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 detergent25,30,31,32,33,34,35. For historical insight into the studies that supported the biochemical isolation of IF proteins please see recent reviews36,37. The current method is based on more recent protocols developed for the study of simple epithelial keratins26. The advantage of the method is that it can serve as an initial step to enable investigators who are new to the IF field to effectively isolate IF proteins from most mammalian tissues. It represents a visual guide of related procedures that are widely used by investigators in the field to study IF protein regulation21,38.
This technique can be used to study the regulation and function of IFs in stress communication mechanisms between mammalian tissues39,40. It is becoming well-appreciated that stress in one tissue can affect the functioning of other tissues, for example under conditions of nutritional stress41, protein misfolding42, metabolic stress43, and other changes. Of note, most of the work currently being done in this area is in Drosophila and C. elegans models, while studies on widespread stress responses in mammalian systems are lacking. As the major regulator of cellular stress10, the IF system can unlock clues to organismal-level stress responses, which can have important disease implications. As such, the current protocol can be used to study global pathophysiological mechanisms in various mouse models of stress, injury and disease.
One limitation of the current technique is that the high salt extracts are considered &#34;crude&#34; IF preparations since they also contain other IF-associated proteins, such as plectin for example44. As shown previously, HSEs can be denatured in 8 M urea buffer to obtain highly pure IF proteins that are capable of in vitro re-assembly in phosphate buffer44. After two such cycles of disassembly and re-assembly, the stoichiometry of IF proteins (isolated from HeLa cells) remained the same, whereas the urea treatment removed plectin44. During desmin purification, actin can be removed by solubilization of the HSE in acetic acid45. Whether additional purification steps should be included depends on the ultimate goal of the experiment. For example, if the goal is to characterize the assembly state and generate in vitro reconstituted IFs, the urea step is required to obtain highly pure IF proteins. On the other hand, if the goal is to identify context-dependent associations between IFs and other cellular proteins, then the &#34;crude&#34; preparations can be used. Interacting proteins can be identified by mass spectrometry using either in-gel digest for high abundance targets visible by gel stain, or in-solution digest for low abundance targets. Previous studies using immunoprecipitation of detergent-soluble IFs and HSEs containing IF proteins identified functionally important specific interaction between keratins 8/18 and heat shock protein 70 (Hsp70), which is potentiated after heat stress46, the adaptor protein 14-3-3 after phosphorylation47, and K8/K18 Raf-1 kinase under normal physiologic conditions48 among others. The current protocol can enable similar studies on other IF proteins.
Use of the lysing matrix is a key modification from previous methods and should allow for rapid extraction of IFs from multiple tissues, with the exception of pancreas and spleen. The automation of the initial steps can be particularly useful for high-volume mouse experiments. For example, isolating IF proteins from 3 different tissues from 10 mice per condition (e.g. normal and some form of systemic stress or disease) will require processing of 60 individual tissues. Using the traditional manual method this would need to be done in batches and can take several days. However, using the automated method with lysing matrix, the procedure can be done in just a few hours. Critical steps of the procedure include: ensuring the sample stays cold throughout the procedure; incubation in high salt buffer for 1 h (step 4.5); using hot buffer and extensive vortexing to ensure complete resuspension of the pellet before analysis (step 4.8); and taking all precautions to avoid contamination of samples to be analyzed by mass spectrometry (step 7.2).
The major drawback to mass spectrometry-based identification of PTM sites is that it works very well for certain PTMs &#8211; phosphorylation and lysine acetylation being prime examples &#8211; but others, such as sumoylation are much more challenging49. Nevertheless, proteomics is a powerful tool for studying protein regulation in normal and diseased tissues. Agnetti et al. compiled a comprehensive up-to-date overview of cutting edge proteomic techniques currently in use to study various PTMs in the context of tissue-expressed proteins50. The purpose of the current method is to generate samples that can be applied for different types of analyses. For example, in vitro sumoylation can be performed on immune-precipitated IF proteins, and overall sumoylation can be assessed on isolated IFs in HSE preparations using sumo-specific antibodies18,21. Therefore, the application of this method is not limited to generating samples only for mass spectrometry analysis but can be used to probe multiple IF properties using a variety of downstream techniques.
IF proteins are extensively regulated by numerous PTMs at conserved sites, resulting in altered solubility, filament assembly, and protein interactions17. Recent technological advances have unveiled the incredible magnitude, complexity, and disease significance of post-translational modifications (PTMs) on cellular proteins51,52,53. Improvement in detection methods for common PTMs, such as phosphorylation54 and acetylation51, have yielded vast catalogs of data, but understanding their biological meaning &#8211; cracking the &#34;PTM code&#34; &#8211; is a major remaining challenge55,56. One way to assess the functional roles and relative significance of each PTM is to determine how well it is conserved across multiple members of the IF protein family. For example, identification of a novel phospho-tyrosine site on K8 revealed that this site is contained within a QYE motif found in essentially all cytoplasmic IF proteins and critical for regulating IF protein solubility and filament dynamics57. Importantly, mutation of the conserved tyrosine residue on GFAP to a negatively-charged &#34;phosphomimic&#34; aspartic acid (Y242D) causes Alexander Disease58. The representative data shown here demonstrate how two different tissue-specific IF protein types (K8 and GFAP) undergo similar pattern of upregulation and increased acetylation during aging, which may be an effect of altered metabolism22,51. This example illustrates the utility of the method in understanding the function of IF proteins on a global physiological scale.

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><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 3 x 32-well PCR System</td><td>ThermoFisher Scientific</td><td>4484073</td><td>PCR system</td></tr><tr><td>PVDF transfer membrane&amp;nbsp;</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 tissue lysis. While 3 manual homogenization steps are required for the traditional method, the modified procedure only has 1 manual homogenization step, which shortens the procedure by several hours, especially when processing more than 6 samples. Figure 1A shows a typical result of HSEs from 9 mouse tissues, while the table of expected proteins in the tissue and the molecular weight of each protein is shown in Figure 1B for comparison. Of note, the new automated method does not work well on pancreas and spleen; the traditional homogenization procedure works best for these tissues. This method should be useful for profiling IF proteins in a universal fashion across several tissues to assess organism-level responses to stress. As specific examples here, we show that IF proteins in liver (K8/K18) and brain (GFAP) from old mice are highly upregulated and undergo unique PTMs compared to the corresponding tissues in young mice.
Liver K8 and K18 are strongly upregulated and undergo increased phosphorylation and lysine acetylation in livers from old mice.
Figure 2 shows typical results from several of the analyses described in this protocol. Panel A depicts expression analysis of the two major IF genes in the liver, keratin 8 (KRT8) and keratin 18 (KRT18), which encode the IF proteins K8 and K18, respectively. Epithelial keratins are strongly upregulated under various stress conditions. In the result shown, this occurs during aging, since KRT8 is significantly upregulated in the livers of 24 m old mice compared to 3 m old mice. The results at the protein level are more striking, as observed by the dramatic increase in K8 monomer as well as high molecular mass complexes in the old (24 m) livers. Coomassie-based protein stain here serves as a loading control to ensure equal loading of protein across samples. Note that with total tissue lysates it is easy to overload protein on a gel, as it is in this case. Loading a smaller volume or diluting the sample further in sodium dodecyl sulfate (SDS) buffer will alleviate this problem (especially if it appears viscous and difficult to pipet). Panel C depicts a typical result from a liver High Salt Extract obtained using the automated protocol, demonstrating the strong enrichment of keratins 8 and 18 on the gel. The red lines demarcate the area that was excised and submitted for mass spectrometry analysis. In panel D the results of the mass spec analysis of the samples in panel C shows that K8 and K18 in the old liver have multiple phosphorylation and acetylation sites that are not present in the young liver.
GFAP is strongly upregulated and lysine acetylated in the brains from old mice.
Figure 3 demonstrates that the methods used to extract IFs from epithelia can also be used on non-epithelial tissue. Furthermore, the results reveal a general pattern for aging-dependent upregulation of IF genes and proteins. The qPCR result in Figure 2A reveals a 5-fold induction of GFAP mRNA in the brains of 24 m old mice compared to 3 m old mice. Figure 2B reveals total proteins present in the Triton X-100 fraction and the IF-enriched HSE. Note the increase in the band intensity at 50 kDa marked by the arrow (corresponding to GFAP) in the old brain. Western blot analysis in Figure 2C further reveals the upregulation of GFAP and significant presence of GFAP monomer and potential high molecular mass complex in the Triton X-100 fraction (both marked by arrows). Western blot analysis of the same samples with a pan-acetyl lysine antibody shows that the antibody recognizes a band at ~50 kDa in the HSE of the old mouse brain and at ~250 kDa in the Triton X-100 fraction (Figure 2D). Immuno-enrichment of acetylated proteins from the Triton X-100 fractions reveals increased presence of GFAP protein in the lysate obtained from the old brain. Reducing and non-reducing conditions are shown to highlight GFAP monomer and high molecular mass complexes. Mass spectrometry analysis (similar to Figure 1) can be performed in this case to determine the site-specificity of the acetylated residues on GFAP in the old brain.
<img alt="Figure 1" src="/files/ftp_upload/55655/55655fig1.jpg" /> 
Figure 1: Automated lysis and extraction of IF proteins from multiple mouse tissues. (A) Coomassie-based gel stain of HSEs from the designated mouse tissues. The tissues shown were obtained from the same adult (3 m old) male CBA mouse. Samples were processed as described in Section 5. Brain: note that NFL, NFM, and NFH are heavily phosphorylated27 and migrate slower than expected, at ~70, 145, and 200 kDa, respectively. Heart: In addition to the 53 kDa band corresponding to desmin and vimentin), heart samples also contain a ~42 kDa band, most likely representing actin, which is known to co-purify with desmin28. Large intestine: The identity of the prominent bands above 25 kDa that are co-extracted with K8/K18 are not known, but these bands are not present if the tissues are processed using the traditional dounce homogenizer method. Pancreas and Spleen: Note that the automated homogenization is not suitable for isolation of keratins from pancreas and vimentin from spleen, presumably because of the sensitivity of the lysate to the slight increase in temperature during the pulse in the tissue lyser. (B) Table showing the major IF protein types present in the different tissues and their predicted molecular weight as a reference for panel A. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55655/55655fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55655/55655fig2.jpg" /> 
Figure 2: Molecular and biochemical differences in liver keratins from young and old mice. (A) Analysis of KRT8 and KRT8 mRNA using standard procedure (Protocol 1) reveals significant induction in KRT8 transcript in the livers from old mice. n = 6 for each condition (3 male and 3 female CBA mice were used per group). **p &#60;0.01; one-way ANOVA. (B) Total liver lysates were obtained from 3 young (3 months old) and 3 old (24 months old) male CBA mice using Protocol 2 and the samples were analyzed under non-reducing conditions. TS1 antibody was used to probe for K8 expression and Coomassie-based protein stain was used as a loading control. (C) High salt extracts from young and old mouse livers, corresponding to mice #1 and #4, respectively from panel B. The two arrows point to K8 and K18 on the gel and the red box indicates the part of the gel that was excised and submitted for mass spectrometry analysis. (D) PTM sites identified by mass spec analysis of the samples shown in C. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55655/55655fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55655/55655fig3.jpg" /> 
Figure 3: Molecular and biochemical differences in GFAP from brain of young and old mice. (A) Analysis of GFAP mRNA using standard procedure (Protocol 1) reveals significant induction in the brains from old mice. n = 6 for each condition (3 male and 3 female CBA mice were used per group). **p &#60;0.01; one-way ANOVA. (B) Coomassie-based protein stain of Triton X-100 and HSE fractions from brain tissue of young (3 months old) and old (24 months old) mouse. Note the increase in ~50 kDa GFAP band in HSE from old brain (arrow). (C) GFAP immunoblot (mouse monoclonal; GA5 clone) of the same samples as panel B. (D) Acetyl-lysine immunoblot (rabbit polyclonal; Abcam ab80178) of the same samples as in panel B. (E) GFAP blot after immunoprecipitation with acetyl-lysine antibody. Samples were analyzed under non-reducing (NR) and reducing (R) conditions and arrows point to increase presence of GFAP in the immunoprecipitate from the old brain. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55655/55655fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Biochemistry

8.4K Views.  University of North Carolina at Chapel Hill.  In deze methode presenteren wij biochemische procedures voor snelle en efficiënte isolatie van intermediaire filament (IF) eiwitten uit meerdere muisweefsels. Geïsoleerde IF's kunnen worden gebruikt om veranderingen te onderzoeken in post-translatie modificaties door massaspectrometrie en andere biochemische analyses. 

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

Characterizing Histone Post-translational Modification Alterations in Yeast Neurodegenerative Proteinopathy Models

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives each year. Effective treatment options able to halt disease progression are lacking. Despite the extensive sequencing efforts in large patient populations, the majority of ALS and PD cases remain unexplained by genetic mutations alone. Epigenetics mechanisms, such as the post-translational modification of histone proteins, may be involved in neurodegenerative disease etiology and progression and lead to new targets for pharmaceutical intervention. Mammalian in vivo and in vitro models of ALS and PD are costly and often require prolonged and laborious experimental protocols. Here, we outline a practical, fast, and cost-effective approach to determining genome-wide alterations in histone modification levels using Saccharomyces cerevisiae as a model system. This protocol allows for comprehensive investigations into epigenetic changes connected to neurodegenerative proteinopathies that corroborate previous findings in different model systems while significantly expanding our knowledge of the neurodegenerative disease epigenome.

This protocol outlines experimental procedures to characterize genome-wide changes in the levels of histone post-translational modifications (PTM) occurring in connection with the overexpression of proteins associated with ALS and Parkinson&#39;s disease in Saccharomyces cerevisiae models. After SDS-PAGE separation, individual histone PTM levels are detected with modification-specific antibodies via Western blotting.

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives ...

Genetics

Gait Analysis of Age-dependent Motor Impairments in Mice with Neurodegeneration

Motor behavior tests are commonly used to determine the functional relevance of a rodent model and to test newly developed treatments in these animals. Specifically, gait analysis allows recapturing disease relevant phenotypes that are observed in human patients, especially in neurodegenerative diseases that affect motor abilities such as Parkinson&#39;s disease (PD), Alzheimer&#39;s disease (AD), amyotrophic lateral sclerosis (ALS), and others. In early studies along this line, the measurement of gait parameters was laborious and depended on factors that were hard to control (e.g., running speed, continuous running). The development of ventral plane imaging (VPI) systems made it feasible to perform gait analysis at a large scale, making this method a useful tool for the assessment of motor behavior in rodents. Here, we present an in-depth protocol of how to use kinematic gait analysis to examine the age-dependent progression of motor deficits in mouse models of neurodegeneration; mouse lines with decreased levels of endophilin, in which neurodegenerative damage progressively increases with age, are used as an example.

In this study, we demonstrate the use of kinematic gait analysis based on ventral plane imaging to monitor the subtle changes in motor coordination as well as the progression of neurodegeneration with advancing age in mouse models (e.g., endophilin mutant mouse lines).

Motor behavior tests are commonly used to determine the functional relevance of a rodent model and to test newly developed treatments in these animals. ...

Developmental Biology

Quantitative Microtubule Fractionation Technique to Separate Stable Microtubules, Labile Microtubules, and Free Tubulin in Mouse Tissues

Microtubules, composed of &#945;/&#946;-tubulin dimers, are a crucial component of the cytoskeleton in eukaryotic cells. These tube-like polymers exhibit dynamic instability as tubulin heterodimer subunits undergo repetitive polymerization and depolymerization. Precise control of microtubule stability and dynamics, achieved through tubulin post-translational modifications and microtubule-associated proteins, is essential for various cellular functions. Dysfunctions in microtubules are strongly implicated in pathogenesis, including neurodegenerative disorders. Ongoing research focuses on microtubule-targeting therapeutic agents that modulate stability, offering potential treatment options for these diseases and cancers. Consequently, understanding the dynamic state of microtubules is crucial for assessing disease progression and therapeutic effects.
Traditionally, microtubule dynamics have been assessed in vitro or in cultured cells through rough fractionation or immunoassay, using antibodies targeting post-translational modifications of tubulin. However, accurately analyzing tubulin status in tissues using such procedures poses challenges. In this study, we developed a simple and innovative microtubule fractionation method to separate stable microtubules, labile microtubules, and free tubulin in mouse tissues.
The procedure involved homogenizing dissected mouse tissues in a microtubule-stabilizing buffer at a 19:1 volume ratio. The homogenates were then fractionated through a two-step ultracentrifugation process following initial slow centrifugation (2,400 &#215; g) to remove debris. The first ultracentrifugation step (100,000 &#215; g) precipitated stable microtubules, while the resulting supernatant was subjected to a second ultracentrifugation step (500,000 &#215; g) to fractionate labile microtubules and soluble tubulin dimers. This method determined the proportions of tubulin constituting stable or labile microtubules in the mouse brain. Additionally, distinct tissue variations in microtubule stability were observed that correlated with the proliferative capacity of constituent cells. These findings highlight the significant potential of this novel method for analyzing microtubule stability in physiological and pathological conditions.

Microtubules, which are tubulin polymers, play a crucial role as a cytoskeleton component in eukaryotic cells and are known for their dynamic instability. This study developed a method for fractionating microtubules to separate them into stable microtubules, labile microtubules, and free tubulin to evaluate the stability of microtubules in various mouse tissues.

Microtubules, composed of &#945;/&#946;-tubulin dimers, are a crucial component of the cytoskeleton in eukaryotic cells. These tube-like polymers exhibit ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Biology

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

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

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

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

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

Single Myofiber Isolation and Culture from a Murine Model of Emery-Dreifuss Muscular Dystrophy in Early Post-Natal Development

Autosomal dominant Emery-Dreifuss muscular dystrophy (EDMD) is caused by mutations in the LMNA gene, which encodes the A-type nuclear lamins, intermediate filament proteins that sustain the nuclear envelope and the components of the nucleoplasm. We recently reported that muscle wasting in EDMD can be ascribed to intrinsic epigenetic dysfunctions affecting muscle (satellite) stem cells regenerative capacity. Isolation and culture of single myofibers is one of the most physiological ex-vivo approaches to monitor satellite cells behavior within their niche, as they remain between the basal lamina surrounding the fiber and the sarcolemma. Therefore, it represents an invaluable experimental paradigm to study satellite cells from a variety of murine models. Here, we describe a re-adapted method to isolate intact and viable single myofibers from post-natal hindlimb muscles (Tibialis Anterior, Extensor Digitorum Longus, Gastrocnemius and Soleus). Following this protocol, we were able to study satellite cells from Lamin &#916;8-11 -/- mice, a severe EDMD murine model, at only 19 days after birth.
We detail the isolation procedure, as well as the culture conditions for obtaining a good amount of myofibers and their associated satellite-cells-derived progeny. When cultured in growth-factors rich medium, satellite cells derived from wild type mice activate, proliferate, and eventually differentiate or undergo self-renewal. In homozygous Lamin &#916;8-11 -/- mutant mice these capabilities are severely impaired.
This technique, if strictly followed, allows to study all processes linked to the myofiber-associated satellite cell even in early post-natal developmental stages and in fragile muscles.

Here, we propose a method to efficiently obtain single muscle fibers at early post-natal developmental stages from homozygous mutant Lamin &#916;8-11 mouse model, a very severe model for Emery-Dreifuss muscular dystrophy (EDMD).

Autosomal dominant Emery-Dreifuss muscular dystrophy (EDMD) is caused by mutations in the LMNA gene, which encodes the A-type nuclear lamins, intermediate ...

Medicine

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

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

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

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

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

Quantifying Tissue-Specific Proteostatic Decline in Caenorhabditis elegans

The ability to maintain proper function and folding of the proteome (protein homeostasis) declines during normal aging, facilitating the onset of a growing number of age-associated diseases. For instance, proteins with polyglutamine expansions are prone to aggregation, as exemplified with the huntingtin protein and concomitant onset of Huntington&#8217;s disease. The age-associated deterioration of the proteome has been widely studied through the use of transgenic Caenorhabditis elegans expressing polyQ repeats fused to a yellow fluorescent protein (YFP). This polyQ::YFP transgenic animal model facilitates the direct quantification of the age-associated decline of the proteome through imaging the progressive formation of fluorescent foci (i.e., protein aggregates) and subsequent onset of locomotion defects that develop as a result of the collapse of the proteome. Further, the expression of the polyQ::YFP transgene can be driven by tissue-specific promoters, allowing the assessment of proteostasis across tissues in the context of an intact multicellular organism. This model is highly amenable to genetic analysis, thus providing an approach to quantify aging that is complementary to lifespan assays. We describe how to accurately measure polyQ::YFP foci formation within either neurons or body wall muscle during aging, and the subsequent onset of behavioral defects. Next, we highlight how these approaches can be adapted for higher throughput, and potential future applications using other emerging strategies for C. elegans genetic analysis.

Quantifying Tissue-Specific Proteostatic Decline in Caenorhabditis elegans

Proteostatic decline is a hallmark of aging, facilitating the onset of neurodegenerative diseases. We outline a protocol to quantifiably measure proteostasis in two different Caenorhabditis elegans tissues through heterologous expression of polyglutamine repeats fused to a fluorescent reporter. This model allows rapid in vivo genetic analysis of proteostasis.

The ability to maintain proper function and folding of the proteome (protein homeostasis) declines during normal aging, facilitating the onset of a ...

Quantification of Insoluble Protein Aggregation in Caenorhabditis elegans during Aging with a Novel Data-Independent Acquisition Workflow

We and others have shown that the aging process results in a proteome-wide accumulation of insoluble proteins. Knocking down genes encoding the insoluble proteins over 40% of the time results in an extension of the lifespan in C. elegans, suggesting that many of these proteins are key determinants of the aging process. Isolation and quantitative identification of these insoluble proteins are crucial to understand key biological processes that occur during aging. Here, we present a modified and improved protocol that details how to extract and isolate the SDS-insoluble proteins (insolublome) from C. elegans more efficiently to streamline mass spectrometric workflows via a novel label-free quantitative proteomics analysis. This improved protocol utilizes a highly efficient sonicator for worm lysis that greatly increases efficiency for protein extraction and allows us to use significantly less starting material (approximately 3,000 worms) than in previous protocols (typically using at least 40,000 worms). Subsequent quantitative proteomic analysis of the insolublome was performed using data-dependent acquisition (DDA) for protein discovery and identification and data-independent acquisition (DIA) for comprehensive and more accurate protein quantification. Bioinformatic analysis of quantified proteins provides potential candidates that can be easily followed up with other molecular methods in C. elegans. With this workflow, we routinely identify more than 1000 proteins and quantify more than 500 proteins. This new protocol enables efficient compound screening with C. elegans. Here, we validated and applied this improved protocol to wild-type C. elegans N2-Bristol strain and confirmed that aged day-10 N2 worms showed greater accumulation of the insolublome than day-2 young worms.

Quantification of Insoluble Protein Aggregation in Caenorhabditis elegans during Aging with a Novel Data-Independent Acquisition Workflow

This novel workflow efficiently extracts and isolates SDS-insoluble proteins (insolublome) from Caenorhabditis elegans with minimal starting material for quantitative differential proteomic analysis. The protocol uses a comprehensive data-independent acquisition mass spectrometry analysis to quantify the insolublome and bioinformatic analysis to gain biological insights into aging mechanisms and pathologies.

We and others have shown that the aging process results in a proteome-wide accumulation of insoluble proteins. Knocking down genes encoding the insoluble ...

Enzymatic Isolation and Characterization of Variable Length Mouse Muscle Fibers

Intact Short, Intermediate, and Long Skeletal Muscle Fibers Obtained by Enzymatic Dissociation of Six Hindlimb Muscles of Mice: Beyond Flexor Digitorum Brevis

Skeletal muscle fibers obtained by enzymatic dissociation of mouse muscles are a useful model for physiological experiments. However, most papers deal with the short fibers of the flexor digitorum brevis (FDB), which restrains the scope of results dealing with fiber types, limits the amount of biological material available, and impedes a clear connection between cellular physiological phenomena and previous biochemical and dynamical knowledge obtained in other muscles.
This paper describes how to obtain intact fibers from six muscles with different fiber type profiles and lengths. Using C57BL/6 adult mice, we show the muscle dissection and fiber isolation protocol and demonstrate the suitability of the fibers for Ca2+ transient studies and their morphometric characterization. The fiber type composition of the muscles is also presented. When dissociated, all muscles rendered intact, living fibers that contract briskly for more than 24 h. FDB gave short (&#60;1 mm), peroneus digiti quarti (PDQA) and peroneus longus (PL) gave intermediate (1-3 mm), while extensor digitorum longus (EDL), extensor hallucis longus (EHL), and soleus muscles released long (3-6 mm) fibers.
When recorded with the fast dye Mag-Fluo-4, Ca2+ transients of PDQA, PL, and EHL fibers showed the fast, narrow kinetics reminiscent of the morphology type II (MT-II), known to correspond to type IIX and IIB fibers. This is consistent with the fact that these muscles have over 90% of type II fibers compared with FDB (~80%) and soleus (~65%). Moving beyond FDB, we demonstrate for the first time the dissociation of several muscles, which render fibers spanning a range of lengths between 1 and 6 mm. These fibers are viable and give fast Ca2+ transients, indicating that the MT-II can be generalized to IIX and IIB fast fibers, regardless of their muscle source. These results increase the availability of models for mature skeletal muscle studies.

Author Spotlight: Isolation of Long Muscle Fibers from Mouse Hindlimb Muscles for Studying Excitation-Contraction Coupling Across Fiber Types

We describe a protocol to obtain enzymatically dissociated fibers of different lengths and types from six muscles of adult mice: three of them already described (flexor digitorum brevis, extensor digitorum longus, soleus) and three of them successfully dissociated for the first time (extensor hallucis longus, peroneus longus, peroneus digiti quarti).

Skeletal muscle fibers obtained by enzymatic dissociation of mouse muscles are a useful model for physiological experiments. However, most papers deal ...

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

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