This work was supported in part by JST the establishment of university fellowships toward the creation of science technology innovation (A.HT.; JPMJFS2145), JST SPRING (A.HT.; JPMJSP2129), Grant-in-Aid for JSPS Fellows (A.HT.; 23KJ2078), a Grant-in-Aid for Scientific Research(B) JSPS KAKENHI (22H02946 for TM), a Grant-in-Aid for Scientific Research on Innovative Areas titled &#34;Brain Protein Aging and Dementia Control&#34; from MEXT (TM; 26117004), and by Uehara Research Fellowship from the Uehara Memorial Foundation (TM; 202020027). The authors declare no competing financial interests.

1. MT fractionation method
NOTE: All experiments performed in this study were approved by the Animal Ethics Committee of Doshisha University. C57BL/6J mice of either sex, 3-4 months of age, were used here. In this protocol, dissected tissues, e.g., brain, liver, or thymus, were immediately homogenized in ice-cold microtubule stabilizing buffer (MSB), which contained Taxol (MT stabilizer) at a concentration that prevented not only depolymerization but also repolymerization of MT. The homogenate was separated into three fractions by a two-step ultracentrifugation process (Figure 1). All steps in this protocol were completed without interruption in a cool-temperature environment, and the tissues and fractions were not frozen until they were dissolved in sodium dodecyl sulfate (SDS)-sample buffer.
<ol>
	<li>Preparation of MSB and microtubes
	<ol>
		<li>To prepare MSB, mix the following reagents: 0.1 M 2-(N-morpholino)ethanesulfonic acid (MES), pH 6.8 (neutralized by KOH), 10% glycerol, 0.1 mM DTT, 1 mM MgSO4, 1 mM EGTA, 0.5% Triton X-100, phosphatase inhibitors (1 mM NaF, 1 mM &#946;-glycerophosphate, 1 mM Na3VO4, 0.5 &#181;M okadaic acid), 1x protease inhibitor cocktail, and protease inhibitors (0.1 mM PMSF, 0.1 mM DIFP, 1 &#181;g/mL pepstatin, 1 &#181;g/mL antipain, 10 &#181;g/mL aprotinin, 10 &#181;g/mL leupeptin, 50 &#181;g/mL TLCK) (see Table of Materials) and denote as MSB(-).</li>
		<li>Just before tissue dissection, add 10 &#181;M Taxol and 2 mM GTP (see Table of Materials) to MSB(-). This buffer is denoted as MSB(+). Prepare the buffer on the day of use and keep it on ice.</li>
		<li>Prepare the microtubes for sampling. Empty 2.0 mL microtube for homogenate storage; empty 1.5 mL microtube for supernatant1 (S1) lysate, precipitate2 (P2) sample, and precipitate3 (P3) sample storage; 1.5 mL microtube with 1 mL of ice-cold phosphate-buffered saline (PBS) for dissected tissue storage; 1.5 mL microtube with 200 &#181;L of 2x SDS-sample buffer (0.16 M Tris pH 6.8; 20% glycerol; 2% 2-mercaptoethanol; 4% SDS) for S1 sample and supernatant3 (S3) sample storage; and centrifugation microtube for TLA55 and TLA120.2 centrifuge rotors (see Table of Materials). Label all tubes and place them on ice.</li>
	</ol>
	</li>
	<li>Mouse tissue homogenization
	<ol>
		<li>Prepare a chilled table for tissue dissection. First, fill a box with crushed ice and place two Petri dishes on ice, one with the inner side up and the other with the outer side. Fill ice-cold PBS in one dish for transient wash and storage of dissected tissues. Lay filter paper moistened with PBS on another dish turned over.</li>
		<li>To sacrifice a mouse, perform cervical dislocation under deep anesthesia with a mixed anesthetic of butorphanol, midazolam, and medetomidine. Then, immediately dissect the tissues, e.g., brain, liver, or thymus, and wash them with ice-cold PBS in a Petri dish. 
		NOTE: Any type of soft tissue can be analyzed by this method. However, the size of the tissue is limited by the recommended volume range of the homogenizer used. For example, if a 2 mL volume of homogenizer is used, 50-100 mg of tissue is recommended.</li>
		<li>After weighing the 1.5 mL microtubes filled with PBS for dissected tissue storage, cut out and store tissues inside the microtubes, and reweigh each microtube. Each tissue&#39;s wet weight can be calculated by subtracting the weight of the tube before and after the tissue is added.</li>
		<li>Immediately homogenize the tissue in ice-cold MSB(+) with a chilled homogenizer (see Table of Materials). The volume of MSB(+) was 19 times (&#181;L) the tissue wet weight (mg). Perform homogenization with 20 strokes until the tissue pieces disappear. 
		NOTE: For example, 1,900 &#181;L of MSB(+) is used for 100 mg of tissue. Since the volume of MSB(+) to be added must be adjusted for each wet weight of the tissue pieces analyzed, it is necessary to weigh each tissue piece accurately.</li>
	</ol>
	</li>
	<li>Centrifugation of the mouse tissue homogenates
	<ol>
		<li>Move the whole homogenate to a 2 mL microtube with a Pasteur pipette and centrifuge at 2,400 &#215; g for 3 min at 2 &#176;C to remove the debris via precipitation.</li>
		<li>Transfer the whole supernatant (S1 fraction) to a new 1.5 mL microtube and vortex. Then, aliquot 200 &#181;L of S1 fraction into a centrifugation microtube and centrifuge at 100,000 &#215; g using a TLA-55 rotor for 20 min at 2 &#176;C to obtain the relatively large molecular weight proteins as a precipitate (P2 fraction). 
		NOTE: The volume of the sample subjected to the ultracentrifugation steps affects the radius of centrifugation and the precipitation efficiency of the molecules. Keep the sample volume at 200 &#181;L or less after this step to prevent inaccurate fractionation.</li>
		<li>Further centrifuge all resultant supernatant (S2 fraction) at 500,000 &#215; g using a TLA-120.2 rotor for 60 min at 2 &#176;C to separate the insoluble protein complexes in the precipitate (P3 fraction) from soluble proteins in the supernatant (S3 fraction).</li>
		<li>Add 400 &#181;L of 1x SDS-sample buffer (0.08 M Tris pH 6.8; 10% glycerol; 1% 2-mercaptoethanol; 2% SDS) to the P2 and P3 fraction tubes and briefly sonicated to dissolve the precipitate. Transfer these fraction samples to an empty 1.5 mL microtube.</li>
		<li>Dissolve the total S3 fraction in 200 &#181;L of 2x SDS-sample buffer.</li>
		<li>Mix the remaining S1 fractions with an equal volume of 2x SDS sample buffer for use as a standard curve for Western blotting.</li>
		<li>Boil all these samples at 100 &#176;C for 3 min. After the samples have cooled to room temperature, store the samples at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Quantification of proteins in each fraction
	<ol>
		<li>Quantify proteins in the P2, P3, and S3 fractions by Western blotting. First, use 10% SDS-polyacrylamide gel electrophoresis (SDS-PAGE) to separate proteins from properly diluted the P2, P3, and S3 fractions, and serially diluted S1 sample from any individual as a standard curve. Then, electroblot the samples onto polyvinylidene fluoride membranes (see Table of Materials). 
		&#8203;NOTE: The dilution ratio of each fraction depends on the concentration of an objective protein and the reactivity of the antibody. (e.g., &#945;-tubulin, TUBB3, &#946;-tub, and tyrosinated tubulin in brain tissue: S3 = 1/400, P3 = 1/2,000, P2 = 1/2,000, S1 = 1/50,000, 1/20,000, 1/10,000, 1/5,000, 1/2,000; acetylated tubulin in brain tissue: S3 = 1/200, P3 = 1/400, P2 = 1/8,000, S1 = 1/100,000, 1/40,000, 1/20,000, 1/10,000, 1/4,000; &#945;-tubulin in liver: S3 = 1/20, P3 = 1/100, P2 = 1/20, S1 = 1/50,000, 1/20,000, 1/10,000, 1/5,000, 1/2,000; &#945;-tubulin in thymus: S3 = 1/100, P3 = 1/400, P2 = 1/20, S1 = 1/50,000, 1/20,000, 1/10,000, 1/5,000, 1/2,000 dilution of tissue concentration).</li>
		<li>Block the membrane with 5% skimmed milk in Tris-buffered saline (50 mM Tris-HCl pH 7.6; 152 mM NaCl)&#160;with 0.1% Tween 20 (TBS-T) for over 30 min.</li>
		<li>Immerse the membrane in TBS-T containing primary antibody (see Table of Materials) for over 2 h. After that, wash the membrane with TBS-T for 3 min (3 times).</li>
		<li>Label primary antibody using HRP-conjugated secondary antibodies (see Table of Materials) in TBS-T for over 1 h. After that, wash the membrane with TBS-T for 3 min (3 times).</li>
		<li>Develop the membranes with Enhanced Chemiluminescence reagent. Then, analyze the bands of interest with a luminescent image analyzer (see Table of Materials).</li>
		<li>Quantify the protein band intensities using image analysis software (see Table of Materials) and create a standard curve by plotting the dilution units of the diluted S1 samples used for the standard curve along the X-axis and the band intensities along the Y-axis.</li>
		<li>Read the protein concentration (unit) corresponding to the diluted fraction samples. Multiply the concentration read with the sample dilution factor to obtain a protein unit in each fraction. Divide the measured unit of each fraction by the total protein unit (P2 + P3 + S3) to obtain a percentage.</li>
	</ol>
	</li>
</ol>
2. Evaluation of the properties of tubulin in each fraction
NOTE: This biochemical method provides three groups of tubulin complexes defined by sedimentation properties. Here, the status of tubulin complexes obtained in these fractions was identified based on the size of the complex and tubulin-PTMs. Complete all steps in this protocol without interruption in a cool-temperature environment but do not freeze the fraction samples until they are dissolved in the SDS-sample buffer.
<ol>
	<li>Filter trap assay
	<ol>
		<li>Filter the S2 and S3 fractions (500 &#181;L each) using a 300 kDa ultrafiltration spin column (see Table of Materials). Perform 14,000 &#215; g centrifugation at 2 &#176;C until the whole supernatant is filtered, eluted proteins are collected into receiver tubes, and trapped proteins remain on the filter of reservoir tubes.</li>
		<li>Translate the whole filtrates (approximately 500 &#181;L) in receiver tubes into new 1.5 mL microtubes and dissolve them in 500 &#181;L of 2x SDS-sample buffer.</li>
		<li>Solubilize the residues on the filter of reservoir tubes in 1,000 &#181;L of 1x SDS-sample buffer by pipetting and transferring the samples to a new 1.5 mL microtube.</li>
		<li>Boil the samples at 100 &#176;C for 3 min. After the samples have cooled to room temperature, store the samples at -20 &#176;C.</li>
		<li>Analyze the amounts of tubulin by Western blotting with DM1A (anti-&#945;-tubulin antibody, see Table of Materials).</li>
	</ol>
	</li>
	<li>Size-exclusion chromatography
	<ol>
		<li>Prepare the carrier buffer (0.1 M MES, pH 6.8; 10% glycerol; 1 mM MgSO4; 1 mM EGTA; 0.1 mM DTT) supplemented with 1/10 concentration of protease and phosphatase inhibitors as described in step 1.1.1. Then, filter the solution and store it in a cold environment.</li>
		<li>Prepare a gel filtration chromatography column equipped with a preparative liquid chromatography system in a chromatography chamber&#160;(see Table of Materials) at 4 &#176;C.</li>
		<li>Before injecting samples onto the column, flow 180 mL of the carrier buffer to wash the column. The flow rate is 1 mL/min for 3 h.</li>
		<li>Inject commercially available purified porcine tubulin (see Table of Materials) as a control or the S3 fraction from mouse brains (500 &#181;L each) onto the column.</li>
		<li>Elute at a flow rate of 1.0 mL/min with carrier buffer. Collect the 1.5 mL fractions for 120 min. Monitor eluted proteins by absorbance at 280 nm. Keep the maximum pressure under 0.3 MPa.</li>
		<li>After vortexing the collected fractions, mix 50 &#181;L of them with 50 &#181;L of 2x SDS-sample buffer in a 1.5 mL microtube. Boil all samples at 100 &#176;C for 3 min. After the samples have cooled to room temperature, store the samples at -20 &#176;C.</li>
		<li>Analyze the amounts of tubulin by Western blotting&#160;with DM1A, anti-&#945;-tubulin antibody and KMX-1, anti-&#946;-tubulin antibody (see Table of Materials).</li>
	</ol>
	</li>
</ol>

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reduction+of+acetylated+alpha-tubulin+immunoreactivity+in+neurofibrillary+tangle-bearing+neurons+in+Alzheimer's+disease.">Reduction of acetylated alpha-tubulin immunoreactivity in neurofibrillary tangle-bearing neurons in Alzheimer's disease.</a> Journal of Neuropathology and Experimental Neurology. 55 (9), 964-972 (1996).</li><li>Miyasaka, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imbalanced+expression+of+tau+and+tubulin+induces+neuronal+dysfunction+in+C.+elegans+models+of+tauopathy.">Imbalanced expression of tau and tubulin induces neuronal dysfunction in C. elegans models of tauopathy.</a> Frontiers in Neuroscience. 12, 415 (2018).</li><li>Boiarska, Z., Passarella, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubule-targeting+agents+and+neurodegeneration.">Microtubule-targeting agents and neurodegeneration.</a> Drug Discovery Today. 26 (2), 604-615 (2021).</li></ol>

The authors have no conflicts of interest to report.

The most significant task when investigating the status of tubulin in tissue from living organisms is preventing accidental MT polymerization or depolymerization during preparation. The stability of MTs in samples is affected by factors such as the concentration of Taxol in MSB, the proportion of tissue amount to buffer, and temperature during the process from tissue removal to homogenization and centrifugation. Therefore, the conditions were optimized in each step of the protocol for analyzing mouse tissue with a 20-fol...

Microtubules (MTs) are elongated tubular structures comprising protofilaments consisting of &#945;/&#946;-tubulin heterodimer subunits. They play essential roles in various cellular processes such as cell division, motility, shape maintenance, and intracellular transport, making them integral components of the eukaryotic cytoskeleton1. The minus-end of MTs, where the &#945;-tubulin subunit is exposed, is relatively stable, whereas the plus-end, where the &#946;-tubulin subunit is exposed, undergoes dynamic depolymerization and polymerization2. This continuous cycle of tubulin dimer addition and dissociation at the plus-end, referred to as dynamic instability, results in a repetitive process of rescue and catastrophe3. MTs exhibit focal domains with localized variations in dynamic instability, including stable and labile domains4.
Precise control of the dynamic instability of MTs&#160;is crucial for numerous cellular functions, particularly in neurons characterized by intricate morphologies. The adaptability and durability of MTs play a vital role in the development and proper functioning of nerve cells5,6,7. The dynamic instability of MTs has been found to be associated with various post-translational modifications (PTMs) of tubulin, such as acetylation, phosphorylation, palmitoylation, detyrosination, delta 2, polyglutamine oxidation, and polyglycylation. Additionally, the binding of microtubule-associated proteins (MAPs) serves as a regulatory mechanism8. PTMs, excluding acetylation, predominantly occur in the tubulin carboxy-terminal region situated on the external surface of MTs. These modifications create diverse surface conditions on MTs, influencing their interaction with MAPs and ultimately governing MT stability9. The presence of a carboxy-terminal tyrosine residue in &#945;-tubulin is indicative of dynamic MTs, which are rapidly replaced by the free tubulin pool. Conversely, detyrosination of the carboxy terminus and acetylation of Lys40 signify stable MTs with reduced dynamic instability9,10.
The PTMs of tubulin have been extensively employed in experiments to assess the dynamics and stability of MTs5,7,11,12,13,14,15. For instance, in cell culture studies, tubulins can be segregated into two pools: the free tubulin pool and the MT pool. This is achieved by releasing free tubulin through cell permeabilization before fixing the remaining MTs15,16,17,18,19. Biochemical methods involve the use of chemical MT stabilizers that safeguard MTs from catastrophe, enabling the separation of MTs and free tubulin through centrifugation20,21,22. However, these procedures do not differentiate between stable and less stable (labile) MTs, thereby rendering it impossible to quantify MTs or soluble tubulin in tissues like the brain. Consequently, evaluating MT stability in organisms under physiological and pathological conditions has proven to be challenging. To address this experimental limitation, we have developed a novel technique for precisely separating MTs and free tubulin in mouse tissue23.
This unique MT fractionation method involves tissue homogenization under conditions that maintain tubulin status in tissues and two-step centrifugation to separate stable MTs, labile MTs, and free tubulin. This simple procedure can be applied to broad studies, including basic research on MTs and MAPs in living organisms, physiological and pathological analyses of health and diseases associated with MT stability, and developing drugs and other therapeutics that target MTs.

<table><tbody><tr><td>1.5 ML TUBE CASE OF 500</td><td>Beckman Coulter</td><td>357448</td><td></td></tr><tr><td>1A2</td><td>Sigma-Aldrich</td><td>T9028</td><td>1:5,000 dilution</td></tr><tr><td>2-(N-morpholino)ethanesulfonic acid (MES)</td><td>Nacalai Tesque</td><td>02442-44</td><td></td></tr><tr><td>300 kDa ultrafiltration spin column</td><td>Aproscience</td><td>PT-1013</td><td></td></tr><tr><td>6-11B1</td><td>Sigma-Aldrich</td><td>T7451</td><td>1:5,000 dilution</td></tr><tr><td>&amp;#196;KTAprime plus</td><td>Cytiva</td><td>11001313</td><td></td></tr><tr><td>anti-mouse IgG</td><td>Jackson ImmunoResearch</td><td>115-035-146</td><td>1:5,000 dilution</td></tr><tr><td>antipain</td><td>Peptide Institute Inc.</td><td>4062</td><td></td></tr><tr><td>aprotinin</td><td>Nacalai Tesque</td><td>03346-84</td><td></td></tr><tr><td>Chemi-Lumi One L</td><td>Nacalai Tesque</td><td>07880-54</td><td></td></tr><tr><td>Corning bottle-top vacuum filter system</td><td>Corning</td><td>430758</td><td>0.22&amp;micro;m 33.2cm&amp;sup2; Nitrocellulose membrane</td></tr><tr><td>DIFP</td><td>Sigma-Aldrich</td><td>55-91-4&amp;nbsp;</td><td></td></tr><tr><td>DIGITAL HOMOGENIZER HK-1</td><td>AS ONE</td><td>1-2050-11</td><td></td></tr><tr><td>DM1A</td><td>Sigma-Aldrich</td><td>T9026</td><td>1:5,000 dilution</td></tr><tr><td>DTT</td><td>Nacalai Tesque</td><td>14128-46</td><td></td></tr><tr><td>EGTA</td><td>Nacalai Tesque</td><td>37346-05</td><td></td></tr><tr><td>FluoroTrans W 3.3 Meter Roll</td><td>Pall Corporation</td><td>BSP0161</td><td></td></tr><tr><td>glycerol</td><td>Nacalai Tesque</td><td>17018-25</td><td></td></tr><tr><td>GTP</td><td>Nacalai Tesque</td><td>17450-61</td><td></td></tr><tr><td>HIGH SPEED REFRIGERATIOED MICRO CENTRIFUGE Kitman</td><td>TOMY</td><td>KITMAN-24</td><td></td></tr><tr><td>HiLoad 16/600 Superdex 200 pg column</td><td>Cytiva</td><td>28-9893-35</td><td></td></tr><tr><td>Image Gauge Software&amp;nbsp;</td><td>FUJIFILUM Wako Pure Chemical Corporation</td><td></td><td></td></tr><tr><td>ImmunoStar LD&amp;nbsp;</td><td>FUJIFILUM Wako Pure Chemical Corporation</td><td>292-69903</td><td></td></tr><tr><td>KMX-1</td><td>Millipore</td><td>MAB3408</td><td>1:5,000 dilution</td></tr><tr><td>LAS-4000 luminescent image analyzer</td><td>FUJIFILUM Wako Pure Chemical Corporation</td><td></td><td></td></tr><tr><td>leupeptin</td><td>Peptide Institute Inc.</td><td>43449-62</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Nacalai Tesque</td><td>21003-75</td><td></td></tr><tr><td>Na&lt;sub&gt;3&lt;/sub&gt;VO&lt;sub&gt;4&lt;/sub&gt;</td><td>Nacalai Tesque</td><td>32013-92</td><td></td></tr><tr><td>NaF</td><td>Nacalai Tesque</td><td>31420-82</td><td></td></tr><tr><td>okadaic acid</td><td>LC Laboratories</td><td>O-2220&amp;nbsp;</td><td></td></tr><tr><td>OPTIMA MAX-XP</td><td>Beckman Coulter</td><td>393315</td><td></td></tr><tr><td>pepstatin</td><td>Nacalai Tesque</td><td>26436-52</td><td></td></tr><tr><td>PMSF</td><td>Nacalai Tesque</td><td>27327-81</td><td></td></tr><tr><td>Polycarbonate Centrifuge Tubes for TLA120.2</td><td>Beckman Coulter</td><td>343778</td><td></td></tr><tr><td>Protease inhibitor cocktail (cOmplete&amp;#8482;, EDTA-free)</td><td>Roche</td><td>11873580001</td><td></td></tr><tr><td>Purified tubulin&amp;nbsp;</td><td>Cytoskeleton</td><td>T240</td><td></td></tr><tr><td>QSONICA Q55</td><td>QSonica</td><td>Q55</td><td></td></tr><tr><td>Taxol</td><td>LC Laboratories</td><td>P-9600</td><td></td></tr><tr><td>TLA-120.2 rotor</td><td>Beckman Coulter</td><td>357656</td><td></td></tr><tr><td>TLA-55 rotor</td><td>Beckman Coulter</td><td>366725</td><td></td></tr><tr><td>TLCK</td><td>Nacalai Tesque</td><td>34219-94</td><td></td></tr><tr><td>Triton X-100</td><td>Nacalai Tesque</td><td>12967-45</td><td></td></tr><tr><td>&amp;beta;-glycerophosphate</td><td>Sigma-Aldrich</td><td>G9422</td><td></td></tr></tbody></table>

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.

Quantification of tubulin in the P2, P3, and S3 fractions from mouse brain by the MT fractionation method 
Tubulin in mouse tissue was separated into the P2, P3, and S3 fractions by the MT fractionation method and quantified by Western blotting (Figure 1A). The precipitate of MTs that remained in the P2 fraction by ultracentrifugation at 100,000 &#215; g for 20 min accounted for 34.86% &#177; 1.68% of total tubulin in a mouse brain. The supernatant (S2) was fur...

Watch this Scientific Journal Video about Quantitative Microtubule Fractionation Technique to Separate Stable Microtubules, Labile Microtubules, and Free Tubulin in Mouse Tissues at JoVE.com

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

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 ...

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Research

JoVE Journal

Biochemistry

1.9K Views.  Doshisha University. 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.

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

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

Developmental Biology

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 ...

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and networks often contain subsets of microtubule arrays that differ in their dynamic properties. For example, in dividing cells, stable bundles of crosslinked microtubules coexist in close proximity to dynamic non-crosslinked microtubules. TIRF-microscopy-based in vitro reconstitution studies enable the simultaneous visualization of the dynamics of these different microtubule arrays. In this assay, an imaging chamber is assembled with surface-immobilized microtubules, which are either present as single filaments or organized into crosslinked&#160;bundles. Introduction of tubulin, nucleotides, and protein regulators allows direct visualization of associated proteins and of dynamic properties of single and crosslinked microtubules. Furthermore, changes that occur as dynamic single microtubules organize into bundles can be monitored in real-time. The method described here allows for a systematic evaluation of the activity and localization of individual proteins, as well as synergistic effects of protein regulators on two different microtubule subsets under identical experimental conditions, thereby providing mechanistic insights that are inaccessible by other methods.

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Here, a TIRF microscopy-based in vitro reconstitution assay is presented to simultaneously quantify and compare the dynamics of two microtubule populations. A method is described to simultaneously view the collective activity of multiple microtubule-associated proteins on crosslinked microtubule bundles and single microtubules.

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and ...

Extracting Modified Microtubules from Mammalian Cells to Study Microtubule-Protein Complexes by Cryo-Electron Microscopy

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the posttranslational modifications, microtubules can form complexes with various interacting proteins. These microtubule-protein complexes are often implicated in human diseases. Understanding the structure of such complexes is useful for elucidating their mechanisms of action and can be studied by cryo-electron microscopy (cryo-EM). To obtain such complexes for structural studies, it is important to extract microtubules containing or lacking specific posttranslational modifications. Here, we describe a simplified protocol to extract endogenous tubulin from genetically modified mammalian cells, involving microtubule polymerization, followed by sedimentation using ultracentrifugation. The extracted tubulin can then be used to prepare cryo-electron microscope grids with microtubules that are bound to a purified microtubule-binding protein of interest. As an example, we demonstrate the extraction of fully tyrosinated microtubules from cell lines engineered to lack the three known tubulin-detyrosinating enzymes. These microtubules are then used to make a protein complex with enzymatically inactive microtubule-associated tubulin detyrosinase on cryo-EM grids.

Here, we describe a protocol to extract endogenous tubulin from mammalian cells, which can lack or contain specific microtubule-modifying enzymes, to obtain microtubules enriched for a specific modification. We then describe how the extracted microtubules can be decorated with purified microtubule-binding proteins to prepare grids for cryo-electron microscopy.

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the ...

Efficient Tubulin Isolation from Neural Tissue

Optimizing Tubulin Yield from Porcine Brain Tissue

Neural tissue from any source is an excellent material for tubulin isolation, as the dendrites and axons of neurons are rich in microtubules. Here, we present a procedure for extracting tubulin that can be employed, with minor modifications, for neural tissue from multiple sources. In the presented protocol, a new clarification step of the crude lysate has been introduced, which led to a significant reduction in the initial amount of insoluble debris before the first polymerization step occurred. This additional step allowed the processing of additional tissue while using the same instrumentation and, thus, increasing the relative volume of the processed homogenate. The newly introduced step has no significant effect on the quality of purified tubulin, as confirmed by in vitro activity assays and transmission electron microscopy. The described procedure contains all crucial steps, including tissue collection, transport, tissue homogenization, tubulin isolation cycles, and final polishing by ion exchange chromatography using FPLC and subsequent activity measurement assays. The homogeneity of purified tubulin was more than 97%, as confirmed by MS/MS analysis utilizing electrospray ionization and MALDI-TOF.

Author Spotlight: Purifying High-Quality Tubulin to Study Protein Dynamics and Therapeutic Applications

This protocol describes a technique for the high-yield isolation of tubulin from the porcine brain optimized for small-scale instrumentation. The isolation procedures are complemented by procedures for determining tubulin polymerization activity in vitro using co-sedimentation assays and transmission electron microscopy.

Neural tissue from any source is an excellent material for tubulin isolation, as the dendrites and axons of neurons are rich in microtubules. Here, we ...

Preparation of Segmented Microtubules to Study Motions Driven by the Disassembling Microtubule Ends

Microtubule depolymerization can provide force to transport different protein complexes and protein-coated beads in vitro. The underlying mechanisms are thought to play a vital role in the microtubule-dependent chromosome motions during cell division, but the relevant proteins and their exact roles are ill-defined. Thus, there is a growing need to develop assays with which to study such motility in vitro using purified components and defined biochemical milieu. Microtubules, however, are inherently unstable polymers; their switching between growth and shortening is stochastic and difficult to control. The protocols we describe here take advantage of the segmented microtubules that are made with the photoablatable stabilizing caps. Depolymerization of such segmented microtubules can be triggered with high temporal and spatial resolution, thereby assisting studies of motility at the disassembling microtubule ends. This technique can be used to carry out a quantitative analysis of the number of molecules in the fluorescently-labeled protein complexes, which move processively with dynamic microtubule ends. To optimize a signal-to-noise ratio in this and other quantitative fluorescent assays, coverslips should be treated to reduce nonspecific absorption of soluble fluorescently-labeled proteins. Detailed protocols are provided to take into account the unevenness of fluorescent illumination, and determine the intensity of a single fluorophore using equidistant Gaussian fit. Finally, we describe the use of segmented microtubules to study microtubule-dependent motions of the protein-coated microbeads, providing insights into the ability of different motor and nonmotor proteins to couple microtubule depolymerization to processive cargo motion.

Microtubules are inherently unstable polymers, and their switching between growth and shortening is stochastic and difficult to control. Here we describe protocols using segmented microtubules with photoablatable stabilizing caps. Depolymerization of segmented microtubules can be triggered with high temporal and spatial resolution, thereby assisting analysis of motions with the disassembling microtubule ends.

Microtubule depolymerization can provide force to transport different protein complexes and protein-coated beads in vitro. The underlying mechanisms are ...

Biology

Subcellular Fractionation for the Isolation of Synaptic Components from the Murine Brain

Synaptic terminals are the primary sites of neuronal communication. Synaptic dysfunction is a hallmark of many neuropsychiatric and neurological disorders. The characterization of synaptic sub-compartments by biochemical isolation is, therefore, a powerful method to elucidate the molecular bases of synaptic processes, both in health and disease. This protocol describes the isolation of synaptic terminals and synaptic sub-compartments from mouse brains by subcellular fractionation. First, sealed synaptic terminal structures, known as synaptosomes, are isolated following brain tissue homogenization. Synaptosomes are neuronal pre- and post-synaptic compartments with pinched-off and sealed membranes. These structures retain a metabolically active state and are valuable for studying synaptic structure and function. The synaptosomes are then subjected to hypotonic lysis and ultracentrifugation to obtain synaptic sub-compartments enriched for synaptic vesicles, synaptic cytosol, and synaptic plasma membrane. Fraction purity is confirmed by electron microscopy and biochemical enrichment analysis for proteins specific to sub-synaptic compartments. The presented method is a straightforward and valuable tool for studying the structural and functional characteristics of the synapse and the molecular etiology of various brain disorders.

This protocol presents a robust, detailed method to obtain highly pure synaptosomes, synaptic vesicles, and other synaptic fractions from the mouse brain. This method enables the evaluation of synaptic processes, including the biochemical analysis of protein localization and function with compartmental resolution.

Synaptic terminals are the primary sites of neuronal communication. Synaptic dysfunction is a hallmark of many neuropsychiatric and neurological ...

Neuroscience

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a genetically encoded marker for the positive ends of microtubules (EB3 labelled with tdTomato fluorescent protein) and high-speed, high-resolution, live-cell imaging using spinning disk confocal microscopy. Cell cycle synchronization and increased density of microtubules are achieved by inhibiting centrosomal separation in mitotic cells, and analysis of growth is performed using open-source U-Track software. The use of a bright and red-shifted fluorescent protein, in combination with the lower laser power and reduced exposure time required for spinning disk microscopy reduce phototoxicity and the probability of light-induced artifacts. This allows for imaging a larger number of cells in the same preparation while maintaining the cells in a growth medium under standard culture conditions. Because the analysis is performed in a supervised automatic fashion, the results are statistically robust and reproducible.

Here we present a robust and detailed method of microtubule dynamics analysis in cells synchronized in prometaphase using live-cell spinning disk confocal microscopy and MATLAB-based image processing.

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a ...

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as specialized arrays during mitosis to regulate chromosome segregation. Proteins that interact with microtubules include motors such as kinesins and dynein, which can generate active forces and directional motion, as well as non-motor proteins that crosslink filaments into higher-order networks or regulate filament dynamics. To date, biophysical studies of microtubule-associated proteins have overwhelmingly focused on the role of single motor proteins needed for vesicle transport, and significant progress has been made in elucidating the force-generating properties and mechanochemical regulation of kinesins and dyneins. However, for processes in which microtubules act both as cargo and track, such as during filament sliding within the mitotic spindle, much less is understood about the biophysical regulation of ensembles of the crosslinking proteins involved. Here, we detail our methodology for directly probing force generation and response within crosslinked microtubule minimal networks reconstituted from purified microtubules and mitotic proteins. Microtubule pairs are crosslinked by proteins of interest, one microtubule is immobilized to a microscope coverslip, and the second microtubule is manipulated by an optical trap. Simultaneous total internal reflection fluorescence microscopy allows for multichannel visualization of all the components of this microtubule network as the filaments slide apart to generate force. We also demonstrate how these techniques can be used to probe pushing forces exerted by kinesin-5 ensembles and how viscous braking forces arise between sliding microtubule pairs crosslinked by the mitotic MAP PRC1. These assays provide insights into the mechanisms of spindle assembly and function and can be more broadly adapted to study dense microtubule network mechanics in diverse contexts, such as the axon and dendrites of neurons and polar epithelial cells.

Here, we present a protocol for reconstituting microtubule bundles in vitro and directly quantifying the forces exerted within them using simultaneous optical trapping and total internal reflection fluorescence microscopy. This assay allows for nanoscale-level measurement of the forces and displacements generated by protein ensembles within active microtubule networks.

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as ...

Forming, Confining, and Observing Microtubule-Based Active Nematics

The formation of biopolymer-based active phases has become an important technique for researchers interested in exploring the emerging field of active liquid crystals and their possible roles in cell biology. These novel systems consist of self-driven sub-units that consume energy locally, producing an out-of-equilibrium dynamic fluid. To form the active liquid crystal phase described in this report, purified protein components including biopolymers and molecular motors are combined, and the active nematic phase spontaneously forms in the presence of adenosine triphosphate (ATP). To observe the nematic state, the material must be confined in a suitable geometry for microscopy at a high enough density. This article describes two different methods for the formation of an active nematic phase using microtubules and kinesin motors: assembly of a two-dimensional active layer at an oil and water interface and assembly under an oil layer using an elastomeric well. Techniques to insert the active material into small wells of different shapes are also described.

Presented here are methods for preparing active nematics from microtubules and kinesin motors, including protein preparation and construction and the use of wells for active nematic confinement.

The formation of biopolymer-based active phases has become an important technique for researchers interested in exploring the emerging field of active ...

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

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Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

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

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Extracting Modified Microtubules from Mammalian Cells to Study Microtubule-Protein Complexes by Cryo-Electron Microscopy

Optimizing Tubulin Yield from Porcine Brain Tissue

Preparation of Segmented Microtubules to Study Motions Driven by the Disassembling Microtubule Ends

Subcellular Fractionation for the Isolation of Synaptic Components from the Murine Brain

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Forming, Confining, and Observing Microtubule-Based Active Nematics