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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#160;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 (CaCl2&#183;2H2O)</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 (MgCl2&#183;6H2O)</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 ((KH2PO4)</td>
			<td>Sigma-Aldrich</td>
			<td>P9791</td>
			<td>Krebs buffer component</td>
		</tr>
		<tr>
			<td>Sodium borohydride (NaBH4)</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 (NaHCO3)</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

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

Simultaneous fMRI and Electrophysiology in the Rodent Brain

To examine the neural basis of the blood oxygenation level dependent (BOLD) magnetic resonance imaging (MRI) signal, we have developed a rodent model in which functional MRI data and in vivo intracortical recording can be performed simultaneously. The combination of MRI and electrical recording is technically challenging because the electrodes used for recording distort the MRI images and the MRI acquisition induces noise in the electrical recording. To minimize the mutual interference of the two modalities, glass microelectrodes were used rather than metal and a noise removal algorithm was implemented for the electrophysiology data. In our studies, two microelectrodes were separately implanted in bilateral primary somatosensory cortices (SI) of the rat and fixed in place. One coronal slice covering the electrode tips was selected for functional MRI. Electrode shafts and fixation positions were not included in the image slice to avoid imaging artifacts. The removed scalp was replaced with toothpaste to reduce susceptibility mismatch and prevent Gibbs ringing artifacts in the images. The artifact structure induced in the electrical recordings by the rapidly-switching magnetic fields during image acquisition was characterized by averaging all cycles of scans for each run. The noise structure during imaging was then subtracted from original recordings. The denoised time courses were then used for further analysis in combination with the fMRI data. As an example, the simultaneous acquisition was used to determine the relationship between spontaneous fMRI BOLD signals and band-limited intracortical electrical activity. Simultaneous fMRI and electrophysiological recording in the rodent will provide a platform for many exciting applications in neuroscience in addition to elucidating the relationship between the fMRI BOLD signal and neuronal activity.

We have developed a method for simultaneous functional magnetic resonance imaging and electrophysiological recording in the rodent brain, providing a platform for the investigation of the relationship between neural activity and the blood oxygenation level dependent (BOLD) MRI signal.

To examine the neural basis of the blood oxygenation level dependent (BOLD) magnetic resonance imaging (MRI) signal, we have developed a rodent model in ...

DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; Gan et al., 2000). Here we provide a detailed description of each step required together with exemplary images of good and bad outcomes that will help when setting up the technique. In our experience, a few steps proved critical for the successful application of DiOLISTICS. These considerations include the quality of the DiI-coated bullets, the extent of fixative exposure, and the concentration of detergent used in the incubation solutions. Tips and solutions for common problems are provided. This is a versatile labeling technique that can be applied to multiple animal species at a wide range of ages. Unlike other fluorescent labeling techniques that are limited to preparations from young animals or restricted to mice because they rely on the expression of a fluorescent transgene, DiOLISTIC labeling can be applied to animals of all ages, species and genotypes and it can be used in combination with immunostaining to identify a specific subpopulation of cells. Here we demonstrate the use of DiOLISTICS to label neurons in brain slices from adult mice and adult non-human primates with the purpose of quantifying dendrite branching and dendritic spine morphology.

We demonstrate the use of the gene gun to introduce fluorescent dyes, such as DiI, into neurons in brain slices from rodents and non-human primates of different ages. In this particular case, we use adult mice (3-6 months old) and adult cynomologus monkeys (9-15 years old). This technique, originally described by the laboratory of Dr. Lichtman (Gan et al., 2000), is well suited for the study of dendritic branching and dendritic spine morphology and can be combined with traditional immunostaining, if detergents are kept at a low concentration.

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; ...

Double Fluorescence in situ Hybridization in Fresh Brain Sections

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh frozen brain sections. Our group has successfully used this approach to study gene co-regulation. More specifically, we have used this dFISH method to explore the anatomical organization, neurochemical properties, and the impact of sensory experience in central sensory circuits, at single cell resolution. This protocol has been validated in brain tissue from mice, rats and songbirds but is expected to be easily adaptable to other vertebrate species, as well as to an array of non-neural tissues. In this film we provide a detailed demonstration of the main steps of this procedure.

Double Fluorescence in situ Hybridization in Fresh Brain Sections

This protocol involves a non-radioactive in-situ hybridization procedure that enables the simultaneous identification of two transcript species, at a single cell resolution, in thin sections of the vertebrate brain.

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh ...

A Rapid Approach to High-Resolution Fluorescence Imaging in Semi-Thick Brain Slices

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are characteristic to neurons in both normal and diseased brain. Towards this, a great deal of effort has been placed on building high-resolution neuroanatomical maps1-3. With the expansion of molecular genetics and advances in light microscopy has come the ability to query not only neuronal morphologies, but also the molecular and cellular makeup of individual neurons and their associated networks4. Major advances in the ability to mark and manipulate neurons through transgenic and gene targeting technologies in the rodent now allow investigators to 'program' neuronal subsets at will5-6. Arguably, one of the most influential contributions to contemporary neuroscience has been the discovery and cloning of genes encoding fluorescent proteins (FPs) in marine invertebrates7-8, alongside their subsequent engineering to yield an ever-expanding toolbox of vital reporters9. Exploiting cell type-specific promoter activity to drive targeted FP expression in discrete neuronal populations now affords neuroanatomical investigation with genetic precision.
Engineering FP expression in neurons has vastly improved our understanding of brain structure and function. However, imaging individual neurons and their associated networks in deep brain tissues, or in three dimensions, has remained a challenge. Due to high lipid content, nervous tissue is rather opaque and exhibits auto fluorescence. These inherent biophysical properties make it difficult to visualize and image fluorescently labelled neurons at high resolution using standard epifluorescent or confocal microscopy beyond depths of tens of microns. To circumvent this challenge investigators often employ serial thin-section imaging and reconstruction methods10, or 2-photon laser scanning microscopy11. Current drawbacks to these approaches are the associated labor-intensive tissue preparation, or cost-prohibitive instrumentation respectively.
Here, we present a relatively rapid and simple method to visualize fluorescently labelled cells in fixed semi-thick mouse brain slices by optical clearing and imaging. In the attached protocol we describe the methods of: 1) fixing brain tissue in situ via intracardial perfusion, 2) dissection and removal of whole brain, 3) stationary brain embedding in agarose, 4) precision semi-thick slice preparation using new vibratome instrumentation, 5) clearing brain tissue through a glycerol gradient, and 6) mounting on glass slides for light microscopy and z-stack reconstruction (Figure 1).
For preparing brain slices we implemented a relatively new piece of instrumentation called the 'Compresstome' VF-200 (http://www.precisionary.com/products_vf200.html). This instrument is a semi-automated microtome equipped with a motorized advance and blade vibration system with features similar in function to other vibratomes. Unlike other vibratomes, the tissue to be sliced is mounted in an agarose plug within a stainless steel cylinder. The tissue is extruded at desired thicknesses from the cylinder, and cut by the forward advancing vibrating blade. The agarose plug/cylinder system allows for reproducible tissue mounting, alignment, and precision cutting. In our hands, the 'Compresstome' yields high quality tissue slices for electrophysiology, immunohistochemistry, and direct fixed-tissue mounting and imaging. Combined with optical clearing, here we demonstrate the preparation of semi-thick fixed brain slices for high-resolution fluorescent imaging.

Here we describe a rapid and simple method to image fluorescently labeled cells in semi-thick brain slices. By fixing, slicing, and optically clearing brain tissue we describe how standard epifluorescent or confocal imaging can be used to visualize individual cells and neuronal networks within intact nervous tissue.

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are ...

A General Method for Evaluating Incubation of Sucrose Craving in Rats

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and relapse to food taking 1. Food craving conceptualized in this way is akin to drug craving in response to drug-paired cues. A rich literature has been developed around understanding the behavioral and neurobiological determinants of drug craving; we and others have been focusing recently on translating techniques from basic addiction research to better understand addiction-like behaviors related to food 2-4.
As done in previous studies of drug craving, we examine sucrose craving behavior by utilizing a rat model of relapse. In this model, rats self-administer either drug or food in sessions over several days. In a session, lever responding delivers the reward along with a tone+light stimulus. Craving behavior is then operationally defined as responding in a subsequent session where the reward is not available. Rats will reliably respond for the tone+light stimulus, likely due to its acquired conditioned reinforcing properties 5. This behavior is sometimes referred to as sucrose seeking or cue reactivity. In the present discussion we will use the term "sucrose craving" to subsume both of these constructs.
In the past decade, we have focused on how the length of time following reward self-administration influences reward craving. Interestingly, rats increase responding for the reward-paired cue over the course of several weeks of a period of forced-abstinence. This "incubation of craving" is observed in rats that have self-administered either food or drugs of abuse 4,6. This time-dependent increase in craving we have identified in the animal model may have great potential relevance to human drug and food addiction behaviors.
Here we present a protocol for assessing incubation of sucrose craving in rats. Variants of the procedure will be indicated where craving is assessed as responding for a discrete sucrose-paired cue following extinction of lever pressing within the sucrose self-administration context (Extinction without cues) or as responding for sucrose-paired cues in a general extinction context (Extinction with cues).

Responding for food or drug-paired cues increases over the course of a period of abstinence and this may relate to an increased susceptibility to relapse behaviors. Here we detail a procedure for evaluating this "incubation of craving" in rats that have self-administered sucrose.

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Personalized Needles for Microinjections in the Rodent Brain

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to deliver gene or cell therapy products. Unfortunately, current microinjection techniques use steel or glass needles that are suboptimal for multiple reasons: in particular, steel needles may cause tissue damage, and glass needles may bend when lowered deeply into the brain, missing the target region. In this article, we describe a protocol to prepare and use quartz needles that combine a number of useful features. These needles do not produce detectable tissue damage and, being very rigid, ensure reliable delivery in the desired brain region even when using deep coordinates. Moreover, it is possible to personalize the design of the needle by making multiple holes of the desired diameter. Multiple holes facilitate the injection of large amounts of solution within a larger area, whereas large holes facilitate the injection of cells. In addition, these quartz needles can be cleaned and re-used, such that the procedure becomes cost-effective.

We describe here a protocol for microinjection in the rodent brain that uses quartz needles. These needles do not produce detectable tissue damage and ensure reliable delivery even in deep regions. Moreover, they can be adapted to research needs by personalized designs and can be re-used.

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to ...

High-throughput Analysis of Locomotor Behavior in the Drosophila Island Assay

Advances in next-generation sequencing technologies contribute to the identification of (candidate) disease genes for movement disorders and other neurological diseases at an increasing speed. However, little is known about the molecular mechanisms that underlie these disorders. The genetic, molecular, and behavioral toolbox of Drosophila melanogaster makes this model organism particularly useful to characterize new disease genes and mechanisms in a high-throughput manner. Nevertheless, high-throughput screens require efficient and reliable assays that, ideally, are cost-effective and allow for the automatized quantification of traits relevant to these disorders. The island assay is a cost-effective and easily set-up method to evaluate Drosophila locomotor behavior. In this assay, flies are thrown onto a platform from a fixed height. This induces an innate motor response that enables the flies to escape from the platform within seconds. At present, quantitative analyses of filmed island assays are done manually, which is a laborious undertaking, particularly when performing large screens.
This manuscript describes the &#34;Drosophila Island Assay&#34; and &#34;Island Assay Analysis&#34; algorithms for&#160;high-throughput, automated data processing and quantification of island assay data. In the setup, a simple webcam connected to a laptop collects an image series of the platform while the assay is performed. The &#34;Drosophila Island Assay&#34; algorithm developed for the open-source software Fiji processes these image series and quantifies, for each experimental condition, the number of flies on the platform over time. The &#34;Island Assay Analysis&#34; script, compatible with the free software R, was developed to automatically process the obtained data and to calculate whether treatments/genotypes are statistically different. This greatly improves the efficiency of the island assay and makes it a powerful readout for basic locomotion and flight behavior. It can thus be applied to large screens investigating fly locomotor ability, Drosophila models of movement disorders, and drug efficacy.

High-throughput Analysis of Locomotor Behavior in the Drosophila Island Assay

The island assay is a relatively new, cost-effective assay that can be used to evaluate the basic locomotor behavior of Drosophila melanogaster. This manuscript describes algorithms for automatic data processing and objective quantification of island assay data, making this assay a sensitive, high-throughput readout for large genetic or pharmacological screens.

Advances in next-generation sequencing technologies contribute to the identification of (candidate) disease genes for movement disorders and other ...

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their locomotory patterns upon detection of diverse environmental stimuli and can modulate their suite of behavioral responses in response to starvation conditions. Nematodes typically exhibit a decreased aversive response when removed from a food source for over 30 min. Observation of behavioral changes in response to a changing nutritional status can provide insight into the mechanisms that regulate the transition from a well-fed to starved state.
We have developed an assay that measures a nematode&#39;s ability to cross an aversive barrier (i.e. copper) then reach a food source over a prolonged period of time. This protocol builds upon previous work by integrating multiple variables in a manner that allows for continued data collection as the organisms shift towards an increasingly starved condition. Moreover, this assay permits an increased sample size so that larger populations of nematodes can be simultaneously evaluated.
Organisms defective for the ability to detect or respond to copper immediately cross the chemical barrier, while wild type nematodes are initially repelled. As wild type worms are increasingly starved, they begin to cross the barrier and reach the food source. We designed this assay to evaluate a mutant that is incapable of responding to diverse environmental cues, including food sensation or detection of aversive chemicals. When evaluated via this protocol, the defective organisms immediately crossed the barrier, but were also incapable of detecting a food source. Hence, these mutants repeatedly cross the chemical barrier despite temporarily reaching a food source. This assay can straightforwardly test populations of worms to evaluate potential pathway defects related to aversion and starvation.

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

Here, we present a Caenorhabditis elegans-specific assay designed to evaluate changes in copper aversion behavior and the ability to locate a common food source, as the organism progresses from a well-fed to starved nutritional state.

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their ...

Effects of Blast-induced Neurotrauma on Pressurized Rodent Middle Cerebral Arteries

Though there have been studies on the histopathological and behavioral effects of blast exposure, fewer have been dedicated to blast&#39;s cerebral vascular effects. Impact (i.e., non-blast) traumatic brain injury (TBI) is known to decrease pressure autoregulation in the cerebral vasculature in both humans and experimental animals. The hypothesis that blast-induced traumatic brain injury (bTBI), like impact TBI, results in impaired cerebral vascular reactivity was tested by measuring myogenic dilatory responses to reduced intravascular pressure in rodent middle cerebral arterial (MCA) segments from rats subjected to mild bTBI using an Advanced Blast Simulator (ABS) shock tube. Adult, male Sprague-Dawley rats were anesthetized, intubated, ventilated and prepared for Sham bTBI (identical manipulation and anesthesia except for blast injury) or mild bTBI. Rats were randomly assigned to receive Sham bTBI or mild bTBI followed by sacrifice 30 or 60 min post-injury. Immediately after bTBI, righting reflex (RR) suppression times were assessed, euthanasia at the time points post-injury was completed, the brain was harvested and the individual MCA segments were collected, mounted and pressurized. As the intraluminal pressure perfused through the arterial segments was reduced in 20 mmHg increments from 100 to 20 mmHg, MCA diameters were measured and recorded. With decreasing intraluminal pressure, MCA diameters steadily increased significantly above baseline in the Sham bTBI groups while MCA dilator responses were significantly reduced (p &#60; 0.05) in both bTBI groups as evidenced by the impaired, smaller MCA diameters recorded for the bTBI groups. In addition, RR suppression in the bTBI groups was significantly (p &#60; 0.05) higher than in the Sham bTBI groups. MCA&#39;s collected from the Sham bTBI groups exhibited typical vasodilatory properties to decreases in intraluminal pressure while MCA&#39;s collected following bTBI exhibited significantly impaired myogenic vasodilatory responses to reduced pressure that persisted for at least 60 min after bTBI.

Here, we present a protocol to describe methods for ex vivo vascular reactivity determination following a primary blast traumatic brain injury (bTBI) using isolated, pressurized, rodent middle cerebral arterial (MCA) segments. bTBI induction is accomplished using a shock tube, also known as an Advanced Blast Simulator (ABS) device.

Though there have been studies on the histopathological and behavioral effects of blast exposure, fewer have been dedicated to blast&#39;s cerebral ...