Name: optimizing tubulin yield from porcine brain tissue
Uploaded: 2024-10-11T12:30:00.000+00:00
Duration: 6 min 30 s
Description: 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 ...

The study was supported by Technology Agency of the Czech Republic (project nr. TN02000017 - National Centre for Biotechnology in Veterinary Medicine - NaCeBiVet).

The composition of all solutions and instrumentation is described in the Table of Materials. All solutions were prepared using FPLC-grade chemicals and filtered through a 0.22 &#181;m filter prior to use. Personal protective equipment, e.g., laboratory coat, gloves, and safety glasses, were used during all operations. All instruments were clean and free of traces of detergents. During the procedures, appropriate animal care guidelines (as approved by the institution) were followed. All biological material, including brain tissue, was bought from a slaughterhouse as raw material. No live animals were used at any step of the protocol.
1. Activation of a phospho-cellulose column for fast protein liquid chromatography
<ol>
	<li>Add 700 mL of 50% ethanol to 15 g of dry phospho-cellulose resin and mix it gently in the beaker with a spoon.</li>
	<li>Pour the suspension into a suitable chromatographic column of appropriate volume (700 mL). Close the column with pistons equipped with frits. Leave at least 10% of empty head space. 
	NOTE: Other vessels can be used, but the FPLC column, in combination with a peristaltic pump, significantly reduces the time necessary for resin activation and resin losses.</li>
	<li>Incubate on a rocker-shaker at 60 rpm for 30 min.</li>
	<li>Let the resin settle and remove the excess of 50% ethanol using the peristaltic pump with the flow set to 3 mL/min. Do not let the resin dry.</li>
	<li>Remove the top piston and add 300 mL of 50% ethanol. Close the column, briefly shake it, and remove the excess ethanol via a peristaltic pump.</li>
	<li>Remove the remaining ethanol. Add 300 mL of ultra-pure water into the column, close the column with the piston, and tilt until the resin is resuspended. Remove excess liquid using a peristaltic pump. Repeat this step three times.</li>
	<li>Open the column at the top, add 500 mL of 0.5 M HCl, close the column, and resuspend the resin by gentle tilting. Incubate on a rocker-shaker at 60 rpm for 30 min.
	<ol>
		<li>Remove excess liquid using a peristaltic pump. Add 300 mL of 0.5 M HCl, resuspend the resin, and remove the excess liquid.</li>
	</ol>
	</li>
	<li>Remove the remaining HCl. Add 300 mL of ultra-pure water into the column, close the column with the piston, and tilt until the resin resuspends. Remove excess liquid using a peristaltic pump. Repeat this step three times.</li>
	<li>Pour the 700 mL of MES buffer (25 mM MES, 0.1 mM EGTA, 0.5 mM MgCl2) into the column and resuspend the resin by tilting. Pour the suspension into the beaker, adjust pH to 6.1 with 1 M NaOH, and let incubate for 2 h under gentle agitation.</li>
	<li>Let the resin settle, decant the liquid, add 300 mL of MES buffer, pH 6.4, and incubate overnight at 4 &#176;C.</li>
	<li>Decant the excess liquid, add 96% ethanol to 20% final concentration, and store at 4 &#176;C until use.</li>
</ol>
2. Phospho-cellulose column packing
<ol>
	<li>Pour 12 mL of activated resin into the clean chromatographic column with a closed bottom plug. Let the resin settle. The volume of settled resin is about 5 mL. Remove the resin upper layer (approximately 2-3 mm) where lighter broken matrix fragments might deposit.</li>
	<li>Remove the bottom plug and let the storage solution drain freely, drop by drop. Do not let the resin dry.</li>
	<li>Close the column and connect it to the FPLC system. Wash the column with PEM buffer (100 mM PIPES pH 6.9, 1 mM EGTA, 1 mM MgSO4) at 1 mL/min (max. pressure 1 bar) for 30 min, adjust the piston height, turn the column upside down and wash the column under the same conditions for the next 30 min in the reverse flow.</li>
	<li>Store the column filled with activated phospho-cellulose at 4 &#176;C for several days.</li>
</ol>
3. Tubulin separation and purification
NOTE: Tubulin is highly prone to degradation, and it is crucial to proceed swiftly. All solutions, instruments, and equipment should be prepared, chilled, or heated up, if necessary, in advance. The procedures are sensitive to shifts in recommended temperatures. The brain tissue should be processed as soon after dissection as possible, considering the amount of biological waste produced during purification. The procedure was optimized for the rotor with six 75 mL ultracentrifugation cuvettes. The amount of processed tissue can be increased or decreased according to the available ultracentrifuge.
<ol>
	<li>Tissue transport
	<ol>
		<li>Put 500 g of freshly dissected porcine brains (6-8 pieces) into a 3 L transport vessel and pour the ice-cold Transport buffer (4.1 mM MES pH 7, 320 mM Saccharose, 1 mM EGTA) until fully immersed. Let stand on ice for 5 min and change the Transport buffer for a fresh one to facilitate heat dissipation. Transport swiftly on ice for further processing.</li>
	</ol>
	</li>
	<li>Tissue homogenization 
	NOTE: Tissue homogenization is carried out on ice in a cold room. All instruments should be prechilled to prevent spontaneous polymerization. The primary source of tubulin is grey matter, and the rest is removed in the following steps.
	<ol>
		<li>Pour 100 mL of Extraction buffer (4.1 mM MES pH 7, 520 mM Saccharose, 1 mM EGTA, 1 mM ATP, and 0.1 mM GTP) into a 1 L plastic beaker and determine the weight W1 (g) including the vessel. ATP and GTP are added just prior to the use.</li>
		<li>Take the porcine brain out of the transport vessel and remove the whole cerebellum, fat, big chunks of white matter, meninges, and blood vessels with fingers by applying modest force. Scissors or tweezers can be used as well. Place porcine brains striped of unwanted tissue into the beaker with 100 mL of cold Extraction buffer. Proceed until all tissue is processed.</li>
		<li>Weigh the beaker with processed brain tissue and determine W2 (g) weight.</li>
		<li>Add Extraction buffer according to the formula, Extraction buffer weight = W2 - W1 (e.g., for 400 g of tissue, 300 g of additional Extraction buffer is added).</li>
		<li>Transfer the brain tissue with Extraction buffer into the prechilled kitchen blender and process in 4-8 short, three-second pulses.</li>
		<li>Process the partially homogenized tissue with a high-speed dispersion homogenizer (18 000 rpm) for 1 min. Leave the suspension on ice for 3-5 min, occasionally mixing with a spoon. Repeat five times or until complete homogenization.</li>
		<li>Pour into prechilled ultracentrifugation tubes and spin for 10 min, at 4 &#176;C and 50,000 x g. Keep the supernatant and discard the pellet. Repeat until 430 mL of clarified extract is collected.</li>
		<li>Pour the clarified extract into prechilled ultracentrifugation tubes (70 mL each) and spin at 4 &#176;C and 75,000 x g for 60 min. Collect the supernatant and determine the volume S1. 
		NOTE: If only one rotor exists, heat it in a warm water bath for the next centrifugation step (37 &#176;C).</li>
	</ol>
	</li>
	<li>First polymerization
	<ol>
		<li>Add 10x MEM buffer (1 M MES pH 6.8, 10 mM EGTA, 10 mM MgCl2) according to formula V = S1/9 mL (for 585 mL of supernatant, add 65 mL of 10x MEM buffer). Mix well and add glycerol to 3.5 M and GTP to 0.1 mM final concentration. Pour the suspension into ultracentrifugation tubes (the volume of supernatant exceeds the volume of tubes; the rest is discarded).</li>
		<li>Incubate the tubes with the supernatant in a water bath for 45 min at 37 &#176;C.</li>
		<li>Spin the tubes for 90 min at 37 &#176;C and 75,000 x g. Measure the volume of the supernatant (S2) to determine the volume of MEM buffer, add it to the pellets in step 3.3.4, and then discard the supernatant. Keep the pellet where the polymerized tubulin is contained.</li>
		<li>Prepare 0.5 x S2 volume of ice-cold 1x MEM buffer. Add an equal amount of 1x MEM buffer into each ultracentrifuge tube with the pellet. Resuspend pellets in the added 1x MEM buffer, transfer suspensions into the graduated cylinder, and determine the total volume.
		<ol>
			<li>Add GTP to 1 mM final concentration. Transfer the suspension into a Dounce glass homogenizer and homogenize the solution every 10 min or so for 45 min on ice. The piston needs to be carefully operated as it can break easily. 
			NOTE: In the meantime, chill the rotor in water with ice.</li>
		</ol>
		</li>
		<li>Pour the homogenized solution into prechilled ultracentrifugation tubes and spin at 75,000 x g and 4 &#176;C for 60 min. After centrifugation, determine the supernatant volume (S3).</li>
	</ol>
	</li>
	<li>Second polymerization
	<ol>
		<li>Mix the supernatant with the 0.35 x S3 mL of glycerol and add GTP to 1 mM final concentration. Pour the supernatant into ultracentrifugation tubes and incubate for 45 min at 37 &#176;C. 
		NOTE: In the meantime, warm up the rotor to 37 &#176;C in the water bath.</li>
		<li>Centrifuge the polymerized supernatant from step 3.4.1 for 60 min at 75,000 x g and 37 &#176;C. Determine the volume of the supernatant (S4). 
		NOTE: This is an optional stopping point. The pellet with tubulin can be shock-frozen in liquid nitrogen and stored at -80 &#176;C.</li>
		<li>Let the frozen pellets slowly thaw on ice for 30 min. Prepare 0.25 x S4 mL of PIPES buffer (500 mM PIPES pH 6.9, 1 mM EGTA, 1 mM MgSO4, 1 mM DTT, 0.1 mM ATP), divide evenly into tubes with pellets, resuspend the pellets with a spoon, and transfer the solution into a Dounce glass homogenizer. Homogenize each 10 min on ice for 45 min. 
		NOTE: In the meantime, chill the rotor in water with ice.</li>
		<li>Pour the solution with depolymerized tubulin into ultracentrifugation tubes and spin for 60 min at 4 &#176;C and 75,000 x g.</li>
	</ol>
	</li>
	<li>Third polymerization
	<ol>
		<li>Determine the volume of supernatant (S5) and add S5/9 mL of DMSO. Pour the suspension into ultracentrifugation tubes and incubate for 20 min at 37 &#176;C to polymerize. 
		NOTE: In the meantime, warm up the rotor to 30 &#176;C in the water bath.</li>
		<li>Centrifuge the polymerized tubulin for 60 min at 75,000 x g and 30 &#176;C. Determine the volume of supernatant (S6) and keep the pellets.</li>
		<li>Dissolve the pellets in 0.25 x S6 mL of PEM buffer (100 mM PIPES pH 6.9, 1 mM EGTA, 1 mM MgSO4, 1 mM DTT, 0.1 mM ATP). Use a Dounce glass homogenizer to enhance solubilization. Keep adding PEM buffer until the pellets are dissolved completely (no opalescent particles are visible anymore).</li>
	</ol>
	</li>
	<li>Ion exchange chromatography purification 
	NOTE: The reverse ion exchange chromatography is employed to remove microtubulin-associated proteins. Tubulin flows freely through the column at the given pH, and contaminating proteins are retained on the phospho-cellulose resin. All steps are performed at temperatures close to 4&#176; C or on ice. Do not clarify the tubulin extract before FPLC purification, either by filtration or centrifugation.</li>
	<li>Equilibrate FPLC and the column with the PEM buffer (100 mM PIPES pH 6.9, 1 mM EGTA, 1 mM MgSO4, 1 mM DTT, 0.1 mM ATP). Use at least 10 column volumes for equilibration. The pump flow should not exceed the flow used for column packing (1 mL/min).</li>
	<li>Load the tubulin suspension slowly (0.5 mL/min) onto the column and collect the unbound protein flowing through the resin.</li>
	<li>Add 10 &#181;L of 100 mM GTP to each 1 mL fraction containing the tubulin protein.</li>
	<li>Freeze fractions containing tubulin in liquid nitrogen and immediately transfer them into the container with liquid nitrogen. Tubulin can be stored for several years under these conditions. The stability decreases rapidly over several months if the tubulin is stored at - 80 &#176;C.</li>
</ol>

Efficient Tubulin Isolation from Neural Tissue

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The authors have nothing to disclose.

The neural tissue from any source is an excellent material for tubulin isolation as the dendrites and axons of neurons are rich in microtubules (up to 40%)23. Brain tissue can be relatively easily obtained in sufficient quantities. The main disadvantage might be an unspecified level of post-translational modifications, which can impact subsequent experiments24,25. Otherwise, the only concern is the rapid degradation of starting material, especially in warm conditions. We employed several transfer-buffer exchanges to enhance heat dissipation prior to the transportation and swift transport for the processing, which begins as soon as possible. The fresh tissue is dissected in conditions preventing heating and homogenized in a buffer containing GTP and glycerol, which further stabilizes tubulin26.
In the presented protocol, the clarification step (step 3.2.7) of the crude lysate has been introduced. Prolonged centrifugation before the first polymerization is generally not recommended due to tubulin susceptibility to irreversible modifications and proteases. On the other hand, the present experiments demonstrated that a short spin in high G-force reduces the debris and thus increases the relative volume of the processed homogenate without significantly impacting tubulin quality. 
The exact protein concentration determination is essential for further experiments, mainly when interactions with tubulin-binding proteins or inhibitors are studied. During the investigations, we encountered significant differences in the results of protein concentration assays. The main reason was the presence of DTT, GTP, and ATP at high concentrations interfering with the assays. The BCA assay was shifted to the upper limits due to the high amount of DTT in the storage buffer, decreasing assay capacity. Moreover, the similar absorbance maxima of proteins and ATP or GTP caused inconsistencies in protein concentration determination using the absorbance at 280 nm. The same issue was noticeable in readouts from the FPLC UV detector. The most reliable assay, with stable results, was the Bradford protein assay, where no influence of buffer compounds was observed. Nevertheless, it is essential to prepare the protein standard dilution series in the storage buffer.
The ability of purified tubulin to create appropriate arrangements is a prerequisite for subsequent experiments. Tubulin is, by nature, highly prone to degradation even in the glycerol-GTP-rich environment, resulting in a reduced capacity to form homogenous filaments. It is critical to follow the purification process and experimentally verify the ability of stored tubulin to polymerize into stable regular filaments. Several independent methods verifying tubulin state have been introduced in vivo and in vitro. Among the most prominent ones, the circular dichroism spectroscopy27, surface plasmon resonance assay28, thermal shift assay29, polymerization inhibition assay30, immunofluorescence staining31,32, coprecipitation22,33 and transmission electron microscope analysis32 can be mentioned. The polymerization assay and coprecipitation of tubulin used in this protocol are easy to perform. They can be swiftly evaluated by either the pellets occurring at the bottom of the tube or using SDS-PAGE. On the other hand, the precipitated tubulin can be in the form of aggregates. More sophisticated methods, such as transmission electron microscopy, must be included to confirm the presence of microtubule fibers for quality control.

Microtubules, hollow protein filaments (24 nm in diameter) formed by alpha- and beta-tubulin heterodimers, are involved in various essential cellular processes. They participate in forming intracellular structures, motility, cell division, cell differentiation, cellular transport, shape maintenance, and secretion1. The cellular functions of microtubules can be affected by direct or indirect interactions with microtubule-associated proteins (MAPs) and other proteins or via complex post-translational modifications defined in the tubulin code2.
Tubulin fibers arise from dynamic non-covalent interactions among alpha- and beta-subunits in the nucleation-elongation mechanism. Short microtubules are formed, and the subsequent growth of tubulin fibers is achieved by reversible elongation at both ends, forming cylinders consisting of tubulin heterodimers arranged in parallel protofilaments2. Dynamic instability refers to the fact that assembled microtubules are often not in equilibrium with their subunits but can undergo phase transition between an extended period of growth and shrinkage while maintaining a steady state1,2.
The dynamic instability of tubulin fibers is principally utilized in many tubulin separation and purification procedures using cycles of high-temperature polymerization and low-temperature depolymerization in the environment of high-purity glycerol, DMSO, GTP/ATP, Mg2+, or other chemical agents (such as taxol or polycations)3. Most of the separation procedures4 are followed by protein chromatography5,6,7,8,9, which ensures the separation of tubulin-associated proteins with nucleoside diphosphokinase and ATPase activity5. Similar results can be obtained by utilizing high-salt buffers10. Multiple sources, including neural11,12,13,14 and non-neural15 tissues, fishes16 (both fresh-water and marine), yeasts, or recombinant variants17 overexpressed in different production strains11,12,13,14, and other sources9,18 were used for fractionation and purification.
The presented protocol utilizes precipitation and protein chromatography to isolate tubulin from the porcine brain into high homogeneity (Figure 1). The main advantage is a relatively high yield achieved with instrumentation available in laboratories equipped for routine molecular biology experiments.

<table><tbody><tr><td>1.5 mL tubes</td><td>---</td><td>---</td><td>Common material</td></tr><tr><td>10 mL tubes</td><td>---</td><td>---</td><td>Common material</td></tr><tr><td>50 mL tubes</td><td>---</td><td>---</td><td>Common material</td></tr><tr><td>ATP</td><td>ROTH</td><td>HN35.2</td><td>Analytical grade</td></tr><tr><td>DMSO</td><td>ROTH</td><td>A994.2</td><td>Analytical grade</td></tr><tr><td>Dounce glass homogenizer</td><td>P-LAB</td><td>H244043</td><td>Homogenizer</td></tr><tr><td>DTT</td><td>ROTH</td><td>6908.1</td><td>Analytical grade</td></tr><tr><td>EGTA</td><td>ROTH</td><td>3054.3</td><td>Analytical grade</td></tr><tr><td>Ethanol</td><td>PENTA</td><td>70390-11001</td><td>Analytical grade</td></tr><tr><td>Glycerol</td><td>ROTH</td><td>6967.2</td><td>Analytical grade</td></tr><tr><td>Graduated beakers</td><td>---</td><td>---</td><td>Common equipment</td></tr><tr><td>Graduated cilinders</td><td>---</td><td>---</td><td>Common equipment</td></tr><tr><td>GTP</td><td>MERCK</td><td>36051-31-7</td><td>Very high quality</td></tr><tr><td>HCl</td><td>PENTA</td><td>19360-11000</td><td>Analytical grade</td></tr><tr><td>Izolated box for tissue transport</td><td>---</td><td>---</td><td>Common equipment</td></tr><tr><td>Kitchen blender</td><td>Waring</td><td>7011HB</td><td>Glass or plastic vessel</td></tr><tr><td>Liquid nitrogen</td><td>---</td><td>---</td><td>Common material</td></tr><tr><td>MES&amp;nbsp;</td><td>ROTH</td><td>6066.4</td><td>Analytical grade</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>MERCK</td><td>814733&amp;nbsp;</td><td>Analytical grade</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>PENTA</td><td>43180-31000</td><td>Analytical grade</td></tr><tr><td>NaOH</td><td>PENTA</td><td>15650-11000</td><td>Analytical grade</td></tr><tr><td>Optima XPN100</td><td>Beckman Coulter&amp;nbsp;</td><td>A94469</td><td>Ultracentrifuge</td></tr><tr><td>Phosphocellulose column</td><td>VWR</td><td>GENO786-1291</td><td>Empty column</td></tr><tr><td>Phosphocellulose resin</td><td>Creative - Biomart, inc</td><td>Phosphate-001C</td><td>Ion exchange resin</td></tr><tr><td>PIPES</td><td>ROTH</td><td>9156.2</td><td>Analytical grade</td></tr><tr><td>Saccharose</td><td>PENTA</td><td>24970-31000</td><td>Analytical grade</td></tr><tr><td>Scales</td><td>---</td><td>---</td><td>Common equipment</td></tr><tr><td>Scissors</td><td>---</td><td>---</td><td>Common equipment</td></tr><tr><td>Spoons&amp;nbsp;</td><td>---</td><td>---</td><td>Plastic or glass</td></tr><tr><td>Ti45 rotor</td><td>Beckman Coulter&amp;nbsp;</td><td>339160</td><td>Rotor for Ultracentrifuge</td></tr><tr><td>Tweezers</td><td>---</td><td>---</td><td>Common equipment</td></tr><tr><td>Ultra turrax IKA T18 basic</td><td>IKA</td><td>356 1000</td><td>Laboratory dispenser</td></tr><tr><td>Water bath 37 &amp;deg;C</td><td>---</td><td>---</td><td>Stirred</td></tr></tbody></table>

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.

During the separation and purification steps, samples for SDS-PAGE electrophoresis were taken and subsequently analyzed using Coomassie-blue staining (Figure 2). 20 &#181;L of each sample was mixed with 10 &#181;L of Laemmli sample buffer (188 mM Tris-HCl pH 6.8, 3% SDS (w/w), 30% glycerol (v/v), 0.01 bromophenol blue (w/w), 15% &#946;-mercaptoethanol) and incubated at 95 &#176;C for 15 min. 4 &#181;L of each sample was loaded on 12.5% acrylamide SDS gel and separated under a constant current of 30 mA per gel in reducing and denaturing conditions.
The results confirmed an incremental increase in the relative tubulin concentration accompanied by a reduction in contaminating proteins. Moreover, there was no significant loss of tubulin in clarified lysate in the first centrifugation (step 3.2.7) compared to omitting this step (Figure 2A,B).
The protein concentration was determined using three independent methods: BCA assay, Bradford protein assay, and SDS-PAGE gel densitometry analysis19 (Figure 3). The overall yield of tubulin using the procedure described was 123 mg of purified tubulin from 250 g of neural tissue. During the measurements, it is necessary to take into account that the high DTT concentration in the storage buffer has a significant impact on the BCA assay. Both the PEM buffer and PBS buffer, with the addition of DTT, increase the background absorbance by about 0.900 A595, which significantly reduces the capacity of the BCA assay (Figure 3A). The negative effect of DTT is detectable even after ten times dilution with pure water (data not shown). The Bradford assay did not seem to be affected by the storage buffer (Figure 3B), as confirmed by densitometry analysis.
The purity of tubulin preparation was verified by mass spectrometry analysis at two independent facilities (VRI Brno, Czech Republic; CEITEC MU Brno, Czech Republic) using electrospray ionization and MALDI-TOF. Both analyses confirmed the presence of porcine tubulins alpha and beta in several isoforms. The overall purity was more than 97.07% (PSMs 1065), with the most prevalent impurities originating from Homo sapiens Keratin Type II (PSMs 246, 2.24% of impurities) which were introduced most probably during the tubulin isolation and sample preparation for MS/MS analysis. Other impurities composed of 322 PSMs and only Serum albumin, Actin gamma, and Trypsinogen originating from Sus scrofa were identified with one peptide resolution (0.0069%).
In subsequent experiments, the preservation of polymerization capacity after snap freezing and storage in liquid nitrogen was verified. The 10 mg/mL aliquot was removed from liquid nitrogen and slowly thawed on ice. Samples of different concentrations (10 mg/mL, 8 mg/mL, 6 mg/mL, 4 mg/mL, and 2 mg/mL) were prepared by diluting the aliquots in PEM buffer containing DTT, ATP, and GTP for the self-assembly experiment. One dilution series was incubated at 37 &#176;C for 60 min, and the second one was incubated on ice for 60 min. Both series were centrifuged for 60 min at 21,000 x g and the corresponding temperature (4 &#176;C or 37 &#176;C). 30 &#181;L of supernatant was removed and used for SDS-PAGE. The remaining supernatant was carefully removed using a pipette and discarded. Pellets were briefly washed by adding 100 &#181;M of PEM buffer, followed by immediate removal using a pipette. Subsequently, the pellets were resuspended in 50 &#181;L of 1x concentrated SDS-loading buffer so that the relative concentration of pellet and supernatant is preserved. 10 &#181;L of SDS-loading buffer was added to each supernatant. All samples were analyzed using the SDS-PAGE and Coomassie Staining (Figure 4). The volume of each sample loaded to SDS-PAGE was adjusted according to the starting concentration, so the difference in precipitation due to the concentration is more obvious. The self-assembly test of tubulin in PEM buffer confirmed the ability to form tubulin fibers in a temperature-dependent manner.
The MAP2c driven tubulin20,21 assembly assay verifying the ability of tubulin to interact with other proteins was performed22. The serial dilutions of MAP2c were prepared where 100 &#181;L of 1 mg/mL tubulin aliquots were mixed with MAP2c to the final concentration ranging from 0 &#181;M to 8 &#181;M. All samples were prepared on ice by diluting in freshly prepared PEM buffer with 1 mM DTT and 1 mM GTP. The tubulin was incubated for 15 min at 37 &#176;C with different concentrations of MAP2c, and centrifuged for 60 min at 21,000 x g and 37 &#176;C. The final wash of the pellet was performed twice with 100 &#181;L of PEM buffer. The experiment confirmed the capacity of the prepared tubulin to undergo MAP2c-driven polymerization, as only the sample without MAP2c did not form a pellet (Figure 5).
Transmission electron microscopy was further used to confirm the presence of tubulin filaments from co-polymerization experiments. Suspensions of purified tubulin precipitated with MAP2c were prepared for transmission electron microscopy using negative staining. The samples were adsorbed onto Formvar-coated, carbon-stabilized copper grids. The grids were then negatively stained with 2% NH4MoO4 and examined under an electron microscope at 18,000x magnification and an accelerating voltage of 80 kV. The presence of homogenous microtubules with a clear filamentous structure and appropriate size showed the ability to form microtubules in native conformation (Figure 6).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64925/64925fig01.jpg" /> 
Figure 1: A schematic diagram of tubulin separation and purification. The number in parentheses refers to the protocol step. <a href="https://www.jove.com/files/ftp_upload/64925/64925fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64925/64925fig02.jpg" /> 
Figure 2: Tubulin separation and purification. (A) The SDS-PAGE analysis of samples taken during the separation of tubulin utilizing temperature-driven polymerization (3 &#181;L per line). The decrease in impurities with a stable increase in relative tubulin abundance (approximately 50 kDa) is visible. The numbers above each line correspond to the step number in the protocol. (B) The SDS-PAGE analysis of fractions obtained after protein chromatography on phospho-cellulose resin (4 &#181;L per line). <a href="https://www.jove.com/files/ftp_upload/64925/64925fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64925/64925fig03.jpg" /> 
Figure 3: Protein concentration. The dilution series of bovine serum albumin of up to 1 mg/mL in PEM buffer or PBS was compared with BSA dilution series containing 0.1 ATP, 1 mM GTP, 1 mM DTT separately or in combination (0.1 mM ATP, 1 mM GTP, and 1 mM DTT) in the PEM or PBS buffer. (A) When the BCA assay was used, there was a significant shift in the background of samples containing DTT (solid symbols). (B) This effect was not detected when concentrations were measured in the Bradford assay. <a href="https://www.jove.com/files/ftp_upload/64925/64925fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64925/64925fig04.jpg" /> 
Figure 4: Tubulin self-assembly assay. The SDS-PAGE analysis of the self-assembly assay confirmed the capacity of stored tubulin incubated at either (A) 4 &#176;C or (B) 37 &#176;C to polymerize in a temperature-dependent manner in a broad spectrum of concentrations. (P - pellet, S - supernatant; the respective concentration of tubulin is denoted above each line pair; the amount of each sample loaded was 10 &#181;g). <a href="https://www.jove.com/files/ftp_upload/64925/64925fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64925/64925fig05.jpg" /> 
Figure 5: Tubulin co-sedimentation assay. The SDS-PAGE analysis of MAP2c-assisted tubulin assembly confirmed the capacity of stored tubulin to undergo polymerization driven by interaction with microtubule-associated protein in a concentration-dependent manner. The molar concentration of MAP2c is indicated above each line. (P - pellet, S - supernatant). <a href="https://www.jove.com/files/ftp_upload/64925/64925fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64925/64925fig06.jpg" /> 
Figure 6: Transmission electron microscopy. The TEM microphotographs showed that tubulin assembles (A) into microtubules of appropriate diameter (B) consisting of homogenous tubulin filaments decorated with MAP2c proteins. The bar corresponds to 1000 nm (A) and 200 nm (B). <a href="https://www.jove.com/files/ftp_upload/64925/64925fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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.

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 ...

optimizing-tubulin-yield-from-porcine-brain-tissue

Research

JoVE Journal

Biochemistry

1.5K Views.  Veterinary Research Institute. 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.

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

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 ...

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 ...

Peering at Brain Polysomes with Atomic Force Microscopy

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, formed by ribosomes, mRNAs, several proteins and non-coding RNAs, represent integrated platforms where translational controls take place. However, while the ribosome has been widely studied, the organization of polysomes is still lacking comprehensive understanding. Thus much effort is required in order to elucidate polysome organization and any novel mechanism of translational control that may be embedded. Atomic force microscopy (AFM) is a type of scanning probe microscopy that allows the acquisition of 3D images at nanoscale resolution. Compared to electron microscopy (EM) techniques, one of the main advantages of AFM is that it can acquire thousands of images both in air and in solution, enabling the sample to be maintained under near physiological conditions without any need for staining and fixing procedures. Here, a detailed protocol for the accurate purification of polysomes from mouse brain and their deposition on mica substrates is described. This protocol enables polysome imaging in air and liquid with AFM and their reconstruction as three-dimensional objects. Complementary to cryo-electron microscopy (cryo-EM), the proposed method can be conveniently used for systematically analyzing polysomes and studying their organization.

While ribosome structure has been extensively characterized, the organization of polysomes is still understudied. To overcome this lack of knowledge, we present here a detailed preparation protocol for accurate imaging of mammalian polysomes by atomic force microscopy (AFM) in air and liquid.

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, ...

Neuroscience

Improved Method for the Establishment of an In Vitro Blood-Brain Barrier Model Based on Porcine Brain Endothelial Cells

The aim of this protocol presents an optimized procedure for the purification and cultivation of pBECs and to establish in vitro blood-brain barrier (BBB) models based on pBECs in mono-culture (MC), MC with astrocyte-conditioned medium (ACM), and non-contact co-culture (NCC) with astrocytes of porcine or rat origin. pBECs were isolated and cultured from fragments of capillaries from the brain cortices of domestic pigs 5-6 months old. These fragments were purified by careful removal of meninges, isolation and homogenization of grey matter, filtration, enzymatic digestion, and centrifugation. To further eliminate contaminating cells, the capillary fragments were cultured with puromycin-containing medium. When 60-95% confluent, pBECs growing from the capillary fragments were passaged to permeable membrane filter inserts and established in the models. To increase barrier tightness and BBB characteristic phenotype of pBECs, the cells were treated with the following differentiation factors: membrane permeant 8-CPT-cAMP (here abbreviated cAMP), hydrocortisone, and a phosphodiesterase inhibitor, RO-20-1724 (RO). The procedure was carried out over a period of 9-11 days, and when establishing the NCC model, the astrocytes were cultured 2-8 weeks in advance. Adherence to the described procedures in the protocol has allowed the establishment of endothelial layers with highly restricted paracellular permeability, with the NCC model showing an average transendothelial electrical resistance (TEER) of 1249 &#177; 80 &#937; cm2, and paracellular permeability (Papp) for Lucifer Yellow of 0.90 10-6 &#177; 0.13 10-6 cm sec-1 (mean &#177; SEM, n=55). Further evaluation of this pBEC phenotype showed good expression of the tight junctional proteins claudin 5, ZO-1, occludin and adherens junction protein p120 catenin. The model presented can be used for a range of studies of the BBB in health and disease and, with the highly restrictive paracellular permeability, this model is suitable for studies of transport and intracellular trafficking.

Improved Method for the Establishment of an In Vitro Blood-Brain Barrier Model Based on Porcine Brain Endothelial Cells

The aim of the protocol is to present an optimized procedure for the establishment of an in vitro blood-brain barrier (BBB) model based on primary porcine brain endothelial cells (pBECs). The model shows high reproducibility, high tightness, and is suitable for studies of transport and intracellular trafficking in drug discovery.

The aim of this protocol presents an optimized procedure for the purification and cultivation of pBECs and to establish in vitro blood-brain barrier (BBB) ...

Medicine

Sampling Strategies and Processing of Biobank Tissue Samples from Porcine Biomedical Models

In translational medical research, porcine models have steadily become more popular. Considering the high value of individual animals, particularly of genetically modified pig models, and the often-limited number of available animals of these models, establishment of (biobank) collections of adequately processed tissue samples suited for a broad spectrum of subsequent analyses methods, including analyses not specified at the time point of sampling, represent meaningful approaches to take full advantage of the translational value of the model. With respect to the peculiarities of porcine anatomy, comprehensive guidelines have recently been established for standardized generation of representative, high-quality samples from different porcine organs and tissues. These guidelines are essential prerequisites for the reproducibility of results and their comparability between different studies and investigators. The recording of basic data, such as organ weights and volumes, the determination of the sampling locations and of the numbers of tissue samples to be generated, as well as their orientation, size, processing and trimming directions, are relevant factors determining the generalizability and usability of the specimen for molecular, qualitative, and quantitative morphological analyses. Here, an illustrative, practical, step-by-step demonstration of the most important techniques for generation of representative, multi-purpose biobank specimen from porcine tissues is presented. The methods described here include determination of organ/tissue volumes and densities, the application of a volume-weighted systematic random sampling procedure for parenchymal organs by point-counting, determination of the extent of tissue shrinkage related to histological embedding of samples, and generation of randomly oriented samples for quantitative stereological analyses, such as isotropic uniform random (IUR) sections generated by the &#34;Orientator&#34; and &#34;Isector&#34; methods, and vertical uniform random (VUR) sections.

The practical application and performance of methods for generation of representative tissue samples of porcine animal models for a broad spectrum of downstream analyses in biobank projects are demonstrated, including volumetry, systematic random sampling, and differential processing of tissue samples for qualitative and quantitative morphologic and molecular analyses types.

In translational medical research, porcine models have steadily become more popular. Considering the high value of individual animals, particularly of ...

Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

The ubiquitin-proteasome system is a key regulator of protein degradation and a variety of other cellular processes in eukaryotes. In the brain, increases in ubiquitin-proteasome activity are critical for synaptic plasticity and memory formation and aberrant changes in this system are associated with a variety of neurological, neurodegenerative and psychiatric disorders. One of the issues in studying ubiquitin-proteasome functioning in the brain is that it is present in all cellular compartments, in which the protein targets, functional role and mechanisms of regulation can vary widely. As a result, the ability to directly compare brain ubiquitin protein targeting and proteasome catalytic activity in different subcellular compartments within the same animal is critical for fully understanding how the UPS contributes to synaptic plasticity, memory and disease. The method described here allows collection of nuclear, cytoplasmic and crude synaptic fractions from the same rodent (rat) brain, followed by simultaneous quantification of proteasome catalytic activity (indirectly, providing activity of the proteasome core only) and linkage-specific ubiquitin protein tagging. Thus, the method can be used to directly compare subcellular changes in ubiquitin-proteasome activity in different brain regions in the same animal during synaptic plasticity, memory formation and different disease states. This method can also be used to assess the subcellular distribution and function of other proteins within the same animal.

This protocol is designed to efficiently quantify ubiquitin-proteasome system (UPS) activity in different cellular compartments of the rodent brain. Users are able to examine UPS functioning in nuclear, cytoplasmic and synaptic fractions in the same animal, reducing the amount of time and number of animals needed to perform these complex analyses.

The ubiquitin-proteasome system is a key regulator of protein degradation and a variety of other cellular processes in eukaryotes. In the brain, increases ...

A Time-Efficient Fluorescence Spectroscopy-Based Assay for Evaluating Actin Polymerization Status in Rodent and Human Brain Tissues

Actin, the major component of cytoskeleton, plays a critical role in the maintenance of neuronal structure and function. Under physiological states, actin occurs in equilibrium in its two forms: monomeric globular (G-actin) and polymerized filamentous (F- actin). At the synaptic terminals, actin cytoskeleton forms the basis for critical pre- and post-synaptic functions. Moreover, dynamic changes in the actin polymerization status (interconversion between globular and filamentous forms of actin) are closely linked to plasticity-related alterations in synaptic structure and function. We report here a modified fluorescence-based methodology to assess polymerization status of actin in ex vivo conditions. The assay employs fluorescently labelled phalloidin, a phallotoxin that specifically binds to actin filaments (F-actin), providing a direct measure of polymerized filamentous actin. As a proof of principle, we provide evidence for the suitability of the assay both in rodent and post-mortem human brain tissue homogenates. Using latrunculin A (a drug that depolymerizes actin filaments), we confirm the utility of the assay in monitoring alterations in F-actin levels. Further, we extend the assay to biochemical fractions of isolated synaptic terminals wherein we confirm increased actin polymerization upon stimulation by depolarization with high extracellular K+.

We report a simple, time-efficient and high-throughput fluorescence spectroscopy-based assay for the quantification of actin filaments in ex vivo biological samples from brain tissues of rodents and human subjects.

Actin, the major component of cytoskeleton, plays a critical role in the maintenance of neuronal structure and function. Under physiological states, actin ...

A High Output Method to Isolate Cerebral Pericytes from Mouse

In recent years, cerebral pericytes have become the focus of extensive research in vascular biology and pathology. The importance of pericytes in blood brain barrier formation and physiology is now demonstrated but its molecular basis remains largely unknown. As the pathophysiological role of cerebral pericytes in neurological disorders is intriguing and of great importance, the in vitro models are not only sufficiently appropriate but also able to incorporate different techniques for these studies. Several methods have been proposed as in vitro models for the extraction of cerebral pericytes, although an antibiotic-free protocol with high output is desirable. Most importantly, a method that has increased output per extraction reduces the usage of more animals.
Here, we propose a simple and efficient method for extracting cerebral pericytes with sufficiently high output. The mouse brain tissue homogenate is mixed with a BSA-dextran solution for the separation of the tissue debris and microvascular pellet. We propose a three-step separation followed by filtration to obtain a microvessel rich filtrate. With this method, the quantity of microvascular fragments obtained from 10 mice is sufficient to seed 9 wells (9.6 cm2 each) of a 6-well plate. Most interestingly with this protocol, the user can obtain 27 pericyte rich wells (9.6 cm2 each) in passage 2. The purity of the pericyte cultures are confirmed with the expression of classical pericyte markers: NG2, PDGFR-&#946; and CD146. This method demonstrates an efficient and feasible in vitro tool for physiological and pathophysiological studies on pericytes.

We present a protocol for the extraction of murine cerebral pericytes. Based on an antibiotic-free enrichment oriented pericyte extraction, this protocol is a valuable tool for in vitro studies providing high purity and high yield, thus decreasing the number of experimental animals used.

In recent years, cerebral pericytes have become the focus of extensive research in vascular biology and pathology. The importance of pericytes in blood ...

Biology

Isolating Tumor-Infiltrating Immune Cells from a Murine Brain

Source: Baker, G. J., et al., Lowenstein, P. R. <a href="https://app.jove.com/v/53676">Isolation and Flow Cytometric Analysis of Glioma-infiltrating Peripheral Blood Mononuclear Cells.</a> J. Vis. Exp. (2015)
The video demonstrates the isolation of tumor-infiltrating peripheral blood mononuclear cells from a mouse brain using mechanical dissociation and enzymatic digestion. This is followed by density gradient centrifugation to collect the immune cells for further analysis.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
Prepare density centrifugation media ...

JoVE Encyclopedia of Experiments

Neuronal Culture Techniques

Co-culture and Interaction Studies

Purifying Capillaries From a Porcine Brain

Source: Nielsen, S. S. E., et al. <a href="https://app.jove.com/v/56277">Improved Method for the Establishment of an In Vitro Blood-Brain Barrier Model Based on Porcine Brain Endothelial Cells.</a> J. Vis. Exp. (2017).
This video demonstrates the isolation of capillaries from fresh porcine brains. The process involves removing the meninges, isolating the grey matter from the brain tissue, homogenizing and filtering the grey matter to isolate the capillaries, and finally using enzymatic digestion to purify the capillaries further.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Purification of Porcine Brain ...

Optimizing Tubulin Yield from Porcine Brain Tissue

Summary

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Chapters in this video

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

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

Peering at Brain Polysomes with Atomic Force Microscopy

Improved Method for the Establishment of an In Vitro Blood-Brain Barrier Model Based on Porcine Brain Endothelial Cells

Sampling Strategies and Processing of Biobank Tissue Samples from Porcine Biomedical Models

Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

A Time-Efficient Fluorescence Spectroscopy-Based Assay for Evaluating Actin Polymerization Status in Rodent and Human Brain Tissues

A High Output Method to Isolate Cerebral Pericytes from Mouse

Isolating Tumor-Infiltrating Immune Cells from a Murine Brain

Purifying Capillaries From a Porcine Brain