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 most widely used instruments to detect actin filaments particularly for subsequent fluorescence microscopy-based analyses. Importantly, phalloidin has been shown to stain even loose, irregular meshwork of short actin filaments51, indicating that phalloidin binding is not dependent on the filament length. Our protocol, on the other hand, relies on fluorescence spectroscopy to analyze actin dynamics in ex vivo biological samples, for example brain tissues from rodents and humans.
The major advantage of the protocol is that it considerably reduces the time taken with respect to the existing protocols that first require a high-speed centrifugation-based biochemical isolation of F-actin (separation from G-actin based on insolubility of actin filaments in certain detergents such as Triton X-100) and subsequent analysis of immunoreactive levels using Western blotting6,28,29. The time-efficiency of our assay is also an advantage with regards to phalloidin-based immunocytochemistry techniques40,52, although there might be some other benefits associated with the latter. Another advantage is that it can be applied to cryopreserved post-mortem human brain tissues, procurement of which is always associated with some post-mortem delay. Further, with respect to the original protocol30 from which this methodology has been modified, we have considerably reduced the requirement for the tissue amount from 1 mL cuvette to a single well of a 96-well plate. Moreover, because of the inclusion of a phalloidin standard curve in each set of experiment, our protocol can quantitate absolute levels of actin filaments in units of phalloidin bound (Figures 2A-B, 3A, 4A-B; see also Supplementary Figure 2), as well as the levels relative to control samples (Figure 5A-B).
It should be noted that the application of phalloidin to assess F-actin levels however is restricted to fixed cells and samples, both for our assay protocol as well as microscopy-based protocols. This is because phalloidin is essentially a toxic bicyclic heptapeptide that binds specifically at the interface between F-actin subunits with high affinity and stabilizes these actin filaments, rendering them incapable to depolymerize and in fact increasing the net conversion of G-actin into F-actin40,53,54. Hence, phalloidin stabilizes actin filaments in vivo and in vitro, and can result in significant changes in the equilibrium status of actin per se47,55. As such evaluation of actin polymerization status mediated by fluorescent phalloidin is based upon arrested filament structures. Moreover, because of its low permeability through the lipid bilayer, phalloidin-based methodologies rely on permeabilization of the cells or biological samples. Infeasibility of a time course live-cell assay is hence a major limitation of the protocol as with immunocytochemistry-based methods employing phalloidin.
Infeasibility of phalloidin-based methods to evaluate dynamic changes in actin filaments in live unfixed cells begs the question of whether there are alternative procedures to do so. Indeed, advances have been made in this regard with exogenous expression of fluorescent-actin analogs prior to analysis using protocols such as video micrography36,56, fluorescence recovering after photobleaching (FRAP)38 or fluorescence resonance energy transfer (FRET)39. Heterologous expression of fluorescently tagged peptides and proteins that bind actin filaments are also employed to study actin dynamics in live cells; however there are disadvantages and limitations associated with them as well as with the heterologous expression of fluorescently tagged actin47,49,57. For example, G-actin that comprises 50-70% of total actin in most cells is soluble and freely diffusible in the cytosol, resulting in a higher background causing challenges in differentiating signals from actin filaments specifically57.
A critical factor in the assay is that as shown in Figure 2 and Figure 3; it is not linear throughout the range of protein amounts tested. Hence, an optimal amount of protein suitable for the assay should be determined first or the amount of phalloidin analog should be adjusted such that it is no longer limiting (at higher amounts of proteins). Another critical aspect of the protocol is that the multiple centrifugation-based steps for removal of fixative, permeabilization agent and unbound phalloidin can lead to varying loss of proteins (and F-actin bound phalloidin) from the samples, particularly when lower amounts of samples are used. Hence it is important to normalize the amount of sample retained by monitoring light scattering at 540 nm. Lastly, since actin is in a dynamic state of interconversion between its F- and G-forms, fixing should be fast. A minor related critical aspect of the assay is that we could not evaluate its efficiency in assessing pharmacological actin polymerization. As opposed to pharmacological actin depolymerization by latrunculin A, jasplakinolide (a reliable and widely used actin polymerizing agent) has overlapping binding sites with phalloidin and competitively inhibits its binding to actin filaments58,59. Nevertheless, employment of KCl-stimulated synaptic terminals as an ex vivo model for increased actin polymerization indicates that our assay can also detect increases in F-actin levels.
In conclusion, we describe a robust time-efficient and high-throughput assay for analysis of actin filaments (F-actin) and its alternations in physiological and pathophysiological states suitable for a 96-well plate format. In combination with other existing methods for evaluation of F-actin in fixed and unfixed samples, the protocol will prove to be an essential tool in actin-related studies in the neuroscience field, as well as other areas of biological science research.

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) were processed. The assay was found to be linear in the range of 50-200 &#956;g of protein as assessed by amounts of labeled phalloidin retained. Light scattering at 540 nm was used to confirm the different amounts of samples (Figure 2C and Figure 3C).
Latrunculin, an actin depolymerizing agent reduces binding of labelled phalloidin 
Latrunculin A is known to depolymerize actin filaments and reduce the levels of F-actin36,37,38,39. Homogenates from either rodent or human brain tissues were incubated with 2 &#956;M latrunculin A for 1 hour at 37 &#176;C to depolymerize actin filaments. Respective untreated control sets were incubated with DMSO for the same duration of 1 hour at 37 &#176;C. The assay robustly measured the loss of F-actin levels from 95.7&#160;&#177; 6.6 (mean&#160;&#177; SEM) in control samples to 72.0&#160;&#177; 3.2 (mean&#160;&#177; SEM) &#956;U of bound phalloidin in latrunculin A-treated samples in rodent brain homogenates (Figure 4A). A similar decrease (from 83.7&#160;&#177; 3.9 to 66.9&#160;&#177; 4.2 &#956;U) in retention of labeled phalloidin was also observed when homogenates from human brain tissues were subjected to latrunculin A treatment (Figure 4B). Noteworthy, total actin levels, as assessed by immunoblotting, did not alter upon treatment with latrunculin A in both rat (Supplementary Figure 1A-B) and human (Supplementary Figure 1C-D) brain homogenates.
Depolarization of isolated synaptic terminalsstimulates actin polymerization and filament formation 
Ex vivo depolarization of isolated synaptic terminals has been shown to result in rapid stimulation of actin polymerization6,30,40, and this phenomenon was used as a further confirmation of the assay reported herein. Biochemical fractions enriched in synaptic terminals were prepared in two different manners; a gradient-based ultracentrifugation method to obtain &#34;synaptosomes&#34;41,42,43,44 and a sequential filtration-based protocol to obtain &#34;synaptoneurosomes&#34;43,45,46. Because the yield for the latter is higher, we used it for human post-mortem brain tissues wherein the tissue amounts are often limiting. On the other hand, we preferred to use synaptosomes with a higher degree of enrichment of synaptic fragments41 for our rat brain tissue experiments.
Depolarization of synaptosomes or synaptoneurosomes and stimulation of actin polymerization were achieved by a short (30 second) burst of increase in extracellular K+ to 50 mM. KCl exposure resulted in increased phalloidin binding by almost 40% in rodent brain synaptosomes compared to the respective mock-stimulated controls (Figure 5A; see also Supplementary Figure 2A). A smaller (around 20%) but consistent increase was also observed in human synaptoneurosomes treated with KCl (Figure 5B; see also Supplementary Figure 2A). These experiments validate the robustness of our assay in determining alterations in F-actin levels in brain tissue samples, including isolated synaptic terminals.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62268/62268fig01.jpg" /> 
Figure 1. Standard curve for fluorescence of Alexa Fluor 647 Phalloidin.&#160; Linearity of fluorescence emission of different amounts (25-500 &#956;U) of labeled phalloidin was confirmed by fluorescence spectroscopy at an excitation of 645 nm and emission of 670 nm (R2 = 0.9942). Data are represented as mean&#160;&#177; SEM (n=3).&#160;<a href="https://www.jove.com/files/ftp_upload/62268/62268fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62268/62268fig02.jpg" /> 
Figure 2. Linearity of phalloidin binding to whole-cell homogenates from rat brain tissue.&#160;&#160;(A)&#160;Binding of phalloidin to different amounts of homogenates (50-300 &#956;g protein) was assessed. (B) Binding was linear in the range of 50-200 &#956;g protein (R2 = 0.9602). (C)&#160;Scattering at 540 nm was used to confirm different amounts of the samples (R2 = 0.8319). Data are represented as mean&#160;&#177; SEM (n=4).&#160;<a href="https://www.jove.com/files/ftp_upload/62268/62268fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62268/62268fig03.jpg" /> 
Figure 3. Phalloidin binding to whole-cell homogenates from post-mortem human brain tissue.&#160;&#160;(A)&#160;Phalloidin retention in a range of amounts of homogenates (50-300 &#956;g protein) was evaluated. (B) Phalloidin binding was found to be linear in the range of 50-200 &#956;g protein (R2 = 0.8832). (C)&#160;Scattering at 540 nm confirmed the varying amounts of the samples (R2 = 0.9730). Data are represented as mean &#177; SEM (n=4).&#160;<a href="https://www.jove.com/files/ftp_upload/62268/62268fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62268/62268fig04.jpg" /> 
Figure 4. Effects of latrunculin A on F-actin levels.&#160;&#160;Treatment of brain homogenates with actin depolymerizing agent Latrunculin A (2 &#956;M, 1 h at 37&#176;C) resulted in significant decrease in the amounts of actin filaments compared to the respective mock-treated control samples as assessed by retention of labeled phalloidin both in rodent (p = 0.0034; paired two-tailed Student&#39;s t-test) (A) and post-mortem human tissues (p = 0.0011; paired two-tailed Student&#39;s t-test) (B). Data are represented as mean&#160;&#177; SEM (n=6 pairs).&#160;<a href="https://www.jove.com/files/ftp_upload/62268/62268fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62268/62268fig05.jpg" /> 
Figure 5. Effects of KCl-mediated depolarization on F-actin amounts in isolated synaptic terminals.&#160; (A) Incubation of synaptosomes from rat brain with 50 mM KCl for 30 s at 37&#176;C stimulated actin polymerization which consequently resulted in an increase in phalloidin binding (p = 0.0014; paired two-tailed Student&#39;s t-test). (B) A smaller increase was also observed in synaptoneurosomal fraction from post-mortem human brain tissues (p = 0.014; paired two-tailed Student&#39;s t-test). Data are represented as mean&#160;&#177; SEM (n=6 pairs).&#160;<a href="https://www.jove.com/files/ftp_upload/62268/62268fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1. Effects of latrunculin A on total actin levels.&#160;&#160;Brain homogenates were incubated with latrunculin A (2 &#956;M, 1 h at 37&#176;C) or equal volume of DMSO (1 h at 37&#176;C). 10 &#956;g protein per sample (latrunculin A treated and DMSO mock-treated controls) were collected prior to fixation. Total actin levels were assessed by immunoblotting. Representative blots are shown for rat (A) and human (C) brain homogenates. Latrunculin A did not alter the total actin levels in both rat (B; p = 0.40; paired two-tailed Student&#39;s t-test) and human (D; p = 0.42; paired two-tailed Student&#39;s t-test) brain homogenates. Data are represented as mean&#160;&#177; SEM (n=3 pairs).&#160;<a href="https://www.jove.com/files/ftp_upload/62268/Supplementary Fig.1.tif" target="_blank">Please click here to download this File.</a>
Supplementary Figure 2. Effects of KCl-mediated depolarization on F-actin levels in rat synaptosomes and human synaptoneurosomes.&#160;&#160;(A) Incubation of synaptosomes from rat brain with 50 mM KCl (30 s at 37 &#176;C) stimulated actin polymerization and a consequent increase in phalloidin binding (p = 0.0019; paired two-tailed Student&#39;s t-test). (B) Increase in phalloidin retention was also observed in human synaptoneurosomal fraction depolarized by KCl compared to the respective unstimulated controls (p = 0.015; paired two-tailed Student&#39;s t-test). Data are represented as mean&#160;&#177; SEM (n=6 pairs).&#160;<a href="https://www.jove.com/files/ftp_upload/62268/Supplementary Fig.2.tif" target="_blank">Please click here to download this File.</a>

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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 San Sebastian

Research

JoVE Journal

Neuroscience

4.1K 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

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 ...

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 ...

Collection of Frozen Rodent Brain Regions for Downstream Analyses

As our understanding of neurobiology has progressed, molecular analyses are often performed on small brain areas such as the medial prefrontal cortex (mPFC) or nucleus accumbens. The challenge in this work is to dissect the correct area while preserving the microenvironment to be examined. In this paper, we describe a simple, low-cost method using resources readily available in most labs. This method preserves nucleic acid and proteins by keeping the tissue frozen throughout the process. Brains are cut into 0.5&#8211;1.0 mm sections using a brain matrix and arranged on a frozen glass plate. Landmarks within each section are compared to a reference, such as the Allen Mouse Brain Atlas, and regions are dissected using a cold scalpel or biopsy punch. Tissue is then stored at -80 &#176;C until use. Through this process rat and mouse mPFC, nucleus accumbens, dorsal and ventral hippocampus and other regions have been successfully analyzed using qRT-PCR and Western assays. This method is limited to brain regions that can be identified by clear landmarks.

This procedure describes the collection of discrete frozen brain regions to obtain high-quality protein and RNA using inexpensive and commonly available tools.

As our understanding of neurobiology has progressed, molecular analyses are often performed on small brain areas such as the medial prefrontal cortex ...

Determining the Functional Status of the Corticospinal Tract Within One Week of Stroke

High interindividual variability in the recovery of upper limb (UL) function after stroke means it is difficult to predict an individual&#39;s potential for recovery based on clinical assessments alone. The functional integrity of the corticospinal tract is an important prognostic biomarker for recovery of UL function, particularly for those with severe initial UL impairment. This article presents a protocol for evaluating corticospinal tract function within 1 week of stroke. This protocol can be used to select and stratify patients in trials of interventions designed to improve UL motor recovery and outcomes after stroke. The protocol also forms part of the PREP2 algorithm, which predicts UL function for individual patients 3 months poststroke. The algorithm sequentially combines a UL strength assessment, age, transcranial magnetic stimulation, and stroke severity, within a few days of the stroke. The benefits of using PREP2 in clinical practice are described elsewhere. This article focuses on the use of a UL strength assessment and transcranial magnetic stimulation to evaluate corticospinal tract function.

This protocol is for evaluating corticospinal tract function within 1 week of stroke. It can be used to select and stratify patients in trials of interventions designed to improve upper limb motor recovery and outcomes and in clinical practice for predicting upper limb functional outcomes 3 months after stroke.

High interindividual variability in the recovery of upper limb (UL) function after stroke means it is difficult to predict an individual&#39;s potential ...

Flow Cytometric Analysis of Multiple Mitochondrial Parameters in Human Induced Pluripotent Stem Cells and Their Neural and Glial Derivatives

Mitochondria are important in the pathophysiology of many neurodegenerative diseases. Changes in mitochondrial volume, mitochondrial membrane potential (MMP), mitochondrial production of reactive oxygen species (ROS), and mitochondrial DNA (mtDNA) copy number are often features of these processes. This report details a novel flow cytometry-based approach to measure multiple mitochondrial parameters in different cell types, including human induced pluripotent stem cells (iPSCs) and iPSC-derived neural and glial cells. This flow-based strategy uses live cells to measure mitochondrial volume, MMP, and ROS levels, as well as fixed cells to estimate components of the mitochondrial respiratory chain (MRC) and mtDNA-associated proteins such as mitochondrial transcription factor A (TFAM). 
By co-staining with fluorescent reporters, including MitoTracker Green (MTG), tetramethylrhodamine ethyl ester (TMRE), and MitoSox Red, changes in mitochondrial volume, MMP, and mitochondrial ROS can be quantified and related to mitochondrial content. Double staining with antibodies against MRC complex subunits and translocase of outer mitochondrial membrane 20 (TOMM20) permits the assessment of MRC subunit expression. As the amount of TFAM is proportional to mtDNA copy number, the measurement of TFAM per TOMM20 gives an indirect measurement of mtDNA per mitochondrial volume. The entire protocol can be carried out within 2-3 h. Importantly, these protocols allow the measurement of mitochondrial parameters, both at the total level and the specific level per mitochondrial volume, using flow cytometry.

This study reports a novel approach to measure multiple mitochondrial functional parameters based on flow cytometry and double staining with two fluorescent reporters or antibodies to detect changes in mitochondrial volume, mitochondrial membrane potential, reactive oxygen species level, mitochondrial respiratory chain composition, and mitochondrial DNA.

Mitochondria are important in the pathophysiology of many neurodegenerative diseases. Changes in mitochondrial volume, mitochondrial membrane potential ...