in a humidity chamber for 5 min at RT. 
	NOTE: For easy handling, the drops of antibody solution can be placed on a sheet of sealing film within another container instead of directly onto a solid surface.</li>
	<li>Collect the coverslips using tweezers, and, after gently draining off the excess Hoechst solution using thin tissue wipes, replace the coverslips in the 24-well plate, and wash three times with PBS and once with de-ionized water (500 &#956;L per wash).</li>
	<li>Carefully drain the excess deionized water using thin tissue wipes, and gently lay down each coverslip (cell side down) onto a 10-20 &#956;L drop of mounting solution (see Table of Materials) spotted onto a microscope slide for immunofluorescence, avoiding the formation of air bubbles. 
	NOTE: The mounting medium used here has the same refractive index as immersion oil and hardens after a few hours at RT or overnight at 4 &#176;C. Non-hardening mounting solutions can be used, but care must be taken to seal the coverslip with nail polish.</li>
	<li>Remove the excess mounting solution by aspiration, and let the samples dry while lying flat at RT overnight in the dark, either in a slide holder or covered with aluminum foil.</li>
</ol>
3. Image acquisition
<ol>
	<li>Turn on the confocal microscope. Open the imaging software (see Table of Materials) to start the image acquisition setup.</li>
	<li>In the Function tab, select the Plan-Apochromat 63x/1.4 Oil DIC M27 objective lens to capture the images for analysis.</li>
	<li>In the Acquisition Function tab, turn on the appropriate lasers in the Laser zone of the Control Panel (Argon, Diode-405-30, DPSS 561-10, and HeNe633).</li>
	<li>In the Imaging Setup zone of the Control Panel, create four tracks, with each track corresponding to one channel, to perform sequential acquisition.</li>
	<li>Set the appropriate resolution of the images in the Acquisition Mode window. Set the resolution at 1,024 x 1,024 (Frame size), and select a Bit depth of 16-bit.</li>
	<li>For each channel, adjust the laser power and gain to obtain a good signal without saturated pixels, which would impede the image intensity analysis. Using the Live button, set the Laser output level and Gain (Master). 
	NOTE: The use of the range indicator option is recommended for saturated pixel detection. Keep the laser power between 1% and 10% and the Gain (Master) below 850 to avoid background noise.</li>
	<li>In the Channels zone of the Control Panel, set the same pinhole aperture for each channel, considering 1 Airy Unit (AU) for the channel with the highest wavelength.</li>
	<li>Acquire 10 random fields containing similar amounts of cells (20-30 cells per field). The acquisition of 100-200 cells per condition will provide good power for the later image analysis.</li>
</ol>
4. Image analysis of ATG9A dispersal
<ol>
	<li>Download FIJI software from the internet (see Table of Materials). Open the image with FIJI by clicking on Plugins &#62; Bio-Formats &#62; BioFormats Importer.</li>
	<li>Click on Analyse &#62; Set measurements, and select the Mean gray value in the measurements window for the intensity analysis.</li>
	<li>Click on Image &#62; Color &#62; Split Channels, and separate the channels into three images (Golgi marker, cytoskeleton, ATG9A signal).</li>
	<li>Go to Image &#62; Adjust &#62; Threshold, and define a threshold in the channel corresponding to the Golgi marker.</li>
	<li>Click on Edit &#62; Selection &#62; Create Selection, and create a selection from the binary image.</li>
	<li>Go to Edit &#62; Selection &#62; Add to Manager &#62; Rename (Golgi), save the selection in the ROI manager, and rename it to Golgi.</li>
	<li>Click on Image &#62; Adjust &#62; Threshold, and define a threshold in the channel corresponding to the cytoskeleton marker to define the cell contour.</li>
	<li>Go to Edit &#62; Selection &#62; Create Selection, and create a selection from the binary image.</li>
	<li>Click on Edit &#62; Selection &#62; Add to Manager &#62; Rename (Total), save the selection in the ROI manager, and rename it Total.</li>
	<li>Select the image corresponding to the ATG9A staining.</li>
	<li>In Analyse &#62; Measure, apply the ROI Golgi by clicking on it, and measure the intensity of the Golgi region.</li>
	<li>Click on Analyse &#62; Measure, apply the ROI Total by clicking on it, and measure the intensity of the Total region.</li>
	<li>Repeat the procedure in all the images, save the results as .csv files, and process the data using the following formula: 
	ATG9A dispersal rate = Golgi intensity/Total intensity</li>
</ol>
5. Live cell imaging of ATG9A constructs
<ol>
	<li>Seed HEK293A cells (or cells of choice) in 2 mL of medium into a 60 mm tissue culture dish so they reach ~65%-70% confluency the next day; this typically gives 1 x 106-2 x 106 cells.</li>
	<li>The following day, prepare Lipofectamine:DNA mixes (or a suitable alternative DNA transfection reagent, see Table of Materials) as follows:
	<ol>
		<li>Depending on construct expression efficiency, dilute 0.5-2 &#181;g of plasmid DNA into 100 &#181;L of a suitable serum-free medium, and gently mix the solution by pipetting up and down. Incubate the mixture for 5 min at RT. 
		NOTE: The plasmid DNA construct is experiment-specific. Researchers may either clone by themselves or obtain from the authors upon request.</li>
		<li>Dilute Lipofectamine 2000 transfection reagent at a ratio of 3:1 Lipofectamine:DNA into 100 &#181;L of a suitable serum-free medium and gently mix the solution by pipetting up and down. Incubate this mix for 5 min at RT.</li>
		<li>Mix both solutions together by gently pipetting up and down and incubating for 20 min at RT.</li>
	</ol>
	</li>
	<li>Add the Lipofectamine:DNA mix to each cell culture plate containing 4 mL of growth medium, and gently rock the plate back and forth to distribute the mix evenly. Incubate the cells at 37 &#176;C in a humidified cell culture incubator at 10% CO2.</li>
	<li>Replace the medium with fresh growth medium 4 h after transfection, and incubate the cells at 37 &#176;C in a humidified cell culture incubator at 10% CO2 overnight.</li>
	<li>The next day, trypsinize, count, and reseed the cells on culture dishes suitable for live-cell microscopy (culture dishes with a No. 1.5 glass coverslip, see Table of Materials). Seed 0.4 x 106-0.7 x 106 cells onto the dishes to reach ~60%-75% confluency on the coverslip at the time of imaging. 
	NOTE: Coating with poly-D-lysine, as explained in step 2.1, is recommended for HEK293A cells.</li>
	<li>The next day (48 h after transfection), image the cells.</li>
</ol>
6. Investigating the glycosylation state of ATG9A
<ol>
	<li>Seed HEK293A cells (or cells of choice) in a 10 cm tissue culture dish so they reach ~80% confluency the next day if planning to investigate endogenous ATG9A, which typically gives 1.5 x 106-2.5 x 106 cells in 10 mL. Alternatively, seed the cells so they reach ~65%-70% confluency if transfecting ATG9A constructs, which typically gives 1 x 106-1.5 x 106 cells in 10 mL.</li>
	<li>Treat the cells with 100 &#181;g/mL cyclohexamide (CHX, see Table of Materials) or vehicle the next day. Incubate for 24 h (or until a desired time point).</li>
	<li>Remove the cells from the incubator, aspirate the medium, and place on ice. Replace it with 5 mL of ice-cold 1x PBS. Physically detach the cells from the dish using a cell scraper, and pipette the PBS-cell solution into a 15 mL conical-bottom tube. Centrifuge at 800 x g for 5 min at 4 &#176;C to pellet the cells.</li>
	<li>Aspirate the supernatant, and resuspend the cells in ~100 &#181;L (depending on the size of the cell pellet) of ice-cold TNTE lysis buffer (1% Triton, 150 mM NaCl, 20&#160;mM Tris-HCl, pH 7.4, 0.5 mM EDTA) supplemented with protease inhibitor cocktail tablets (see Table of Materials), and transfer to a 1.5 mL microcentrifuge tube.
	<ol>
		<li>Incubate the lysate on ice for 15 min. After this, centrifuge the lysate at 20,000 x g for 10 min at 4 &#176;C to sediment the nuclei and insoluble debris, and transfer the supernatant to a fresh microcentrifuge tube.</li>
	</ol>
	</li>
	<li>Quantify the protein concentration of the lysate using the Bradford method20 and a spectrophotometer that can measure at a wavelength of 595 nm.</li>
	<li>Normalize the protein amounts, and combine 16 &#181;L of the lysate (containing no more than 100 &#181;g of protein in total) with 5x PNGase F (Peptide:N-glycosidase F) buffer according to the manufacturer&#39;s instructions before adding 1 &#181;L of PNGase F enzyme (see Table of Materials). Incubate as instructed by the PNGase F manufacturer.</li>
	<li>Add a volume of 3x Laemmli sample buffer to achieve a 1x concentration, and incubate at 65 &#176;C for 5 min before loading for electrophoresis using a Tris-acetate gel to maximize the separation of proteins. Transfer the proteins from the gel to a suitable membrane (i.e., PVDF [polyvinylidene difluoride]) using standard western blot protocols21 (see Table of Materials). 
	NOTE: Boiling the samples at 95 &#176;C will cause ATG9A to be aggregated, thus reducing the detection of ATG9A.</li>
	<li>Perform the western blot using specific antibodies for ATG9A (STO-215 antibody, produced in-house1) (see Table of Materials). Leave the higher-molecular weight section of the membrane uncut to visualize the higher ATG9A molecular weight species.</li>
</ol>

Investigating ATG9A Localization in Cells

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coordinated+regulation+of+autophagy+by+p38alpha+MAPK+through+mAtg9+and+p38IP.">Coordinated regulation of autophagy by p38alpha MAPK through mAtg9 and p38IP.</a> EMBO Journal. 29 (1), 27-40 (2010).</li><li>Claude-Taupin, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ATG9A+protects+the+plasma+membrane+from+programmed+and+incidental+permeabilization.">ATG9A protects the plasma membrane from programmed and incidental permeabilization.</a> Nature Cell Biology. 23 (8), 846-858 (2021).</li><li>Aaron, J. S., Taylor, A. B., Chew, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Image+co-localization+-+Co-occurrence+versus+correlation.">Image co-localization - Co-occurrence versus correlation.</a> Journal of Cell Science. 131 (3), 211847 (2018).</li><li>D'Antuono, R., Nechyporuk-Zloy, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+digital+image+acquisition,+design,+processing,+analysis,+management,+and+presentation.">Basic digital image acquisition, design, processing, analysis, management, and presentation.</a> Principles of Light Microscopy: From Basic to Advanced. , 77-104 (2022).</li></ol>

S.A.T. serves on the scientific advisory board of Casma Therapeutics.

This study illustrates the various tools that can be used to investigate ATG9A localization. Firstly, this study describes how ATG9A can be visualized by immunofluorescence and how this can be quantified. Secondly, strategies that can be used to tag ATG9A with a fluorescent marker for visualization in either fixed or live cells are compared. Finally, this work describes how to investigate and use the glycosylation state of ATG9A to determine if ATG9A has exited the ER and trafficked through the Golgi.
Regarding the characterization of endogenous ATG9A localization by immunofluorescence, care must be taken with the fixation and permeabilization methods employed for the experiment. According to the standard procedures here described, paraformaldehyde fixation coupled with digitonin permeabilization are good conditions to visualize both Golgi-associated ATG9A and ATG9A-positive vesicles7. Together with fixation and permeabilization, the timing of incubation with the primary antibody solution is also critical. We have observed, but not documented, that higher concentrations of primary antibody solution and longer incubation times can lead to a misrepresentative increase in the Golgi staining of ATG9A, which eventually compromises the detection of ATG9A redistribution to other membrane compartments. Additionally, since ATG9A is present in many intracellular compartments1,13,17,22,23,24,27,28,it is important to use specific membrane markers, together with ATG9A, to identify where ATG9A is located. Several approaches have been used in the past to quantify ATG9A localization, including Pearson&#39;s correlation coefficient for colocalization29. However, the partial overlap of ATG9A with the Golgi and the distinct vesicular compartment leads to a high number of pixel outliers, which may bias the interpretation of the correlation coefficient. For this reason, a more simplistic approach based on the ratio of the mean fluorescence in the two compartments to be analyzed is preferred, and this approach is less sensitive to cell-by-cell variability. For further information on image analysis through microscopy, readers are directed to this book chapter30.
When investigating the glycosylation status of ATG9A, the selection of gels for running the western blots is important. For this protocol, 3%-8% Tris-acetate gels are preferred because they offer the highest resolution for larger proteins, but alternative gel compositions or running buffers that offer a good separation of high-molecular weight proteins can also be used. The experimenter can ensure the maximal separation of proteins by increasing the time of electrophoresis.
When preparing the samples to visualize ATG9A on western blot, care should be taken not to boil the samples after adding the Laemmli buffer; boiling at 95 &#176;C induces the formation of ATG9A aggregates, and subsequently, ATG9A does not migrate efficiently into the gel1. Heating the samples at 65 &#176;C for 5 min is recommended27.
High levels of transfection usually lead to higher accumulation of ATG9A in the ER, while moderate expression levels help the physiological localization of the protein. Anecdotally, incubation times of 72 h instead of 48 h often help reduce ER localization artifacts. Notably, mRFP-ATG9A can accurately report on ATG9A trafficking and function if the levels are controlled through expression levels or by using stable cell lines8,9,22,27.
The failure of a population of overexpressed ATG9A to acquire mature N-linked glycans can be used as a read-out for perturbed ATG9A trafficking. When mutating or deleting certain regions of ATG9A, there is a risk of increased ER retention, which can lead to failure to acquire mature N-linked glycans and, thus, a faster-migrating ATG9A band on western blot. Researchers working with truncated ATG9A constructs should check for ER retention, glycosylation states, and Golgi localization.
For the live-cell imaging of ATG9A, an Airyscan microscope, relying on the fast Airyscan function, provides optimal resolution of typically about 120 nm. For localization accuracy, frame rates of around 1-2 frames per second (fps) in super-resolution mode are optimal depending on how many channels are imaged. Similar confocal microscopes that can image at high speed can also be used for the imaging of ATG9A vesicles; however, it should be noted that the imaging speed can directly affect the detection of events and, therefore, affect the interpretation of the data.
In summary, the presented protocols describe ways to quantify and characterize ATG9A localization by immunofluorescence, live-cell microscopy, and its glycosylation status. These protocols can aid researchers working with ATG9A and help avoid some pitfalls.

ATG9A is the only transmembrane protein of the core autophagy machinery and is trafficked between the Golgi and a cytosolic ATG9A vesicle compartment, transiting through the endosomal compartment1. Having long been enigmatic, ATG9A has recently been described to function as a lipid scramblase, as it equilibrates lipids across membrane bilayers2,3. It is now clear that ATG9A resides at the top of the hierarchy in autophagosome formation, and its study is, thus, vital for understanding autophagy4,5. As such, ATG9A vesicles have been recently proposed as the &#34;seed&#34; of the autophagosome6,7. However, previous studies have demonstrated that ATG9A only transiently interacts with the forming autophagosome at different steps of its maturation and does not integrate into the autophagic membrane6,8,9,10,11. Thus, further investigations are needed to completely unravel the role and potential multiple functions of ATG9A in autophagosome formation. However, the discrepancy between the current models and the previous data can only be resolved through targeted experiments addressing the trafficking of ATG9A using validated quantitative approaches and intracellular markers.
There are various tools in use to study ATG9A, each with advantages and disadvantages, and the use of these tools is complicated by the structure of ATG9A, its molecular function, and cellular trafficking2,8,12. ATG9A forms a homotrimer, is glycosylated, and is trafficked throughout the cell to compartments such as the Golgi, the endosomes, and the plasma membrane13,14. Given its complex itinerary, there are several challenges in interpreting readouts such as ATG9A dispersal from the Golgi upon specific treatments or stimuli (such as nutrient and serum starvation). ATG9A is extremely dynamic in terms of vesicular trafficking; indeed, ATG9A-containing vesicles have been defined as the ATG9A compartment in the context of starvation-induced autophagy. The ATG9A compartment, formed by these dynamic vesicles, transiently interacts with several intracellular organelles8,15,16,17. The techniques described here, including immunofluorescence, live imaging, and glycosylation assays, should aid in the detection and understanding of ATG9A biology. In particular, the approaches described in this article will help to address questions about localization to specific cellular compartments and interactions with specific protein partners and/or membrane compartments. As the ATG9A hydrophobic conserved core domain (PFAM domain PF04109) has a unique topology and ATG9A cycles between several membrane compartments, researchers should be aware of certain pitfalls and artifacts when transiently overexpressing ATG9A, including, but not restricted to, endoplasmic reticulum (ER) retention. Other possible issues may arise due to misfolding of the protein, artifactual aggregation in normal growing conditions, or insufficient detection of the vesicular compartment due to suboptimal permeabilization protocols for immunofluorescence.
When imaging endogenous ATG9A, care must be taken in the sample preparation and image acquisition to ensure the quality of the subsequent quantitative analysis and the correct interpretation of the data. Combining the techniques described in this article with standard biochemical approaches (such as immunoprecipitation or pull-down experiments not described here) should improve our understanding of ATG9A function. This experimental toolkit is intended to help new researchers navigate some of the assays required to determine the function of ATG9A in their biological system.

<table><tbody><tr><td>24 multiwell plates&amp;nbsp;</td><td>Falcon</td><td>353047</td><td>For tissue culture</td></tr><tr><td>35 mm Dish | No. 1.5 Coverslip | 14 mm Glass Diameter | Uncoated</td><td>MATTEK</td><td>P35G-1.5-14-C</td><td>Cell culture dish for live-cell microscopy</td></tr><tr><td>4x Laemmli Sample Buffer&amp;nbsp;</td><td>Bio-Rad</td><td>1610747</td><td></td></tr><tr><td>60 mm tissue culture dish&amp;nbsp;</td><td>Thermofisher Scientific</td><td>10099170</td><td>For tissue culture</td></tr><tr><td>Alexa Fluor 647 Phalloidin</td><td>Thermofisher Scientific</td><td>A22287</td><td>Actin stain</td></tr><tr><td>anti-ATG9A antibody</td><td>home made</td><td>STO-215</td><td>Rabbit anti N-terminal peptide ATG9A</td></tr><tr><td>anti-Rabbit IgG, peroxidase-linked&amp;nbsp;</td><td>Invitrogen</td><td>10794347/NA934-1ml</td><td>Secondary antibody for rabbit polyclonal STO-215</td></tr><tr><td>anti-RFP antibody</td><td>Evrogen</td><td>AB233</td><td>for Western Blot</td></tr><tr><td>ATG9A Monoclonal Antibody (14F2 8B1), Invitrogen</td><td>Invitrogen</td><td>15232826</td><td>Antibody for immunofluorescence</td></tr><tr><td>ATG9A-eGFP</td><td>home made</td><td></td><td>Construct which expresses tagged ATG9A</td></tr><tr><td>Bemis Parafilm</td><td>Thermofisher Scientific</td><td>11747487</td><td>self-sealing thermoplastic film&amp;nbsp;</td></tr><tr><td>Bio-Rad Protein Assay Dye Reagent Concentrate</td><td>Bio-Rad</td><td>5000006</td><td>For determining protein concentration</td></tr><tr><td>Bovine serum albumin (BSA)&amp;nbsp;</td><td>Merck</td><td>10735086001</td><td>For blocking non-specific labelling</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;.2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>/</td><td></td><td>For PBS and EBSS</td></tr><tr><td>cOmplete Protease Inhibitor Cocktail</td><td>Roche</td><td>11697498001</td><td>Supplement in lysis buffer to prevent protein degradation</td></tr><tr><td>Cy3 AffiniPure Goat Anti-Armenian Hamster IgG</td><td>Jackson ImmunoResearch</td><td>127-165-099</td><td>Secondary antibody for i14F2 8B1 antibody for mmunofluorescence</td></tr><tr><td>Cyclohexamide</td><td>Sigma Aldrich</td><td>66-81-9</td><td>To stop protein translation</td></tr><tr><td>D-Glucose</td><td>/</td><td></td><td>For EBSS</td></tr><tr><td>Digitonin&amp;nbsp;</td><td>Merck</td><td>300410</td><td>For permeabilizing cells</td></tr><tr><td>DMEM</td><td>Merck</td><td>D6546-6x500ml</td><td>For tissue culture</td></tr><tr><td>eGFP-ATG9A</td><td>home made</td><td></td><td>Construct which expresses tagged ATG9A</td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco</td><td>10270-106</td><td>Supplement for DMEM for cell culture</td></tr><tr><td>FIJI (ImageJ)</td><td>/</td><td>https:/fiji.sc/</td><td>Open source image analysis software</td></tr><tr><td>Hoechst&amp;nbsp;</td><td>Thermofisher Scientific</td><td>H3570</td><td>Stains the nucleus</td></tr><tr><td>KCl</td><td>/</td><td></td><td>For EBSS</td></tr><tr><td>L-glutamine&amp;nbsp;</td><td>Sigma</td><td>67513</td><td>For tissue culture</td></tr><tr><td>Lipofectamine 2000&amp;nbsp;</td><td>Invitrogen</td><td>11668-019</td><td>For Cell Transfection</td></tr><tr><td>LSM880 Airyscan microscope&amp;nbsp;</td><td>Zeiss</td><td>/</td><td>Confocal microscopy</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>/</td><td></td><td>For PBS</td></tr><tr><td>MgSO4.7H2O</td><td>/</td><td></td><td>For EBSS</td></tr><tr><td>Mowiol mounting solution</td><td>Millipore</td><td>475904</td><td>for permanent mounting glass coverslips</td></tr><tr><td>NaCl</td><td>/</td><td></td><td>For EBSS</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;.2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>/</td><td></td><td>For EBSS</td></tr><tr><td>NaHCO3</td><td>/</td><td></td><td>For EBSS</td></tr><tr><td>NuPAGE 3 to 8%, Tris-Acetate, 1.5 mm, Mini Protein Gels</td><td>Thermofisher Scientific</td><td>EA0378BOX</td><td>for Western Blotting</td></tr><tr><td>NuPAGE MES SDS Running Buffer (20x)</td><td>Life Tech</td><td>NP0002</td><td>for Western Blotting</td></tr><tr><td>Opti-MEM I Reduced Serum Medium</td><td>Thermofisher Scientific</td><td>31985062</td><td>For Cell Transfection</td></tr><tr><td>Paraformaldehyde&amp;nbsp;</td><td>Agar Scientific</td><td>R1026</td><td>For fixing cells</td></tr><tr><td>pcDNA3.1-mCherry-3xFlag-ATG9A</td><td>home made</td><td></td><td>Construct which expresses tagged ATG9A</td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>/</td><td></td><td>For tissue culture</td></tr><tr><td>PNGaseF</td><td>NEB</td><td>&amp;nbsp;P0710S</td><td>To remove N-linked glycans</td></tr><tr><td>Poly-D-lysine hydrobromide mol wt 70,000-150,000</td><td>Merck</td><td>P0899</td><td>For coating coverslips</td></tr><tr><td>Rapid PNGase F enzyme</td><td>NEB</td><td>P07105</td><td>To remove N-linked glycans</td></tr><tr><td>RFP-ATG9A</td><td>home made</td><td></td><td>Construct which expresses tagged ATG9A</td></tr><tr><td>Triton X-100</td><td>Thermofisher Scientific</td><td>13454259</td><td>Detergent for Cell lysis</td></tr><tr><td>Trypsin-EDTA solution&amp;nbsp;</td><td>Sigma</td><td>T4049</td><td>For tissue culture</td></tr><tr><td>Whatman Filter Paper</td><td>Merck</td><td>WHA1001325</td><td>For Western Blot and IF</td></tr><tr><td>XCell SureLock Mini-Cell Electrophoresis System</td><td>Invitrogen</td><td>EI0001</td><td>For Western Blotting</td></tr><tr><td>Zen Black edition</td><td>Zeiss</td><td>/</td><td>Used to operate the LSM 880</td></tr></tbody></table>

imaging atg9a, a multi-spanning membrane protein

Autophagy is a highly conserved pathway that the cell uses to maintain homeostasis, degrade damaged organelles, combat invading pathogens, and survive pathological conditions. A set of proteins, called ATG proteins, comprise the core autophagy machinery and work together in a defined hierarchy. Studies in recent years have improved our knowledge of the autophagy pathway. Most recently, it has been proposed that ATG9A vesicles are at the heart of autophagy, as they control the rapid de novo synthesis of an organelle called the phagophore. The study of ATG9A has proven challenging, since ATG9A is a transmembrane protein, and it is present in different membrane compartments. As such, understanding its trafficking is an important element for understanding autophagy. Here, detailed methods are presented that can be used to study ATG9A and, in particular, its localization using immunofluorescence techniques, which can be assessed and quantified. The pitfalls of transient overexpression are also addressed. The correct characterization of ATG9A function and the standardization of techniques to analyze its trafficking are crucial to further characterize the events governing autophagy initiation.

ATG9A is a transmembrane protein associated with several intracellular membrane compartments8,17,22,23,24. In basal conditions, ATG9A is mainly localized at the trans-Golgi network (TGN), as indicated by the immunofluorescence of the endogenous protein and the overlaps with GM130, a cis-Golgi marker (Figure 1A), as well as in small vesicles that partially overlap with the endocytic recycling compartment (ERC)23. ATG9A localization at the Golgi can be detected using different immunofluorescence protocols. However, the vesicular fraction of ATG9A, as well as its change of localization, in particular the increase in the vesicular pool, in response to specific stimuli such as nutrient and serum starvation, can be quite variable in intensity and difficult to visualize with conventional imaging approaches. The ratio between ATG9A localized at the Golgi and ATG9A localized to a vesicular fraction is termed the ATG9A dispersal rate. To detect changes in the ATG9A dispersal rate, for instance upon EBSS treatment, which is used to deplete both serum and amino acids, a Golgi marker such as GM130 or TGN46 and a cytoskeleton marker such as Phalloidin, which stains the cell contour25, are useful to readily quantify the ATG9A dispersal (Figure 1B). Importantly, the mean fluorescence ratio analysis can only be interpreted as a comparative measure between conditions rather than as a fixed rate of dispersal. The ratio between compartments is highly dependent on biological and non-biological factors such as the cell line used, the staining quality, or the thresholding methods applied (Figure 1B). For this reason, the researcher needs to set up a pipeline that is able to detect ATG9A Golgi enrichment in their specific experimental conditions and then extend the analysis with the same parameters to all the images in the set to be analyzed. Representative binary images and areas selected for the analysis of ATG9A mean fluorescence are shown as a guide in Figure 1B.
ATG9A harbors several transmembrane domains flanked by two relatively flexible and unstructured N- and C-terminal domains, of which the C-terminal sequence encompasses almost half the protein12. Importantly, the localization pattern of overexpressed ATG9A can be influenced by which protein end is tagged (Figure 2A). In particular, when using transient expression systems and tagging ATG9A directly on its N-terminus with a fluorescent tag (e.g., eGFP, mRFP, or derivatives), its Golgi localization can be partially compromised, with less enrichment seen in basal (i.e., fed) conditions, while the ATG9A vesicles are still readily visible (Figure 2A). Tagging ATG9A on its C-terminus seems to slightly induce larger GFP positive clusters that could be aggregated. Finally, a monomeric version of mRFP-ATG9A also shows similar fluorescent clusters of vesicles and little Golgi staining in overexpressing cells (Figure 2A).
ATG9A folds in the ER membrane before being trafficked to the Golgi and ATG9A vesicles. During its residence in the ER, ATG9A becomes modified by N-linked glycans on Asparagine 99, and then upon reaching the Golgi, it acquires complex, mature N-linked glycans1,14. This modification by glycosylation can be detected through western blot by the appearance of a double band14. Consistent with its intracellular localization, most endogenous ATG9A harbors complex N-linked glycans, and, therefore, the higher-molecular weight band is predominant, with a faint lower-molecular weight band also visible (Figure 2B). The presence of a double band is most readily seen when using Tris-acetate gels to improve the resolution of higher-molecular weight proteins (Figure 2B,&#160;control, t = 0). When the endogenous protein is subjected to PNGase F (Peptide:N-glycosidase F) treatment, which removes most of the complex N-linked glycans, the protein runs as a single band (Figure 2B, PNGase F, t = 0). Therefore, the N-linked glycosylation status of ATG9A can be used as a proxy to monitor the exiting of ATG9A from the ER to the Golgi, which is reflected by the relative ratio between the two bands.
When transfecting mRFP-ATG9A constructs transiently, the overexpressed protein initially accumulates in the ER, potentially because the trafficking machinery is unable to fold and traffic all the ATG9A, and the lower molecular weight band is predominant (Figure 2C, control t = 0). Notably, after 24 h of expression of mRFP-ATG9A, there is approximately an equal distribution between the upper and lower bands, suggesting that the mRFP-ATG9A pool is moving into the Golgi (Figure 2C,&#160;Control, t = 24). If the cells are treated with cycloheximide (CHX), which blocks de novo protein synthesis26, the folding and exit of ATG9A from the ER can be clarified. As the endogenous protein is folded, glycosylated, and resident in the Golgi, treatment with CHX does not significantly alter the ratio of the lower- and higher-molecular weight bands (Figure 2B,&#160;Control). However, using the transient expression of mRFP-ATG9A, the CHX treatment promotes the accumulation of the higher-molecular weight band (Figure 2C,&#160;Control, CHX t = 24). The higher-molecular weight overexpressed mRFP-ATG9A band collapses into the lower band after treatment with PNGase F (Figure 2C,&#160;PNGase F, t = 24). These data show that the endogenous protein rapidly acquires mature glycans, as reflected by the predominance of the higher-molecular weight band, and the CHX chase does not affect the ratio of the double bands (Figure 2B). In the case of transiently overexpressed mRFP-ATG9A, CHX treatment induces the accumulation of the upper band, indicating that more mature glycans are acquired as the ER pool folds and exits the ER to the Golgi (Figure 2C).
The addition of a linker between the ATG9A sequence and the fluorescent tags can be helpful in promoting a more physiological localization and trafficking of the protein. Fusing a 3x-FLAG sequence (24 amino acids) between an N-terminal fluorophore and ATG9A helps the overexpressed protein behave similarly to the endogenous one (Figure 3). Indeed, overexpressed mCherry-3xFLAG-ATG9A colocalizes with the Golgi marker GM130 in fed conditions (Figure 3A). Importantly, this localization and the ATG9A vesicular compartment are preserved over time, allowing the spatiotemporal study of the trafficking of ATG9A (Figure 3B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65349/65349fig01.jpg" /> 
Figure 1: Image analysis of endogenous ATG9A localization. (A) Representative immunofluorescence image of endogenous ATG9A (red), GM130 as a Golgi marker (green), and Phalloidin to visualize the actin cytoskeleton (cyan). Scale bar = 10 &#956;m. (B) Workflow of the image analysis to determine the fraction of endogenous ATG9A that localizes at the Golgi area. Scale bar = 10 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65349/65349fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65349/65349fig02.jpg" /> 
Figure 2: Analysis of fluorescently tagged-ATG9A constructs by localization and glycosylation. (A) eGFP N-terminally tagged ATG9A is less localized at the Golgi and primarily resides in the vesicles. eGFP C-terminally tagged ATG9A exhibits aggregates within the cell (some examples are marked by white arrowheads; the eGFP-ATG9A and ATG9A-eGFP are in green). mRFP N-terminally tagged ATG9A is less localized at the Golgi and primarily resides in the vesicles. N denotes the approximate location of the cell nucleus, and the mRFP-ATG9A is in red. Scale bar = 5 &#956;m. (B) Endogenous ATG9A appears as two bands when analyzed by western blot (arrowheads): an upper band (complex N-linked glycans) and a lower band (no mature N-linked glycans). Treatment with cyclohexamide (CHX) does not affect the ratio between the upper and lower bands. Treatment with PNGase F causes the disappearance of the upper band. (C) After transient transfection of mRFP-tagged ATG9A in HEK293A cells, two prominent bands are visible on western blot (arrowheads). Treatment with PNGase F causes the disappearance of the upper band. Treatment with CHX after transfection leads to increased glycosylation as the pool of transfected ATG9A is trafficked from the ER to the Golgi. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65349/65349fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65349/65349fig03.jpg" /> 
Figure 3: Analysis of mCherry-3xFLAG-ATG9A localization by immunofluorescence and live imaging. (A)&#160;Immunofluorescence experiments of HEK293A cells transiently overexpressing mCherry-3xFLAG-ATG9A and stained with the Golgi marker GM130. Scale bar = 10 &#956;m. The mCherry-3xFLAG-ATG9A is in red, and the GM130 Golgi marker is in green. (B) Montage from live-imaging experiments in HEK293A cells transiently overexpressing mCherry-3xFLAG-ATG9A. N denotes the approximate location of the nucleus. Time frame = 1 fps. Scale bar = 10 &#956;m. The mCherry-3xFLAG-ATG9A is in red. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65349/65349fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Imaging ATG9A, a Multi-Spanning Membrane Protein at JoVE.com

Author Spotlight: Imaging ATG9A, a Multi-Spanning Membrane Protein  

This protocol describes various methods that can help in the study of ATG9A biology, including immunofluorescence followed by image analysis, transient overexpression considerations, and investigating the ATG9A glycosylation status using western blot.

Imaging ATG9A, a Multi-Spanning Membrane Protein

Autophagy is a highly conserved pathway that the cell uses to maintain homeostasis, degrade damaged organelles, combat invading pathogens, and survive ...

imaging-atg9a-a-multi-spanning-membrane-protein

Nanyang Technological University

Research

JoVE Journal

Biology

1.9K Views.  The Francis Crick Institute. This protocol describes various methods that can help in the study of ATG9A biology, including immunofluorescence followed by image analysis, transient overexpression considerations, and investigating the ATG9A glycosylation status using western blot.

Video: Author Spotlight: Imaging ATG9A, a Multi-Spanning Membrane Protein  

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient association of pore-forming and auxiliary subunits, lateral diffusion, and clustering with other proteins. However, the relationship between lateral diffusion and function is poorly understood. To approach this problem, we describe how lateral mobility and activity of individual channels in supported lipid membranes can be monitored and correlated using total internal reflection fluorescence (TIRF) microscopy. Membranes are fabricated on an ultrathin hydrogel substrate using the droplet interface bilayer (DIB) technique. Compared to other types of model membranes, these membranes have the advantage of being mechanically robust and suitable for highly sensitive analytical techniques. This protocol measures Ca2+ ion flux through single channels by observing the fluorescence emission of a Ca2+-sensitive dye in close proximity to the membrane. In contrast to classical single-molecule tracking approaches, no fluorescent fusion proteins or labels, which can interfere with lateral movement and function in the membrane, are required. Possible changes in ion flux associated with conformational changes of the protein are only due to protein lateral motion in the membrane. Representative results are shown using the mitochondrial protein translocation channel TOM-CC and the bacterial channel OmpF. In contrast to OmpF, the gating of TOM-CC is very sensitive to molecular confinement and the nature of lateral diffusion. Hence, supported droplet-interface bilayers are a powerful tool to characterize the link between lateral diffusion and the function of ion channels.

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

This protocol describes how to use TIRF microscopy to track individual ion channels and determine their activity in supported lipid membranes, thereby defining the interplay between lateral membrane movement and channel function. It describes how to prepare the membranes, record the data, and analyze the results.

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient ...

Biochemistry

Imaging Protein-protein Interactions in vivo

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, preventing their identification and analysis through traditional biochemical methods such as co-immunoprecipitation. In this regard, the genetically encodable fluorescent proteins (GFP, RFP, etc.) and their associated overlapping fluorescence spectrum have revolutionized our ability to monitor weak interactions in vivo using F&ouml;rster resonance energy transfer (FRET)1-3. Here, we detail our use of a FRET-based proximity assay for monitoring receptor-receptor interactions on the endothelial cell surface.

Imaging Protein-protein Interactions in vivo

This protocol describes how to image protein-protein interactions using a FRET-based proximity assay.

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, ...

Monitoring GTPase Atlastin in Different Nucleotide Loading States

Single-Molecule FRET Imaging for Observing the Conformational Dynamics of Dynamin-Like GTPase Atlastin

The rigid-body rotation of the three-helical middle domain (3HB) relative to the GTPase domain of dynamin-like protein atlastin (ATL) is a crucial driver of homotypic membrane fusion within the endoplasmic reticulum (ER). Disruptions in this process have been associated with hereditary spastic paraplegia (HSP), a neurodegenerative disorder. Structural and biochemical studies suggest that the conformational changes in ATL are linked to GTP hydrolysis, but real-time visualization of these conformational dynamics during the GTP hydrolysis cycle remains challenging. To better understand the mechanical mechanisms behind ATL function, single-molecule F&#246;rster resonance energy transfer (smFRET) was utilized. Three specific strategies were employed to immobilize the N-terminal cytosolic region of human ATL1 (ATL1cyto) in a streptavidin-coated microfluidic chamber, facilitating the application of intramolecular and intermolecular smFRET imaging. This allowed precise monitoring of protein conformations in various nucleotide-loading states, providing direct insights into individual molecular behaviors. This method can be applied to study other mechanochemical proteins as well.

The functions of dynamin superfamily proteins depend on conformational changes coupled with GTP hydrolysis. A system is described using the single-molecule FRET (smFRET) technique to monitor the conformational dynamics of dynamin-like GTPase atlastin in different nucleotide-loading states.

The rigid-body rotation of the three-helical middle domain (3HB) relative to the GTPase domain of dynamin-like protein atlastin (ATL) is a crucial driver ...

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 ...

Imaging of Intracellular ATP in Organotypic Tissue Slices of the Mouse Brain using the FRET-based Sensor ATeam1.03YEMK

Neuronal activity in the central nervous system (CNS) evokes a high demand on cellular energy provided by the breakdown of adenosine triphosphate (ATP). A large share of ATP is needed to re-install ion gradients across plasma membranes degraded by electrical signaling of neurons. There is evidence that astrocytes - while not generating fast electrical signals themselves - undergo increased production of ATP in response to neuronal activity and support active neurons by providing energy metabolites to them. The recent development of genetically encoded sensors for different metabolites now enables the study of such metabolic interactions between neurons and astrocytes. Here, we describe a protocol for cell-type specific expression of the ATP-sensitive Fluorescence Resonance Energy Transfer- (FRET-) sensor ATeam1.03YEMK in organotypic tissue slice cultures of the mouse hippocampus and cortex using adeno-associated viral vectors (AAV). Furthermore, we demonstrate how this sensor can be employed for dynamic measurement of changes in cellular ATP levels in neurons and astrocytes upon increases in extracellular potassium and following induction of chemical ischemia (i.e., an inhibition of cellular energy metabolism).

Imaging of Intracellular ATP in Organotypic Tissue Slices of the Mouse Brain using the FRET-based Sensor ATeam1.03YEMK

We describe a protocol for cell-type specific expression of the genetically encoded FRET-based sensor ATeam1.03YEMK in organotypic slice cultures of the mouse forebrain. Furthermore, we show how to use this sensor for dynamic imaging of cellular ATP levels in neurons and astrocytes.

Neuronal activity in the central nervous system (CNS) evokes a high demand on cellular energy provided by the breakdown of adenosine triphosphate (ATP). A ...

Neuroscience

Imaging Plasma Membrane Deformations With pTIRFM

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during the fusion of vesicle and plasma membranes. These rapid and localized changes are achieved by dynamic interactions between lipids and specialized proteins that control membrane curvature. The absence of such interactions would not only have devastating consequences for vesicle fusion, but a host of other cellular functions that involve control of membrane shape. In recent years, the identity of a number of proteins with membrane-shaping properties has been determined. What remains missing is a roadmap of when, where, and how they act as fusion and content release progress.
Our understanding of the molecular events that enable membrane remodeling has historically been limited by a lack of analytical methods that are sensitive to membrane curvature or have the temporal resolution to track rapid changes. PTIRFM satisfies both of these criteria. We discuss how pTIRFM is implemented to visualize and interpret rapid, submicron changes in the orientation of chromaffin cell membranes during dense core vesicle (DCV) fusion. The chromaffin cells we use are isolated from bovine adrenal glands. The membrane is stained with a lipophilic carbocyanine dye,1,1&#39;-dioctadecyl-3,3,3&#39;,3&#39;-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate, or&#160;diD. DiD intercalates in the membrane plane with a &#34;fixed&#34; orientation and is therefore sensitive to the polarization of the evanescent field. The diD-stained cell membrane is sequentially excited with orthogonal polarizations of a 561&#160;nm laser (p-pol, s-pol). A 488&#160;nm laser is used to visualize vesicle constituents and time the moment of fusion. Exocytosis is triggered by locally perfusing cells with a depolarizing KCl solution. Analysis is performed offline using custom-written software to understand how diD emission intensity changes relate to fusion pore dilation.

Polarization-based Total Internal Reflection Fluorescence Microscopy (pTIRFM) enables real-time detection of cell membrane dynamics. This article describes the implementation of pTIRFM for the study of membrane remodeling during regulated exocytosis. The technique is generalizable to other processes in cell biology that directly or indirectly involve changes in membrane shape.

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during ...

Aip1p Dynamics Are Altered by the R256H Mutation in Actin

Mutations in actin cause a range of human diseases due to specific molecular changes that often alter cytoskeletal function. In this study, imaging of fluorescently tagged proteins using total internal fluorescence (TIRF) microscopy is used to visualize and quantify changes in cytoskeletal dynamics. TIRF microscopy and the use of fluorescent tags also allows for quantification of the changes in cytoskeletal dynamics caused by mutations in actin. Using this technique, quantification of cytoskeletal function in live cells valuably complements in vitro studies of protein function. As an example, missense mutations affecting the actin residue R256 have been identified in three human actin isoforms suggesting this amino acid plays an important role in regulatory interactions. The effects of the actin mutation R256H on cytoskeletal movements were studied using the yeast model. The protein, Aip1, which is known to assist cofilin in actin depolymerization, was tagged with green fluorescent protein (GFP) at the N-terminus and tracked in vivo using TIRF microscopy. The rate of Aip1p movement in both wild type and mutant strains was quantified. In cells expressing R256H mutant actin, Aip1p motion is restricted and the rate of movement is nearly half the speed measured in wild type cells (0.88 &#177; 0.30 &#956;m/sec in R256H cells compared to 1.60 &#177; 0.42 &#956;m/sec in wild type cells, p &#60; 0.005).

Disease-causing mutations in actin can alter cytoskeletal function. Cytoskeletal dynamics are quantified through imaging of fluorescently tagged proteins using total internal fluorescence microscopy. As an example, the cytoskeletal protein, Aip1p, has altered localization and movement in cells expressing the mutant actin isoform, R256H.

Mutations in actin cause a range of human diseases due to specific molecular changes that often alter cytoskeletal function. In this study, imaging of ...

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal transport have devastating consequences for individual neurons and their networks, and contribute to a plethora of neurological disorders. As many of these conditions involve both cell autonomous and non-autonomous mechanisms, and often display a spectrum of pathology across neuronal subtypes, methods to accurately identify and analyze neuronal subsets are imperative.
This paper details protocols to assess in vivo axonal transport of signaling endosomes and mitochondria in sciatic nerves of anesthetized mice. Stepwise instructions are provided to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice that selectively express fluorescent proteins within cholinergic motor neurons; and 2) separately or concurrently assess in vivo axonal transport of signaling endosomes and mitochondria. These complementary intravital approaches facilitate the simultaneous imaging of different cargoes in distinct peripheral nerve axons to quantitatively monitor axonal transport in health and disease.

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals.

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal ...

Imaging the Intracellular Trafficking of APP with Photoactivatable GFP

Beta-amyloid (A&#946;) is the major constituent of senile plaques found in the brains of Alzheimer&#8217;s disease patients. A&#946; is derived from the sequential cleavage of Amyloid Precursor Protein (APP) by &#946; and &#947;-secretases. Despite the importance of A&#946; to AD pathology, the subcellular localization of these cleavages is not well established. Work in our laboratory and others implicate the endosomal/lysosomal system in APP processing after internalization from the cell surface. However, the intracellular trafficking of APP is relatively understudied.
While cell-surface proteins are amendable to many labeling techniques, there are no simple methods for following the trafficking of membrane proteins from the Golgi. To this end, we created APP constructs that were tagged with photo-activatable GFP (paGFP) at the C-terminus. After synthesis, paGFP has low basal fluorescence, but it can be stimulated with 413 nm light to produce a strong, stable green fluorescence. By using the Golgi marker Galactosyl transferase coupled to Cyan Fluorescent Protein (GalT-CFP) as a target, we are able to accurately photoactivate APP in the trans-Golgi network. Photo-activated APP-paGFP can then be followed as it traffics to downstream compartments identified with fluorescently tagged compartment marker proteins for the early endosome (Rab5), the late endosome (Rab9) and the lysosome (LAMP1). Furthermore, using inhibitors to APP processing including chloroquine or the &#947;-secretase inhibitor L685, 458, we are able to perform pulse-chase experiments to examine the processing of APP in single cells.
We find that a large fraction of APP moves rapidly to the lysosome without appearing at the cell surface, and is then cleared from the lysosome by secretase-like cleavages. This technique demonstrates the utility of paGFP for following the trafficking and processing of intracellular proteins from the Golgi to downstream compartments.

While the transport of cell surface proteins is relatively easily studied, visualizing the trafficking of intracellular proteins is much more difficult. Here, we use constructs incorporating photoactivatable GFP and demonstrate a method to accurately follow the amyloid precursor protein from the Golgi apparatus to down-stream compartments and follow its clearance.

Beta-amyloid (A&#946;) is the major constituent of senile plaques found in the brains of Alzheimer&#8217;s disease patients. A&#946; is derived from the ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Imaging ATG9A, a Multi-Spanning Membrane Protein

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Chapters in this video

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

Imaging Protein-protein Interactions in vivo

Single-Molecule FRET Imaging for Observing the Conformational Dynamics of Dynamin-Like GTPase Atlastin

4D Imaging of Protein Aggregation in Live Cells

Imaging of Intracellular ATP in Organotypic Tissue Slices of the Mouse Brain using the FRET-based Sensor ATeam1.03^YEMK

Imaging Plasma Membrane Deformations With pTIRFM

Aip1p Dynamics Are Altered by the R256H Mutation in Actin

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Imaging the Intracellular Trafficking of APP with Photoactivatable GFP

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast