The authors thank Dr. Percy Tumbale and Dr. Melissa Wells for their critical reading of this manuscript. This work was supported by the US National Institute of Health Intramural Research Program; US National Institute of Environmental Health Sciences (NIEHS; ZIA ES103247 to R.E.S).

1. Preparation of 7% - 47% sucrose gradients
NOTE: The linear range of the sucrose gradient can be modified to achieve better separation depending on the cell type used. This protocol is optimized for polysome profiles for S. cerevisiae.
<ol>
	<li>Prepare stock solutions of 7% and 47% sucrose in sucrose gradient buffer (20 mM Tris-HCl pH 7.4, 60 mM KCl, 10 mM MgCl2, and 1 mM DTT). Filter sterilize the sucrose stock solutions through a 0.22 &#956;m filter and store at 4 &#176;C.</li>
	<li>Prepare 14 mL of 17%, 27%, and 37% sucrose solutions by dispensing and mixing the 7% and 47% sucrose stock solutions in the manner described in Table 1.</li>
	<li>Place six polypropylene centrifuge tubes (14 x 89 mm) into a full view test tube rack. Ensure enough space between the tubes so that actions with one tube do not disturb the others.</li>
	<li>Attach a long needle to a 3 mL syringe. For this protocol, use a 9 in, 22 G needle with a blunt tip (Figure 1), but any needle long enough to reach the bottom of the centrifuge tube will suffice. 
	NOTE: Perform a test fill and dispensation to ensure that the syringe can hold the sucrose solution without any dripping prior to setting up the gradients.</li>
	<li>Add 2 mL of the 7% sucrose to the bottom of each centrifuge tube.</li>
	<li>Add 2 mL of the 17% sucrose beneath the 7% solution by positioning the needle tip within the immediate vicinity of the tube bottom and dispensing the solution slowly and carefully.</li>
	<li>Repeat with 2 mL each of the 27%, 37%, and 47% sucrose solutions. Ensure that each layer is distinguishable from one another by a line marking the separation of densities (Figure 2). 
	NOTE: If the gradients are not going to be used within 48 hours then at this point, prior to their settlement into a continuous gradient, flash freeze the tubes with layered sucrose solutions in liquid nitrogen and store in a -80 &#176;C freezer for long-term storage.</li>
	<li>Store gradients at 4 &#176;C overnight to allow gradients to settle into a continuous, increasing percentage of sucrose. It takes 8-12 h for the layered sucrose solutions to settle into a linear sucrose gradient. Linear gradients are stable for up to 48 h. Overnight storage at 4 &#176;C provides sufficient time for thawing and settlement into a linear gradient for using frozen, layered sucrose solutions. Use a densitometer to assess the gradient quality. 
	NOTE: It is critical to store the gradients in a stable place where they will not be disturbed, as any movements or vibrations will disrupt the gradient.</li>
</ol>
2. Preparation of yeast cell extracts
<ol>
	<li>Inoculate the yeast strain of interest into 50 mL of yeast extract peptone dextrose (YPD) media and grow overnight at 30 &#176;C in a shaking incubator to stationary phase. 
	NOTE: The temperature may vary depending upon yeast strain requirements.</li>
	<li>Transfer 10 mL of stationary phase culture into 1 L of fresh YPD media. Incubate the cells with vigorous shaking at 30 &#176;C (or other suitable temperature) until the culture reaches the mid-exponential growth phase (OD600 = 0.4-0.6).</li>
	<li>At the mid-exponential growth phase, add cycloheximide to the culture at a final concentration of 0.1 mg/mL. Incubate on ice for 5 min.</li>
	<li>Harvest the cells by centrifugation at 3,000 x g for 10 min at 4 &#176;C. 
	NOTE: At this point, cells can be frozen and stored at -80 &#176;C.</li>
	<li>Resuspend the cells in chilled 700 &#956;L of polysome extraction buffer (20 mM Tris-HCl pH 7.4, 60 mM KCl, 10 mM MgCl2, 1 mM DTT, 1% Triton X-100, 0.1 mg/mL of cycloheximide, 0.2 mg/mL of heparin). Add 100 units of RNase Inhibitor and transfer it to a 1.5 mL centrifuge tube.</li>
	<li>Add ~400 &#956;L of pre-chilled glass beads with a size range of 425-600 &#181;m to the centrifuge tube. Disrupt the yeast cells by vigorous agitation in a bead-beater for 5 min.</li>
	<li>Clarify the lysate by centrifugation at 8,000 x g for 5 min at 4 &#176;C.</li>
	<li>Determine the concentration of the RNA in the clarified lysate by measuring the absorbance at 260 nm with a spectrophotometer or using a fluorescence-based RNA detection system. 
	NOTE: For very accurate RNA concentration measurements, use fluorescence-based RNA detection kits.</li>
	<li>Ensure that the RNA concentration is 0.5-1 &#181;g/&#956;L. If the RNA concentration is too low, reduce the volume of polysome extraction buffer used to resuspend the cells in future experiments. 
	&#8203;NOTE: Load the lysate onto the gradients immediately after lysis and RNA quantitation. If necessary, flash freeze the lysates in liquid nitrogen and store at -80 &#176;C for a few days.</li>
</ol>
3. Centrifugation of gradients
<ol>
	<li>Carefully load the lysate onto the top of the gradients. Place the pipette tip against the inner wall, at the top of the polypropylene tube. Gently angle the tube and slowly dispense the lysate onto the top of the gradient by dribbling against the wall. Take great care not to disrupt or disturb the gradient when loading the lysate. 
	NOTE: The amount of lysate loaded will vary per cell type. The content of the RNA will also vary. It may be necessary to perform a number of experiments to determine the amount of lysate needed to generate an optimal polysome profile. For yeast, 300 &#956;g&#160;of RNA is a good starting point for optimization.</li>
	<li>Gently place the tubes into the pre-chilled buckets of a swinging bucket rotor. 
	NOTE: Each polypropylene centrifuge tube should have equal volumes of gradient and the amount of the lysate loaded. This can vary from tube to tube. Ensure that the variation does not cause an imbalance error during ultracentrifugation.</li>
	<li>Centrifuge the gradients at 260,110 x g for 150 min at 4 &#176;C.</li>
</ol>
4. Fraction and data collection
<ol>
	<li>Carefully remove the centrifuge tubes from the swinging bucket rotor and place them in a tube holder.</li>
	<li>Label the 96-well plates to store the fractions and pre-chill on ice. 
	NOTE: Use a 96-well plate suitable for a spectrophotometer that has an optical window down to 230 nm for nucleic acid determinations at 260 nm/280 nm.</li>
	<li>Collect 100 &#181;L or 200 &#181;L fractions starting from the top of the gradient by carefully inserting a pipet tip into the top of the gradient. Collect fractions until the entire gradient is aliquoted. The number of fractions will depend on total gradient volume. 
	NOTE: Collect the fractions in a manner that does not disrupt the rest of the gradient. Ensure that all the fractions have equal volume. In addition to manual fractionation, another low-cost method for fractionation is to use a small peristaltic pump.</li>
	<li>Transfer each fraction to the 96-well plate to reach the bottom of the centrifuge tube. Keep the collected fractions on the ice at all times.</li>
	<li>Measure the absorbance of each fraction at 254 nm with a spectrophotometer. Use the 7% and 47% sucrose solutions as blanks. 
	NOTE: When measuring the absorbance of fractions, bear in mind that for most spectrometers and colorimeters, the most effective absorbance range is 0.1-1. If the absorbance measurements are out of range (&#8805;1.0), the fractions have too much material. Dilute the sample, recollect the data, and then account for the dilution factor when plotting the profile.</li>
	<li>Create the polysome profile by plotting the fraction number versus absorbance.</li>
</ol>

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The authors have nothing to disclose.

Here a method to create polysome profiles without the use of expensive automated fractionation systems has been described. The advantage of this method is that it makes polysome profiling accessible to labs that do not have automated fractionation systems. The major disadvantages of this protocol are tedious hand fractionation and reduced sensitivity compared to the dedicated density fractionation system.
This protocol entails careful preparation of sucrose gradients with resolution sufficient...

Ribosomes are mega-Dalton ribonucleoprotein complexes that perform the fundamental process of translating mRNA into proteins. Ribosomes are responsible for carrying out the synthesis of all proteins within a cell. Eukaryotic ribosomes comprise two subunits designated as the small ribosomal subunit (40S) and the large ribosomal subunit (60S) according to their sedimentation coefficients. The fully assembled ribosome is designated as the 80S monosome. Polysomes are groups of ribosomes engaged in translating a single mRNA molecule. Polysome fractionation by sucrose density gradient centrifugation is a powerful method used to create ribosome profiles, identify specific mRNAs associated with translating ribosomes and analyze polysome associated factors1,2,3,4,5,6,7,8,9,10,11,12,13. This technique is often used to separate polysomes from single ribosomes, ribosomal subunits, and messenger ribonucleoprotein particles. The profiles obtained from fractionation can provide valuable information regarding the translation activity of polysomes14 and the assembly status of the ribosomes15,16,17.
Ribosome assembly is a very complex process facilitated by a group of proteins known as ribosome assembly factors18,19,20,21. These factors perform a wide range of functions during ribosome biogenesis through interactions with many other proteins, including ATPases, endo- and exo-nucleases, GTPases, RNA helicases, and RNA binding proteins22. Polysome fractionation has been a powerful tool used to investigate the role of these factors in ribosome assembly. For example, this method has been utilized to demonstrate how mutations in the polynucleotide kinase Grc3, a pre-rRNA processing factor, can negatively affect the ribosome assembly process17,23. Polysome profiling has also highlighted and shown how the conserved motifs within the ATPase Rix7 are essential to ribosome production16.
The procedure for polysome fractionation begins with making soluble cell lysates from cells of interest. The lysate contains RNA, ribosomal subunits, and polysomes, as well as other soluble cellular components. A continuous, linear sucrose gradient is made within an ultracentrifuge tube. The soluble fraction of cell lysate is gently loaded onto the top of the sucrose gradient tube. The loaded gradient tube is then subjected to centrifugation, which separates the cellular components by size within the sucrose gradient by the force of gravity. The larger components travel further into the gradient than the smaller components. The top of the gradient houses the smaller, slower traveling cellular components, whereas the larger, faster traveling cellular components are found at the bottom. After centrifugation, the contents of the tube are collected as fractions. This method effectively separates ribosomal subunits, monosomes, and polysomes. The optical density of each fraction is then determined by measuring the spectral absorbance at a wavelength of 254 nm. Plotting absorbance vs. fraction number yields a polysome profile.
Linear sucrose density gradients can be generated utilizing a gradient maker. Following centrifugation, gradients are often fractionated, and absorbances measured using an automated density fractionation system3,7,13,24,25. While these systems work very well to produce polysome profiles, they are expensive and can be cost-prohibitive to some laboratories. Here a protocol to generate polysome profiles without the use of these instruments is presented. Instead, this protocol utilizes equipment typically available in most molecular biology laboratories.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Automatic Fractionator</td>
			<td>Brandel</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Clariostar Multimode Plate Reader</td>
			<td>BMG Labtech</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cycloheximide</td>
			<td>Sigma Aldrich</td>
			<td>C7698</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithiothreitol</td>
			<td>Invitrogen</td>
			<td>15508-013</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Beads, acid washed</td>
			<td>Sigma Aldrich</td>
			<td>G8772</td>
			<td>425&#8211;600 &#956;m</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Sigma Aldrich</td>
			<td>H4784</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Chloride, 1 M</td>
			<td>KD Medical</td>
			<td>CAC-5290</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle, 22 G, Metal Hub</td>
			<td>Hamilton Company</td>
			<td>7748-08</td>
			<td>custom length 9 inches, point style 3</td>
		</tr>
		<tr>
			<td>Optima XL-100K Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Polypropylene Centrifuge tubes</td>
			<td>Beckman Coulter</td>
			<td>331372</td>
			<td></td>
		</tr>
		<tr>
			<td>Polypropylene Test Tube Peg Rack</td>
			<td>Fisher Scientific</td>
			<td>14-810-54A</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>Sigma Aldrich</td>
			<td>P9541</td>
			<td></td>
		</tr>
		<tr>
			<td>Qubit 4 Fluorometer</td>
			<td>Thermo Fisher Scientific</td>
			<td>Q33228</td>
			<td></td>
		</tr>
		<tr>
			<td>Qubit RNA HS Assay Kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>Q32855</td>
			<td></td>
		</tr>
		<tr>
			<td>RNAse Inhibitor</td>
			<td>Applied Biosystems</td>
			<td>N8080119</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma Aldrich</td>
			<td>S0389</td>
			<td></td>
		</tr>
		<tr>
			<td>SW41 Swinging Bucket Rotor Pkg</td>
			<td>Beckman Coulter</td>
			<td>331336</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe, 3 mL</td>
			<td>Coviden</td>
			<td>888151394</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris, 1 M,&#160; pH 7.4</td>
			<td>KD Medical</td>
			<td>RGF-3340</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma Aldrich</td>
			<td>X100</td>
			<td></td>
		</tr>
		<tr>
			<td>UV-Star Microplate, 96 wells</td>
			<td>Greiner Bio-One</td>
			<td>655801</td>
			<td></td>
		</tr>
	</tbody>
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polysome profiling without gradient makers or fractionation systems

Polysome fractionation by sucrose density gradient centrifugation is a powerful tool that can be used to create ribosome profiles, identify specific mRNAs being translated by ribosomes, and analyze polysome associated factors. While automated gradient makers and gradient fractionation systems are commonly used with this technique, these systems are generally expensive and can be cost-prohibitive for laboratories that have limited resources or cannot justify the expense due to their infrequent or occasional need to perform this method for their research. Here, a protocol is presented to reproducibly generate polysome profiles using standard equipment available in most molecular biology laboratories without specialized fractionation instruments. Moreover, a comparison of polysome profiles generated with and without a gradient fractionation system is provided. Strategies to optimize and produce reproducible polysome profiles are discussed. Saccharomyces cerevisiae is utilized as a model organism in this protocol. However, this protocol can be easily modified and adapted to generate ribosome profiles for many different organisms and cell types.

Three representative polysome profiles are shown in Figure 3. All profiles are from the same yeast strain. A typical polysome profile will have well-resolved peaks for the 40S, 60S, and 80S ribosomal subunits as well as polysomes. The crest of each ribosomal subunit and polysome peak will be apparent on each profile (Figure 3). A representative profile from an automated density fractionation system is shown in Figure 3A. The sucrose...

Watch this Scientific Journal Video about Polysome Profiling without Gradient Makers or Fractionation Systems at JoVE.com

Polysome Profiling without Gradient Makers or Fractionation Systems

This protocol describes how to generate a polysome profile without using automated gradient makers or gradient fractionation systems.

Polysome fractionation by sucrose density gradient centrifugation is a powerful tool that can be used to create ribosome profiles, identify specific mRNAs ...

polysome-profiling-without-gradient-makers-or-fractionation-systems

Research

JoVE Journal

Biochemistry

4.8K Views.  National Institutes of Health. This protocol describes how to generate a polysome profile without using automated gradient makers or gradient fractionation systems.

Video: Polysome Profiling without Gradient Makers or Fractionation Systems

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate neurotransmission that occurs within synapses because it is highly regulated by dynamic protein-protein interactions and phosphorylation events. Accordingly, when any method is used to study synaptic transmission, a major goal is to preserve these transient physiological modifications. Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments. In other words, the protocol describes a biochemical methodology to carry out protein enrichment from presynaptic, postsynaptic, and extrasynaptic compartments. First, synaptosomes, or synaptic terminals, are obtained from neurons that contain all synaptic compartments by means of a discontinuous sucrose gradient. Of note, the quality of this initial synaptic membrane preparation is critical. Subsequently, the isolation of the different subsynaptic compartments is achieved with light solubilization using mild detergents at differential pH conditions. This allows for separation by gradient and isopycnic centrifugations. Finally, protein enrichment at the different subsynaptic compartments (i.e., pre-, post- and extrasynaptic membrane fractions) is validated by means of immunoblot analysis using well-characterized synaptic protein markers (i.e., SNAP-25, PSD-95, and synaptophysin, respectively), thus enabling a direct assessment of the synaptic distribution of any particular neuronal protein.

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments.

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate ...

Fractionation for Resolution of Soluble and Insoluble Huntingtin Species

The accumulation of misfolded proteins is central to pathology in Huntington's disease (HD) and many other neurodegenerative disorders. Specifically, a key pathological feature of HD is the aberrant accumulation of mutant HTT (mHTT) protein into high molecular weight complexes and intracellular inclusion bodies composed of fragments and other proteins. Conventional methods to measure and understand the contributions of various forms of mHTT-containing aggregates include fluorescence microscopy, western blot analysis, and filter trap assays. 
However, most of these methods are conformation specific, and therefore may not resolve the full state of mHTT protein flux due to the complex nature of aggregate solubility and resolution.
For the identification of aggregated mHTT and various modified forms and complexes, separation and solubilization of the cellular aggregates and fragments is mandatory. Here we describe a method to isolate and visualize soluble mHTT, monomers, oligomers, fragments, and an insoluble high molecular weight (HMW) accumulated mHTT species. HMW mHTT tracks with disease progression, corresponds with mouse behavior readouts, and has been beneficially modulated by certain therapeutic interventions1. This approach can be used with mouse brain, peripheral tissues, and cell culture but may be adapted to other model systems or disease contexts.

A method is described for fractionation of insoluble and soluble mutant huntingtin species from mouse brain and cell culture. The method described is useful for characterization and quantification of huntingtin protein flux and aids in analyzing protein homeostasis in disease pathogenesis and in the presence of perturbations

The accumulation of misfolded proteins is central to pathology in Huntington's disease (HD) and many other neurodegenerative disorders. Specifically, a ...

Polysome Profiling in Leishmania, Human Cells and Mouse Testis

Proper protein expression at the right time and in the right amounts is the basis of normal cell function and survival in a fast-changing environment. For a long time, the gene expression studies were dominated by research on the transcriptional level. However, the steady-state levels of mRNAs do not correlate well with protein production, and the translatability of mRNAs varies greatly depending on the conditions. In some organisms, like the parasite Leishmania, the protein expression is regulated mostly at the translational level. Recent studies demonstrated that protein translation dysregulation is associated with cancer, metabolic, neurodegenerative and other human diseases. Polysome profiling is a powerful method to study protein translation regulation. It allows to measure the translational status of individual mRNAs or examine translation on a genome-wide scale. The basis of this technique is the separation of polysomes, ribosomes, their subunits and free mRNAs during centrifugation of a cytoplasmic lysate through a sucrose gradient. Here, we present a universal polysome profiling protocol used on three different models - parasite Leishmania major, cultured human cells and animal tissues. Leishmania cells freely grow in suspension and cultured human cells grow in adherent monolayer, while mouse testis represents an animal tissue sample. Thus, the technique is adapted to all of these sources. The protocol for the analysis of polysomal fractions includes detection of individual mRNA levels by RT-qPCR, proteins by Western blot and analysis of ribosomal RNAs by electrophoresis. The method can be further extended by examination of mRNAs association with the ribosome on a transcriptome level by deep RNA-seq and analysis of ribosome-associated proteins by mass spectroscopy of the fractions. The method can be easily adjusted to other biological models.

Polysome Profiling in Leishmania, Human Cells and Mouse Testis

The overall goal of polysome profiling technique is analysis of translational activity of individual mRNAs or transcriptome mRNAs during protein synthesis. The method is important for studies of protein synthesis regulation, translation activation and repression in health and multiple human diseases.

Proper protein expression at the right time and in the right amounts is the basis of normal cell function and survival in a fast-changing environment. For ...

Detection of Small GTPase Prenylation and GTP Binding Using Membrane Fractionation and  GTPase-linked Immunosorbent Assay

The Rho GTPase family belongs to the Ras superfamily and includes approximately 20 members in humans. Rho GTPases are important in the regulation of diverse cellular functions, including cytoskeletal dynamics, cell motility, cell polarity, axonal guidance, vesicular trafficking, and cell cycle control. Changes in Rho GTPase signaling play an essential regulatory role in many pathological conditions, such as&#160;cancer, central nervous system diseases, and immune system-dependent diseases. The posttranslational modification of Rho GTPases (i.e., prenylation by mevalonate pathway intermediates) and GTP binding are key factors which affect the activation of this protein. In this paper, two essential and simple methods are provided to detect a broad range of Rho GTPase prenylation and GTP binding activities. Details of the technical procedures that have been used are explained step by step in this manuscript.

Here we describe a protocol to investigate the prenylation and guanosine-5'-triphosphate (GTP)-loading of Rho GTPase. This protocol consists of two detailed methods, namely membrane fractionation and a GTPase-linked immunosorbent assay. The protocol can be used for measuring the prenylation and GTP loading of different other small GTPases.

The Rho GTPase family belongs to the Ras superfamily and includes approximately 20 members in humans. Rho GTPases are important in the regulation of ...

Manipulating Living Cells to Construct Stable 3D Cellular Assembly Without Artificial Scaffold

Regenerative medicine and tissue engineering offer several advantages for the treatment of intractable diseases, and several studies have demonstrated the importance of 3-dimensional (3D) cellular assemblies in these fields. Artificial scaffolds have often been used to construct 3D cellular assemblies. However, the scaffolds used to construct cellular assemblies are sometimes toxic and may change the properties of the cells. Thus, it would be beneficial to establish a non-toxic method for facilitating cell-cell contact. In this paper, we introduce a novel method for constructing stable cellular assemblies by using optical tweezers with dextran. One of the advantages of this method is that it establishes stable cell-to-cell contact within a few minutes. This new method allows the construction of 3D cellular assemblies in a natural hydrophilic polymer and is expected to be useful for constructing next-generation 3D single-cell assemblies in the fields of regenerative medicine and tissue engineering.

We demonstrate a novel method for constructing a single-cell-based 3-dimensional (3D) assembly without an artificial scaffold.

Regenerative medicine and tissue engineering offer several advantages for the treatment of intractable diseases, and several studies have demonstrated the ...

The Identification of Sea Lamprey Pheromones Using Bioassay-Guided Fractionation

Bioassay-guided fractionation is an iterative approach that uses the results of physiological and behavioral bioassays to guide the isolation and identification of an active pheromone compound. This method has resulted in the successful characterization of the chemical signals that function as pheromones in a wide range of animal species. Sea lampreys rely on olfaction to detect pheromones that mediate behavioral or physiological responses. We use this knowledge of fish biology to posit functions of putative pheromones and to guide the isolation and identification of active pheromone components. Chromatography is used to extract, concentrate, and separate compounds from the conditioned water. Electro-olfactogram (EOG) recordings are conducted to determine which fractions elicit olfactory responses. Two-choice maze behavioral assays are then used to determine if any of the odorous fractions are also behaviorally active and induce a preference. Spectrometric and spectroscopic methods provide the molecular weight and structural information to assist with the structure elucidation. The bioactivity of the pure compounds is confirmed with EOG and behavioral assays. The behavioral responses observed in the maze should ultimately be validated in a field setting to confirm their function in a natural stream setting. These bioassays play a dual role to 1) guide the fractionation process and 2) confirm and further define the bioactivity of isolated components. Here, we report the representative results of a sea lamprey pheromone identification that exemplify the utility of the bioassay-guided fractionation approach. The identification of sea lamprey pheromones is particularly important because a modulation of its pheromone communication system is among the options considered to control the invasive sea lamprey in the Laurentian Great Lakes. This method can be readily adapted to characterize the chemical communication in a broad array of taxa and shed light on waterborne chemical ecology.

Here, we present a protocol to isolate and characterize the structure, olfactory potency, and behavioral response of putative pheromone compounds of sea lampreys.

Bioassay-guided fractionation is an iterative approach that uses the results of physiological and behavioral bioassays to guide the isolation and ...

Cell Fractionation of U937 Cells in the Absence of High-speed Centrifugation

In this protocol we detail a method to obtain subcellular fractions of U937 cells without the use of ultracentrifugation or indiscriminate detergents. This method utilizes hypotonic buffers, digitonin, mechanical lysis and differential centrifugation to isolate the cytoplasm, mitochondria and plasma membrane. The process can be scaled to accommodate the needs of researchers, is inexpensive and straightforward. This method will allow researchers to determine protein localization in cells without specialized centrifuges and without the use of commercial kits, both of which can be prohibitively expensive. We have successfully used this method to separate cytosolic, plasma membrane and mitochondrial proteins in the human monocyte cell line U937.

Here, we present a protocol to isolate the plasma membrane, cytoplasm and mitochondria of U937 cells without the use of high-speed centrifugation. This technique can be used to purify subcellular fractions for subsequent examination of protein localization via immunoblotting.

In this protocol we detail a method to obtain subcellular fractions of U937 cells without the use of ultracentrifugation or indiscriminate detergents. ...

Nitropeptide Profiling and Identification Illustrated by Angiotensin II

Protein nitration is one of the most important post-translational modifications (PTM) on tyrosine residues and it can be induced by chemical actions of reactive oxygen species (ROS) and reactive nitrogen species (RNS) in eukaryotic cells. Precise identification of nitration sites on proteins is crucial for understanding the physiological and pathological processes related to protein nitration, such as inflammation, aging, and cancer. Since the nitrated proteins are of low abundance in cells even under induced conditions, no universal and efficient methods have been developed for the profiling and identification of protein nitration sites. Here we describe a protocol for nitropeptide enrichment by using a chemical reduction reaction and biotin labeling, followed by high resolution mass spectrometry. In our method, nitropeptide derivatives can be identified with high accuracy. Our method exhibits two advantages compared to the previously reported methods. First, dimethyl labeling is used to block the primary amine on nitropeptides, which can be used to generate quantitative results. Second, a disulfide bond containing NHS-biotin reagent is used for the enrichment, which can be further reduced and alkylated to enhance the detection signal on a mass spectrometer. This protocol has been successfully applied to the model peptide Angiotensin II in the current paper.

Proteomic profiling of tyrosine-nitrated proteins has been a challenging technique due to the low abundance of the 3-nitrotyrosine modification. Here we describe a novel approach for nitropeptide enrichment and profiling by using Angiotensin II as the model. This method can be extended for other in vitro or in vivo systems.

Protein nitration is one of the most important post-translational modifications (PTM) on tyrosine residues and it can be induced by chemical actions of ...

Measuring Interactions between Fluorescent Probes and Lignin in Plant Sections by sFLIM Based on Native Autofluorescence

In lignocellulosic biomass (LB), the activity of enzymes is limited by the appearance of non-specific interactions with lignin during the hydrolysis process, which maintains enzymes far from their substrate. Characterization of these complex interactions is thus a challenge in complex substrates such as LB. The method here measures molecular interactions between fluorophore-tagged molecules and native autofluorescent lignin, to be revealed by F&#246;rster resonance energy transfer (FRET). Contrary to FRET measurements in living cells using two exogenous fluorophores, FRET measurements in plants using lignin is not trivial due to its complex autofluorescence. We have developed an original acquisition and analysis pipeline with correlated observation of two complementary properties of fluorescence: fluorescence emission and lifetime. sFLIM (spectral and fluorescent lifetime imaging microscopy) provides the quantification of these interactions with high sensitivity, revealing different interaction levels between biomolecules and lignin.

This protocol describes an original setup combining spectral and fluorescence lifetime measurements to evaluate F&#246;rster resonance energy transfer (FRET) between rhodamine-based fluorescent probes and lignin polymer directly in thick plant sections.

In lignocellulosic biomass (LB), the activity of enzymes is limited by the appearance of non-specific interactions with lignin during the hydrolysis ...

Phosphoproteomic Strategy for Profiling Osmotic Stress Signaling in Arabidopsis

Protein phosphorylation is crucial for the regulation of enzyme activity and gene expression under osmotic condition. Mass spectrometry (MS)-based phosphoproteomics has transformed the way of studying plant signal transduction. However, requirement of lots of starting materials and prolonged MS measurement time to achieve the depth of coverage has been the limiting factor for the high throughput study of global phosphoproteomic changes in plants. To improve the sensitivity and throughput of plant phosphoproteomics, we have developed a stop and go extraction (stage) tip based phosphoproteomics approach coupled with Tandem Mass Tag (TMT) labeling for the rapid and comprehensive analysis of plant phosphorylation perturbation in response to osmotic stress. Leveraging the simplicity and high throughput of stage tip technique, the whole procedure takes approximately one hour using two tips to finish phosphopeptide enrichment, fractionation, and sample cleaning steps, suggesting an easy-to-use and high efficiency of the approach. This approach not only provides an in-depth plant phosphoproteomics analysis (&#62; 11,000 phosphopeptide identification) but also demonstrates the superior separation efficiency (&#60; 5% overlap) between adjacent fractions. Further, multiplexing has been achieved using TMT labeling to quantify the phosphoproteomic changes of wild-type and snrk2 decuple mutant plants. This approach has successfully been used to reveal the phosphorylation events of Raf-like kinases in response to osmotic stress, which sheds light on the understanding of early osmotic signaling in land plants.

Phosphoproteomic Strategy for Profiling Osmotic Stress Signaling in Arabidopsis

Presented here is a phosphoproteomic approach, namely stop and go extraction tip based phosphoproteomic, which provides high-throughput and deep coverage of Arabidopsis phosphoproteome. This approach delineates the overview of osmotic stress signaling in Arabidopsis.