Name: in vivo quantification of protein turnover in aging c. elegans using photoconvertible dendra2
Uploaded: 2020-06-13T14:30:00
Description: Watch this Scientific Journal Video about In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2 at JoVE.com

We acknowledge the DFG (KI-1988/5-1 to JK, NeuroCure PhD fellowship by the NeuroCure Cluster of Excellence to MLP) for funding. We also acknowledge the Imaging Core Facility of the Leibniz Research Institute for Molecular Pharmacology Berlin (FMP) for providing the imaging set up. In addition, we would like to thank Diogo Feleciano who established the Dendra2 system in the lab and provided instructions.

1. Generation of&#160;&#160;C. elegans expressing neuronal Huntingtin-Dendra2 fusion protein
<ol>
	<li>Clone the gene encoding the POI in a nematode expression vector (i.e., pPD95_75, Addgene #1494), by traditional restriction enzyme digest10, Gibson assembly11, or any method of choice. Insert a promoter to drive expression in a desired tissue or at a desired developmental stage. Insert the Dendra2 fluorophore either N- or C-terminally in frame with the POI.</li>...

<ol>
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To comprehend a protein&#39;s function it is important to understand its synthesis, location, and degradation. With the development of novel, stable, and bright FPs, visualizing and monitoring POIs has become easier and more efficient. Genetically expressed fusion PAFPs such as Dendra2 are uniquely positioned to study the stability of a POI. Upon exposure to purple-blue light, Dendra2 breaks at a precise location within a triad of conserved amino acids. The fluorophore undergoes a small structural change, resulting in a ...

The proteome of a living organism is constantly renewing itself. Proteins are continuously degraded and synthesized according to the physiological demand of a cell. Some proteins are quickly eliminated, whereas others are longer lived. Monitoring protein dynamics is a simpler, more accurate, and less invasive task when using genetically encoded fluorescent proteins (FPs). FPs form autocatalytically and can be fused to any protein of interest (POI), but do not require enzymes to fold or need cofactors save for oxygen1. A newer generation of FPs has recently been engineered to switch color upon irradiation with a light pulse of determined wavelength. These photoactivatable FPs (PAFPs) allow for labeling and tracking of POIs, or the organelles or cells they reside in, and to examine their quantitative and/or qualitative parameters2. FPs make it possible to track any POI&#39;s movement, directionality, rate of locomotion, coefficient of diffusion, mobile versus immobile fractions, the time it resides in one cellular compartment, as well as its turnover rate. For specific organelles, locomotion and transport, or fission and fusion events can be determined. For a particular cell type, a cell&#39;s position, rate of division, volume, and shape can be established. Crucially, the use of PAFPs allows tracking without continuous visualization and without interference from any newly synthetized probe. Studies both in cells and in whole organisms have successfully employed PAFPs to address biological questions in vivo, such as the development of cancer and metastasis, assembly or disassembly of the cytoskeleton, and RNA-DNA/protein interactions3. In this manuscript, light microscopy and PAFPs are used to uncover the turnover rates of the aggregation-prone protein huntingtin (HTT) in vivo in a C. elegans model of neurodegenerative disease.
The protocol described here quantifies the stability and degradation of the fusion protein huntingtin-Dendra2 (HTT-D2). Dendra2 is a second-generation monomeric PAFP4 that irreversibly switches its emission/excitation spectra from green to red in response to either UV or visible blue light, with an increase in its intensity of up to 4,000-fold5,6. Huntingtin is the protein responsible for causing Huntington&#39;s disease (HD), a fatal hereditary neurodegenerative disorder. Huntingtin exon-1 contains a stretch of glutamines (CAG, Q). When the protein is expressed with over 39Q, it misfolds into a mutant, toxic, and pathogenic protein. Mutant HTT is prone to aggregation and leads to neuronal cell death and degeneration, either as&#160;short oligomeric species or as larger highly structured amyloids7.
The nematode is a model system for studying aging and neurodegeneration thanks to its ease of manipulation, isogenic nature, short lifespan, and its optical transparency8. To study the stability of HTT in vivo, a fusion construct was expressed in the nervous system of C. elegans. A HTT-D2 transgene containing either a physiological stretch of 25Qs (HTTQ25-D2) or a pathological stretch of 97Qs (HTTQ97-D2) is overexpressed pan-neuronally throughout the nematode&#39;s lifetime9. By subjecting live C. elegans to a brief and focused point of light, a single neuron is photoswitched and the converted HTT-D2 is tracked over time. To establish the amount of HTT-D2 degraded, the difference between the red signal of the freshly converted HTT-D2 is compared to the remaining red signal of HTT-D2 after a determined period of time. Therefore, it becomes possible to investigate how huntingtin is degraded when found in its expanded and toxic form compared to its physiological form; how anterior or posterior neurons respond differently to the presence of Q97 versus Q25, especially over prolonged time periods; and how the collapse of the proteostasis network (PN) during aging contribute to the differences in degradation rates. These results only describe a small set of observations on the turnover of HTT-D2. However, many more biological questions relevant to both the field of protein aggregation and proteostasis can be addressed with this in vivo application.

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			<td>Agarose, Universal Grade</td>
			<td>Bio &#38; Sell GmbH</td>
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			<td>Deckgl&#228;ser-18x18mm</td>
			<td>Carl Roth GmbH + Co. KG</td>
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			<td>Cover slips</td>
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			<td>EC Plan-Neufluar 20x/0.50 Ph2 M27</td>
			<td>Carl Zeiss AG</td>
			<td></td>
			<td>Objective</td>
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			<td>Fiji/ImageJ 1.52p</td>
			<td>NIH</td>
			<td></td>
			<td>Analysis Software</td>
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			<td>Levamisole Hydrochloride</td>
			<td>AppliChem GmbH</td>
			<td>A4341</td>
			<td>Anesthetic</td>
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			<td>LSM710-ConfoCor3</td>
			<td>Carl Zeiss AG</td>
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			<td>Leica Camera AG</td>
			<td></td>
			<td>Mounting microscope</td>
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			<td>neuronal-HTTQ25-Dendra2</td>
			<td>this paper</td>
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			<td>neuronal-HTTQ97-Dendra2</td>
			<td>this paper</td>
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			<td>OP50 Escherichia coli</td>
			<td>CAENORHABDITIS GENETICS CENTER (CGC)</td>
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			<td>Carl Zeiss AG</td>
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			<td>Confocal acquisition software</td>
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in vivo quantification of protein turnover in aging c. elegans using photoconvertible dendra2

Proteins are synthesized and degraded constantly within a cell to maintain homeostasis. Being able to monitor the degradation of a protein of interest is key to understanding not only its life cycle, but also to uncover imbalances in the proteostasis network. This method shows how to track the degradation of the disease-causing protein huntingtin. Two versions of huntingtin fused to Dendra2 are expressed in the C. elegans nervous system: a physiological version or one with an expanded and pathogenic stretch of glutamines. Dendra2 is a photoconvertible fluorescent protein; upon a short ultraviolet (UV) irradiation pulse, Dendra2 switches its excitation/emission spectra from green to red. Similar to a pulse-chase experiment, the turnover of the converted red-Dendra2 can be monitored and quantified, regardless of the interference from newly synthesized green-Dendra2. Using confocal-based microscopy and due to the optical transparency of C. elegans, it is possible to monitor and quantify the degradation of huntingtin-Dendra2 in a living, aging organism. Neuronal huntingtin-Dendra2 is partially degraded soon after conversion and cleared further over time. The systems controlling degradation are deficient in the presence of mutant huntingtin and are further impaired with aging. Neuronal subtypes within the same nervous system exhibit different turnover capacities for huntingtin-Dendra2. Overall, monitoring any protein of interest fused to Dendra2 can provide important information not only on its degradation and the players of the proteostasis network involved, but also on its location, trafficking, and transport.

Two nematode strains expressing the huntingtin exon-1 protein fragment in frame with the photoconvertible protein Dendra2 were obtained via microinjection and the plasmids were kept as an extrachromosomal array. The fusion construct was expressed in the whole C. elegans nervous system from development throughout aging. Here, HTT-D2 contained either the physiological 25 polyglutamine stretch (HTTQ25-D2, Figure 1A) or a fully penetrant...

Watch this Scientific Journal Video about In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2 at JoVE.com

In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2

Presented here is a protocol to monitor degradation of the protein huntingtin fused to the photoconvertible fluorophore Dendra2.

In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2

Proteins are synthesized and degraded constantly within a cell to maintain homeostasis. Being able to monitor the degradation of a protein of interest is ...

This protocol allows for tracking and quantification of the degradation of a protein of interest in live and aging C.elegans. This in vivo and noninvasive technique requires only little microscopy set up to reliably convert Dendra2 into a bright and stable fluorophore. The method is adaptable to mammalian systems, as well as zebrafish.To study, put in turnover within the proteostatis network as well as transport and trafficking. Take care not to provoke unwanted conversion of Dendra2 while scanning with the blue light laser, and to test and optimize the acquisition settings and conversion settings for each system and fusion protein. Begin by growing age-matched nematodes at 20 degrees Celsius to the desired age for the experiment.On the day of the imaging experiment, melt general grade agarose at a 3%concentration in double distilled water. While the agarose is cooling, cut the tip of a one milliliter pipette tip to facilitate aspiration of approximately 400 microliters of the melted agarose. Carefully add a few drops of agarose onto a clean glass slide and immediately place another slide on top of the drops to create a thin pad of agarose between the two glass pieces.When the agarose has dried, carefully remove the top slide. When enough slides have been prepared, open the confocal microscope software and set the light path for excitation and emission. For green Dendra2 add 486 to 553 nanometers, and for red Dendra2 add 580 to 740 nanometers.Adjust the power and gain of both of the channels according to the intensity of the fluorophore and set the pinhole to fully open. Select a sequential channel mode and set the track to switch every frame. Set the scan mode as frame and the frame size as 1024 by 1024 with a line step of one.Set the averaging to two and to average by the mean method and mode of unidirectional line. Set the bit depth to eight bits. Define the multidimensional acquisition setting for the conversion of Dendra2.For conversion and bleaching, use the 405 nanometer diode laser set to 60%energy power. Select a time series of two cycles with a 0.0 millisecond interval between the cycles and a normal start and stop. Set the bleaching to start after scan one of two, to repeat for 30 iterations and to stop when the intensity drops to 50%Then, define the speed of the acquisition pixel dwell to fast for conversion and to medium for capturing a snap image.To mount the nematodes onto the slides, use a permanent marker to draw and number a window with four squares on the opposite of the agarose pad on each slide. Add 15 microliters of levamisole to the middle of the agarose pad and use a stereo microscope and a wire pick to transfer for age-matched nematodes into the droplet. Use an eyelash pick to gently move each nematode into one window of the square.When all of the nematodes have been repositioned, wait until the worms have stopped moving before placing a cover slip over the liquid to immobilize the nematodes during imaging. Then, place the slide onto the confocal microscope stage. For green Dendra2 conversion, use the eyepiece of the 20 X objective under green fluorescence to locate the first nematode and focus on the head or tail.Switch to confocal mode and increase or decrease the gain and laser power to obtain a saturated, but not overexposed image. Draw a region of interest around the selected neuron and draw a second larger region of interest around the tail that includes the first region of interest. Set the first region to be acquired, bleached, and analyzed.Set the second region of interest to be acquired and analyzed, but not bleached. Set the speed of scanning to maximum and start the scan to convert the selected Dendra2 neurons. Immediately after conversion, begin live scanning with the green 561 nanometer laser to visualize Dendra2 in the red channel and find the focus and respective maximum projection of the converted neuron.Quickly set the scan rate to a lower pixel dwell speed and acquire a snapshot image of both channels at a higher resolution. This image is considered at time zero after conversion. Then, save the scan with an identifiable name and or number, followed by the time zero label.To track Dendra2 degradation over time, open the times zero image of the respective nematode neuron. Using the same image setting, begin scanning live in the red channel and bring the converted red neuron into focus. Then, without changing any acquisition parameters, obtain a snapshot at the same speed as the first image.Be sure to define your conversion setting carefully to convert Dendra2 efficiently and sufficiently and to maintain the same acquisition parameters throughout different time points. To analyze the converted Dendra2 images, open the time zero and second time point images in Fiji and open Analyze and Set Measurements. Select the Area and Integrated Density functions and select the image obtained with the red channel.Using the Polygon Selection Tool, draw a region of interest around the time zero converted neuron. To properly identify the contours of the neuron, select Image, Adjust, and Threshold to highlight the intensity thresholds. Once the selection has been defined, click Analyze and Measure.A pop up results window will appear, including the area, integrated density, and raw integrated density values for the selected region of interest. Perform the same process of selection and measurement for the image from the second time point. Then, copy the values into the spreadsheet.In this representative analysis before conversion, no red signal was visible when the sample was excited in the red channel. Upon irradiation, the green signal diminished as HTT-D2 was converted and a red signal subsequently appeared. HTT-D2 expression degraded over time, resulting in a reduction in the levels of red HTT-D2 signal and a possible increase of the green HTT-D2 signal as more fusion protein was newly synthesized.Notably, the neurons of the tail region were determined to be significantly more active compared with those of the head. Further, there was significantly more degradation of control HTTQ25-D2 24 hours after conversion, then after two hours in both the head and tail neurons. This difference was not observed in pathogenic HTTQ97-D2, however, suggesting that the proteostasis network was unable to remove HTT-D2 containing longer glutamine stretches throughout the nervous system.HTTQ97-D2 was not degraded as efficiently as HTTQ25-D2 in very old nematodes, highlighting the inability of the proteostasis network to remove aggregated and possibly toxic species of Huntingtin in older animals. Importantly, tail neurons were able to cope with toxic HTTQ97-D2 in young nematodes, suggesting that various concomitant proteostasis network mechanisms might be at work to remove HTT-D2. For each sample acquire and non-overexposed and non-saturated image immediately after conversion and reuse the same setting for that sample throughout time.With this protocol, it is possible to test changes in a proteostatis network, and also, with the help of super-resolution microscopy, to track the spreading of disease associated proteins.

in-vivo-quantification-protein-turnover-aging-c-elegans-using

Research

JoVE Journal

Biology

Generation of Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. F&ouml;rster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 &alpha;-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

By employing a spectrally resolved two-photon microscopy imaging system, pixel-level maps of F&ouml;rster Resonance Energy Transfer (FRET) efficiencies are obtained for cells expressing membrane receptors hypothesized to form homo-oligomeric complexes. From the FRET efficiency maps, we are able to estimate stoichiometric information about the oligomer complex under study.

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron (dgn)-destabilized Green Fluorescent Protein (GFP)-based Reporter Protein

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. Maintenance of proteasome activity was implicated in many key cellular processes, like cell's stress response 3, cell cycle regulation and cellular differentiation 4 or in immune system response 5. The dysfunction of the ubiquitin-proteasome system has been related to the development of tumors and neurodegenerative diseases 4, 6. Additionally, a decrease in proteasome activity was found as a feature of cellular senescence and organismal aging 7, 8, 9, 10. Here, we present a method to measure ubiquitin-proteasome activity in living cells using a GFP-dgn fusion protein. To be able to monitor ubiquitin-proteasome activity in living primary cells, complementary DNA constructs coding for a green fluorescent protein (GFP)&#x2013;dgn fusion protein (GFP&#x2013;dgn, unstable) and a variant carrying a frameshift mutation (GFP&#x2013;dgnFS, stable 11) are inserted in lentiviral expression vectors. We prefer this technique over traditional transfection techniques because it guarantees a very high transfection efficiency independent of the cell type or the age of the donor. The difference between fluorescence displayed by the GFP&#x2013;dgnFS (stable) protein and the destabilized protein (GFP-dgn) in the absence or presence of proteasome inhibitor can be used to estimate ubiquitin-proteasome activity in each particular cell strain. These differences can be monitored by epifluorescence microscopy or can be measured by flow cytometry.

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron dgn-destabilized Green Fluorescent Protein GFP-based Reporter Protein

A method to monitor ubiquitin-proteasome activity in living cells is described. A degron-destabilized GFP- (GFP-dgn) and a stable GFP-dgnFS fusion protein are generated and transduced into the cell using a lentiviral expression vector. This technique allows to generate a stable GFP-dgn/GFP-dgnFS expressing cell line in which ubiquitin-proteasome activity can be easily assessed using epifluorescence or flow cytometry.

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. ...

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. "Freeze-cracking" is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

"Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

To stain C.  elegans with antibodies, the relatively impermeable cuticle must be  bypassed by chemical or mechanical methods.  "Freeze-cracking" is one ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These age-associated disorders are characterized by the appearance of protein aggregates with fibrillary structure in the brains of these patients. Exactly why normally soluble proteins undergo an aggregation process remains poorly understood. The discovery that protein aggregation is not limited to disease processes and instead part of the normal aging process enables the study of the molecular and cellular mechanisms that regulate protein aggregation, without using ectopically expressed human disease-associated proteins. Here we describe methodologies to examine inherent protein aggregation in Caenorhabditis elegans through complementary approaches. First, we examine how to grow large numbers of age-synchronized C. elegans to obtain aged animals and we present the biochemical procedures to isolate highly-insoluble-large aggregates. In combination with a targeted genetic knockdown, it is possible to dissect the role of a gene of interest in promoting or preventing age-dependent protein aggregation by using either a comprehensive analysis with quantitative mass spectrometry or a candidate-based analysis with antibodies. These findings are then confirmed by in vivo analysis with transgenic animals expressing fluorescent-tagged aggregation-prone proteins. These methods should help clarify why certain proteins are prone to aggregate with age and ultimately how to keep these proteins fully functional.

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

The goal of the method presented here is to explore protein aggregation during normal aging in the model organism C. elegans. The protocol represents a powerful tool to study the highly insoluble large aggregates that form with age and to determine how changes in proteostasis impact protein aggregation.

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These ...

Quantification of Site-specific Protein Lysine Acetylation and Succinylation Stoichiometry Using Data-independent Acquisition Mass Spectrometry

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein functions, for example, by changing enzymatic activity or by mediating interactions. Precise quantification of site-specific protein acylation occupancy, or stoichiometry, is essential for understanding the functional consequences of both global low-level stoichiometry and individual high-level acylation stoichiometry of specific lysine residues. Other groups have reported measurement of lysine acetylation stoichiometry by comparing the ratio of peptide precursor isotopes from endogenous, natural abundance acylation and exogenous, heavy isotope-labeled acylation introduced after quantitative chemical acetylation of proteins using stable isotope-labeled acetic anhydride. This protocol describes an optimized approach featuring several improvements, including: (1) increased chemical acylation efficiency, (2) the ability to measure protein succinylation in addition to acetylation, and (3) improved quantitative accuracy due to reduced interferences using fragment ion quantification from data-independent acquisitions (DIA) instead of precursor ion signal from data-dependent acquisition (DDA). The use of extracted peak areas from fragment ions for quantification also uniquely enables differentiation of site-level acylation stoichiometry from proteolytic peptides containing more than one lysine residue, which is not possible using precursor ion signals for quantification. Data visualization in Skyline, an open source quantitative proteomics environment, allows for convenient data inspection and review. Together, this workflow offers unbiased, precise, and accurate quantification of site-specific lysine acetylation and succinylation occupancy of an entire proteome, which may reveal and prioritize biologically relevant acylation sites.

Here, we present unbiased quantification of site-specific protein acetylation and/or succinylation occupancy (stoichiometry) of an entire proteome through a ratiometric analysis of endogenous modifications to modifications introduced after quantitative chemical acylation using stable isotope-labeled anhydrides. In combination with sensitive data-independent acquisition mass spectrometry, accurate site occupancy measurements are obtained.

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein ...

Measurements of Physiological Stress Responses in C. Elegans

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and physiology. Thus, organisms have evolved targeted and specific stress responses dedicated to repair damage and maintain homeostasis. These mechanisms include the unfolded protein response of the endoplasmic reticulum (UPRER), the unfolded protein response of the mitochondria (UPRMT), the heat shock response (HSR), and the oxidative stress response (OxSR). The protocols presented here describe methods to detect and characterize the activation of these pathways and their physiological consequences in the nematode, C. elegans. First, the use of pathway-specific fluorescent transcriptional reporters is described for rapid cellular characterization, drug screening, or large-scale genetic screening (e.g., RNAi or mutant libraries). In addition, complementary, robust physiological assays are described, which can be used to directly assess sensitivity of animals to specific stressors, serving as functional validation of the transcriptional reporters. Together, these methods allow for rapid characterization of the cellular and physiological effects of internal and external proteotoxic perturbations.

Measurements of Physiological Stress Responses in C. Elegans

Here, we characterize cellular proteotoxic stress responses in the nematode C. elegans by measuring the activation of fluorescent transcriptional reporters and assaying sensitivity to physiological stress.

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and ...

Measuring RAN Peptide Toxicity in C. elegans

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis (ALS) and Huntington&#8217;s disease. Recently, repeat expansion-containing RNA was shown to be the substrate for a novel type of protein translation called repeat-associated non-AUG-dependent (RAN) translation. Unlike canonical translation, RAN translation does not require a start codon and only occurs when repeats exceed a threshold length. Because there is no start codon to determine the reading frame, RAN translation occurs in all reading frames from both sense and antisense RNA templates that contain a repeat expansion sequence. Therefore, RAN translation expands the number of possible disease-associated toxic peptides from one to six. Thus far, RAN translation has been documented in eight different repeat expansion-based neurodegenerative and neuromuscular diseases. In each case, deciphering which RAN products are toxic, as well as their mechanisms of toxicity, is a critical step towards understanding how these peptides contribute to disease pathophysiology. In this paper, we present strategies to measure the toxicity of RAN peptides in the model system C. elegans. First, we describe procedures for measuring RAN peptide toxicity on the growth and motility of developing C. elegans. Second, we detail an assay for measuring postdevelopmental, age-dependent effects of RAN peptides on motility. Finally, we describe a neurotoxicity assay for evaluating the effects of RAN peptides on neuron morphology. These assays provide a broad assessment of RAN peptide toxicity and may be useful for performing large-scale genetic or small molecule screens to identify disease mechanisms or therapies.

Measuring RAN Peptide Toxicity in C. elegans

Repeat-associated non-ATG-dependent translational products are emerging pathogenic features of several repeat expansion-based diseases. The goal of the protocol described is to evaluate toxicity caused by these peptides using behavioral and cellular assays in the model system C. elegans.

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis ...

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits obesity and many associated diseases including cardiovascular disorders, type II diabetes, and cancer. To advance the current understanding of lipid metabolic regulation, quantitative methods to precisely measure in vivo lipid storage levels in time and space have become increasingly important and useful. Traditional approaches to analyze lipid storage are semi-quantitative for microscopic assessment or lacking spatio-temporal information for biochemical measurement. Stimulated Raman scattering (SRS) microscopy is a label-free chemical imaging technology that enables rapid and quantitative detection of lipids in live cells with a subcellular resolution. As the contrast is exploited from intrinsic molecular vibrations, SRS microscopy also permits four-dimensional tracking of lipids in live animals. In the last decade, SRS microscopy has been widely used for small molecule imaging in biomedical research and overcome the major limitations of conventional fluorescent staining and lipid extraction methods. In the laboratory, we have combined SRS microscopy with the genetic and biochemical tools available to the powerful model organism, Caenorhabditis elegans, to investigate the distribution and heterogeneity of lipid droplets across different cells and tissues and ultimately to discover novel conserved signaling pathways that modulate lipid metabolism. Here, we present the working principles and the detailed setup of the SRS microscope and provide methods for its use in quantifying lipid storage at distinct developmental timepoints of wild-type and insulin signaling deficient mutant C. elegans.

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows selective, label-free imaging of specific chemical moieties and it has been effectively employed to image lipid molecules in vivo. Here, we provide a brief introduction to the principle of SRS microscopy and describe methods for its use in imaging lipid storage in Caenorhabditis elegans.

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits ...