This work was supported by the Neurological Foundation of New Zealand (1835-PG), the New Zealand Health Research Council (#16-597) and the Department of Anatomy, University of Otago, New Zealand. We are indebted to the Neurological Tissue Bank of HCB-IDIBAPS BioBank (Spain) for human brain tissues. We thank Jiaxian Zhang for her help in recording and editing of the video.

All experimental procedures were carried out in accordance with the regulations of the University of Otago Committee on Ethics in the Care and Use of Laboratory Animals (Ethics Protocol No. AUP95/18 and AUP80/17) and New Zealand legislature. Human brain tissues were obtained from the Neurological Tissue Bank of Hospital Cl&#237;nic-IDIBAPS BioBank in Barcelona, Spain. All tissue collection protocols were approved by the Ethics Committee of Hospital Cl&#237;nic, Barcelona, and informed consent was obtained from the families.
1. Preparation of buffers and reagents
<ol>
	<li>Prepare the following buffers for the homogenization of brain tissue and the preparation of enriched fraction of synaptic terminals: 
	Homogenization buffer: 5 mM HEPES, pH 7.4 supplemented with 0.32 M sucrose 
	Resuspension buffer: 5 mM Tris, pH 7.4 supplemented with 0.32 M sucrose 
	Washing buffer: 5 mM Tris, pH 8.1 
	1.2 M sucrose 
	1.0 M sucrose 
	0.85 M sucrose</li>
	<li>Add homemade or commercial mix of protease and phosphatase inhibitors. 
	NOTE: We have used EDTA-free version of Complete Protease Inhibitor mix (1 tablet per 10 mL buffer) and Phosphatase inhibitor cocktail IV (1:100; volume: volume).</li>
	<li>Prepare the following buffers and reagents for the fixation, permeabilization and binding of phalloidin: 
	Krebs buffer: 118.5 mM NaCl, 4.7 mM KCl, 1.2 mM MgCl2, 2 mM CaCl2, 0.1 mM KH2PO4, 5 mM NaHCO3, 10 mM glucose, 20 mM HEPES, pH 7.4 
	1 M KCl 
	25% glutaraldehyde (stock solution) 
	Krebs buffer containing 0.1 % Triton X-100 and 1 mg/mL NaBH4 
	400x Alexa Fluor 647 Phalloidin in DMSO 
	Krebs buffer containing 0.32 M sucrose 
	50&#160;&#956;M latrunculin A in DMSO (stock solution) 
	1&#160;M KCl (stock solution) 
	CAUTION: Phalloidin is toxic (LD50 = 2 mg/kg) and must be handled with care. Inhalation of glutaraldehyde is toxic and it should be handled in a fume hood.</li>
	<li>Store the buffers at 4 &#176;C and phalloidin and latrunculin A at -20 &#176;C for long-term storage. 
	&#8203;NOTE: Long-term storage of buffers is not recommended. The protocol can be paused here.</li>
</ol>
2. Brain tissue homogenization
<ol>
	<li>Homogenize the cryopreserved or freshly dissected rat brain tissue in 10 volumes of homogenization buffer in a Potter-Elvehjem glass tube and pestle on ice. 
	NOTE: Optimal homogenization is achieved by 15-20 strokes of the pestle by hand for brain tissues. Successful homogenization can be confirmed by smooth flow of the suspension through the glass tube. The protocol can be paused here.</li>
	<li>Determine the protein concentration in the homogenate using a Bradford assay31. 
	NOTE: Alternate homemade or commercial assays for protein estimation can also be used.</li>
	<li>Dilute homogenate samples in Krebs buffer at a concentration of 2-3 mg protein/mL in a volume of 50 &#181;L. 
	&#8203;NOTE: In some of our experiments, we incubate homogenates with 2 &#956;M latrunculin A or DMSO (controls) at 37 &#176;C for 1 h. For this, homogenates are resuspended in 48 &#181;L of Krebs buffer, and 2 &#181;L of 50 &#956;M latrunculin A or 2 &#181;L of DMSO is added. Further for immunoblotting, a small amount of sample (10&#160;&#956;g) is collected after the incubation.</li>
	<li>Fix the homogenate samples (section 5).</li>
</ol>
3. Preparation of isolated nerve terminals
<ol>
	<li>Preparation of synaptosomes 
	NOTE: Alternative protocols32,33 for synaptosomes can also be used.
	<ol>
		<li>Centrifuge brain homogenate at 1,200 x g for 10 min at 4 &#176;C.</li>
		<li>Discard the pellet, which is the crude nuclear fraction.</li>
		<li>Further centrifuge the supernatant (S1) obtained in step 3.1.1 at 12,000 x g for 15 min at 4 &#176;C.</li>
		<li>Remove the supernatant (S2), which is the soluble cytosolic fraction.</li>
		<li>Resuspend the pellet (P2) obtained in step 3.1.3, which is the crude synaptosomal fraction in resuspension buffer. 
		NOTE: The volume of resuspension buffer depends on the amount of starting tissue and the amount of pellet obtained. For example, when starting with 150-300 mg of brain tissues, the pellet obtained can be resuspended in 200 &#181;L of resuspension buffer.</li>
		<li>Load the resuspended crude synaptosomes onto a discontinuous sucrose gradient made of equal volumes of 0.85-1.0-1.2 M sucrose. 
		NOTE: We typically use 1 mL each of the sucrose solution (for 150-300 mg tissue). This can be changed according for larger tissue amounts. Discontinuous gradients can be made using a 25G syringe pressed against the internal wall of the ultracentrifuge tube and gentle layering of the sucrose layers.</li>
		<li>Centrifuge at 85,000 x g for 2 h at 4 &#176;C. 
		NOTE: Because of the high-speed of centrifugation, an ultracentrifuge capable of creating a vacuum to reduce heating is required.</li>
		<li>Collect the synaptosomal fraction at the interface between 1.0 and 1.2 M sucrose using a 200 &#181;L pipet tip.</li>
		<li>Wash the synaptosomal fraction with ice-cold washing buffer by centrifugation at 18,000 x g for 10 minutes at 4 &#176;C. 
		NOTE: For washing, the synaptosomal fraction obtained at the interface of 1.0 and 1.2 M sucrose is collected in a fresh 1.5 mL tube, and an equal volume of washing buffer is added, ensuring the removal of high sucrose from the medium.</li>
		<li>Wash the synaptosomal pellet again with ice-cold homogenization buffer at 12,000 g for 10 minutes at 4 &#176;C.</li>
		<li>Resuspend the synaptosomes in homogenization buffer on ice. 
		NOTE: The protocol can be briefly paused here.</li>
		<li>Determine the protein concentration using a Bradford assay.</li>
		<li>Resuspend the synaptosomes in Krebs buffer at a concentration of 2-3 mg protein/mL in a volume of 50 &#181;L (47.5 &#181;L if synaptosomes are to be depolarized by KCl; see section 4). 
		NOTE: For resuspension, the synaptosomal fraction is first spun at 12,000 x g for 5 min at 4 &#176;C and the supernatant (buffer) is removed. The synaptosomal pellet is then resuspended in Krebs buffer by gentle pipetting using a 200 &#181;L pipet tip.</li>
		<li>Proceed with depolarization (section 4).</li>
	</ol>
	</li>
	<li>Preparation of synaptoneurosomes 
	NOTE: Alternative protocols34,35 for synaptoneurosomes can also be used.
	<ol>
		<li>Pass the brain homogenate through a pre-wetted net filter of 100 &#956;m pore size in a filter holder using a 1 mL syringe. 
		NOTE: Pre-wetting of all filters is important to avoid loss of sample and should be done using homogenization buffer. For this, homogenization buffer is passed through the filters in the filter holder using a 1 mL syringe until the&#160;buffer flow out.</li>
		<li>Collect the filtrate (F1) in a pre-chilled 1.5 mL tube on ice.</li>
		<li>Repeat the process (steps 3.2.1 and 3.2.2)&#160;for F1 fraction to obtain the second filtrate (F2).</li>
		<li>Pass F2 filtrate through a net filter of 5 &#956;m pore size.</li>
		<li>Collect the filtrate (F3) in a pre-chilled 1.5 mL tube on ice.</li>
		<li>Centrifuge filtrate F3 at 1,500 x g for 10 min at 4 &#176;C.</li>
		<li>Resuspend the pellet (synaptoneurosomes) in Krebs buffer on ice at a concentration of 2-3 mg protein/mL in a volume of 50 &#181;L (47.5 &#181;L if synaptosomes are to be depolarized by KCl; see section 4). 
		NOTE: The protocol can be briefly paused here.</li>
		<li>Estimate the protein concentration using a Bradford assay.</li>
		<li>Proceed with depolarization (section 4).</li>
	</ol>
	</li>
</ol>
4. KCl-mediated depolarization of isolated synaptic terminals
<ol>
	<li>Equilibrate synaptosomes/synaptoneurosomes at 37 &#176;C for 5-10 min.</li>
	<li>Stimulate synaptosomes/synaptoneurosomes by adding KCl to increase extracellular K+ to 50 mM for 30 s at 37 &#176;C and add equal volume of Krebs buffer to the respective unstimulated control set. 
	NOTE: For example, add 2.5 &#181;L of 1 M KCl to synaptosomes resuspended in 47.5 &#181;L of Krebs buffer; and add 2.5 &#181;L of Krebs buffer to the respective unstimulated control synaptosome. For experiments wherein a large number of samples are involved, proceed with no more than 2 samples at a time so that the depolarization time does not exceed 30 s.</li>
	<li>Terminate stimulation by adding glutaraldehyde (section 5).</li>
</ol>
5. Fixation and phalloidin staining of samples
<ol>
	<li>Add glutaraldehyde to homogenate/synaptosomal/synaptoneurosomal samples to a final concentration of 2.5% for 2-3 min at room temperature. 
	NOTE: We added 6 &#181;L of 25% glutaraldehyde solution so that the final concentration of glutaraldehyde in the 50 &#181;L samples (homogenates/synaptosomes/synaptoneurosomes) was ca. 2.5%. Fixation is critical and should be fast and hence immediately after adding glutaraldehyde, the sample should be vigorously vortexed.</li>
	<li>Sediment the samples at 20,000 x g for 5 min.</li>
	<li>Remove the supernatant. 
	NOTE: Discard the supernatant in a fume hood as glutaraldehyde is toxic.</li>
	<li>Permeabilize the pellet by resuspension in 100 &#181;L of Krebs buffer containing 0.1% Triton X-100 and 1 mg/mL NaHB4 for 2-3 min at room temperature.</li>
	<li>Sediment the samples at 20,000 x g for 5 min.</li>
	<li>Remove the permeabilization buffer.</li>
	<li>Wash the pellet with 200 &#181;L of Krebs buffer by centrifugation&#160;at 20,000 x g for 5 min.</li>
	<li>Resuspend and stain the pellet with 1x Alexa Fluor 647 Phalloidin (corresponding to 500 &#956;U) in 100 &#181;L of Krebs buffer for 10 minutes in dark at room temperature. 
	NOTE: Other varieties of fluorescent phalloidin analogs are commercially available and can be replaced for the assay. Concentration of phalloidin and total sample volume of incubation may have to be modified according to the amount and type of tissue/sample being tested and optimal conditions should be standardized accordingly.</li>
	<li>Centrifuge the stained samples at 20,000 x g for 5 min.</li>
	<li>Remove the unbound phalloidin (supernatant).</li>
	<li>Wash the sample with 200 &#181;L of Krebs buffer by centrifugation at 20,000 x g for 5 min.</li>
	<li>Resuspend in 200 &#181;L of Krebs buffer containing 0.32 M sucrose. 
	&#8203;NOTE: The protocol can be briefly paused here.</li>
</ol>
6. Fluorometric analysis and light scattering
<ol>
	<li>Dispense Alexa Fluor 647 Phalloidin-stained samples in a black 96-well plate.</li>
	<li>Measure the fluorescence intensity at an excitation wavelength of 645 nm and an emission wavelength of 670 nm in a plate reader at room temperature.</li>
	<li>Transfer the samples from the black 96-well plate to the transparent 96-well plate using 200 &#181;L pipet tips.</li>
	<li>Measure the light scattering at 540 nm to correct for any losses that might have occurred during the previous steps of fixation, permeabilization and staining. 
	NOTE: Variations in biological material retained in the stained samples might be more prominent for smaller amounts of starting material (see Discussion).</li>
	<li>Include a set of Alexa Fluor 647 Phalloidin in Krebs buffer at different concentrations (0.05x, 0.1x, 0.25x, 0.35x, 0.75x, 0.5x and 1x corresponding to 25, 50, 125, 175, 250, 375 and 500 &#956;U) for each batch of the assay as a standard curve. 
	&#8203;NOTE: This is an optional step and does not affect the results of the assay particularly when F-actin levels are being expressed in a relative manner (Section 7). The protocol can be paused here.</li>
</ol>
7. Data analysis
<ol>
	<li>The amount of F-actin in the samples is directly proportional to the fluorescence intensity of bound phalloidin. Express in absolute terms of units of phalloidin bound calculated from the linear curve of the tagged phalloidin standard. 
	NOTE: As an example, see Figures 2A-B, 3A, 4A-B and Supplementary Figure 2.</li>
	<li>Express F-actin levels as a fraction of the control samples. 
	NOTE: As an example, see Figures 5A-B.</li>
</ol>

The authors have nothing to disclose.

The assay described here, essentially adapted from a previous study30 with modifications, employs a phallotoxin, phalloidin tagged with a fluorescent label. Fluorescent phalloidin analogs are considered to be the gold standard for staining actin filaments in fixed tissues47,48,49. In fact, they are the oldest tools to specifically identify actin filaments50 and still remain the mos...

Cytoskeletal protein actin is involved in multiple cellular functions, including structural support, cellular transport, cell motility and division. Actin occurs in equilibrium in two forms: monomeric globular actin (G-actin) and polymerized filamentous actin (F-actin). Rapid changes in the polymerization status of actin (interconversion between its G- and F- forms) result in rapid filament assembly and disassembly and underlie its regulatory roles in cellular physiology. Actin forms the major component of the neuronal cytoskeletal structure and influences a wide range of neuronal functions1,2. Of note, the actin cytoskeleton forms an integral part of the structural platform of the synaptic terminals. As such, it is a major determinant of synaptic morphogenesis and physiology and plays a fundamental role in control of the size, number and morphology of synapses3,4,5. In particular, dynamic actin polymerization-depolymerization is a key determinant of the synaptic remodelling associated with synaptic plasticity underlying the memory and learning processes. Indeed, both presynaptic (such as neurotransmitter release6,7,8,9,10) and postsynaptic functions (plasticity related dynamic remodeling11,12,13,14) critically rely on dynamic changes in the polymerization status of the actin cytoskeleton.
Under physiological conditions, F-actin levels are dynamically and tightly regulated through a multimodal pathway involving posttranslational modification4,15,16 as well as actin-binding proteins (ABPs)4,17. ABPs can influence actin dynamics at multiple levels (such as initiating or inhibiting polymerization, inducing filament branching, severing of filaments to smaller pieces, promoting depolymerization, and protecting against depolymerization), and are in-turn under a stringent modulatory control sensitive to various extra- and intracellular signals18,19,20. Such regulatory checks at multiple levels dictate a strict regulation of actin dynamics at the synaptic cytoskeleton, fine-tuning pre- and postsynaptic aspects of neuronal physiology both at the basal and activity-induced states.
Given the important roles of actin in neuronal physiology, it is not surprising that several studies have provided evidence for alterations in actin dynamics as critical pathogenic events linked to a wide range of neurological disorders including neurodegeneration, psychological diseases as well as neurodevelopmental ailments3,21,22,23,24,25,26,27. In spite of the wealth of research data pointing to key roles of actin in neuronal physiology and pathophysiology, however, significant gaps still remain in the understanding of actin dynamics, particularly at the synaptic cytoskeleton. More research studies are needed to have a better comprehension of neuronal actin and its alterations under pathological conditions. One major area of focus in this context is the assessment of actin polymerization status. There are Western blotting-based commercial kits (G-Actin/F-Actin in vivo assay biochemical kit; Cytoskeleton SKU BK03728,29) and home-made assays for the assessment of F-actin levels6. However, because these require biochemical isolation of F-actin and G-actin and because their subsequent quantification is based upon immunoblotting protocols, they can be time consuming. We herein report a fluorescence spectroscopy-based assay adapted from a previous study30 with modifications that can be used to evaluate both basal levels of F-actin, as well as dynamic changes in its assembly-disassembly. Notably, we have efficiently modified the original protocol that requires samples suitable for a 1 mL cuvette to the current 96-well plate format. The modified protocol has therefore significantly reduced the tissue/sample amount required for the assay. Further, we provide evidence that the protocol is suitable for not only brain tissue homogenates, but also subcellular fractions such as isolated synaptic terminals (synaptosomes and synaptoneurosomes). Lastly, the assay can be employed for freshly dissected rodent brain tissues and long-term stored post-mortem human brain samples. Of note, while the assay is presented in a neuronal context, it can be suitably extended to other cell-types and physiological processes associated with them.

<table><tbody><tr><td>3.5 mL, open-top thickwall polycarbonate tube</td><td>Beckman Coulter</td><td>349622</td><td>For gradient centrifugation (synaptosome prep)</td></tr><tr><td>Alexa Fluor 647 Phalloidin</td><td>Thermo Fisher Scientific</td><td>A22287</td><td>F-actin specific ligand</td></tr><tr><td>Antibody against &amp;nbsp;b-actin</td><td>Santa Cruz Biotechnology</td><td>Sc-47778</td><td>For evaluation of total actin levels by immunoblotting</td></tr><tr><td>Antibody against GAPDH</td><td>Abcam</td><td>Ab181602</td><td>For evaluation of GAPDH levels by immunoblotting</td></tr><tr><td>Bio-Rad Protein Assay Dye Reagent Concentrate</td><td>Bio-Rad</td><td>5000006</td><td>Bradford based protein estimation</td></tr><tr><td>Calcium chloride dihydrate (CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>C3306</td><td>Krebs buffer component</td></tr><tr><td>cOmplete, Mini, EDTA-free Protease Inhibitor Cocktail</td><td>Sigma-Aldrich</td><td>4693159001</td><td>For inhibition of endogenous protease activity during sample preparation</td></tr><tr><td>Corning 96-well Clear Flat Bottom Polystyrene</td><td>Corning</td><td>3596</td><td>For light-scattering measurements</td></tr><tr><td>D-(+)-Glucose</td><td>Sigma-Aldrich</td><td>G8270</td><td>Krebs buffer component</td></tr><tr><td>Dimethyl sulfoxide</td><td>Sigma-Aldrich</td><td>D5879</td><td>Solvent for phalloidin and latrunculin A</td></tr><tr><td>Fluorescent flatbed scanner (Odyssey Infrared Scanner)</td><td>Li-Cor Biosciences</td><td></td><td>For detection of immunoreactive signals on immunoblots</td></tr><tr><td>Glutaraldehyde solution (25% in water) Grade II</td><td>Sigma-Aldrich</td><td>G6257</td><td>Fixative</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375</td><td>Buffer ingredient for sample preparation and Krebs buffer component</td></tr><tr><td>Latrunculin A</td><td>Sigma-Aldrich</td><td>L5163</td><td>Depolymerizer of actin filaments</td></tr><tr><td>Magnesium chloride hexahydrate (MgCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>M2670</td><td>Krebs buffer component</td></tr><tr><td>Microplates</td><td></td><td></td><td></td></tr><tr><td>Mitex membrane filter 5 mm</td><td>Millipore</td><td>LSWP01300</td><td>Preparation of synaptoneurosomes</td></tr><tr><td>Nunc F96 MicroWell Black Plate</td><td>Thermo Fisher Scientific</td><td>237105</td><td>For fluorometric measurements</td></tr><tr><td>Nylon net filter 100 mm</td><td>Millipore</td><td>NY1H02500</td><td>Preparation of synaptoneurosomes</td></tr><tr><td>Phosphatase Inhibitor Cocktail IV</td><td>Abcam</td><td>ab201115</td><td>For inhibition of endogenous phosphatase activity during sample preparation</td></tr><tr><td>Potassium chloride (KCl)</td><td>Sigma-Aldrich</td><td>P9541</td><td>Krebs buffer component and for depolarization of synaptic terminals</td></tr><tr><td>Potassium phosphate monobasic ((KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>P9791</td><td>Krebs buffer component</td></tr><tr><td>Sodium borohydride (NaBH&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>71320</td><td>Component of Permeabilization buffer</td></tr><tr><td>Sodium chloride (NaCl)</td><td>LabServ (Thermo Fisher Scientific)</td><td>BSPSL944</td><td>Krebs buffer component</td></tr><tr><td>Sodium hydrogen carbonate (NaHCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>LabServ (Thermo Fisher Scientific)</td><td>BSPSL900</td><td>Krebs buffer component</td></tr><tr><td>SpectraMax i3x</td><td>Molecular Devices</td><td></td><td>For fluorometric measurements</td></tr><tr><td>Sucrose</td><td>Fisher Chemical</td><td>S/8600/60</td><td>Buffer ingredient for sample preparation</td></tr><tr><td>Swimnex Filter Holder</td><td>Millipore</td><td>Sx0001300</td><td>Preparation of synaptoneurosomes</td></tr><tr><td>Tissue grinder 5 mL Potter-Elvehjem</td><td>Duran Wheaton Kimble</td><td>358034</td><td>For tissue homogenization</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100</td><td>Component of Permeabilization buffer</td></tr><tr><td>Trizma base</td><td>Sigma-Aldrich</td><td>T6066</td><td>Buffer ingredient for sample preparation</td></tr></tbody></table>

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+.

Linearity of the assay for evaluation of F-actin levels 
First, a standard curve for the linear increase in fluorescence of Alexa Fluor 647 Phalloidin was ascertained and was repeated for each set of experiments (Figure 1). To investigate the linear range of the assay, different amounts of brain homogenates from rodents (Figures 2A and 2B) and post-mortem human subjects (Figure 3A and 3B) w...

Watch this Scientific Journal Video about A Time-Efficient Fluorescence Spectroscopy-Based Assay for Evaluating Actin Polymerization Status in Rodent and Human Brain Tissues at JoVE.com

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

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-time-efficient-fluorescence-spectroscopy-based-assay-for-evaluating

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

4.2K Views.  University of Otago. 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.

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

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

Biochemistry

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

Modified Roller Tube Method for Precisely Localized and Repetitive Intermittent Imaging During Long-term Culture of Brain Slices in an Enclosed System

Cultured rodent brain slices are useful for studying the cellular and molecular behavior of neurons and glia in an environment that maintains many of their normal in vivo interactions. Slices obtained from a variety of transgenic mouse lines or use of viral vectors for expression of fluorescently tagged proteins or reporters in wild type brain slices allow for high-resolution imaging by fluorescence microscopy. Although several methods have been developed for imaging brain slices, combining slice culture with the ability to perform repetitive high-resolution imaging of specific cells in live slices over long time periods has posed problems. This is especially true when viral vectors are used for expression of exogenous proteins since this is best done in a closed system to protect users and prevent cross contamination. Simple modifications made to the roller tube brain slice culture method that allow for repetitive high-resolution imaging of slices over many weeks in an enclosed system are reported. Culturing slices on photoetched coverslips permits the use of fiducial marks to rapidly and precisely reposition the stage to image the identical field over time before and after different treatments. Examples are shown for the use of this method combined with specific neuronal staining and expression to observe changes in hippocampal slice architecture, viral-mediated neuronal expression of fluorescent proteins, and the development of cofilin pathology, which was previously observed in the hippocampus of Alzheimer's disease (AD) in response to slice treatment with oligomers of amyloid-&#946; (A&#946;) peptide.

Presented here is a modified roller tube method for culturing and intermittent high-resolution imaging of rodent brain slices over many weeks with precise repositioning on photoetched coverslips. Neuronal viability and slice morphology are well maintained. Applications of this fully enclosed system using viruses for cell-type specific expression are provided.

Cultured rodent brain slices are useful for studying the cellular and molecular behavior of neurons and glia in an environment that maintains many of ...

Using Near-infrared Fluorescence and High-resolution Scanning to Measure Protein Expression in the Rodent Brain

Neuroscience is the study of how cells in the brain mediate various functions. Measuring protein expression in neurons and glia is critical for the study of neuroscience as cellular function is determined by the composition and activity of cellular proteins. In this article, we describe how immunocytochemistry can be combined with near-infrared high-resolution scanning to provide a semi-quantitative measure of protein expression in distinct brain regions. This technique can be used for single or double protein expression in the same brain region. Measuring proteins in this fashion can be used to obtain a relative change in protein expression with an experimental manipulation, molecular signature of learning and memory, activity in molecular pathways, and neural activity in multiple brain regions. Using the correct proteins and statistical analysis, functional connectivity among brain regions can be determined as well. Given the ease of implementing immunocytochemistry in a laboratory, using immunocytochemistry with near-infrared high-resolution scanning can expand the ability of the neuroscientist to examine neurobiological processes at a systems level.

Here, we present a protocol that uses near-infrared dyes in conjunction with immunohistochemistry and high-resolution scanning to assay proteins in brain regions.

Neuroscience is the study of how cells in the brain mediate various functions. Measuring protein expression in neurons and glia is critical for the study ...

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study of this movement in situ was only possible using radioisotopic pulse-labeling, which permits analysis of axonal transport in whole nerves with a temporal resolution of days and a spatial resolution of millimeters. To study neurofilament transport in situ with higher temporal and spatial resolution, we developed a hThy1-paGFP-NFM transgenic mouse that expresses neurofilament protein M tagged with photoactivatable GFP in neurons. Here we describe fluorescence photoactivation pulse-escape and pulse-spread methods to analyze neurofilament transport in single myelinated axons of tibial nerves from these mice ex vivo. Isolated nerve segments are maintained on the microscope stage by perfusion with oxygenated saline and imaged by spinning disk confocal fluorescence microscopy. Violet light is used to activate the fluorescence in a short axonal window. The fluorescence in the activated and flanking regions is analyzed over time, permitting the study of neurofilament transport with temporal and spatial resolution on the order of minutes&#160;and microns, respectively. Mathematical modeling can be used to extract kinetic parameters of neurofilament transport including the velocity, directional bias and pausing behavior from the resulting data. The pulse-escape and pulse-spread methods can also be adapted to visualize neurofilament transport in other nerves. With the development of additional transgenic mice, these methods could also be used to image and analyze the axonal transport of other cytoskeletal and cytosolic proteins in axons.

We describe fluorescence photoactivation methods to analyze the axonal transport of neurofilaments in single myelinated axons of peripheral nerves from transgenic mice that express a photoactivatable neurofilament protein.

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study ...

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

Analyses of Actin Dynamics, Clutch Coupling and Traction Force for Growth Cone Advance

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes depend on the advances of growth cones that located at the tips of neurites. Axonal growth cones generate driving forces by sensing their local microenvironment and modulating cytoskeletal dynamics and actin-adhesion coupling (clutch coupling). Decades of research have led to the identification of guidance molecules, their receptors, and downstream signaling cascades for regulating neuronal migration and axonal guidance; however, the molecular machineries required for generating forces to drive growth cone advance and navigation are just beginning to be elucidated. At the leading edge of neuronal growth cones, actin filaments undergo retrograde flow, which is powered by actin polymerization and actomyosin contraction. A clutch coupling between F-actin retrograde flow and adhesive substrate generates traction forces for growth cone advance. The present study describes a detailed protocol for monitoring F-actin retrograde flow by single speckle imaging. Importantly, when combined with an F-actin marker Lifeact, this technique can quantify 1) the F-actin polymerization rate and 2) the clutch coupling efficiency between F-actin retrograde flow and the adhesive substrate. Both are critical variables for generating forces for growth cone advance and navigation. In addition, the present study describes a detailed protocol of traction force microscopy, which can quantify 3) traction force generated by growth cones. Thus, by coupling the analyses of&#160;single speckle imaging and traction force microscopy, investigators can monitor the molecular mechanics underlying growth cone advance and navigation.

To advance, growth cones must exert traction forces against the external environment. The generation of traction forces is dependent on actin dynamics and clutch coupling. The present study describes methods for analyzing actin dynamics, clutch coupling and traction forces for growth cone advance.

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

Biology

Quantification of Filamentous Actin Puncta in Rat Cortical Neurons

Source: Li, H., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/53697">Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons.</a> J. Vis. Exp. (2016)
This video demonstrates the procedure for fluorescently staining rat cortical neurons for microscopy. It targets F-actin and dendritic proteins to analyze structural integrity and calculate F-actin density, which is crucial for studying neuronal function and health.

Kwantyfikacja nitkowatej aktyny punktowej w neuronach kory mózgowej szczura

1. Fluorescent Labeling and Immunocytochemistry
Note: The immunofluorescent labeling of primary cortical cell cultures was carried out in glass bottom 35 ...