This work was supported by the NIH grants NIH R01 DK110355,&#160;DK093776 [N.T.S.], DK102450 [N.T.S.], and P30 DK034987 [to UNC-Chapel Hill]. The authors thank Deekshita Ramanarayanan for assistance with qPCR and western blot experiments.

The protocol is approved and performed in accordance with the Institutional Animal Care and Use Committee (IACUC) at the University of North Carolina.
1. Preparations
<ol>
	<li>Prepare Triton-X buffer (1% Triton X-100, 5 mM ethylenediaminetetraacetic acid (EDTA), bring up volume in phosphate-buffered saline (PBS), pH 7.4). To make 500 mL: stir 5 mL each of Triton X-100 and 500 mM EDTA into 490 mL of PBS, pH 7.4. Store Triton-X buffer solution at 4 &#176;C.</li>
	<li>Prepare High Salt Buffer (10 mM Tris-HCl, pH 7.6, 140 mM NaCl, 1.5 M KCl, 5 mM EDTA, 0.5% Triton X-100, bring up volume in double distilled (dd) H2O). To make 500 mL: stir 10 mL of 0.5 M Tris-HCl (pH 7.6), 14 mL of 5 M NaCl, 55.9 g KCl, 5 mL of 0.5 M EDTA, and 2.5 mL of Triton X-100, adjust volume to 500 mL using double distilled water). Store High Salt Buffer solution at 4 &#176;C.</li>
	<li>Just prior to use, supplement an appropriate amount of Triton-X and High Salt Buffer (e.g. 1 mL for each 25 mg tissue sample) with a protease or protease/phosphatase inhibitor cocktail and discard any unused buffer containing the inhibitors.</li>
	<li>Prepare 5 mM EDTA in 1x PBS, pH 7.4. To make 500 mL: stir 5 mL of 0.5 M EDTA into 495 mL of 1x PBS, pH 7.4.</li>
	<li>Isolate different mouse organs (brain, heart, lung, liver, pancreas, colon, intestine, kidney, spleen) using approved protocols that comply with veterinary guidelines and institutional standards23. The tissue collection procedure should take no more than 5 min to preserve RNA and protein integrity of protease-rich tissues (e.g. pancreas should be processed first).</li>
	<li>Cut a small amount of tissue (~5-20 mg) and place in RNA storage solution, for subsequent RNA extraction, cDNA synthesis, and quantitative real-time PCR analysis for IF gene expression. Place RNA storage solution tubes with tissue at 4 &#176;C overnight and follow manufacturer protocol for further storage and isolation steps.</li>
	<li>Cut the rest of the tissue into smaller fragments (e.g. 0.5 cm) and place in cryovial. Snap-freeze and store vials at -80 &#176;C or liquid nitrogen for longer term storage.</li>
</ol>
2. IF Gene Expression Analysis
<ol>
	<li>Extract RNA from tissues preserved in the RNA storage solution reagent. Use any suitable/preferred RNA extraction method according to manufacturer&#39;s protocol.</li>
	<li>Quantify RNA concentration and use 2 &#956;g of RNA to generate cDNA using a suitable reverse transcription kit according to manufacturer&#39;s protocol.</li>
	<li>Using the generated cDNA and mouse IF gene-specific primers set up qPCR reactions, including three technical replicates of each sample as well as blank control according to manufacturer&#39;s protocol.</li>
	<li>Quantify IF gene expression as fold change comparing different conditions (e.g. young versus old tissue).</li>
</ol>
3. Preparation of Total Tissue Lysates for Immunoblot
<ol>
	<li>Homogenize 25 mg of tissue in 1 mL of 2x non-reducing SDS sample buffer. Omit bromophenol blue dye from the sample buffer if a colorimetric protein assay is to be performed to quantify protein concentration.</li>
	<li>Add 5% (v/v) of 2-mercaptoethanol (2-ME) to make reduced samples. To 200 &#956;L of the non-reducing samples, add 10 &#956;L of 2-ME.</li>
	<li>Determine protein concentration using detergent- and reducing agent-compatible protein assay. If dye is already included in the sample buffer, the protein amount can be estimated after running a gel via a number of techniques, including Coomassie-based stain24.</li>
	<li>Vortex all samples and heat at 95 &#176;C for 5 min.</li>
	<li>Perform western blotting under both reducing and non-reducing conditions. Exposure membrane briefly (&#60;1 min) to reveals monomeric species, and longer (&#62;1 min) to reveal high molecular mass complexes containing IF proteins. If monitoring aggregation, examine entire gel/membrane (including the bottoms of the gel wells).</li>
	<li>Run a parallel gel and stain with a protein stain as a loading control24. In disease models or injury experiments total protein stain should be used as a loading control, as opposed to immunoblots for &#39;housekeeping&#39; proteins (e.g. actin, GAPDH) because the latter change under different stress conditions.</li>
</ol>
4. Preparation of Detergent-soluble and High-salt Extracts of Tissue-specific IFs
<ol>
	<li>Add 1 mL of ice-cold Triton X-100 buffer into a glass tube homogenizer and place it on ice.</li>
	<li>Remove a small piece of tissue (~25 mg) from liquid nitrogen storage and place directly into the glass homogenizer. Use a polytetrafluoroethylene pestle to homogenize (50 strokes) and avoid making bubbles. Keep the homogenizer and lysate cold at all times.</li>
	<li>Transfer lysate to a 1.5 mL microcentrifuge tube on ice and centrifuge at 20,000 x g for 10 min in a pre-chilled centrifuge (4 &#176;C).</li>
	<li>Collect the supernatant fraction into a separate tube. This is the Triton X-soluble fraction, which can be used for immunoprecipitation (i.p.) and analysis of the detergent-soluble pool of IF proteins. Note that steps 4.1-4.4 may be repeated to achieve a cleaner IF extract from brain tissue.</li>
	<li>Add 1 mL of High Salt Buffer to the tissue pellet, transfer to a clean homogenizer and dounce 100 strokes. Transfer the homogenate back to the microcentrifuge tube and place the tube on a rotating shaker in the cold room for 1 h.</li>
	<li>Centrifuge the homogenates at 20,000 x g for 20 min at 4 &#176;C. Discard the supernatant.</li>
	<li>Add 1 mL of ice-cold PBS/EDTA buffer to the pellet and homogenize the pellet (20 strokes) in a clean homogenizer as a final clean-up step*. Transfer to a new tube and centrifuge at 20,000 x g for 10 min at 4 &#176;C to obtain the IF protein-rich high salt extract (HSE). 
	* Optionally, vortex instead of homogenization at this step.</li>
	<li>Discard the supernatant and dissolve the pellets in 300 &#956;L of non-reducing SDS sample buffer that has been pre-heated. Break up the pellet initially by pipetting and vortexing, and then heat the samples for 5 min at 95 &#176;C.</li>
	<li>Vortex and pipet as needed to ensure the pellet is dissolved. It may take several minutes to fully dissolve the pellets.</li>
	<li>Store all samples at -20 &#176;C until analysis.</li>
</ol>
5. Automated Tissue Lysis for IF Protein Extraction in High-volume Experiments
<ol>
	<li>For RNA extraction, place lysis buffer (600 &#956;L buffer per 25 mg of tissue) in a tube containing lysing matrix D (uses small ceramic spheres) and pulse twice for 25 s in the tissue lyser. Separate lysate from the matrix by centrifugation at 20,000 x g and proceed to the next step in isolation.</li>
	<li>For protein extraction, place Triton X-100 (or SDS sample buffer if preparing total lysate) in a lysing tube with lysing matrix SS (use a single stainless steel bead). After testing multiple matrices, this was selected because it produces IF protein extracts that are in similar quality as the traditional douncing method. Note that the automated method is not optimal for pancreas and spleen, and the standard homogenization protocol should be used for these tissues.</li>
	<li>To proceed with preparation of High Salt Extract, remove the stainless steel bead from the tube using a magnet, and centrifuge the tubes at 20,000 x g for 10 min at 4 &#176;C.</li>
	<li>Continue with step 4.4 (above) of the manual protocol.</li>
	<li>Store all samples at -20 &#176;C until analysis.</li>
</ol>
6. Immuno-enrichment of Post-translationally Modified IF Proteins
<ol>
	<li>Prepare PBST buffer (0.02% Tween-20 in PBS). To make 50 mL, add 10 &#956;L of Tween-20 to 50 mL of PBS, pH 7.4.</li>
	<li>Prepare PTM antibody solution (1-10 &#956;g of antibody in 200 &#956;L of PBST). In general, 3 &#956;g of antibody/reaction is a good starting condition that can be further optimized if needed.</li>
	<li>For each reaction, aliquot 50 &#956;L of magnetic beads into a microcentrifuge tube, place on the magnet and aspirate the bead storage solution.</li>
	<li>Conjugate the beads to the immunoprecipitation antibody by re-suspending in the antibody solution and incubating on rotator (end-over-end, to ensure mixing of small volumes) at room temperature for 20 min.</li>
	<li>Place the tubes on magnet and aspirate antibody solution.</li>
	<li>Rinse the antibody-conjugated beads once in 200 &#956;L PBST and remove wash buffer.</li>
	<li>Add 0.6-1 mL of the tissue lysate to the beads, mix by gentle pipetting and incubate for 3 h on rotator in a cold room.</li>
	<li>Place tubes on magnet, remove lysate, and wash the beads five times with 200 &#956;L of PBST. After the last washing step, collect the beads in 100 &#956;L of PBS (no Tween-20) and transfer to a clean new tube. Place the tube on the magnet.</li>
	<li>Aspirate the PBS and add 100 &#956;L of non-reducing sample buffer. Remove 50 &#956;L and add 2-ME (5%) to make reducing samples. Heat the samples to 95 &#176;C for 5 min.</li>
	<li>Separate the i.p. fraction from the beads on the magnet and collect it into a new tube.</li>
	<li>Store samples at -20 &#176;C until analysis.</li>
</ol>
7. Preparation of IF Protein Samples for Mass Spectrometry Analysis
<ol>
	<li>Schedule a consultation with a proteomics expert prior to initiating a study since there is significant time and cost involved with mass spectrometry analysis.</li>
	<li>Take special precautions to avoid contamination. Handle all gels with clean gloves and incubate in clean containers, washed only using ddH2O (avoid soap).</li>
	<li>Run 20-50 &#956;L of the HSE sample (from Sections 4 and 5) on an SDS-PAGE gel according to standard conditions.</li>
	<li>Stain with a protein stain for 1 h. Rinse multiple times and de-stain in ddH2O overnight. The IF protein bands should be easily visible after de-staining.</li>
	<li>Place the gel between plastic sheet protectors, scan and mark the bands that will be excised and sent for analysis.</li>
	<li>Excise the IF protein bands using a new clean razor.</li>
	<li>Place the gel bands in clean microcentrifuge tubes and transfer to a mass spectrometry facility.</li>
</ol>

The authors declare that they have no competing financial interests.

Methods that enable biochemical characterization of IF proteins can be useful to understand numerous pathophysiological phenomena in mammalian systems, since IF proteins are both markers and modulators of cellular and tissue stress29. The principle behind the current method is based on the initial procedures developed in the 1970s and 1980s to isolate, separate and reconstitute IF proteins from cells and tissues, generally employing low and high salt solutions and Triton-X100 detergent<sup class="...

IFs are a family of proteins that in humans are encoded by 73 genes and categorized into six major types: types I-IV are cytoplasmic (e.g. epithelial and hair keratins (K), myocyte desmin, neurofilaments, glial fibrillary acidic protein (GFAP), and others); type V are the nuclear lamins; and type VI are IFs in the eye lens1. In terms of their molecular organization, IF proteins have three common domains: a highly conserved coiled-coil &#34;rod&#34; domain, and globular &#34;head&#34; and &#34;tail&#34; domains. IF protein tetramers assemble to form short filament precursors, which are ultimately incorporated into mature filaments that shape dynamic cytoskeletal and nucleoskeletal structures involved in mechanical protection2, stress sensing3,4, regulation of transcription5 and growth, and other critical cellular functions1,6,7.
The functional importance of the IF system is highlighted by the existence of many human diseases caused by missense mutations in IF genes, including neuropathies, myopathies, skin fragility disorders, metabolic dysfunctions, and premature aging syndromes8. Some IF gene mutations do not cause, but predispose their carriers to disease progression, such as the simple epithelial keratins in liver disease9. The latter is due to the critical stress-protective functions of IFs in epithelia. IFs in general are among the most abundant cellular proteins under basal conditions, but are further strongly induced during various types of stress10. For example, recent studies evaluating proteome-wide changes in the nematode C. elegans demonstrated that multiple IFs are highly upregulated and prone to aggregation during organismal aging11,12. Since maintenance of a proper IF structure is essential for cellular resistance to various forms of stress10, IF aggregation may also contribute to the functional decline during aging. However, organismal-level studies examining multiple mammalian IF proteins across different tissues undergoing stress are lacking.
IFs are highly dynamic structures that adapt to meet cellular demands. Keratins, for example, undergo a biosynthesis-independent cycling between soluble (non-filamentous) and insoluble (filamentous) protein pool13. Under normal physiologic conditions approximately 5% the total K8/K18 pool can be extracted in detergent-free buffer, in comparison to approximately 20% that can be solubilized in the non-ionic detergent Nonidet P-40, which is biochemically comparable to Triton-X10014,15. During mitosis there is a notable increase in the solubility of simple-type epithelial K8 and K1814, which is less apparent in epidermal keratins but more apparent in vimentin and other type III IF proteins15,16. Solubility properties of IF proteins are tightly regulated by phosphorylation, a key post-translational modification (PTM) for filament rearrangement and solubility17,18,19,20. Most IFs undergo extensive regulation by a number of PTMs at conserved sites, resulting in functional changes17.
The purpose of this method is to introduce investigators who are new to the IF field to biochemical extraction and analytical methods for the study of IF proteins across multiple mouse tissues. Specifically, we focus on isolation of IF proteins using a high-salt extraction method and assessment of changes in PTMs via mass-spectrometry and by PTM-targeting antibodies. These methods build upon previously published procedures21 but include modifications for extracting different IF protein types to uncover common mechanism for regulation across the IF family. For example, K8 acetylation at a specific lysine residue regulates filament organization, while hyperacetylation promotes K8 insolubility and aggregate formation22. Recent global proteomic profiling studies have additionally revealed that most tissue-specific IF proteins are also targets for acetylation and that most IF acetylation sites are confined to the highly conserved rod domain. This highlights the need for methods suitable for global profiling of the IF system. We also introduce a rapid method of isolating IF proteins from multiple tissues using automated homogenization in optimized lysing matrix. The resulting preparations are suitable for downstream PTM analysis via mass spectrometry and other methods.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Dynabeads Protein G</td>
			<td>ThermoFisher Scientific</td>
			<td>10009</td>
			<td>immunoprecipitation beads</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>ThermoFisher Scientific</td>
			<td>10010049</td>
			<td>for buffers</td>
		</tr>
		<tr>
			<td>Purelink RNA mini kit</td>
			<td>ThermoFisher Scientific</td>
			<td>12183018</td>
			<td>RNA extraction from tissue</td>
		</tr>
		<tr>
			<td>Purelink DNAse set</td>
			<td>ThermoFisher Scientific</td>
			<td>12185010A</td>
			<td>on column DNA digestion</td>
		</tr>
		<tr>
			<td>Dynamag-2</td>
			<td>ThermoFisher Scientific</td>
			<td>12321D</td>
			<td>magnet for use with dynabeads</td>
		</tr>
		<tr>
			<td>GelCode Blue Stain Reagent</td>
			<td>ThermoFisher Scientific</td>
			<td>24592</td>
			<td>mass spectrometry-compatible gel stain</td>
		</tr>
		<tr>
			<td>Pierce ECL Western Blotting Substrate</td>
			<td>ThermoFisher Scientific</td>
			<td>32106</td>
			<td>for use in western blot</td>
		</tr>
		<tr>
			<td>High Capacity cDNA reverse transcription kit</td>
			<td>ThermoFisher Scientific</td>
			<td>4368813</td>
			<td>for use in gene expression analysis</td>
		</tr>
		<tr>
			<td>Proflex 3x32-well PCR System</td>
			<td>ThermoFisher Scientific</td>
			<td>4484073</td>
			<td>PCR system</td>
		</tr>
		<tr>
			<td>PVDF transfer membrane&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>88520</td>
			<td>for western blot</td>
		</tr>
		<tr>
			<td>Power Up SYBR master mix</td>
			<td>ThermoFisher Scientific</td>
			<td>A25778</td>
			<td>for qPCR analysis</td>
		</tr>
		<tr>
			<td>RNAlater</td>
			<td>ThermoFisher Scientific</td>
			<td>AM7020</td>
			<td>solution for tissue storage prior to RNA isolation</td>
		</tr>
		<tr>
			<td>Novex 4-20% Tris Glycine Gel</td>
			<td>ThermoFisher Scientific</td>
			<td>XP04205BOX</td>
			<td>Precast protein gel</td>
		</tr>
		<tr>
			<td>Anti-Keratin 8 antibody (TS1)</td>
			<td>ThermoFisher Scientific</td>
			<td>MA514428</td>
			<td>for western blot detection of K8</td>
		</tr>
		<tr>
			<td>Anti-Vimentin</td>
			<td>ThermoFisher Scientific</td>
			<td>MA511883</td>
			<td>for western blot detection of vimentin</td>
		</tr>
		<tr>
			<td>Tris Glycine Transfer Buffer (25X)</td>
			<td>ThermoFisher Scientific</td>
			<td>LC3675</td>
			<td>for wet transfer of protein gels</td>
		</tr>
		<tr>
			<td>2X SDS Sample Buffer</td>
			<td>ThermoFisher Scientific</td>
			<td>LC2676</td>
			<td>for preparing protein gel samples</td>
		</tr>
		<tr>
			<td>Tris Glycine SDS Running Buffer (10X)</td>
			<td>ThermoFisher Scientific</td>
			<td>LC26755</td>
			<td>for running protein gels</td>
		</tr>
		<tr>
			<td>Lysing beads - Matrix D</td>
			<td>MP Biomedicals</td>
			<td>116913100</td>
			<td>Lysis beads and matrix tubes for tissue disruption and RNA extraction</td>
		</tr>
		<tr>
			<td>Lysing beads - Matrix SS</td>
			<td>MP Biomedicals</td>
			<td>116921100</td>
			<td>Lysis beads and matrix tubes for tissue disruption and protein extraction</td>
		</tr>
		<tr>
			<td>NanoDrop Lite Spectrophotometer</td>
			<td>ThermoFisher Scientific</td>
			<td>ND-LITE</td>
			<td>measurement of protein and nucleic acid</td>
		</tr>
		<tr>
			<td>Precellys 24 homogenizer</td>
			<td>Bertin Instruments</td>
			<td>EQ03119</td>
			<td>Automated tissue homogenizer</td>
		</tr>
	</tbody>
</table>

isolation of intermediate filament proteins from multiple mouse tissues to study aging-associated post-translational modifications

Intermediate filaments (IFs), together with actin filaments and microtubules, form the cytoskeleton &#8211; a critical structural element of every cell. Normal functioning IFs provide cells with mechanical and stress resilience, while a dysfunctional IF cytoskeleton compromises cellular health and has been associated with many human diseases. Post-translational modifications (PTMs) critically regulate IF dynamics in response to physiological changes and under stress conditions. Therefore, the ability to monitor changes in the PTM signature of IFs can contribute to a better functional understanding, and ultimately conditioning, of the IF system as a stress responder during cellular injury. However, the large number of IF proteins, which are encoded by over 70 individual genes and expressed in a tissue-dependent manner, is a major challenge in sorting out the relative importance of different PTMs. To that end, methods that enable monitoring of PTMs on IF proteins on an organism-wide level, rather than for isolated members of the family, can accelerate research progress in this area. Here, we present biochemical methods for the isolation of the total, detergent-soluble, and detergent-resistant fraction of IF proteins from 9 different mouse tissues (brain, heart, lung, liver, small intestine, large intestine, pancreas, kidney, and spleen). We further demonstrate an optimized protocol for rapid isolation of IF proteins by using lysing matrix and automated homogenization of different mouse tissues. The automated protocol is useful for profiling IFs in experiments with high sample volume (such as in disease models involving multiple animals and experimental groups). The resulting samples can be utilized for various downstream analyses, including mass spectrometry-based PTM profiling. Utilizing these methods, we provide new data to show that IF proteins in different mouse tissues (brain and liver) undergo parallel changes with respect to their expression levels and PTMs during aging.

A new rapid method for high salt-based extraction of IF proteins from multiple mouse tissues using lysing matrix. 
The traditional method25,26 of isolating the bulk of the intermediate filament protein fraction from epithelial tissue was modified here to include 9 different organs and a more rapid procedure for tiss...

Watch this Scientific Journal Video about Isolation of Intermediate Filament Proteins from Multiple Mouse Tissues to Study Aging-associated Post-translational Modifications at JoVE.com

Isolation of Intermediate Filament Proteins from Multiple Mouse Tissues to Study Aging-associated Post-translational Modifications

In this method, we present biochemical procedures for rapid and efficient isolation of intermediate filament (IF) proteins from multiple mouse tissues. Isolated IFs can be used to study changes in post-translational modifications by mass spectrometry and other biochemical assays.

Intermediate filaments (IFs), together with actin filaments and microtubules, form the cytoskeleton &#8211; a critical structural element of every cell. ...

isolation-intermediate-filament-proteins-from-multiple-mouse-tissues

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

8.4K Views.  University of North Carolina at Chapel Hill. In this method, we present biochemical procedures for rapid and efficient isolation of intermediate filament (IF) proteins from multiple mouse tissues. Isolated IFs can be used to study changes in post-translational modifications by mass spectrometry and other biochemical assays.

Video: Isolation of Intermediate Filament Proteins from Multiple Mouse Tissues to Study Aging-associated Post-translational Modifications

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies have revealed that these reversible RNA modifications affect RNA splicing, translation, degradation, and localization. Multiple physiological processes, like circadian rhythms, stem cell pluripotency, fibrosis, triglyceride metabolism, and obesity are also controlled by m6A modifications. Immunoprecipitation/sequencing, mass spectrometry, and modified northern blotting are some of the methods commonly employed to measure m6A modifications. Herein, we present a northeastern blotting technique for measuring m6A modifications. The current protocol provides good size separation of RNA, better accommodation and standardization for various experimental designs, and clear delineation of m6A modifications in various sources of RNA. While m6A modifications are known to have a crucial impact on human physiology relating to circadian rhythms and obesity, their roles in other (patho)physiological states are unclear. Therefore, investigations on m6A modifications have immense possibility to provide key insights into molecular physiology.

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

A modified northern blotting method for measuring N6-methyladenosine (m6A) modifications in RNA is described. The current method can detect modifications in diverse RNAs and controls under various experimental designs.

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies ...

Nuclear Magnetic Resonance Spectroscopy for the Identification of Multiple Phosphorylations of Intrinsically Disordered Proteins

Aggregates of the neuronal Tau protein are found inside neurons of Alzheimer's disease patients. Development of the disease is accompanied by increased, abnormal phosphorylation of Tau. In the course of the molecular investigation of Tau functions and dysfunctions in the disease, nuclear magnetic resonance (NMR) spectroscopy is used to identify the multiple phosphorylations of Tau. We present here detailed protocols of recombinant production of Tau in bacteria, with isotopic enrichment for NMR studies. Purification steps that take advantage of Tau's heat stability and high isoelectric point are described. The protocol for in vitro phosphorylation of Tau by recombinant activated ERK2 allows for generating multiple phosphorylations. The protein sample is ready for data acquisition at the issue of these steps. The parameter setup to start recording on the spectrometer is considered next. Finally, the strategy to identify phosphorylation sites of modified Tau, based on NMR data, is explained. The benefit of this methodology compared to other techniques used to identify phosphorylation sites, such as immuno-detection or mass spectrometry (MS), is discussed.

We describe here a method to identify multiple phosphorylations of an intrinsically disordered protein by Nuclear Magnetic Resonance Spectroscopy (NMR), using Tau protein as a case study. Recombinant Tau is isotopically enriched and modified in vitro by a kinase prior to data acquisition and analysis.

Aggregates of the neuronal Tau protein are found inside neurons of Alzheimer's disease patients. Development of the disease is accompanied by increased, ...

Utilizing a Comprehensive Immunoprecipitation Enrichment System to Identify an Endogenous Post-translational Modification Profile for Target Proteins

It is now well-appreciated that post-translational modifications (PTMs) play an integral role in regulating a protein&#39;s structure and function, which may be essential for a given protein&#39;s role both physiologically and pathologically. Enrichment of PTMs is often necessary when investigating the PTM status of a target protein, because PTMs are often transient and relatively low in abundance. Many pitfalls are encountered when enriching for a PTM of a target protein, such as buffer incompatibility, the target protein antibody is not IP-compatible, loss of PTM signal, and others. The degree of difficulty is magnified when investigating multiple PTMs like acetylation, ubiquitination, SUMOylation 2/3, and tyrosine phosphorylation for a given target protein. Studying a combination of these PTMs may be necessary, as crosstalk between PTMs is prevalent and critical for protein regulation. Often, these PTMs are studied in different lysis buffers and with unique inhibitor compositions. To simplify the process, a unique denaturing lysis system was developed that effectively isolates and preserves these four PTMs; thus, enabling investigation of potential crosstalk in a single lysis system. A unique filter system was engineered to remove contaminating genomic DNA from the lysate, which is a problematic by-product of denaturing buffers. Robust affinity matrices targeting each of the four PTMs were developed in concert with the buffer system to maximize the enrichment and detection of the endogenous states of these four PTMs. This comprehensive PTM detection toolset streamlines the process of obtaining critical information about whether a protein is modified by one or more of these PTMs.

Investigating multiple, endogenous post-translational modifications for a target protein can be extremely challenging. Techniques described here utilize an optimized lysis buffer and filter system developed with specific PTM-targeting, affinity matrices to detect the acetylation, ubiquitination, SUMOylation 2/3, and tyrosine phosphorylation modifications for a target protein using a single, streamlined system.

It is now well-appreciated that post-translational modifications (PTMs) play an integral role in regulating a protein&#39;s structure and function, which ...

Leaf Spray Mass Spectrometry: A Rapid Ambient Ionization Technique to Directly Assess Metabolites from Plant Tissues

Plants produce thousands of small molecules that are diverse in their chemical properties. Mass spectrometry (MS) is a powerful technique for analyzing plant metabolites because it provides molecular weights with high sensitivity and specificity. Leaf spray MS is an ambient ionization technique where plant tissue is used for direct chemical analysis via electrospray, eliminating chromatography from the process. This approach to sampling metabolites allows for a wide range of chemical classes to be detected simultaneously from intact plant tissues, minimizing the amount of sample preparation needed. When used with a high-resolution, accurate mass MS, leaf spray MS facilitates the rapid detection of metabolites of interest. It is also possible to collect tandem mass fragmentation data with this technique to facilitate a compound identification. The combination of accurate mass measurements and fragmentation is beneficial in confirming compound identities. The leaf spray MS technique requires only minor modifications to a nanospray ionization source and is a useful tool to further expand the capabilities of a mass spectrometer. Here, fresh leaf tissue from Sceletium tortuosum (Aizoaceae), a traditional medicinal plant from South Africa, is analyzed; numerous mesembrine alkaloids are detected with leaf spray MS.

Leaf spray mass spectrometry is a direct chemical analysis technique that minimizes the sample preparation and eliminates chromatography, allowing for the rapid detection of small molecules from plant tissues.

Plants produce thousands of small molecules that are diverse in their chemical properties. Mass spectrometry (MS) is a powerful technique for analyzing ...

Isolation of Glomeruli and In Vivo Labeling of Glomerular Cell Surface Proteins

Proteinuria results from the disruption of the glomerular filter that is composed of the fenestrated endothelium, glomerular basement membrane, and podocytes with their slit diaphragms. The delicate structure of the glomerular filter, especially the slit diaphragm, relies on the interplay of diverse cell surface proteins. Studying these cell surface proteins has so far been limited to in vitro studies or histologic analysis. Here, we present a murine in vivo biotinylation labeling method, which enables the study of glomerular cell surface proteins under physiologic and pathophysiologic conditions. This protocol contains information on how to perfuse mouse kidneys, isolate glomeruli, and perform endogenous immunoprecipitation of a protein of interest. Semi-quantitation of glomerular cell surface abundance is readily available with this novel method, and all proteins accessible to biotin perfusion and immunoprecipitation can be studied. In addition, isolation of glomeruli with or without biotinylation enables further analysis of glomerular RNA and protein as well as primary glomerular cell culture (i.e., primary podocyte cell culture).

Isolation of Glomeruli and In Vivo Labeling of Glomerular Cell Surface Proteins

Here we present a protocol for murine in vivo labeling of glomerular cell surface proteins with biotin. This protocol contains information on how to perfuse mouse kidneys, isolate glomeruli, and perform endogenous immunoprecipitation of the protein of interest.

Proteinuria results from the disruption of the glomerular filter that is composed of the fenestrated endothelium, glomerular basement membrane, and ...

Looking Outwards: Isolation of Cyanobacterial Released Carbohydrate Polymers and Proteins

Cyanobacteria can actively secrete a wide range of biomolecules into the extracellular environment, such as heteropolysaccharides and proteins. The identification and characterization of these biomolecules can improve knowledge about their secretion pathways and help to manipulate them. Furthermore, some of these biomolecules are also interesting in terms of biotechnological applications. Described here are two protocols for easy and rapid isolation of cyanobacterial released carbohydrate polymers and proteins. The method for isolation of released carbohydrate polymers is based on conventional precipitation techniques of polysaccharides in aqueous solutions using organic solvents. This method preserves the characteristics of the polymer and simultaneously avoids the presence of contaminants from cell debris and culture medium. At the end of the process, the lyophilized polymer is ready to be used or characterized or can be subjected to further rounds of purification, depending on the final intended use. Regarding the isolation of the cyanobacterial exoproteome, the technique is based on the concentration of the cell-free medium after removal of the major contaminants by centrifugation and filtration. This strategy allows for reliable isolation of proteins that reach the extracellular milieu via membrane transporters or outer membrane vesicles. These proteins can be subsequently identified using standard mass spectrometry techniques. The protocols presented here can be applied not only to a wide range of cyanobacteria, but also to other bacterial strains. Furthermore, these procedures can be easily tailored according to the final use of the products, purity degree required, and bacterial strain.

Here, protocols for the isolation of cyanobacterial released carbohydrate polymers and isolation of their exoproteomes are described. Both procedures embody key steps to obtain polymers or proteins with high purity degrees that can be used for further analysis or applications. They can also be easily adapted according to specific user needs.

Cyanobacteria can actively secrete a wide range of biomolecules into the extracellular environment, such as heteropolysaccharides and proteins. The ...

Profiling Ubiquitin and Ubiquitin-like Dependent Post-translational Modifications and Identification of Significant Alterations

Ubiquitin (ub) and ubiquitin-like (ubl) dependent post-translational modifications of proteins play fundamental biological regulatory roles within the cell by controlling protein stability, activity, interactions, and intracellular localization. They enable the cell to respond to signals and to adapt to changes in its environment. Alterations within these mechanisms can lead to severe pathological situations such as neurodegenerative diseases and cancers. The aim of the technique described here is to establish ub/ubls dependent PTMs profiles, rapidly and accurately, from cultured cell lines. The comparison of different profiles obtained from different conditions allows the identification of specific alterations, such as those induced by a treatment for example. Lentiviral mediated cell transduction is performed to create stable cell lines expressing a two-tags (6His and Flag) version of the modifier (ubiquitin or a ubl such as SUMO1 or Nedd8). These tags permit the purification of ubiquitin and therefore of ubiquitinated proteins from the cells. This is done through a two-step purification process: The first one is performed in denaturing conditions using the 6His tag, and the second one in native conditions using the Flag tag. This leads to a highly specific and pure isolation of modified proteins which are subsequently identified and semi-quantified by liquid chromatography followed by tandem mass spectrometry (LC-MS/MS) technology. Easy informatics analysis of MS data using Excel software enables the establishment of PTM profiles by eliminating background signals. These profiles are compared between each condition in order to identify specific alterations which will then be studied more specifically, starting with their validation by standard biochemistry techniques.

This protocol aims at establishing ubiquitin (Ub) and ubiquitin-likes (Ubls) specific proteomes in order to identify alterations of these kind of post-translational modifications (PTMs), associated with a specific condition such as a treatment or a phenotype.

Ubiquitin (ub) and ubiquitin-like (ubl) dependent post-translational modifications of proteins play fundamental biological regulatory roles within the ...

Transverse Sectioning of Mature Rice (Oryza sativa L.) Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm phenotyping can be used to classify genotypes based on SG morphotype using scanning electron microscopic (SEM) analysis. SGs can be visualized using SEM by slicing through the kernel (pericarp, aleurone layers, and endosperm) and exposing the organellar contents. Current methods require the rice kernel to be embedded in plastic resin and sectioned using a microtome or embedded in a truncated pipette tip and sectioned by hand using a razor blade. The former method requires specialized equipment and is time-consuming, while the latter introduces a new host of problems depending on rice genotype. Chalky rice varieties, particularly, pose a problem for this type of sectioning due to the friable nature of their endosperm tissue. Presented here is a technique for preparing translucent and chalky rice kernel sections for microscopy, requiring only pipette tips and a scalpel blade. Preparing the sections within the confines of a pipette tip prevents rice kernel endosperm from shattering (for translucent or &#8216;vitreous&#8217; phenotypes) and crumbling (for chalky phenotypes). Using this technique, endosperm cell patterning and the structure of intact SGs can be observed.

Transverse Sectioning of Mature Rice Oryza sativa L. Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

This protocol allows for the preparation of transverse sections of cereal seeds (e.g., rice) for the analysis of endosperm and starch granule morphology using scanning electron microscopy.

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm ...

Purification of Tubulin with Controlled Posttranslational Modifications and Isotypes from Limited Sources by Polymerization-Depolymerization Cycles

One important aspect of studies of the microtubule cytoskeleton is the investigation of microtubule behavior in in vitro reconstitution experiments. They allow the analysis of the intrinsic properties of microtubules, such as dynamics, and their interactions with microtubule-associated proteins (MAPs). The &#8220;tubulin code&#8221; is an emerging concept that points to different tubulin isotypes and various posttranslational modifications (PTMs) as regulators of microtubule properties and functions. To explore the molecular mechanisms of the tubulin code, it is crucial to perform in vitro reconstitution experiments using purified tubulin with specific isotypes and PTMs.
To date, this was technically challenging as brain tubulin, which is widely used in in vitro experiments, harbors many PTMs and has a defined isotype composition. Hence, we developed this protocol to purify tubulin from different sources and with different isotype compositions and controlled PTMs, using the classical approach of polymerization and depolymerization cycles. Compared to existing methods based on affinity purification, this approach yields pure, polymerization-competent tubulin, as tubulin resistant to polymerization or depolymerization is discarded during the successive purification steps.
We describe the purification of tubulin from cell lines, grown either in suspension or as adherent cultures, and from single mouse brains. The method first describes the generation of cell mass in both suspension and adherent settings, the lysis step, followed by the successive stages of tubulin purification by polymerization-depolymerization cycles. Our method yields tubulin that can be used in experiments addressing the impact of the tubulin code on the intrinsic properties of microtubules and microtubule interactions with associated proteins.

This protocol describes tubulin purification from small/medium-scale sources such as cultured cells or single mouse brains, using polymerization and depolymerization cycles. The purified tubulin is enriched in specific isotypes or has specific posttranslational modifications and can be used in in vitro reconstitution assays to study microtubule dynamics and interactions.

One important aspect of studies of the microtubule cytoskeleton is the investigation of microtubule behavior in in vitro reconstitution experiments. They ...

Purification of Ubiquitinated p53 Proteins from Mammalian Cells

Ubiquitination is a type of posttranslational modification that regulates not only the stability but also the localization and function of a substrate protein. The ubiquitination process occurs intracellularly in eukaryotes and regulates almost all basic cellular biological processes. Purification of ubiquitinated proteins aids the investigation of the role of ubiquitination in controlling the function of substrate proteins. Here, a step-by-step procedure to purify ubiquitinated proteins in mammalian cells is described with the p53 tumor suppressor protein as an example. Ubiquitinated p53 proteins were purified under stringent nondenaturing and denaturing conditions. Total cellular Flag-tagged p53 protein was purified with anti-Flag antibody-conjugated agarose under nondenaturing conditions. Alternatively, total cellular His-tagged ubiquitinated protein was purified using nickel-charged resin under denaturing conditions. Ubiquitinated p53 proteins in the eluates were successfully detected with specific antibodies. Using this procedure, the ubiquitinated forms of a given protein can be efficiently purified from mammalian cells, facilitating studies on the roles of ubiquitination in regulating protein function.

The protocol describes a step-by-step method to purify ubiquitinated proteins from mammalian cells using the p53 tumor suppressor protein as an example. Ubiquitinated p53 proteins were purified from cells under stringent nondenaturing and denaturing conditions.

Ubiquitination is a type of posttranslational modification that regulates not only the stability but also the localization and function of a substrate ...

Isolation of Intermediate Filament Proteins from Multiple Mouse Tissues to Study Aging-associated Post-translational Modifications

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Chapters in this video

A Method for Measuring RNA N⁶-methyladenosine Modifications in Cells and Tissues

Nuclear Magnetic Resonance Spectroscopy for the Identification of Multiple Phosphorylations of Intrinsically Disordered Proteins

Utilizing a Comprehensive Immunoprecipitation Enrichment System to Identify an Endogenous Post-translational Modification Profile for Target Proteins

Leaf Spray Mass Spectrometry: A Rapid Ambient Ionization Technique to Directly Assess Metabolites from Plant Tissues

Isolation of Glomeruli and In Vivo Labeling of Glomerular Cell Surface Proteins

Looking Outwards: Isolation of Cyanobacterial Released Carbohydrate Polymers and Proteins

Profiling Ubiquitin and Ubiquitin-like Dependent Post-translational Modifications and Identification of Significant Alterations

Transverse Sectioning of Mature Rice (Oryza sativa L.) Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

Purification of Tubulin with Controlled Posttranslational Modifications and Isotypes from Limited Sources by Polymerization-Depolymerization Cycles

Purification of Ubiquitinated p53 Proteins from Mammalian Cells