The muscle-Q40-mRFP strain provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). The neuronal-Q40-CFP was a kind gift of the Morimoto Lab. We acknowledge the DFG (KI-1988/5-1 to JK, NeuroCure PhD fellowship by the NeuroCure Cluster of Excellence to MLP), EMBO (Short term fellowship to MLP) and the Company of Biologists (travel grants to CG and MLP) for funding. We also acknowledge the Advanced Light Microscopy imaging facility at the Max Delbr&#252;ck Centre for Molecular Medicine, Berlin, for providing the setup to image the YFP constructs.

1. Synchronization of C. elegans
<ol>
	<li>Synchronize C. elegans either via alkaline hypochlorite solution treatment or via simple egg laying for 4 h at 20 &#176;C8.</li>
	<li>Grow and maintain nematodes at 20 &#176;C on nematode growth medium (NGM) plates seeded with OP50 E. coli according to standard procedures9. Age the nematodes until the desired developmental stage or day. 
	NOTE: In this protocol, young adults are imaged on day 4 and old nematodes are imaged on day 8 of life.</li>
</ol>
2. RNAi-mediated knockdown of chaperone machinery via feeding
NOTE: Perform knockdown of heat shock protein 1 (hsp-1) chaperone by feeding the corresponding RNAi vector to the nematodes10. The hsp-1 RNAi plasmid was obtained from the Ahringer library (clone ID: F26D10.3).
<ol>
	<li>Grow the HT115 (DE3) E. coli expressing the hsp-1 RNAi plasmid for 6 h to overnight in Luria Bertani (LB) medium containing 50 &#181;g/mL ampicillin.</li>
	<li>Prepare fresh NGM agar plates containing isopropyl &#946;-D-1-thiogalactopyranoside (IPTG; 1 mM) and ampicillin (25 &#181;g/mL) and seed with the hsp-1 RNAi bacteria. Leave plates to dry and induce at room temperature for 1-3 days.</li>
	<li>Place synchronized eggs on an siRNA plate and leave to hatch, or place gravid nematodes and allow to lay eggs for 4 h at 20 &#176;C before removing. Grow the nematodes until the desired age or stage. 
	NOTE: The second generation of nematodes might present a stronger phenotype of the knockdown. The siRNA protocol and conditions described here are general and adapted to hsp1. RNA interference via siRNA of a specific clone/gene needs to be established and optimized by the end user. It is important to note that not all siRNA have the same efficiency, and it is therefore recommended to test the efficacy of the knockdown by quantification by either quantitative reverse transcription polymerase chain reaction (RT-qPCR) or Western blot.</li>
</ol>
3. Preparation of microscopy slides
<ol>
	<li>On the day of imaging, start by preparing the imaging slides. Melt agarose in ddH2O at a concentration of 3% (w/v) and let cool slightly.</li>
	<li>Cut the tip of a 1 mL pipette tip and take roughly 200 &#181;L of melted agarose. Pipette the agarose onto a clean glass slide and immediately place a second one on top, avoiding the formation of any bubbles. Leave to dry and gently remove the top glass slide. The result is a glass slide with an even agarose surface where the nematodes will be positioned. 
	NOTE: Each slide will be used to image between 5-10 nematodes. Slides can be prepared and stored for a few hours in a humified box to prevent the agarose from drying out.</li>
</ol>
4. Mounting nematodes onto microscopy slides
NOTE: FLIM requires the nematodes to be immobilized. Perform this step once the imaging setup (e.g., microscopes, lasers, detectors) is ready to use.
<ol>
	<li>Using a platinum wire pick, place nematodes of the desired age onto a fresh unseeded plate and let them crawl to remove the excess OP50 bacteria from their bodies.</li>
	<li>Prepare the anesthetic compound (sodium azide or levamisole) to immobilize the nematode. Keep a 500 mM NaN3 stock in the dark at 4 &#176;C and dilute in fresh ddH2O to a final concentration of 250 mM. If using levamisole, dilute a 20 mM stock to 2 mM working solution in ddH2O. 
	CAUTION: Sodium azide (NaN3) is highly toxic. Use gloves and protective eyewear and work under a ventilated hood.</li>
	<li>Working under a stereomicroscope, place a 10 &#181;L drop of anesthetic compound onto an agarose pad and gently transfer 5-10 nematodes into it. Use an eyelash tip to separate the nematodes. Keep them close together but not touching to allow for easier localization of the nematodes during image acquisition.</li>
	<li>Carefully overlay the nematodes with a coverslip. Take measurements within 1 h after mounting. 
	NOTE: Both anesthetics will eventually kill C. elegans. The nematodes must be completely immobile during imaging, because the map of the lifetime is recorded from each pixel. Any movement of the x,y parameters prevents the reading of the lifetime in the same excited pixel.</li>
</ol>
5. Acquisition of FLIM data
NOTE: In this protocol, the lifetime of the fluorophore is acquired via the time-domain TCSPC method. FLIM requires a pulse of light to be generated by the laser at a set and constant repetition rate. The repetition rate varies according to the laser type and needs to be known by the user. Lifetime measurements are achieved by detectors and electronic equipment installed alongside a conventional microscope. In this protocol, measurements are performed on three different laser scanning confocal microscopes with detectors and software provided by two different companies (Table of Materials) for acquisition of mRFP, CFP, and YFP lifetimes, respectively. Check that the correct filters of emission/excitation are in place and minimize any background or monitor backlight before starting. Before starting any experiment, establish the photostability of the chosen fluorophore. If the fluorophore bleaches within a short time within the nematode tissues, it is not suitable for FLIM measurements in C. elegans.
<ol>
	<li>Open the FLIM acquisition software. The FLIM software also allows control of the confocal microscope. Locate the tab/button to allow for the detector&#39;s outputs to be enabled and press Enable Outputs.</li>
	<li>Acquire the instrument response function (IRF), which describes the timing precision of the instrumental setup. 
	NOTE: This step should be performed preferably before mounting the nematodes.
	<ol>
		<li>If available, remove the excitation/emission filters.</li>
		<li>Place an empty coverslip above the objective and find its surface. Record the scatter signal obtained from the coverslip for a minimum of 30 s. 
		NOTE: For lifetimes of several nanoseconds, the acquisition software can automatically estimate the IRF shift. Acquiring an IRF is always recommended.</li>
	</ol>
	</li>
	<li>Place the slide with the mounted C. elegans on the stage. Using a 10x magnification lens in transmission mode and localize the position of the nematodes on the slide.</li>
	<li>Remove the slide, switch the objective to a 63x magnification lens, and apply the required immersion medium (e.g., oil). Replace the slide on the stage and localize the nematodes.</li>
	<li>Locate the Pinhole Manager on the acquisition software and open it to the Maximum. Start scanning the sample, select a region of interest (e.g., head, upper body), and focus on its maximum projection plane.</li>
	<li>Monitor the laser pulse rate and the three other values present on the interface of the software: The Constant Fraction Discriminator (CFD), the Time-to-Amplitude (TAC), and the Analogue-to-Digital Converter (ADC). 
	NOTE: The laser should have a maximum gate of 1 x 108 single photon counts. This number represents the maximum number of photons supplied by the laser. The CFD provides information on the receipt of the single photon pulse in reference to the laser pulse by the detector. This value should be roughly 1 x 105. The TAC discriminates between the time one photon was detected and the next laser pulse. Finally, the ADC converts the TAC voltage into a storable memory signal11. The CFD, TAC, and ADC should all have similar values to ensure that photons emitted by the fluorophore are not lost. Correct evaluation of these parameters ensures that enough photons are being collected to create an accurate lifetime map.</li>
	<li>On the interface of the FLIM software, preview the number of photons detected: the ADC value should be between 1 x 104 and 1 x 105. If necessary, shift the focus on a different plane or increase the laser power to collect more photons. 
	NOTE: In general, the number of recorded photons per second should not exceed 1% of the laser&#39;s repetition rate.</li>
	<li>In the menu bar, select the tab to set the acquisition parameters. Select scan sync in to allow for single photon detection.</li>
	<li>Set the acquisition to a fixed amount of time or a fixed number of photons. For example, acquire a lifetime decay curve for 2 min or until a single pixel reaches a photon count of 2,000 single events. Press Start to begin acquisition. 
	NOTE: Different fluorophores will require different excitation and emission lasers and filters. According to the brightness of the sample, the laser power can also be adjusted, which will not interfere with the lifetime. These protocols use the following excitation/emission settings: YFP ex500/em520-50 nm, mRFP ex561/em580-620 nm. A pulsed two photon laser was employed for CFP measurements using ex800/em440 nm. The amount of time and photon count required for acquisition of a FLIM map will need to be empirically established for each setup and each experimental purpose.</li>
</ol>
6. Analysis of FLIM data using FLIMfit software
NOTE: Perform data analysis using the FLIMfit software tool developed at Imperial College London12 (see Figure 1).
<ol>
	<li>Open the software and import FLIM data files via File | Load FLIM Data. Load all samples from one condition, even if obtained in different sessions and from different biological repeats.</li>
	<li>If necessary, segment a single nematode from any FLIM picture via Segmentation | Segmentation Manager. Drag the cropping tool around the area of interest until it is highlighted. Once completed, press OK. 
	NOTE: Segmentation must be done for all images.</li>
	<li>Select a small region where the intensity-based image of a C. elegans appears (Figure 1, Arrows 1). The decay curve of that region will appear in the large decay window on the right side of the interface (Figure 1, Arrow 2). 
	NOTE: The decay can be displayed linearly or logarithmically.</li>
	<li>Set the correct parameters to extrapolate the lifetime via the software&#39;s algorithm as described in steps 6.5-6.8.</li>
	<li>On the Data tab (Figure 1, Arrow 3):
	<ol>
		<li>Set an arbitrary Integrated Minimum value to exclude any pixels that are too dim to produce a good fit. Depending on the C. elegans sample this value varies from 40-300. Input different values until a satisfactory preview is achieved.</li>
		<li>Select a Time Min and a Time Max number to limit the FLIM signal to these values. All events that appear before and after this threshold will be excluded. 
		NOTE: For example, for the analysis of mRFP, the events prior to 800 ps and after 4,000 ps were excluded. These values depend on the lifetime of the fluorophore and need to be determined by the end user.</li>
		<li>Do not change the preset Counts/Photon of 1.</li>
		<li>Input the Repetition Rate, in MHz, of the laser utilized during acquisition. 
		NOTE: For the current protocol, different lasers were utilized with various repetition rates. The two photon laser used for acquisition of CFP lifetimes possesses a repetition rate of 80 MHz, for YFP the laser repeats at 40 MHz, and for mRFP the value is 78.01 MHz. These values were inputted into FLIMfit according to the sample analyzed.</li>
		<li>Input a Gate Max value to exclude all saturated pixels. 
		NOTE: For lifetime measurements in C. elegans, this value is set to any large number (e.g., 1 x 108).</li>
	</ol>
	</li>
	<li>On the Lifetime tab, select a global fitting to be used (e.g., a pixel-wise fitting). See Figure 1, Arrow 4. 
	NOTE: A Pixel-wise fitting will produce a decay fitted to each individual pixel. An Image-wise fitting will produce a global fitting of each individual image and display a single lifetime value per image. A Global-wise fitting will produce a single fitting across the whole dataset. A single lifetime value is provided for all images.</li>
	<li>Do not change any other parameter except for the No. Exp selection if it is known that the chosen fluorescence decay is multiexponential and exhibits more than a single lifetime. 
	NOTE: In the present protocol, this function was utilized to calculate the lifetimes of the biexponential CFP fluorophore.</li>
	<li>Upload the IRF via the IRF menu: IRF | Load IRF. To estimate the IRF shift, select IRF | Estimate IRF Shift. A set of values will automatically appear on the IRF tab. Once this is established, do not change any other parameters of this tab.</li>
	<li>Once all parameters are set, press Fit Dataset (Figure 1, Arrow 5). The algorithm will produce a fit for the decay curve and establish a lifetime value for each image. 
	NOTE: The resulting fit, highlighted in a blue line, should overlap with all the events. A good fit is obtained when all events are aligned along the fit.</li>
	<li>Click the Parameters tab (Figure 1, Arrow 6), located within the top right menus of the software&#39;s interface, and select Statistic: w_mean (weighted mean) and check that the chi2 value is as close as possible to 1. 
	NOTE: A chi2 close to one ensures the accuracy of the fit. The lifetime value of the selected image is thus revealed as tau_1.</li>
	<li>Export any information of interest: File | Export Intensity Images/Fit Result Table/Images/Histograms. Save the data settings used to calculate the lifetime: File | Save Data Settings. 
	NOTE: The parameters employed will be saved for future analysis of the selected samples.</li>
</ol>
7. Graphical representations of FLIM data
NOTE: The lifetimes collected from different samples can be visually represented in various ways. Select to denote the lifetime values either in nanoseconds or picoseconds.
<ol>
	<li>Show the quality of the fit and the accuracy of the curve by exporting the decay curve directly from FLIMfit.</li>
	<li>Represent the distribution of the photons by plotting the frequency of the photon count versus the lifetime value in a histogram.</li>
	<li>Finally, for statistical comparison, if comparing two or more samples, place lifetime values plus standard deviation of the mean in a scatter plot bar graph. Perform any desired statistical analysis.</li>
</ol>

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The authors have nothing to disclose.

The protocol presented here describes a microscopy-based technique to identify aggregated species in the C. elegans model system. FLIM can accurately characterize the presence of both aggregated and soluble species fused to a fluorophore via measurement of their fluorescence lifetime decays. When a fusion protein starts to aggregate its recorded average lifetime will shift from a higher to a lower value16. The propensity of aggregation can then be deduced by the drop in lifetime: the lowe...

Protein aggregation occurs both in aging and disease. The pathways that lead to the formation and deposition of large amyloids or amorphous inclusions are difficult to follow and their kinetics are similarly challenging to unravel. Proteins can misfold due to intrinsic mutations within their coding sequences, as in the case of genetic diseases. Proteins also misfold because the proteostasis network (PN) that keeps them soluble and properly folded is impaired, as happens during aging. The PN includes molecular chaperones and degradation machineries and is responsible for the biogenesis, folding, trafficking, and degradation of proteins1.
C. elegans has emerged as a model to study aging and disease due to its short lifespan, isogenic nature, and ease of genetic manipulation. Several C. elegans transgenic strains that express human disease-causing proteins in vulnerable tissues have been created. Importantly, many of the strains containing aggregation-prone proteins recapitulate the hallmark of amyloid disorders, the formation of large inclusions. Thanks to C. elegans&#39; transparent body, these aggregates can be visualized in vivo, noninvasively and nondestructively2. Generating any protein of interest (POI) in fusion with a fluorophore allows to investigate its locations, trafficking, interaction network, and general fate.
We present a protocol to monitor the aggregation of disease-causing proteins in living and aging C. elegans via fluorescence lifetime imaging microscopy (FLIM). FLIM is a powerful technique based on the lifetime of a fluorophore, rather than its emission spectra. The lifetime (tau, &#964;) is defined as the average time required by a photon to decay from its excited state back to its ground state. The lifetime of a given molecule is calculated with the time-domain technique of time-correlated single photon counting (TCSPC). In TCSPC-FLIM, the fluorescent decay function is obtained by exciting the fluorophore with short, high-frequency laser pulses and measuring the emitted photon&#39;s arrival times to a detector in respect to the pulses. When scanning a sample, a three-dimensional data array is created for each pixel: the array includes information on the distribution of the photons in their x,y spatial coordinates and their temporal decay curve. A given sample therefore becomes a map of lifetimes revealing information on the protein&#39;s structure, binding, and environment3,4. Each fluorescent protein possesses an intrinsic and precisely defined lifetime, usually of a few nanoseconds (ns), dependent on its physiochemical properties. Importantly, the lifetime of a fluorophore is independent of its concentration, fluorescent intensity, and of the imaging methodology. However, within a biological system, it can be affected by environmental factors such as pH, temperature, ion concentrations, oxygen saturation, and its interaction partners. Lifetimes are also sensitive to internal structural changes and orientation. Fusing a fluorophore to a POI results in a change in its lifetime and consequently information on the behavior of the fused protein. When a fluorophore is surrounded or encapsulated in a tightly bound environment, such as the antiparallel beta sheets of an amyloid structure, it loses energy non-radiatively, a process known as quenching5. Quenching of the fluorophore results in a shortening of its apparent lifetime. When soluble, a protein&#39;s lifetime will stay closer to its original, higher value. In contrast, when a protein starts to aggregate, its lifetime will inevitably shift to a lower value6,7. Therefore, it becomes possible to monitor the aggregation propensity of any amyloid-forming protein at different ages in living C. elegans.
Here we describe a protocol to analyze the aggregation of a fusion protein comprising different polyglutamine (CAG, Q) stretches (Q40, Q44, and Q85). We illustrate how the technique can be applied equally to different fluorophores, such as cyan fluorescent protein (CFP), yellow fluorescent protein (YFP) and monomeric red fluorescent protein (mRFP); and in all tissues of C. elegans, including the neurons, muscles, and the intestine. Moreover, in the context of proteostasis, FLIM is a very useful tool to observe changes upon depletion of molecular chaperones. Knocking down one of the key molecular chaperones, heat shock protein 1 (hsp-1), via RNA interference produces premature misfolding of proteins. The increase in aggregation load as a result of aging, disease, or deficient chaperones, is then measured as a decrease in fluorescence lifetime.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agar-Agar Kobe I</td>
			<td>Carl Roth GmbH + Co. KG</td>
			<td>5210.2</td>
			<td>NGM component</td>
		</tr>
		<tr>
			<td>Ahringer Library hsp-1 siRNA</td>
			<td>Source BioScience UK Limited</td>
			<td>F26D10.3</td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin</td>
			<td>Carl Roth GmbH + Co. KG</td>
			<td>K029.3</td>
			<td>Antibiotic</td>
		</tr>
		<tr>
			<td>B&#38;H DCS-120 SPC-150</td>
			<td>Becker &#38; Hickl GmbH</td>
			<td></td>
			<td>FLIM Aquisition software</td>
		</tr>
		<tr>
			<td>B&#38;H SPC830-SPC Image</td>
			<td>Becker &#38; Hickl GmbH</td>
			<td></td>
			<td>FLIM Aquisition software</td>
		</tr>
		<tr>
			<td>BD Bacto Peptone</td>
			<td>BD-Bionsciences</td>
			<td>211677</td>
			<td>NGM component</td>
		</tr>
		<tr>
			<td>C. elegans iQ44-YFP</td>
			<td>CAENORHABDITIS GENETICS CENTER (CGC)</td>
			<td>OG412</td>
			<td></td>
		</tr>
		<tr>
			<td>C. elegans iQ85-YFP</td>
			<td></td>
			<td></td>
			<td>Kind gift from Morimoto Lab</td>
		</tr>
		<tr>
			<td>C. elegans mQ40-RFP</td>
			<td></td>
			<td></td>
			<td>Kind gift from Morimoto Lab</td>
		</tr>
		<tr>
			<td>C. elegans nQ40-CFP</td>
			<td></td>
			<td></td>
			<td>Kind gift from Morimoto Lab</td>
		</tr>
		<tr>
			<td>Deckgl&#228;ser-18x18mm</td>
			<td>Carl Roth GmbH + Co. KG</td>
			<td>0657.2</td>
			<td>Cover slips</td>
		</tr>
		<tr>
			<td>Isopropyl-&#946;-D-thiogalactopyranosid (IPTG)</td>
			<td>Carl Roth GmbH + Co. KG</td>
			<td>2316.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica M165 FC</td>
			<td>Leica Camera AG</td>
			<td></td>
			<td>Mounting Stereomicroscope</td>
		</tr>
		<tr>
			<td>Leica TCS SP5</td>
			<td>Leica Camera AG</td>
			<td></td>
			<td>Confocal Microscope</td>
		</tr>
		<tr>
			<td>Levamisole Hydrochloride</td>
			<td>AppliChem GmbH</td>
			<td>A4341</td>
			<td>Anesthetic</td>
		</tr>
		<tr>
			<td>OP50 Escherichia coli</td>
			<td>CAENORHABDITIS GENETICS CENTER (CGC)</td>
			<td>OP50</td>
			<td></td>
		</tr>
		<tr>
			<td>PicoQuant PicoHarp300</td>
			<td>PicoQuant GmbH</td>
			<td></td>
			<td>FLIM Aquisition software</td>
		</tr>
		<tr>
			<td>Sodium Azide</td>
			<td>Carl Roth GmbH + Co. KG</td>
			<td>K305.1</td>
			<td>Anesthetic</td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Carl Roth GmbH + Co. KG</td>
			<td>3957.2</td>
			<td>NGM component</td>
		</tr>
		<tr>
			<td>Standard-Objekttr&#228;ger</td>
			<td>Carl Roth GmbH + Co. KG</td>
			<td>0656.1</td>
			<td>Glass slides</td>
		</tr>
		<tr>
			<td>Universal Agarose</td>
			<td>Bio &#38; Sell GmbH</td>
			<td>BS20.46.500</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeiss AxioObserver.Z1</td>
			<td>Carl Zeiss AG</td>
			<td></td>
			<td>Confocal Microscope</td>
		</tr>
		<tr>
			<td>Zeiss LSM510-Meta NLO</td>
			<td>Carl Zeiss AG</td>
			<td></td>
			<td>Confocal Microscope</td>
		</tr>
	</tbody>
</table>

characterization of amyloid structures in aging c. elegans using fluorescence lifetime imaging

Amyloid fibrils are associated with a number of neurodegenerative diseases such as Huntington&#39;s, Parkinson&#39;s, or Alzheimer&#39;s disease. These amyloid fibrils can sequester endogenous metastable proteins as well as components of the proteostasis network (PN) and thereby exacerbate protein misfolding in the cell. There are a limited number of tools available to assess the aggregation process of amyloid proteins within an animal. We present a protocol for fluorescence lifetime microscopy (FLIM) that allows monitoring as well as quantification of the amyloid fibrilization in specific cells, such as neurons, in a noninvasive manner and with the progression of aging and upon perturbation of the PN. FLIM is independent of the expression levels of the fluorophore and enables an analysis of the aggregation process without any further staining or bleaching. Fluorophores are quenched when they are in close vicinity of amyloid structures, which results in a decrease of the fluorescence lifetime. The quenching directly correlates with the aggregation of the amyloid protein. FLIM is a versatile technique that can be applied to compare the fibrilization process of different amyloid proteins, environmental stimuli, or genetic backgrounds in vivo in a non-invasive manner.

The protocol shows how to accurately monitor the formation of aggregated species in living C. elegans, both during its natural aging and when subjected to stress. We selected four different strains of transgenic nematodes expressing polyglutamine proteins of either 40Q, 44Q, or 85Q repeats. These proteins are synthesized in different tissues and were fused to different fluorophores. The C. elegans strains either expressed Q40-mRFP in the body wall muscles (mQ40-RFP), Q40...

Watch this Scientific Journal Video about Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging at JoVE.com

Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Fluorescence lifetime imaging monitors, quantifies and distinguishes the aggregation tendencies of proteins in living, aging, and stressed C. elegans disease models.

Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Amyloid fibrils are associated with a number of neurodegenerative diseases such as Huntington&#39;s, Parkinson&#39;s, or Alzheimer&#39;s disease. These ...

characterization-amyloid-structures-aging-c-elegans-using

Research

JoVE Journal

Biochemistry

7.0K Views.  Leibniz Research Institute for Molecular Pharmacology im Forschungsverbund Berlin. Fluorescence lifetime imaging monitors, quantifies and distinguishes the aggregation tendencies of proteins in living, aging, and stressed C. elegans disease models.

Video: Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Imaging Amyloid Tissues Stained with Luminescent Conjugated Oligothiophenes by Hyperspectral Confocal Microscopy and Fluorescence Lifetime Imaging

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease afflicting primarily the central nervous system, and systemic amyloidosis where serum amyloid A, transthyretin and IgG light chains deposit as amyloid in liver, carpal tunnel, spleen, kidney, heart, and other peripheral tissues. Amyloid has been known and studied for more than a century, often using amyloid specific dyes such as Congo red and Thioflavin T (ThT) or Thioflavin (ThS). In this paper, we present heptamer-formyl thiophene acetic acid (hFTAA) as an example of recently developed complements to these dyes called luminescent conjugated oligothiophenes (LCOs). hFTAA is easy to use and is compatible with co-staining in immunofluorescence or with other cellular markers. Extensive research has proven that hFTAA detects a wider range of disease associated protein aggregates than conventional amyloid dyes. In addition, hFTAA can also be applied for optical assignment of distinct aggregated morphotypes to allow studies of amyloid fibril polymorphism. While the imaging methodology applied is optional, we here demonstrate hyperspectral imaging (HIS), laser scanning confocal microscopy and fluorescence lifetime imaging (FLIM). These examples show some of the imaging techniques where LCOs can be used as tools to gain more detailed knowledge of the formation and structural properties of amyloids. An important limitation to the technique is, as for all conventional optical microscopy techniques, the requirement for microscopic size of aggregates to allow detection. Furthermore, the aggregate should comprise a repetitive &#946;-sheet structure to allow for hFTAA binding. Excessive fixation and/or epitope exposure that modify the aggregate structure or conformation can render poor hFTAA binding and hence pose limitations to accurate imaging.

Amyloid deposition is a hallmark of different diseases and afflicts many different organs. This paper describes the application of luminescent conjugated oligothiophene fluorescence staining in combination with fluorescence microscopy techniques. This staining method represents a powerful tool for detection and exploration of protein aggregates in both clinical and scientific setups.

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find ...

Structural Characterization of Mannan Cell Wall Polysaccharides in Plants Using PACE

Plant cell wall polysaccharides are notoriously difficult to analyze, and most methods require expensive equipment, skilled operators, and large amounts of purified material. Here, we describe a simple method for gaining detailed polysaccharide structural information, including resolution of structural isomers. For polysaccharide analysis by gel electrophoresis (PACE), plant cell wall material is hydrolyzed with glycosyl hydrolases specific to the polysaccharide of interest (e.g., mannanases for mannan). Large format polyacrylamide gels are then used to separate the released oligosaccharides, which have been fluorescently labeled. Gels can be visualized with a modified gel imaging system (see Table of Materials). The resulting oligosaccharide fingerprint can either be compared qualitatively or, with replication, quantitatively. Linkage and branching information can be established using additional glycosyl hydrolases (e.g., mannosidases and galactosidases). Whilst this protocol describes a method for analyzing glucomannan structure, it can be applied to any polysaccharide for which characterized glycosyl hydrolases exist. Alternatively, it can be used to characterize novel glycosyl hydrolases using defined polysaccharide substrates.

A protocol for the structural analysis of polysaccharides by gel electrophoresis (PACE), using mannan as an example, is described.

Plant cell wall polysaccharides are notoriously difficult to analyze, and most methods require expensive equipment, skilled operators, and large amounts ...

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages for identifying biologically active chemical structures that can modify complex phenotypes such as lifespan. Described here is a simple protocol for producing hundreds of 96-well culture plates with fairly consistent numbers of C. elegans in each well. Next, we specified how to use these cultures to screen thousands of chemicals for effects on the lifespan of the nematode C. elegans. This protocol makes use of temperature sensitive sterile strains, agar plate conditions, and simple animal handling to facilitate the rapid and high throughput production of synchronized animal cultures for screening.

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Here we describe a simple protocol for rapidly producing hundreds of nematode growth media agar, 96-well culture plates with consistent numbers of Caenorhhabditis elegans per well. These cultures are useful for the phenotypic screening of whole organisms. We focus here on using these cultures to screen chemicals for pro-longevity effects.

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages ...

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to precisely maneuver the nucleation and crystal growth towards desired sizes of protein crystals. The controlled crystallization is based on sample investigation with in situ Dynamic Light Scattering (DLS) while all visual changes in the droplet are monitored online with the help of a microscope coupled to a CCD camera, thus enabling a full investigation of the protein droplet during all stages of crystallization. The use of in situ DLS measurements throughout the entire experiment allows a precise identification of the highly supersaturated protein solution transitioning to a new phase &#8211; the formation of crystal nuclei. By identifying the protein nucleation stage, the crystallization can be optimized from large protein crystals to the production of protein microcrystals. The experimental protocol shows an interactive crystallization approach based on precise automated steps such as precipitant addition, water evaporation for inducing high supersaturation, and sample dilution for slowing induced homogeneous nucleation or reversing phase transitions.

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

Here we present a protocol for controlled production of protein microcrystals. The process uses an automated device allowing controlled manipulation of several crystallization parameters. The protein crystallization is carried-out by controlled and automated addition of crystallization solutions while monitoring and investigating the radius distribution of particles in the crystallization droplet.

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to ...

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be employed using a conventional confocal microscope equipped with digital detectors. A protocol for the use of the technique in vitro is shown by means of a use case where number and brightness can be seen to accurately quantify the oligomeric state of mVenus-labelled FKBP12F36V before and after the addition of the dimerizing drug AP20187. The importance of using the correct microscope acquisition parameters and the correct data preprocessing and analysis methods are discussed. In particular, the importance of the choice of photobleaching correction is stressed. This inexpensive method can be employed to study protein-protein interactions in many biological contexts.

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

This protocol describes a calibration-free approach for quantifying protein homo-oligomerization in vitro based on fluorescence fluctuation spectroscopy using commercial light scanning microscopy. The correct acquisition settings and analysis methods are shown.

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be ...

Rapid Fluorescence-based Characterization of Single Extracellular Vesicles in Human Blood with Nanoparticle-tracking Analysis

Extracellular vesicles (EVs), including exosomes, are specialized membranous nano-sized vesicles found in bodily fluids that are constitutively released from many cell types and play a pivotal role in regulating cell-cell communication and a diverse range of biological processes. Many different methods for the characterization of EVs have been described. However, most of these methods have the disadvantage that the preparation and characterization of the samples are very time-consuming, or it is extremely difficult to analyze specific markers of interest due to their small size and due to the lack of discrete populations. While methods for analysis of EVs have been considerably improved over the last decade, there is still no standardized method for characterization of single EVs. Here, we demonstrate a semi-automated method for characterization of single EVs by fluorescence-based nanoparticle-tracking analysis. The protocol that is presented addresses the common problem of many researchers in this field and provides the complete workflow for rapid isolation of EVs and characterization with PKH67, a general cell membrane linker, as well as with specific surface markers such as CD63, CD9, vimentin, and lysosomal-associated membrane protein 1 (LAMP-1). The presented results show a high level of reproducibility, as confirmed by other methods, such as Western blotting. In the conducted experiments, we exclusively used EVs isolated from human serum samples, but this method is also suitable for plasma or other body fluids and can be adjusted for characterization of EVs from cell culture supernatants. Irrespective of the future progress of research on EV biology, the protocol that is presented here provides a rapid and reliable method for rapid characterization of single EVs with specific markers.

In this protocol, we describe the complete workflow for rapid isolation of extracellular vesicles from human whole blood and characterization of specific markers by fluorescence-based nanoparticle-tracking analysis. The presented results show a high level of reproducibility and can be adjusted to cell culture supernatants.

Extracellular vesicles (EVs), including exosomes, are specialized membranous nano-sized vesicles found in bodily fluids that are constitutively released ...

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for the targeted expression of genetically encoded calcium sensors. This protocol outlines the use of these sensors for the in vivo imaging of calcium dynamics in targeted cells, specifically the body wall muscles of the worms. By utilizing the co-expression of presynaptic channelrhodopsin, stimulation of acetylcholine release from excitatory motor neurons can be induced using blue light pulses, resulting in muscle depolarization and reproducible changes in cytoplasmic calcium levels. Two worm immobilization techniques are discussed with varying levels of difficulty. Comparison of these techniques demonstrates that both approaches preserve the physiology of the neuromuscular junction and allow for the reproducible quantification of calcium transients. By pairing optogenetics and functional calcium imaging, changes in postsynaptic calcium handling and homeostasis can be evaluated in a variety of mutant backgrounds. Data presented validates both immobilization techniques and specifically examines the roles of the C. elegans sarco(endo)plasmic reticular calcium ATPase and the calcium-activated BK potassium channel in the body wall muscle calcium regulation.

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

This method provides a way to couple optogenetics and genetically encoded calcium sensors to image baseline cytosolic calcium levels and changes in evoked calcium transients in the body wall muscles of the model organism C. elegans.

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for ...

Interactions with and Membrane Permeabilization of Brain Mitochondria by Amyloid Fibrils

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary mechanism of amyloid aggregate-induced toxicity in neurodegenerative diseases. However, most reports describing the mechanisms of membrane disruption are based on phospholipid model systems, and studies directly targeting events occurring at the level of biological membranes are rare. Described here is a model for studying the mechanisms of amyloid toxicity at the membrane level. For mitochondrial isolation, density gradient medium is used to obtain preparations with minimal myelin contamination. After mitochondrial membrane integrity confirmation, the interaction of amyloid fibrils arising from &#945;-synuclein, bovine insulin, and hen egg white lysozyme (HEWL) with rat brain mitochondria, as an in vitro biological model, is investigated. The results demonstrate that treatment of brain mitochondria with fibrillar assemblies can cause different degrees of membrane permeabilization and ROS content enhancement. This indicates structure-dependent interactions between amyloid fibrils and mitochondrial membrane. It is suggested that biophysical properties of amyloid fibrils and their specific binding to mitochondrial membranes may provide explanations for some of these observations.

Provided here is a protocol for investigating the interactions between native form, prefibrillar, and mature amyloid fibrils of different peptides and proteins with mitochondria isolated from different tissues and various areas of the brain.

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary ...

SUMO-Binding Entities (SUBEs) as Tools for the Enrichment, Isolation, Identification, and Characterization of the SUMO Proteome in Liver Cancer

Post-translational modification is a key mechanism regulating protein homeostasis and function in eukaryotic cells. Among all ubiquitin-like proteins in liver cancer, the modification by SUMO (Small Ubiquitin MOdifier) has been given the most attention. Isolation of endogenous SUMOylated proteins in vivo is challenging due to the presence of active SUMO-specific proteases. Initial studies of SUMOylation in vivo were based on the molecular detection of specific SUMOylated proteins (e.g., by western blot). However, in many cases, antibodies, generally made with non-modified recombinant protein, did not immunoprecipitate SUMOylated forms of the protein of interest.&#160;Nickel chromatography has been the other approach to study SUMOylation by capturing histidine-tagged versions of SUMO molecules. This approach is mainly used in cells stably expressing or transiently transfected with His-SUMO molecules. To overcome these limitations, SUMO-binding entities (SUBEs) were developed to isolate endogenous SUMOylated proteins. Herein, we describe all the steps required for the enrichment, isolation, and identification of SUMOylated substrates from human hepatoma cells and hepatic tissues from a liver cancer mouse model by using SUBEs. Firstly, we describe methods involved in the preparation and lysis of the human hepatoma cells and liver tumor tissue samples. Then, a thorough explanation of the preparation of SUBEs and controls is detailed along with the protocol for the protein pull-down assays. Finally, some examples are provided regarding the options available for the identification and characterization of the SUMOylated proteome, namely the use of western-blot analysis for the detection of a specific SUMOylated substrate from liver tumors or the use of proteomics by mass spectrometry for high-throughput characterization of the SUMOylated proteome and interactome in hepatoma cells.

SUMO-Binding Entities SUBEs as Tools for the Enrichment, Isolation, Identification, and Characterization of the SUMO Proteome in Liver Cancer

Here, we present a protocol to enrich, isolate, identify, and characterize proteins modified by SUMO in vivo both from human hepatoma cells and liver tumors obtained from mouse models of hepatocellular carcinoma by using SUMO-binding entities (SUBEs).

Post-translational modification is a key mechanism regulating protein homeostasis and function in eukaryotic cells. Among all ubiquitin-like proteins in ...

Functional Characterization of RING-Type E3 Ubiquitin Ligases In Vitro and In Planta

Ubiquitination, as a posttranslational modification of proteins, plays an important regulatory role in homeostasis of eukaryotic cells. The covalent attachment of 76 amino acid ubiquitin modifiers to a target protein, depending on the length and topology of the polyubiquitin chain, can result in different outcomes ranging from protein degradation to changes in the localization and/or activity of modified protein. Three enzymes sequentially catalyze the ubiquitination process: E1 ubiquitin-activating enzyme, E2 ubiquitin-conjugating enzyme, and E3 ubiquitin ligase. E3 ubiquitin ligase determines substrate specificity and, therefore, represents a very interesting study subject. Here we present a comprehensive approach to study the relationship between the enzymatic activity and function of the RING-type E3 ubiquitin ligase. This four-step protocol describes 1) how to generate an E3 ligase deficient mutant through site-directed mutagenesis targeted at the conserved RING domain; 2&#8211;3) how to examine the ubiquitination activity both in vitro and in planta; 4) how to link those biochemical analysis to the biological significance of the tested protein. Generation of an E3 ligase-deficient mutant that still interacts with its substrate but no longer ubiquitinates it for degradation facilitates the testing of enzyme-substrate interactions in vivo. Furthermore, the mutation in the conserved RING domain often confers a dominant negative phenotype that can be utilized in functional knockout studies as an alternative approach to an RNA-interference approach. Our methods were optimized to investigate the biological role of the plant parasitic nematode effector RHA1B, which hijacks the host ubiquitination system in plant cells to promote parasitism. With slight modification of the in vivo expression system, this protocol can be applied to the analysis of any RING-type E3 ligase regardless of its origins.

The goal of this manuscript is to present an outline for the comprehensive biochemical and functional studies of the RING-type E3 ubiquitin ligases. This multistep pipeline, with detailed protocols, validates an enzymatic activity of the tested protein and demonstrates how to link the activity to function.

Ubiquitination, as a posttranslational modification of proteins, plays an important regulatory role in homeostasis of eukaryotic cells. The covalent ...