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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>AKTA prime plus</td>
			<td>Cytiva</td>
			<td></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&#181;m 33.2cm&#178; Nitrocellulose membrane</td>
		</tr>
		<tr>
			<td>DIFP</td>
			<td>Sigma-Aldrich</td>
			<td>55-91-4&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>DIGITAL HOMOGENIZER</td>
			<td></td>
			<td>AS ONE</td>
			<td>HOM</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></td>
			<td>Nacalai Tesque</td>
			<td>17450-61</td>
		</tr>
		<tr>
			<td>HIGH SPEED REFRIGERATIOED MICRO CENTRIFUGE Kitman</td>
			<td></td>
			<td>TOMY</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&#160;</td>
			<td>FUJIFILUM Wako Pure Chemical Corporation</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImmunoStar LD&#160;</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>MgSO4</td>
			<td>Nacalai Tesque</td>
			<td>21003-75</td>
			<td></td>
		</tr>
		<tr>
			<td>Na3VO4</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&#160;</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, EDTA-free)</td>
			<td></td>
			<td>Roche</td>
			<td>5056489001</td>
		</tr>
		<tr>
			<td>Purified tubulin&#160;</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></td>
			<td>LC Laboratories</td>
			<td>P-9600</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>&#946;-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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Biochemistry

1.8K 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

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.

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. ...

Quantitative and Qualitative Method for Sphingomyelin by LC-MS Using Two Stable Isotopically Labeled Sphingomyelin Species

We present a method of analyzing sphingomyelin (SM) qualitatively and quantitatively by liquid chromatography-electrospray Ionization-tandem mass spectrometry (LC-ESI-MS/MS). SM is a common sphingolipid composed of a phosphorylcholine and a ceramide as the hydrophilic and hydrophobic component, respectively. A number of SM species are present in mammalian cells due to a variety in the sphingoid long chain base (LCB) and an N-acyl moiety in the ceramide. In this report, we show a method of estimating the number of carbon and double bonds in a LCB and an N-acyl moiety based on their corresponding product ions in MS/MS/MS (MS3) experiments. In addition, we present a quantitative analysis method for SM using two stable isotopically labeled SM species, which facilitates determining the range used in SM quantitation. The present method will be useful in characterizing a variety of SM species in biological samples and industrial products such as cosmetics.

Here, we present a protocol to quantify and qualify each sphingomyelin species using multiple reaction monitoring and MS/MS/MS mode, respectively.

We present a method of analyzing sphingomyelin (SM) qualitatively and quantitatively by liquid chromatography-electrospray Ionization-tandem mass ...

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

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

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

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

Robust Comparison of Protein Levels Across Tissues and Throughout Development Using Standardized Quantitative Western Blotting

Western blotting is a technique that is commonly used to detect and quantify protein expression. Over the years, this technique has led to many advances in both basic and clinical research. However, as with many similar experimental techniques, the outcome of Western blot analyses is easily influenced by choices made in the design and execution of the experiment. Specific housekeeping proteins have traditionally been used to normalize protein levels for quantification, however, these have a number of limitations and have therefore been increasingly criticized over the past few years. Here, we describe a detailed protocol that we have developed to allow us to undertake complex comparisons of protein expression variation across different tissues, mouse models (including disease models), and developmental timepoints. By using a fluorescent total protein stain and introducing the use of an internal loading standard, it is possible to overcome existing limitations in the number of samples that can be compared within experiments and systematically compare protein levels across a range of experimental conditions. This approach expands the use of traditional western blot techniques, thereby allowing researchers to better explore protein expression across different tissues and samples.

This method describes a robust and reproducible approach for the comparison of protein levels in different tissues and at different developmental timepoints using a standardized quantitative western blotting approach.

Western blotting is a technique that is commonly used to detect and quantify protein expression. Over the years, this technique has led to many advances ...

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

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

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

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

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 ...

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

The Peel-Blot Technique: A Cryo-EM Sample Preparation Method to Separate Single Layers From Multi-Layered or Concentrated Biological Samples

The peel-blot cryo-EM grid preparation technique is a significantly modified back-injection method with the objective of achieving a reduction in layers of multi-layered samples. This removal of layers prior to plunge freezing can aid in reducing sample thickness to a level suitable for cryo-EM data collection, improving sample flatness, and facilitating image processing. The peel-blot technique allows for the separation of multilamellar membranes into single layers, of layered 2D crystals into individual crystals, and of stacked, sheet-like structures of soluble proteins to likewise be separated into single layers. The high sample thickness of these types of samples frequently poses insurmountable problems for cryo-EM data collection and cryo-EM image processing, especially when the microscope stage must be tilted for data collection. Furthermore, grids of high concentrations of any of these samples can be prepared for efficient data collection since sample concentration prior to grid preparation can be increased and the peel-blot technique adjusted to result in a dense distribution of single-layered specimen.

The peel-blot technique is a cryo-EM grid preparation method that allows for the separation of multilayered and concentrated biological samples into single layers to reduce thickness, increase sample concentration, and facilitate image processing.

The peel-blot cryo-EM grid preparation technique is a significantly modified back-injection method with the objective of achieving a reduction in layers ...

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 ...

Stable Isotope Tracing & LC-MS Analysis to Measure DHODH and SDH Activities

Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in mammalian cells. As oxygen (O2) is the most ubiquitous terminal electron acceptor for the mammalian ETC, the O2 consumption rate is frequently used as a proxy for mitochondrial function. However, emerging research demonstrates that this parameter is not always indicative of mitochondrial function, as fumarate can be employed as an alternative electron acceptor to sustain mitochondrial functions in hypoxia. This article compiles a series of protocols that allow researchers to measure mitochondrial function independently of the O2 consumption rate. These assays are particularly useful when studying mitochondrial function in hypoxic environments. Specifically, we describe methods to measure mitochondrial ATP production, de novo pyrimidine biosynthesis, NADH oxidation by complex I, and superoxide production. In combination with classical respirometry experiments, these orthogonal and economical assays will provide researchers with a more comprehensive assessment of mitochondrial function in their system of interest.

Author Spotlight: Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals  

Here, we present a compilation of assays to directly measure mitochondrial function in mammalian cells independently of their ability to consume molecular oxygen.

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in ...