This work was funded by a grant from the Canadian Institute for Health Research MOP 82890 to SHP. The authors wish to thank G.H. Patterson and J. Lippincott Schwartz for the paGFP construct. SN56 cells were a gift of Dr. Jane Rylett.

1. Cell Plating and Transfection
<ol>
	<li>In a cell culture hood, plate ~300,000 to ~500,000 SN56 cells (a kind gift of Dr. Jane Rylett) on a 35 mm glass-bottom confocal dish and cover with pre-transfection media (Dulbecco&#8217;s Modified Eagle Medium, DMEM, supplemented with 10% FBS).</li>
	<li>Incubate cells overnight at 37 &#176;C in a 5% CO2 incubator to proliferate. 
	Note: Cells should be approximately 50%-70% confluent before transfection.</li>
	<li>In a cell culture hood, transfect cells with plasmids expressing a fluorescent protein tagged TGN marker (Galactosyl transferase coupled to Cyan Fluorescent Protein, GalT-CFP), a fluorescent protein tagged compartment marker (here, lysosome associated membrane protein 1, LAMP1, tagged with mRFP), and the protein of interest tagged with paGFP (&#946;APP-paGFP). (These constructs have previously been described 30)</li>
	<li>Incubate cells with transfection reagent and plasmid DNA for 24 hr at 37 &#176;C in a 5% CO2 incubator. Use a ratio of ~2 &#181;g of DNA to 5 &#181;l of transfection agent (e.g. Lipofectamine 2000) to best balance cytotoxicity and transfection efficiency. Actual concentrations may need to be optimized for different plasmids. In the experiments described here, use 1 &#181;g/confocal dish of &#946;APP-paGFP, 0.3 &#181;g/confocal dish of LAMP1-mRFP, and 0.2 &#181;g/confocal dish of GalT-CFP. 
	Note: The amount of plasmid used will depend on transfection efficiency.</li>
	<li>For neuronal cell lines: After step 1.4, in a cell culture hood, remove pre-transfection media and differentiate SN56 cells in 2 ml DMEM supplemented with 0.1% penicillin/streptomycin with 1 mM dibutyryl cyclic AMP (dbcAMP) for 24 hr.</li>
</ol>
2. Microscope Stage Preparation
<ol>
	<li>Pre-warm phosphate-buffered saline (PBS) and Hank&#8217;s Balanced Salt Solution (HBSS) to 37 &#176;C. Warm-up microscope stage before imaging to 37 &#176;C.</li>
	<li>Wash cells twice with PBS, to remove differentiation media and replace with 2 ml of HBSS at 37 &#176;C. Place cells on microscope stage warmed to 37 &#176;C. 
	Note: Cells are viable for approximately 1.5-2 hr in HBSS.</li>
	<li>Allow 5-10 min for the confocal plate temperature to equilibrate with the microscope stage.</li>
</ol>
3. Photo-activation of ROIs over the Golgi
<ol>
	<li>With the microscope eyepiece, find cells transfected with GalT-CFP, LAMP1-mRFP, and &#946;APP-paGFP.
	<ol>
		<li>Use an oil-immersion lens with high magnification (i.e. 63X or 100X). For visual conformation of fluorescence, use a filter set capable of visualizing FITC (for CFP and GFP fluorescence) and rhodamine (for mRFP fluorescence).</li>
	</ol>
	</li>
	<li>Set the confocal slice for each channel to 1 &#181;m. Take a low-resolution image.</li>
	<li>Crop to image only the cell of interest.</li>
	<li>Using the adjustment knob, manually set the focal plane to the middle of the cell. This is typically the part of the cell where the nucleus is the largest.</li>
	<li>Draw 3-5 circular Regions of Interests (ROIs) within the TGN (GalT-CFP fluorescence).
	<ol>
		<li>To draw ROIs (i.e. in the Zeiss LSM program), select the &#8216;Edit ROI&#8217; button in the command console.</li>
		<li>Select the circular ROI button.</li>
		<li>Click and drag over GalT-CFP positive regions of the cell. 
		Note: The photo-bleaching packages for many microscopes have this capability.</li>
	</ol>
	</li>
	<li>Take an image of the cell with the overlaid ROIs. Save a copy of the ROIs for later reference.
	<ol>
		<li>To save the ROIs to the image, select the &#8216;Macro&#8217; button.</li>
		<li>Press the &#8216;Load&#8217; button to start the &#8216;CopyRoisToOverlay&#8217; (file path: c:/AIM/Macros/AdvancedTimeSeries/Utility.lvb). 
		Note: A confocal microscope equipped with 475-525 nm (CFP) band pass (BP), a 500-530 nm BP (GFP), and a 560 nm long pass (LP) filter sets are required. An argon laser for 458 nm and 488 nm emission and a HeNe laser for 540 nm emission is also required.</li>
	</ol>
	</li>
	<li>Set-up the microscope for a bleach time course.
	<ol>
		<li>Set the laser diode (25 mW 405 nm laser) to maximum power. 
		Caution: Excessive photo-activation could lead to photo-toxicity or damage to cellular membranes. Use eye protection to avoid retinal damage.</li>
		<li>Set-up the microscope for 120 cycles of imaging. 
		Note: One imaging cycle consists of bleaching within the ROIs and imaging of the entire cell. This is a bleach time course.</li>
		<li>Set the microscope to bleach (photo-activate) only the ROIs for 20-30 iterations after every image. The time to image and bleach an image will vary. Set a time delay between images so each cycle is approximately 30 sec. 
		Note: The imaging portion of each cycle takes approximately 15-30 sec depending on image size. The bleaching for each ROI is about 50 msec. Bleaching of all 4 ROIs for 20 iterations will take 4 sec at the end of each bleach/image cycle.</li>
	</ol>
	</li>
	<li>Start the bleach time course.</li>
	<li>Turn off the bleaching after 15 min (30 cycles of imaging and bleaching), but continue capturing images of the cell. 
	Note: Time 0 to 15 min is the &#8216;pulse&#8217; period.</li>
	<li>Continue capturing images for 45 min (without bleaching), to follow the clearance of &#946;APP-paGFP. 
	Note: Time 15 to 45 min is the &#8216;chase&#8217; period.</li>
	<li>Using the analysis method described in step 6, co-localize the protein of interest (here, &#946;APP-paGFP) with the compartment of interest (here, LAMP1-mRFP) by making surfaces to each channel.</li>
</ol>
4. Determine the Downstream Compartment
<ol>
	<li>To determine if lysosomes (in these experiments) are the final compartment, add membrane-permeable protease inhibitors to cause proteins to accumulate in the lysosome.
	<ol>
		<li>For &#946;APP, use 0.5 &#181;M L685, 458 (specific &#947;-secretase inhibitor) overnight or 100 &#181;M chloroquine (deacidifies lysosomes) for 30 min before imaging.</li>
	</ol>
	</li>
	<li>Find cells and photoactivate as in step 3.1 &#8211; 3.8.</li>
	<li>Image the cell for an additional 45-min (without photo-activation) to determine where &#946;APP-paGFP accumulates in absence of cleavage.</li>
	<li>Analyze the delivery of protein to lysosomes (or other downstream compartment) and subsequent clearance according to the method described in step 6.</li>
</ol>
5. Ensure Accuracy of Photo-activation within the Golgi
<ol>
	<li>Warm-up HBSS to 37 &#176;C.</li>
	<li>Prepare a 2 ml solution, for every confocal plate to be imaged, of 66 &#181;M nocodazole (treatment) or DMSO (control) in HBSS.
	<ol>
		<li>For one confocal plate, pipette 2 ml of 37 &#176;C HBSS and add 7.96 &#181;l of nocodazole from a 16.60 mM nocodazole stock.</li>
		<li>As a control solution, pipette 2 ml of 37 &#176;C HBSS into a 2 ml tube and add 7.96 &#181;l of DMSO.</li>
	</ol>
	</li>
	<li>Incubate cells in HBSS with nocodazole or DMSO for 5 min before imaging.</li>
	<li>Find cells as in step 3.1.</li>
	<li>Before photoactivation, prepare the microscope to take a Z-stack after the photo-activation period.
	<ol>
		<li>Click on the &#8216;Z Stack&#8217; button. Start scanning using Fast XY.</li>
		<li>Adjust the focal plane with the microscope adjustment knob to the top of the cell. Press &#8216;Mark First&#8217; to set the first position in the stack.</li>
		<li>Adjust the focal plane with the microscope adjustment knob to the bottom (closest to the glass) of the cell. Press &#8216;Mark Last&#8217; to set the last position in the stack.</li>
		<li>In the Z stack panel, press the &#8216;Z sectioning&#8217; button. Set the &#8216;Interval&#8217; to 1 &#181;m. 
		Note: For a higher spatial resolution stack a smaller stack interval, but more images will need to be taken lengthening the imaging period for the stack.</li>
	</ol>
	</li>
	<li>Bleach the cells, as described in step 3.7-3.8. Photo-activate and image from 0 min to 15 min (30 frames).</li>
	<li>Stop imaging after 30 frames. Immediately save an image of the video. 
	Caution: Some microscopes may overwrite first video if it is not saved before starting a second imaging sequence.</li>
	<li>After saving the video, immediately acquire a Z-stack, using the parameters from step 5.5.</li>
	<li>Co-localize &#946;APP-paGFP with GalT-CFP (or other Golgi marker), as described in step 6. 
	Note: CFP photobleaches rapidly and may not be visible by the end of the imaging period. Colocalization may not be possible with GalT-CFP. If colocalization is required, use mRFP to tag GalT.</li>
</ol>
6. Vesicle Filtering and Colocalization
<ol>
	<li>Use an analysis program (e.g. Imaris) to make surfaces of the compartment (e.g. LAMP1-mRFP) and a protein of interest (e.g. &#946;APP-paGFP).</li>
	<li>Enter the &#8216;Surpass&#8217; section of the analysis program by clicking on the Surpass button on the top menu bar.</li>
	<li>Select the Surface wizard button at the top of the left most panel (5th button from the left).</li>
	<li>Select a channel to filter vesicles (i.e. &#946;APP-paGFP fluorescence, protein of interest).</li>
	<li>Perform background subtraction by submitting the diameter of the largest vesicle in the field to the wizard and press next. The analysis program automatically selects &#8212; based on a threshold derived from the largest vesicle in the field &#8212; an area in the channel of interest.
	<ol>
		<li>Manually adjust the threshold background to refine the selection if needed.</li>
	</ol>
	</li>
	<li>At the bottom of the Threshold selection screen, check the &#8216;Split touching objects&#8217; option to separate the combined Surfaces. 
	Note: If many vesicles are closely grouped together, the analysis program will group these objects under one surface.
	<ol>
		<li>Choose 10-15 representative vesicles and calculate the average vesicle diameter and enter the result into the wizard.</li>
	</ol>
	</li>
	<li>Refine the selected seed points by increasing or decreasing the threshold on &#8216;Quality&#8217; (intensity of the channel in the middle of each spot). 
	Note: Surfaces of the channel of interest will be created. Further refinement of the selected surfaces can be performed at this point. Threshold vesicles based on the number of pixels in each vesicle.</li>
	<li>Mask the original channel with this surface. To mask, select the 4th tab from the left in the surface creation wizard (pencil).</li>
	<li>Select &#8216;Mask All&#8217;. A new channel with the vesicles of interest will be created. 
	Note: During the mask process, an option to duplicate the original channel is offered. Select this option if the original data needs to be saved.</li>
	<li>Repeat steps 6.1 through 6.7 for the unfiltered channel (i.e. LAMP1 mRFP, compartment).</li>
	<li>At the top tool bar, select the colocalization tool.
	<ol>
		<li>In the top panel, as the histograms of the channels to be colocalized appear, select the recently created masked channels.</li>
		<li>Select all of available data. There are dark pixels that are sometimes present after analysis. Do not select these pixels in the histogram, they will skew the results.</li>
		<li>In the far right panel, select &#8216;Build Coloc Channel&#8217;. This creates a new channel containing only the colocalized pixels. The data will be in the same panel under the &#8216;Channel Statistics&#8217; button. The data can be exported as a .CSV file.</li>
		<li>Use the &#8216;Percent of Material colocalized&#8217; statistic. This option takes into account the intensity of the pixels and size of the object.</li>
	</ol>
	</li>
</ol>

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A., Das, U., Tang, Y., Roy, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanistic+logic+underlying+the+axonal+transport+of+cytosolic+proteins.">Mechanistic logic underlying the axonal transport of cytosolic proteins.</a> Neuron. 70 (3), 441-454 (2011).</li><li>Herl, L., 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=Mutations+in+amyloid+precursor+protein+affect+its+interactions+with+presenilin/gamma-secretase.">Mutations in amyloid precursor protein affect its interactions with presenilin/gamma-secretase.</a> Mol. Cell. Neurosci. 41 (2), 166-174 (2009).</li><li>Lorenzen, 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=Rapid+and+Direct+Transport+of+Cell+Surface+APP+to+the+Lysosome+defines+a+novel+selective+pathway.">Rapid and Direct Transport of Cell Surface APP to the Lysosome defines a novel selective pathway.</a> Mol. Brain. 3 (1), 11 (2010).</li><li>Thinakaran, G., Teplow, D. B., Siman, R., Greenberg, B., Sisodia, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metabolism+of+the+'Swedish'+amyloid+precursor+protein+variant+in+neuro2a+(N2a)+cells.+Evidence+that+cleavage+at+the+beta-secretase";site+occurs+in+the+golgi+apparatus.">Metabolism of the 'Swedish' amyloid precursor protein variant in neuro2a (N2a) cells. Evidence that cleavage at the beta-secretase";site occurs in the golgi apparatus.</a> J. Biol. Chem. 271 (16), 9390-9397 (1996).</li><li>Thinakaran, G., Koo, E. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amyloid+Precursor+Protein+Trafficking,+Processing,+and+Function.">Amyloid Precursor Protein Trafficking, Processing, and Function.</a> J. Biol. Chem. 283 (44), 29615-29619 (2008).</li><li>Schneider, M., Barozzi, S., Testa, I., Faretta, M., Diaspro, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-Photon+Activation+and+Excitation+Properties+of+PA-GFP+in+the+720-920-nm+Region.">Two-Photon Activation and Excitation Properties of PA-GFP in the 720-920-nm Region.</a> Biophys. J. 89 (2), 1346-1352 (2005).</li><li>Sevier, C. S., Weisz, O. A., Davis, M., Machamer, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+export+of+the+vesicular+stomatitis+virus+G+protein+from+the+endoplasmic+reticulum+requires+a+signal+in+the+cytoplasmic+tail+that+includes+both+tyrosine-based+and+di-acidic.">Efficient export of the vesicular stomatitis virus G protein from the endoplasmic reticulum requires a signal in the cytoplasmic tail that includes both tyrosine-based and di-acidic.</a> 11 (1), 13-22 (2000).</li><li>Muresan, V., Varvel, N. H., Lamb, B. T., Muresan, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+cleavage+products+of+amyloid-beta+precursor+protein+are+sorted+to+distinct+carrier+vesicles+that+are+independently+transported+within+neurites.">The cleavage products of amyloid-beta precursor protein are sorted to distinct carrier vesicles that are independently transported within neurites.</a> J. Neurosci. 29, 3565-3578 (2009).</li></ol>

The authors have no competing financial interests or conflicts of interest.

This technique describes the accurate photo-activation of membrane proteins tagged with paGFP in the TGN to visualize subsequent trafficking and cleavage. While paGFP was created over a decade ago 25, this is the first example of accurate photo-activation to follow the trafficking of nascent proteins to downstream compartments. Previous studies using APP-paGFP constructs photo-activated a peri-nuclear region of the cell, which was considered to be the Golgi 11. However, as evident in our micrographs...

The hallmark of Alzheimer&#8217;s disease (AD) is the presence of senile plaques and neurofibrillary tangles (NFTs) in the brain. The major constituent of senile plaques is &#946;-amyloid (A&#946;). A&#946; is derived from its precursor; amyloid precursor protein (APP) 1. The amyloidogenic cleavage of APP begins with removal of the ectodomain from APP by &#946;-secretase 2. The remaining 99-residue carboxyl terminal fragment (CTF) can be cleaved by &#947;-secretase to produce A&#946; 3-7. While many experiments have documented the cleavage of cell surface APP after internalization from the cell surface into the endosomal/ lysosomal system, a number of recent studies have suggested that intracellular trafficking of APP is also important in regulating its processing 8-11.
There have been a number of attempts at modulating the levels of A&#946; with &#947;-secretase inhibitors and A&#946; immunotherapies. However, recent clinical trials with these therapies showed no benefit, and, in some cases, caused harm 12. An unexploited strategy to modulate A&#946; production is to alter the sub-cellular localization of APP and &#947;-secretase interaction. The Golgi, plasma membrane, and endosomes/lysosomes have all been suggested as possible locales for &#947;-cleavage of APP. Research from our laboratory suggests that APP and the &#947;-secretase are resident proteins of the lysosomal membrane 13. Furthermore, we have found that the lysosomal &#947;-secretase has an acidic optimal pH 13. In addition, alkalization of the endosomal/lysosomal system with chloroquine or NH4Cl, has been shown to decrease the production of A&#946; 14. &#947;-secretase inhibition or knockout or Presenilin leads to APP-CTFs accumulation in the lysosome 15-17. Moreover, disrupting APP endocytosis lowers A&#946; production 18-20.
Despite the importance of the Golgi as a sorting station for nascent proteins and proteins recycled from the endosomal/lysosomal system, the intracellular trafficking of APP has not been studied in detail 21. Recent work has shown that APP can be recycled to the trans-Golgi network (TGN) via interaction with the retromer complex. Down regulation of the retromer complex decreases A&#946; production 8,22-24. However, the egress of APP from the Golgi has not been well studied.
While following the endocytosis of cell-surface proteins, such as the transferrin receptor, are easily labeled and followed, following the trafficking of intracellular proteins is more challenging. In fact, few proteins have had their intracellular trafficking imaged. The advent of fluorescent protein tags, such as photo-activatable-GFP (paGFP), has provided new tools to examine intracellular trafficking. Photo-activatable-GFP is a form of GFP that is nearly invisible after synthesis, but develops strong green (GFP) fluorescence after being activated by 413 nm laser light and this signal is stable for days 25,26. Constructs using paGFP have been used to demonstrate the intralysosomal trafficking of the Lysosomal Protein membrane protein 1 (LAMP1) 25, the cell surface delivery of the Vesicular Stomatitis Virus Glycoprotein (VSVG, a marker of the secretory pathway) 27, and the turnover of peroxisomes 28 and autophagosomes 29.
In order to visualize the sorting of APP from the TGN in live cells, we designed plasmid expressing the last 112 amino acids of APP coupled to photo-activatable (paGFP) on the C-terminal (referred to as &#946;APP-paGFP) 30. We have also performed these experiments with full length APP and achieved similar results; the &#914;APP construct is used here because it provides brighter images. We then photo-activate &#946;APP-paGFP only in the TGN, as demarcated by the TGN marker Galactosyltransferase (GalT). Although APP has been tagged with paGFP to visualize rapid axonal transport of APP and clearance from the perinuclear region, this is the first demonstration of APP trafficking from one carefully defined compartment to another 11,31,32. Here, we demonstrate accurate photo-activation within the TGN and the egress of APP into downstream compartments and subsequent cleavage and clearance from lysosomes 30. The accurate photo-activation within the Golgi is widely applicable to other protein systems.

<table><tbody><tr><td>35 mm glass bottom culture dishes&amp;nbsp;</td><td>Matek</td><td>P-35G-1.5-20-C</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline</td><td>Life Technologies</td><td>14190-144</td><td></td></tr><tr><td>Hanks Balanced Salt Solution</td><td>Life Technologies</td><td>14025-092</td><td></td></tr><tr><td>Lipofectamine 2000</td><td>Life Technologies</td><td>11668019</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium</td><td>Life Technologies</td><td>11995-092</td><td></td></tr><tr><td>Penicilin/Streptomycin</td><td>Life Technologies</td><td>15140-122</td><td></td></tr><tr><td>dibutyrl cyclic AMP</td><td>Sigma&amp;nbsp;</td><td>D0627</td><td></td></tr><tr><td>Heat in activated Fetal Bovine Serum</td><td>Life Technologies</td><td>10082147</td><td></td></tr><tr><td>Heated Microscopy stage insert P</td><td>PeCon GmbH</td><td></td><td></td></tr><tr><td>Tempcontrol 37&amp;ndash;2 digital 2-channel</td><td>PeCon GmbH</td><td></td><td></td></tr><tr><td>Zeiss LSM-510 META laser- scanning microscope&amp;nbsp;</td><td>Carl Zeiss</td><td></td><td>with laser diode, argon laser, and HeNe1 laser</td></tr><tr><td>Zeiss 63X 1.4 numerical aperture oil immersion lens</td><td>Carl Zeiss</td><td></td><td></td></tr><tr><td>L685, 458&amp;nbsp;</td><td>EMD Millipore</td><td>565771</td><td>dissolved in DMSO</td></tr><tr><td>cholorquine&amp;nbsp;</td><td>Sigma</td><td>C6628&amp;nbsp;</td><td>dissolved in water</td></tr><tr><td>Nocodazole</td><td>Sigma</td><td>M1404</td><td>dissolved in DMSO</td></tr><tr><td>Dimethyl sulphoxide&amp;nbsp;</td><td>Sigma</td><td>472301</td><td></td></tr><tr><td>Imaris&amp;nbsp;</td><td>Bitplane</td><td></td><td>with colocalization package&amp;nbsp;</td></tr></tbody></table>

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.

Typical results show &#946;APP leaving the TGN and appears to traffic rapidly to LAMP1 (Figure 1a and b). During the photo-activation period, vesicles can be seen departing the Golgi destined for lysosomes (Figure 1b). Without inhibitor treatment, paGFP fluorescence will be visible in lysosomes while there is photo-activation in the TGN. After stopping photoactivation, the &#946;APP-paGFP is rapidly cleared from the lysosome (compare Figure 1a to <strong...

Watch this Scientific Journal Video about Imaging the Intracellular Trafficking of APP with Photoactivatable GFP at JoVE.com

Imaging the Intracellular Trafficking of APP with Photoactivatable GFP

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 ...

imaging-the-intracellular-trafficking-of-app-with-photoactivatable-gfp

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JoVE Journal

Biology

11.7K Views.  Western University. 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.

Video: Imaging the Intracellular Trafficking of APP with Photoactivatable GFP

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

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

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

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

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Neuroscience

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane proteins become exposed at the cell surface. One of these proteins is Synaptotagmin-1 (Syt1). An antibody directed against the luminal domain of Syt1, once added to the culture medium, is taken up during the exo-endocytotic cycle. This uptake is proportional to the amount of SV recycling and can be quantified through immunofluorescence. Here, we combine Syt1 antibody uptake with double transfection of cultured hippocampal neurons. This allows us to (1) localize presynaptic sites based on expression of recombinant presynaptic marker Synaptophysin, (2) determine their functionality using Syt1 uptake, and (3) characterize the targeting and effects of a protein of interest, GFP-Rogdi.

We describe an optical assay for synaptic vesicle (SV) recycling in cultured neurons. Combining this protocol with double transfection to express a presynaptic marker and protein of interest allows us to locate presynaptic sites, their synaptic vesicle recycling capacity, and determine the role of the protein of interest.

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane ...

Visualizing Axonal Growth Cone Collapse and Early Amyloid &#946; Effects in Cultured Mouse Neurons

Amyloid-&#946; (A&#946;) causes memory impairments in Alzheimer's disease (AD). Although therapeutics have been shown to reduce A&#946; levels in the brains of AD patients, these do not improve memory functions. Since A&#946; aggregates in the brain before the appearance of memory impairments, targeting A&#946; may be inefficient for treating AD patients who already exhibit memory deficits. Therefore, downstream signaling due to A&#946; deposition should be blocked before AD development. A&#946; induces axonal degeneration, leading to the disruption of neuronal networks and memory impairments. Although there are many studies on the mechanisms of A&#946; toxicity, the source of A&#946; toxicity remains unknown. To help identify the source, we propose a novel protocol that uses microscopy, gene transfection, and live cell imaging to investigate early changes caused by A&#946; in axonal growth cones of cultured neurons. This protocol revealed that A&#946; induced clathrin-mediated endocytosis in axonal growth cones followed by growth cone collapse, demonstrating that inhibition of endocytosis prevents A&#946; toxicity. This protocol will be useful in studying the early effects of A&#946; and may lead to more efficient and preventative AD treatment.

Here a protocol to investigate the early effects of amyloid-&#946; (A&#946;) in the brain is presented. This shows that A&#946; induces clathrin-mediated endocytosis and collapse of axonal growth cones. The protocol is useful in studying early effects of A&#946; on axonal growth cones and may facilitate prevention of Alzheimer's disease.

Amyloid-&#946; (A&#946;) causes memory impairments in Alzheimer's disease (AD). Although therapeutics have been shown to reduce A&#946; levels in the ...

A High-content Assay for Monitoring AMPA Receptor Trafficking

Postsynaptic trafficking of receptors to and from the cell surface is an important mechanism by which neurons modulate their responsiveness to different stimuli. The &#945;-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors, which are responsible for fast excitatory synaptic transmission in neurons, are trafficked to and from the postsynaptic surface to dynamically alter neuronal excitability. AMPA receptor trafficking is essential for synaptic plasticity and can be disrupted in neurological disease. However, prevalent approaches for quantifying receptor trafficking ignore entire receptor pools, are overly time- and labor-intensive, or potentially disrupt normal trafficking mechanisms and therefore complicate the interpretation of resulting data. We present a high-content assay for the quantification of both surface and internal AMPA receptor populations in cultured primary hippocampal neurons using dual fluorescent immunolabeling and a near-infrared fluorescent 96-well microplate scanner. This approach facilitates the rapid screening of bulk internalized and surface receptor densities while minimizing sample material. However, our method has limitations in obtaining single-cell resolution or conducting live cell imaging. Finally, this protocol may be amenable to other receptors and different cell types, provided proper adjustments and optimization.

Neuronal excitability can be modulated through a dynamic process of endo- and exocytosis of excitatory ionotropic glutamate receptors. Described here is an accessible, high-content assay for quantifying surface and internal receptor population pools.

Postsynaptic trafficking of receptors to and from the cell surface is an important mechanism by which neurons modulate their responsiveness to different ...

Visualizing and Analyzing Intracellular Transport of Organelles and Other Cargos in Astrocytes

Astrocytes are among the most abundant cell types in the adult brain, where they play key roles in a multiplicity of functions. As a central player in brain homeostasis, astrocytes supply neurons with vital metabolites and buffer extracellular water, ions, and glutamate. An integral component of the &#8220;tri-partite&#8221; synapse, astrocytes are also critical in the formation, pruning, maintenance, and modulation of synapses. To enable these highly interactive functions, astrocytes communicate among themselves and with other glial cells, neurons, the brain vasculature, and the extracellular environment through a multitude of specialized membrane proteins that include cell adhesion molecules, aquaporins, ion channels, neurotransmitter transporters, and gap junction molecules. To support this dynamic flux, astrocytes, like neurons, rely on tightly coordinated and efficient intracellular transport. Unlike neurons,&#160;where intracellular trafficking has been extensively delineated, microtubule-based transport in astrocytes has been less studied. Nonetheless, exo- and endocytic trafficking of cell membrane proteins and intracellular organelle transport orchestrates astrocytes&#8217; normal biology, and these processes are often affected in disease or in response to injury. Here we present a straightforward protocol to culture high quality murine astrocytes, to fluorescently label astrocytic proteins and organelles of interest, and to record their intracellular transport dynamics using time-lapse confocal microscopy. We also demonstrate how to extract and quantify relevant transport parameters from the acquired movies using available image analysis software (i.e., ImageJ/FIJI) plugins.

Here we describe an in vitro live-imaging method to visualize intracellular transport of organelles and trafficking of plasma membrane proteins in murine astrocytes. This protocol also presents an image analysis methodology to determine cargo transport itineraries and kinetics.

Astrocytes are among the most abundant cell types in the adult brain, where they play key roles in a multiplicity of functions. As a central player in ...

Visualizing Proteins in Primary Hippocampal Neurons by Live&#45;Cell Imaging

Using Live&#45;Cell Imaging to Measure the Effects of Pathological Proteins on Axonal Transport in Primary Hippocampal Neurons

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal transport is disrupted in multiple neurodegenerative diseases, and projection neurons are particularly vulnerable because of the need to transport cellular materials over long distances and sustain substantial axonal mass. Pathological modifications of several disease-related proteins negatively affect transport, including tau, amyloid-&#946;, &#945;-synuclein, superoxide dismutase, and huntingtin, providing a potential common mechanism by which pathological proteins exert toxicity in disease. Methods to study these toxic mechanisms are necessary to understand neurodegenerative disorders and identify potential therapeutic interventions.
Here, cultured primary rodent hippocampal neurons are co-transfected with multiple plasmids to study the effects of pathological proteins on fast axonal transport using live-cell confocal imaging of fluorescently-tagged cargo proteins. We begin with the harvest, dissociation, and culturing of primary hippocampal neurons from rodents. Then, we co-transfect the neurons with plasmid DNA constructs to express fluorescent-tagged cargo protein and wild-type or mutant tau (used as an exemplar of pathological proteins). Axons are identified in live cells using an antibody that binds an extracellular&#160;domain of neurofascin, an axon initial segment protein, and an axonal region of interest is imaged to measure fluorescent cargo transport.
Using KymoAnalyzer, a freely available ImageJ macro, we extensively characterize the velocity, pause frequency, and directional cargo density of axonal transport, all of which may be affected by the presence of pathological proteins. Through this method, we identify a phenotype of increased cargo pause frequency associated with the expression of pathological tau protein. Additionally, gene-silencing shRNA constructs can be added to the transfection mix to test the role of other proteins in mediating transport disruption. This protocol is easily adaptable for use with other neurodegenerative disease-related proteins and is a reproducible method to study the mechanisms of how those proteins disrupt axonal transport.

Author Spotlight: Understanding the Impact of Pathological Proteins on Axonal Transport in Neurodegenerative Diseases

Here, we demonstrate how to combine transfection of primary hippocampal rodent neurons with live-cell confocal imaging to analyze pathological protein-induced effects on axonal transport and identify mechanistic pathways mediating these effects.

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal ...

Live Cell Imaging and 3D Analysis of Angiotensin Receptor Type 1a Trafficking in Transfected Human Embryonic Kidney Cells Using Confocal Microscopy

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in transfected human embryonic kidney-293 (HEK) cells following stimulation with angiotensin II (Ang II). HEK cells are transiently transfected with plasmid DNA containing AT1aR tagged with enhanced green fluorescent protein (EGFP). Lysosomes are identified with a red fluorescent dye. Live-cell images are captured on a laser scanning confocal microscope after Ang II stimulation and analyzed by software in three dimensions (3D, voxels) over time. Live-cell imaging enables investigations into receptor trafficking and avoids confounds associated with fixation, and in particular, the loss or artefactual displacement of EGFP-tagged membrane receptors. Thus, as individual cells are tracked through time, the subcellular localization of receptors can be imaged and measured. Images must be acquired sufficiently rapidly to capture rapid vesicle movement. Yet, at faster imaging speeds, the number of photons collected is reduced. Compromises must also be made in the selection of imaging parameters like voxel size in order to gain imaging speed. Significant applications of live-cell imaging are to study protein trafficking, migration, proliferation, cell cycle, apoptosis, autophagy and protein-protein interaction and dynamics, to name but a few.

Here we present a protocol to image cells expressing green fluorescent protein-tagged angiotensin type 1a receptors during endocytosis initiated by angiotensin II treatment. This technique includes labeling lysosomes with a second fluorescent marker, and then utilizing software to analyze the co-localization of receptor and lysosome in three dimensions over time.

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

Live Fluorescence, Inverse Imaging of Cell Ruffling, and Macropinocytosis

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase nutrients and other material in cells. The dramatic extension of cell surface ruffles, their closure to form macropinosomes, and the maturation of internalized macropinosomes are key events in this pathway that can be difficult to capture using conventional confocal imaging based on tracking a bolus of fluorescent cargo. Fluorescent dextrans are commonly used experimentally as fluid phase markers for macropinosomes and for other endocytic pathways. A method the lab has adopted to optimize the imaging of dextran uptake involves using live imaging of cells bathed in high concentrations of fluorescent dextran in the medium, with the unlabeled cells appearing in relief (as black). The cell ruffles are highlighted to visualize ruffle closure, and internalized macropinosomes appear as fluorescent vacuoles in the cell interior. This method is optimal for visualizing macropinosome features and allows for easy segmentation and quantification. This paper describes dual-labeling of pathways with different sized dextrans and the co-expression of lipid probes and fluorescent membrane proteins to demark macropinosomes and other endosomes. The detection of internalized dextran at an ultrastructural level using correlative light and electron microscopy (CLEM) is also demonstrated. These cell processes can be imaged using multiple live imaging modalities, including in 3D. Taken together, these approaches optimize macropinosome imaging for many different settings and experimental systems.

This protocol demonstrates dextran imaging in live cells using continuous uptake and inverse images to optimize visualization of ruffling, macropinosome maturation, and analysis of dextran and other cell labelings.

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase ...

Imaging the Intracellular Trafficking of APP with Photoactivatable GFP

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