This work was supported in part by a MOST 111-2311-B-001-019-MY3 research grant from the National Science and Technology Council in Taiwan.

1. Preparation of cells expressing diKillerRed or self-labeling proteins (SLPs) in the peroxisome lumen
<ol>
	<li>Seed the desired cells on glass-bottomed cell culture dishes. For the experiment here, seed 2 x 105 human SHSY5Y cells in 840 &#181;L of culture medium (DMEM/F-12 supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin) or 6 x 104 mouse NIH3T3 cells in 840 &#181;L of culture medium (DMEM supplemented with 10% bovine serum and 1% penicillin/streptomycin) on a 35 mm culture dish with a 20 mm diameter glass microwell. 
	NOTE: The number of cells seeded should be adjusted proportionally according to the glass microwell area when using glass bottom dishes of other specifications.</li>
	<li>Grow the cells under 5% CO2/37 &#176;C for 24 h.</li>
	<li>Transfect the cells with desired plasmids using a transfection reagent.
	<ol>
		<li>Use PMP34-TagBFP to label the peroxisomes in live cells. Use either peroxisome-targeting diKillerRed-VKSKL or SLP-VKSKL (+ dye-labeled SLP ligands) for ROS generation in the peroxisome lumen (through light-activated ROS production). In this protocol, we use the self-labeling protein HaloTag-VKSKL21. To monitor the translocation of Stub1/Hsp70, the accumulation of ubiquitinated proteins, and the formation of autophagosomes on ROS-stressed peroxisomes, transfect EGFP-Stub1, EGFP-Hsp70, EGFP-Ub, or EGFP-LC3B, respectively. To check ROS generation in the peroxisomal lumen, co-transfect diKillerRed-VKSKL with the peroxisome-localized fluorescent redox probe roGFP2-VKSKL22.</li>
		<li>For SHSY5Y cell transfection, dilute plasmids in 55 &#181;L of serum-free DMEM/F-12, and dilute 1 &#181;L of transfection reagent in 55 &#181;L of serum-free DMEM/F12. Combine the diluted DNA with the diluted reagent. Mix gently, and incubate for 30 min at room temperature.</li>
		<li>Add the complex to the 20 mm microwell previously plated with 100 &#181;L of culture medium for SHSY5Y without antibiotics (DMEM/F-12 with 10% fetal bovine serum). Gently shake the dish back and forth.</li>
		<li>For NIH3T3 cell transfection, substitute reduced serum medium for the serum-free DMEM/F-12 for diluting the plasmids and transfection reagent. Replace the plating culture medium with SHSY5Y with culture medium for NIH3T3 cells without antibiotics (DMEM with 10% bovine serum).</li>
	</ol>
	</li>
	<li>After 2 h of transfection, remove the transfection reagent-containing medium, and add 1 mL of fresh culture medium. Incubate the NIH3T3 or SHSY5Y cells at 37&#176;C and 5% CO2 for 24 h. 
	NOTE: Other cell lines can also be used to study Stub1-mediated pexophagy (e.g., HeLa cells).</li>
</ol>
2. Staining peroxisomes with dye-labeled SLP ligands (for light-activated ROS production)
<ol>
	<li>At 24 h post-transfection, remove the culture medium, and add 100 &#181;L of 200 nM HaloTag TMR ligand or 200 nM Janelia Fluor 646 HaloTag ligand onto the 20 mm glass microwell. 
	NOTE: The dye-labeled SLP ligands should be diluted with the respective culture medium for each cell line. Choosing Janelia Fluor 646-labeled ligands will free up the blue, green, and red channels for downstream fluorescence imaging.</li>
	<li>Transfer the dish to a 37&#176;C, 5% CO2 incubator. Stain for 1 h. After 1 h, remove the staining medium thoroughly, and add 1 mL of fresh culture medium.</li>
</ol>
3. ROS-stressing of peroxisomes on a laser-scanning confocal microscope
<ol>
	<li>Place a glass-bottomed dish containing the desired cells on a laser-scanning confocal microscope equipped with a stage-top incubator.</li>
	<li>Click on Tool Window, choose Ocular, and click on the DIA button (Figure 2A) to turn on the transmitted light. View the dish through the microscope eyepiece, and bring the cells into focus by turning the focus knob.</li>
	<li>Choose LSM, and click on the Dye &#38; Detector Select button (Figure 2B). Double-click on the desired dyes in the pop-up window to select the appropriate laser and filter settings for imaging the cells (Figure 2C). Click on Livex4 in the Live panel to initiate fast laser scanning to start screening for the fluorescence signals of the cells (Figure 2D).</li>
	<li>Move the stage around using the joystick controller, and locate the medium diKillerRed-VKSKL- or SLP-VKSKL-expressing cells to apply ROS stress (on the peroxisomes). Acquire an image of the desired cell.</li>
	<li>Click on Tool Window, and choose LSM Stimulation (Figure 2E). Click on the ellipse button, and in the live image window, use the mouse to draw a circular ROI of 15 &#181;m in diameter (Figure 2F) containing the desired peroxisomes acquired in step 3.4 to which ROS stress will be applied (see Figure 3 for an example).</li>
	<li>To apply ROS stress, use 561 nm for cells with the diKillerRed-PTS1- or TMR-labeled SLP ligand, and use 640 nm for cells stained with the Janelia Fluor 646-labeled SLP ligand.</li>
	<li>Select the laser wavelength for applying the ROS stress. Enter the desired laser percentage and the duration (Figure 2G).</li>
	<li>Click on Acquire, and click on the Stimulation button (Figure 2H) to initiate the scanning with 561/640 nm light through the ROIs for 30 s each. This corresponds to a ~5% laser power setting for both 561 nm and 640 nm on the confocal microscope used here. This operation will lead to ROS production in the peroxisomes. 
	NOTE: One can select ROIs of different sizes. One should proportionally adjust the illumination time for each ROI according to its area.</li>
	<li>After 30 s of laser illumination, peroxisomes within the ROIs will have lost their diKillerRed-PTS1/TMR/Janelia Fluor 646 signals. This signal loss allows one to distinguish the illuminated from the non-illuminated peroxisomes (damaged vs. healthy) within a cell.</li>
</ol>
4. Monitoring pexophagy in live cells by time-lapse imaging
<ol>
	<li>Follow the translocation of EGFP-tagged Stub1, Hsp70, Ub, p62, or LC3B onto the illuminated/ROS-stressed peroxisomes by time-lapse imaging using 10 min intervals between frames (for examples, see Figures 3-7). 
	NOTE: EGFP-Stub1 or EGFP-Hsp70 signals start to appear 10-20 min post illumination and stay on all the damaged peroxisomes until ~40 min post illumination. The EGFP-Ub signals appear 30 min post illumination and last for 2-3 h. EGFP-p62 translocation appears 60 min post illumination, and EGFP-LC3B signals become visible ~3 h after illumination.</li>
	<li>Quantify the EGFP signals on the damaged peroxisomes (from Stub1, Ub, p62, or LC3B) using ImageJ. Perform the following steps for using ImageJ to quantify a three-channel image (PMP34-TagBFP/EGFP-LC3B/HaloTag-VKSKL).
	<ol>
		<li>Open the image, and apply Elliptical selections or Freehand selections to enclose the region containing all the illuminated peroxisomes (PMP34-TagBFP signal positive and HaloTag ligand signal bleached; Figure 8A).</li>
		<li>Click on Duplicate to pick out the PMP34-TagBFP channel (Figure 8A). Set a threshold using the drop-down menu Image &#62; Adjust &#62; Threshold to mark the peroxisomes (Figure 8B).</li>
		<li>Select Analyze &#62; Analyze particles to determine the exact regions of the illuminated peroxisomes. ImageJ records these regions of interest (ROIs) in the ROI manager (Figure 8C).</li>
		<li>Click on Duplicate to pick out the EGFP channel for analyzing the fluorescence signal from EGFP-conjugated Ub, p62, or LC3B (Figure 8D). Select all the ROIs in the ROI Manager (Figure 8E).</li>
		<li>Apply Measure in the ROI Manager to measure the EGFP signals on the peroxisomes (Figure 8E). Find the value of the area and integrated density (the sum of all the pixel intensities) for each ROI in the Results table (Figure 8E).</li>
		<li>To obtain the average EGFP fluorescence intensities at different time points after illumination, divide the integrated EGFP intensity by the total area of illuminated peroxisomes (PMP34-TagBFP signal positive and HaloTag ligand signal negative areas). To obtain the accumulated signal on the peroxisomes, subtract the averaged intensity at a time point by the averaged intensity at time = 0, and then normalize by the averaged intensity at time = 0.</li>
	</ol>
	</li>
</ol>
5. Globally damaging all the peroxisomes (in every cell) on a culture dish
<ol>
	<li>Perform step 1 (preparation of cells expressing peroxisome-targeted diKillerRed [diKillerRed-VKSKL] or SLP [SLP-VKSKL]) and step 2 (staining peroxisomes with the SLP ligand [for light-activated ROS production]).</li>
	<li>After 1 h of TMR-labeled ligand staining, remove the staining medium, and add 1 mL of culture medium containing 30 mM 3-AT (3-amino-1,2,4-triazole; a catalase inhibitor). 
	NOTE: Catalase is a ROS scavenger expressed in the peroxisome lumen, so 3-AT treatment helps to augment the light-elicited oxidative damage on the peroxisomes.</li>
	<li>Place the culture dish above an LED (555-570 nm). Illuminate all the cells with a power density of 1.8 W/cm2 for 9 h in an incubator. 
	NOTE: To perform this step outside an incubator, place the culture dish on a 37&#176;C heating plate. Add 20 mM HEPES to the 3-AT-containing medium, and cover the culture dish with a transparent film to minimize gas exchange. HEPES is a buffering agent that maintains physiological pH in cell culture outside a CO2 incubator during LED illumination.</li>
	<li>Validate the success of LED-elicited damage by imaging the EGFP-Ub, EGFP-p62, or EGFP-LC3B signals on the peroxisomes. Using the illumination condition outlined above, 82% of SHSY5Y cells stably expressing HaloTag-VKSKL displayed Stub1-mediated pexophagy.</li>
	<li>Harvest the cells for downstream biochemical analysis.</li>
</ol>

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The authors declare no competing financial interests.

This protocol details how to trigger Stub1-mediated pexophagy within cell cultures by elevating peroxisomal ROS levels with light. As the protocol relies on dye-assisted ROS generation, one needs to ensure sufficient expression of diKillerRed-VKSKL or dye-labeled SLP ligand staining within the cells of interest. Given that different cell types or cells of different genetic backgrounds can harbor peroxisomes with slightly different properties, one may need to tune the exact illumination conditions to ensure the induction ...

Peroxisomes are single-membrane-bound organelles present in most eukaryotic cells. Peroxisomes are a metabolic compartment essential for carrying out the beta-oxidation of very long-chain fatty acids, purine catabolism, and ether phospholipid and bile acid synthesis1. Peroxisome-derived acetyl-CoA controls lipid homeostasis by regulating the central signaling in metabolism2. Therefore, it is no surprise that compromised peroxisomal functions are implied in various diseases, including neurodegenerative disorders, aging, cancers, obesity, and diabetes3,4,5. An essential process in the maintenance of peroxisomal operation is pexophagy. Pexophagy is a catabolic process for the selective turnover of peroxisomes by autophagy. Cells use pexophagy to help control the quantity and quality of peroxisomes, thereby ensuring proper peroxisomal function. A recent study demonstrated that peroxisome loss caused by mutations in PEX1 peroxisomal biogenesis factors 1 and 6 results from uncontrolled pexophagy6. Notably, 65% of all peroxisome biogenesis disorder (PBD) patients harbor deficiencies in the peroxisomal AAA ATPase complex, composed of PEX1, PEX6, and PEX26 in mammalian cells7.
A number of methods can be used to initiate and study pexophagy. In yeast, pexophagy is triggered when the supplied nutrients are switched from peroxisome-dependent carbon sources to peroxisome-independent carbon sources (to lower cellular peroxisome numbers)8. For example, the transfer of methanol-grown Pichia pastoris cells from methanol medium to glucose medium and ethanol medium induces micropexophagy and macropexophagy, respectively8,9,10. Micropexophagy sequesters clustered peroxisomes for degradation by remodeling the vacuole to form cup-like vacuolar sequestration membranes and a lid-like structure termed the micropexophagy-specific membrane apparatus (MIPA). In macropexophagy, individual peroxisomes are engulfed by double-membrane structures known as pexophagosomes, followed by fusion with the vacuole for degradation8,9,10. The phosphorylation of pexophagy receptors, such as Atg36p in Saccharomyces cerevisiae and Atg30p in Pichia pastoris, is critical for the receptors to recruit core autophagy machinery and to facilitate peroxisomal targeting to autophagosomes8,11.
In mammalian cells, pexophagy can be induced by ubiquitination. Tagging the peroxisomal membrane proteins PMP34 or PEX3 with ubiquitin on the cytosolic side induces pexophagy12. The overexpression of PEX3 induces peroxisome ubiquitination and peroxisome elimination by lysosomes13. In addition, the fusion of PEX5 with a C-terminal EGFP impairs the export of monoubiquitinated PEX5 and results in pexophagy14. On the other hand, pexophagy can also be triggered by H2O2 treatment. Peroxisomes produce reactive oxygen species (ROS); specifically, the peroxisomal enzyme Acox1, which catalyzes the initial step of beta-oxidation of very long-chain fatty acids (&#62; 22 carbon), produces not only acetyl-CoA but also peroxisomal ROS. In response to the heightened ROS levels under H2O2 treatment, mammalian cells activate pexophagy to lower the ROS production and alleviate stress. It has been reported that H2O2 treatment drives the recruitment of ataxia-telangiectasia mutated (ATM) to peroxisomes. ATM then phosphorylates PEX5 to promote peroxisomal turnover by pexophagy15.
Since peroxisomes are ROS-generating centers, they are also prone to ROS damage. ROS-elicited peroxisomal injuries force cells to activate pexophagy to initiate peroxisome quality control pathways (removal of the damaged peroxisome by autophagy). Here, we outline an approach for the on-demand triggering of ROS-elicited peroxisomal injury. The protocol takes advantage of light-activated ROS production within organelles16,17,18,19,20 (Figure 1). Dye-labeled peroxisomes are illuminated, leading to ROS production within the peroxisomal lumen, which specifically triggers peroxisomal injury. Using this protocol, it is shown that ROS-stressed peroxisomes are removed through a ubiquitin-dependent degradation pathway. ROS-stressed peroxisomes recruit the ubiquitin E3 ligase Stub1 to allow their engulfment into autophagosomes for individual removal by pexophagy16. One can use this protocol to compare the fate of injured and healthy peroxisomes within the same cell by time-lapse microscopy. The method can also be used to globally damage all the peroxisomes (in all cells) on a culture dish, allowing for the biochemical analysis of the pexophagy pathway.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>35 mm culture dish with a 20 mm diameter glass microwell&#160;</td>
			<td>MatTek</td>
			<td>P35G-1.5-20-C</td>
			<td>20 mm glass bottomed</td>
		</tr>
		<tr>
			<td>3-amino-1,2,4-triazole (3-AT)</td>
			<td>Sigma Aldrich</td>
			<td>A8056</td>
			<td></td>
		</tr>
		<tr>
			<td>bovine serum</td>
			<td>ThermoFisher Scientific</td>
			<td>16170060</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture incubator</td>
			<td>Nuaire</td>
			<td>NU-4750</td>
			<td></td>
		</tr>
		<tr>
			<td>diKillerRed-PTS1</td>
			<td>Academia Sinica</td>
			<td></td>
			<td>made by appending the KillerRed tandem dimer with PTS1(VKSKL)</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium (DMEM)</td>
			<td>ThermoFisher Scientific</td>
			<td>11965092</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12)</td>
			<td>ThermoFisher Scientific</td>
			<td>11330032</td>
			<td></td>
		</tr>
		<tr>
			<td>EGFP-C1</td>
			<td>Clontech</td>
			<td>pEGFP-C1</td>
			<td>The backbone of EGFP-C1 was used for cloning EGFP-Stub1, EGFP-Hsp70, EGFP-p62</td>
		</tr>
		<tr>
			<td>EGFP-Hsp70</td>
			<td>Academia Sinica</td>
			<td></td>
			<td>Hsp70 gene (HSPA1A) PCR amplified from HeLa cDNA and cloned into EGFP-C1</td>
		</tr>
		<tr>
			<td>EGFP-LC3B</td>
			<td>Addgene</td>
			<td>11546</td>
			<td></td>
		</tr>
		<tr>
			<td>EGFP-p62</td>
			<td>Academia Sinica</td>
			<td></td>
			<td>generated by inserting the human SQSTM1 gene (through PCR amplification of the HeLa cell cDNA) into EGFP-C1</td>
		</tr>
		<tr>
			<td>EGFP-Stub1</td>
			<td>Academia Sinica</td>
			<td></td>
			<td>generated by inserting the mouse Stub1 gene (through PCR amplification of the total mouse kidney cDNA) into EGFP-C1</td>
		</tr>
		<tr>
			<td>EGFP-Ub</td>
			<td>Addgene</td>
			<td>11928</td>
			<td></td>
		</tr>
		<tr>
			<td>fetal bovine serum</td>
			<td>ThermoFisher Scientific</td>
			<td>10437028</td>
			<td></td>
		</tr>
		<tr>
			<td>HaloTag TMR ligand&#160;</td>
			<td>Promega</td>
			<td>G8252</td>
			<td></td>
		</tr>
		<tr>
			<td>HaloTag-PTS1</td>
			<td>Academia Sinica</td>
			<td></td>
			<td>PTS1 appended and cloned into EGFP-C1 backbone</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>ThermoFisher Scientific</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted Confocal Microscope&#160;</td>
			<td>Olympus</td>
			<td>FV3000RS</td>
			<td>405 nm Ex, 488 nm Ex, 561 nm Ex,&#160; microscope with a TOKAI HIT chamber incubator and the UNIV2-D35 dish attachment</td>
		</tr>
		<tr>
			<td>Janelia Fluor 646 HaloTag Ligand</td>
			<td>Promega</td>
			<td>GA1120</td>
			<td></td>
		</tr>
		<tr>
			<td>LED</td>
			<td>VitaStar</td>
			<td>PAR64</td>
			<td>80 W, 555-570 nm</td>
		</tr>
		<tr>
			<td>lipofectamine 2000&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>11668</td>
			<td>transfection reagent</td>
		</tr>
		<tr>
			<td>NIH3T3 cell</td>
			<td>ATCC</td>
			<td>CRL-1658</td>
			<td>adherent</td>
		</tr>
		<tr>
			<td>Opti-MEM</td>
			<td>ThermoFisher Scientific</td>
			<td>319850</td>
			<td>reduced serum media</td>
		</tr>
		<tr>
			<td>penicillin/streptomycin</td>
			<td>ThermoFisher Scientific</td>
			<td>15140</td>
			<td></td>
		</tr>
		<tr>
			<td>PMP34-TagBFP</td>
			<td>Academia Sinica</td>
			<td></td>
			<td>PMP34 PCR amplified from HeLa cDNA and cloned intoTagBFP-C (Evrogen FP171)</td>
		</tr>
		<tr>
			<td>roGFP2-PTS1</td>
			<td>Academia Sinica</td>
			<td></td>
			<td>generated by appending eroGFP (taken from Addgene plasmid 20131) with the amino acid sequence VKSKL, and cloned into the EGFP-C1</td>
		</tr>
		<tr>
			<td>SHSY5Y cell</td>
			<td>ATCC</td>
			<td>CRL-2266</td>
			<td>adherent</td>
		</tr>
	</tbody>
</table>

monitoring stub1-mediated pexophagy

Mammalian cells can turn over peroxisomes through Stub1-mediated pexophagy. The pathway potentially permits cellular control of the quantity and quality of peroxisomes. During this process, heat shock protein 70 and the ubiquitin E3 ligase, Stub1, translocate onto peroxisomes to be turned over to initiate pexophagy. The Stub1 ligase activity allows the accumulation of ubiquitin and other autophagy-related modules on targeted peroxisomes. Elevating reactive oxygen species (ROS) levels within the peroxisomal lumen can activate Stub1-mediated pexophagy. One can, therefore, use dye-assisted ROS generation to trigger and monitor this pathway. This article outlines the procedures for using two classes of dyes, fluorescent proteins and synthetic fluorophores, to initiate pexophagy within mammalian cell cultures. These dye-assisted ROS generation-based protocols can not only be used to target all the peroxisomes within a cell population globally but can also permit the manipulation of individual peroxisomes within single cells. We also describe how Stub1-mediated pexophagy can be followed using live-cell microscopy.

The Stub1-mediated pexophagy induction scheme shown here takes advantage of dye-assisted ROS generation within the peroxisome lumen. This operation requires minimal light intensities. Peroxisomes containing fluorescent proteins or dyes can, therefore, be illuminated using standard laser-scanning confocal microscopes. Focal illumination leads to instantaneous and localized ROS production within individual peroxisomes, as indicated by the fluorescent reporter roGFP2-VKSKL (Figure 9).&#160;We d...

Watch this Scientific Journal Video about Monitoring Stub1-Mediated Pexophagy at JoVE.com

Monitoring Stub1-Mediated Pexophagy

This protocol provides instructions for triggering and monitoring Stub1-mediated pexophagy in live cells.

Mammalian cells can turn over peroxisomes through Stub1-mediated pexophagy. The pathway potentially permits cellular control of the quantity and quality ...

Research

JoVE Journal

Biology

1.4K Views.  Academia Sinica. This protocol provides instructions for triggering and monitoring Stub1-mediated pexophagy in live cells.

Video: Monitoring Stub1-Mediated Pexophagy

Monitoring Actin Disassembly with Time-lapse Microscopy

Okay, so what I'd like to do today is to show you how to construct a flow chamber, a fusion chamber for imaging on an upright microscope. What then I then do is I create two spaces using film. So what I do is I take the para film, I cut it into small strips.Next I'm going to, this will basically form the channels for the, the walls, and I dry them with a tilted air. I put the cover slip on top of the para, And here's the finished. Then I heat it on a heat block to melt the param and basically create a seal.So what this is, is the, the cover glass here, and then on top of the cover glass is two layers of phim as you see right here. And then on top of this, again, there's a, it's a 22 by two two mil meter cover slip. This one right here, which I pressed against the param to fall the seal.Okay, so the liquid goes in in one direction here and I can suck it out with a filter paper here in this direction. And I'll show you how to do that in a bit. So what I like to do is I like to put it in a humidified chamber, which is very simple.Take two kind of elevated strips and have a filter of paper underneath. And to keep it noisy as I, I put the cover, cover slip on the flow chamber on. Now the way you introduce a solution to the chamber is that you just put the pipet tip at the entrance of the chamber and you just split.Now what I do is I incubate the chamber with the lid closed for 10 minutes. What I usually do to suck the liquid out from the other side is to use it, get a filter paper and cut it into thin strips like this. And I take one of these thin strips and I simultaneously apply the solution into the entrance while putting the filter paper at the exit to suck the exchange, the solution.So you'll see the liquid flowing through the solution. And then I wait for five minutes for the mid close. Now what I'm ready to do is to actually put that the actin it and the actin, I actually have to polymerize, I polymerize inside the flow chamber and it's just the same procedures before.So because like this, and then now that the action is polymerized, I wash out all the un excess un polymerized action using a large volume of just buffer. So here I have about a hundred microliters and it's a few reaction chamber volumes worth of buffer. Just, you know, when the filter paper becomes saturated, I have to change the direction.Okay?And so I'm ready to do imaging with this. So perfusion chamber with actin attached to it. Now I take the slide, mount it under the microscope, and since I'm imaging with an oil objective, I'm gonna put some immersion oil on the top surface of the cover, grass touch, such so assess solution or protein or anything on acting inside.So what I do is I can actually exchange the solution side while I'm doing the time lapse imaging. As such.

Functional Imaging with Reinforcement, Eyetracking, and Physiological Monitoring

We use functional brain imaging (fMRI) to study neural circuits that underlie decision-making.  To understand how outcomes affect decision processes, simple perceptual tasks are combined with appetitive and aversive reinforcement.  However, the use of reinforcers such as juice and airpuffs can create challenges for fMRI.  Reinforcer delivery can cause head movement, which creates artifacts in the fMRI signal.  Reinforcement can also lead to changes in heart rate and respiration that are mediated by autonomic pathways.  Changes in heart rate and respiration can directly affect the fMRI (BOLD) signal in the brain and can be confounded with signal changes that are due to neural activity.  In this presentation, we demonstrate methods for administering reinforcers in a controlled manner, for stabilizing the head, and for measuring pulse and respiration.

This presentation demonstrates the use of fMRI to study neural circuits that underlie decision-making. Simple perceptual tasks are combined with appetitive and aversive reinforcements to investigate how outcomes affect decision processes.

We use functional brain imaging (fMRI) to study neural circuits that underlie decision-making.  To understand how outcomes affect decision processes, ...

Monitoring Plant Hormones During Stress Responses

Plant hormones and related signaling compounds play an important role in the regulation of plant responses to various environmental stimuli and stresses. Among the most severe stresses are insect herbivory, pathogen infection, and drought stress. For each of these stresses a specific set of hormones and/or combinations thereof are known to fine-tune the responses, thereby ensuring the plant's survival. The major hormones involved in the regulation of these responses are jasmonic acid (JA), salicylic acid (SA), and abscisic acid (ABA). To better understand the role of individual hormones as well as their potential interaction during these responses it is necessary to monitor changes in their abundance in a temporal as well as in a spatial fashion. For the easy, sensitive, and reproducible quantification of these and other signaling compounds we developed a method based on vapor phase extraction and gas chromatography/mass spectrometry (GC/MS) analysis (1, 2, 3, 4). After extracting these compounds from the plant tissue by acidic aqueous 1-propanol mixed with dichloromethane the carboxylic acid-containing compounds are methylated, volatilized under heat, and collected on a polymeric absorbent. After elution into a sample vial the analytes are separated by gas chromatography and detected by chemical ionization mass spectrometry. The use of appropriate internal standards then allows for the simple quantification by relating the peak areas of analyte and internal standard.

A simple method is provided that allows for the rapid extraction and analysis of multiple plant hormones from small tissue samples. The procedure uses vapor phase extraction as the solemn purification step. Samples are analyzed by GC/MS with chemical ionization that produces mainly (M+1)+ ions.

Plant hormones and related signaling compounds play an important role in the regulation of plant responses to various environmental stimuli and stresses. ...

Monitoring Acupuncture Effects on Human Brain by fMRI

Functional MRI is used to study the effects of acupuncture on the BOLD response and the functional connectivity of the human brain.  Results demonstrate that acupuncture mobilizes a limbic-paralimbic-neocortical network and its anti-correlated sensorimotor/paralimbic network at multiple levels of the brain and that the hemodynamic response is influenced by the psychophysical response.  Physiological monitoring may be performed to explore the peripheral response of the autonomic nerve function. This video describes the studies performed at LI4 (hegu), ST36 (zusanli) and LV3 (taichong), classical acupoints that are commonly used for modulatory and pain-reducing actions. Some issues that require attention in the applications of fMRI to acupuncture investigation are noted.

FMRI and physiological monitoring is used to study the effects of Acupuncture on the central and peripheral nervous systems. Acupuncture mobilizes a limbic-paralimbic-neocortical network, with great overlap with the default mode network, to modulate neurological activity, possibly related to its autonomic effect in the peripheral nervous system.

Functional MRI is used to study the effects of acupuncture on the BOLD response and the functional connectivity of the human brain.  Results demonstrate ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Bioassays for Monitoring Insecticide Resistance

Pest resistance to pesticides is an increasing problem because pesticides are an integral part of high-yielding production agriculture. When few products are labeled for an individual pest within a particular crop system, chemical control options are limited. Therefore, the same product(s) are used repeatedly and continual selection pressure is placed on the target pest. There are both financial and environmental costs associated with the development of resistant populations. The cost of pesticide resistance has been estimated at approximately $ 1.5 billion annually in the United States. This paper will describe protocols, currently used to monitor arthropod (specifically insects) populations for the development of resistance. The adult vial test is used to measure the toxicity to contact insecticides and a modification of this test is used for plant-systemic insecticides. In these bioassays, insects are exposed to technical grade insecticide and responses (mortality) recorded at a specific post-exposure interval. The mortality data are subjected to Log Dose probit analysis to generate estimates of a lethal concentration that provides mortality to 50% (LC50) of the target populations and a series of confidence limits (CL's) as estimates of data variability. When these data are collected for a range of insecticide-susceptible populations, the LC50 can be used as baseline data for future monitoring purposes. After populations have been exposed to products, the results can be compared to a previously determined LC50 using the same methodology.

This manuscript demonstrates and discusses techniques used to survey pesticide susceptibility and detect resistance to contact and systemic pesticides in arthropod pests.

Pest resistance to pesticides is an increasing problem because pesticides are an integral part of high-yielding production agriculture. When few products ...

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must be maintained to avoid deleterious consequences 1. External or internal factors that disrupt this balance can lead to protein aggregation, toxicity and cell death. In humans this is a major contributing factor to the symptoms associated with neurodegenerative disorders such as Huntington's, Parkinson's, and Alzheimer's diseases 10. It is therefore essential that the proteins involved in maintenance of proteostasis be identified in order to develop treatments for these debilitating diseases. This article describes techniques for monitoring in vivo protein folding at near-real time resolution using the model protein firefly luciferase fused to green fluorescent protein (FFL-GFP). FFL-GFP is a unique model chimeric protein as the FFL moiety is extremely sensitive to stress-induced misfolding and aggregation, which inactivates the enzyme 12. Luciferase activity is monitored using an enzymatic assay, and the GFP moiety provides a method of visualizing soluble or aggregated FFL using automated microscopy. These coupled methods incorporate two parallel and technically independent approaches to analyze both refolding and functional reactivation of an enzyme after stress. Activity recovery can be directly correlated with kinetics of disaggregation and re-solubilization to better understand how protein quality control factors such as protein chaperones collaborate to perform these functions. In addition, gene deletions or mutations can be used to test contributions of specific proteins or protein subunits to this process. In this article we examine the contributions of the protein disaggregase Hsp104 13, known to partner with the Hsp40/70/nucleotide exchange factor (NEF) refolding system 5, to protein refolding to validate this approach.

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

This article describes the use of a firefly luciferase-GFP fusion protein to investigate in vivo protein folding in Saccharomyces cerevisiae. Using this reagent, refolding of a model heat-denatured protein can be monitored simultaneously by fluorescence microscopy and an enzymatic assay to probe the roles of proteostasis network components in protein quality control.

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must ...

Selected Reaction Monitoring Mass Spectrometry for Absolute Protein Quantification

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This protocol provides step-by-step instructions for the development and application of quantitative assays using selected reaction monitoring (SRM) mass spectrometry (MS). First, likely quantotypic target peptides are identified based on numerous criteria. This includes identifying proteotypic peptides, avoiding sites of posttranslational modification, and analyzing the uniqueness of the target peptide to the target protein. Next, crude external peptide standards are synthesized and used to develop SRM assays, and the resulting assays are used to perform qualitative analyses of the biological samples. Finally, purified, quantified, heavy isotope labeled internal peptide standards are prepared and used to perform isotope dilution series SRM assays. Analysis of all of the resulting MS data is presented. This protocol was used to accurately assay the absolute abundance of proteins of the chemotaxis signaling pathway within RAW 264.7 cells (a mouse monocyte/macrophage cell line). The quantification of Gi2 (a heterotrimeric G-protein &#945;-subunit) is described in detail.

This protocol describes how to perform absolute quantification assays of target proteins within complex biological samples using selected reaction monitoring. It was used to accurately quantify proteins of the mouse macrophage chemotaxis signaling pathway. Target peptide selection, assay development, and qualitative and quantitative assays are described in detail.

Absolute quantification of target proteins within complex biological samples is critical to a wide range of research and clinical applications. This ...

Detection of Intracellular Gene Expression in Live Cells of Murine, Human and Porcine Origin Using Fluorescence-labeled Nanoparticles

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, rat, human, pig and others. The verification of iPS clones mainly relies on the detection of the endogenous expression of different pluripotency genes. These genes mostly represent transcription factors which are located in the cell nucleus. Traditionally, the proof of their endogenous expression is supplied by immunohistochemical staining after fixation of the cells. This approach requires replicate cultures of each clone at this early stage to preserve validated clones for further experiments. The present protocol describes an approach with gene-specific nanoparticles which allows the evaluation of intracellular gene expression directly in live cells by fluorescence. The nanoparticles consist of a central gold particle coupled to a capture strand carrying a sequence complementary to the target mRNA as well as a quenched reporter strand. These nanoparticles are actively endocytosed and the target mRNA displaces the reporter strand which then start to fluoresce. Therefore, specific target gene expression can be detected directly under the microscope. In addition, the emitted fluorescence allows the identification, isolation and enrichment of cells expressing a specific gene by flow cytometry. This method can be applied directly to live cells in culture without any manipulation of the target cells.

This manuscript describes a novel technique which allows the detection of intracellular gene expression after endocytosis of fluorescence-labeled nanoparticles directly in live cells. The method does not require manipulation of the cells and is not restricted with respect to the target gene or species.

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, ...