We would like to thank P. Colosi for EM image preparation. This work was supported by the National Institutes of Health (R01 NS064963, SSC; R01 NS110354, SSC; R01 NS083846, SSC; R21 NS094971, SSC; T32 NS007224, SMT; T32 NS041228, SMT), the United States Department of Defense (W81XWH-17-1-0564, SSC; W81XWH-19-1-0264, VDJ), Aligning Science Across Parkinson&#39;s (ASAP) Collaborative Research Network (SSC), and the Michael J. Fox Foundation Target Advancement Program (MJFF-020160, SSC &#38; VDJ). We created graphical illustrations using BioRender.com.

All experiments with mice were approved by the Institutional Animal Care and Use Committee (IACUC) at Yale University (Protocol 2021-11117) and performed in a facility accredited by the Association for the Assessment and Accreditation of Laboratory Animal Care International (AAALAC). Animal care and housing complied with the Guide for the Care and Use of Laboratory Animals33 and were provided by the Yale Animal Resource Center (YARC). Animals were maintained in a 12 h light/dark cycle with ad libitum access to food and water. Five to eight mice or two to four rats per genotype or condition are required for the following protocol. Fewer rats are necessary due to their larger brain volumes. Similarly, the age of the experimental animals may affect fraction yield; additional mice may be required for ages less than 2 months. Otherwise, the outlined procedures apply to both murine species and healthy adult animals of any age. The representative data presented in this study utilized wild-type (C57BL/6J) mice (age = 2 months; four males and four females per replicate) obtained from a commercial source (see Table of Materials).
1. Experimental preparation
NOTE: This protocol requires ~11 h for a single researcher to complete. It is highly recommended to complete benchtop setup (Figure 1), buffer preparation (Table 1), the precooling of centrifuges and rotors to 4 &#176;C, and the collection and labeling of necessary materials and equipment (see Table of Materials) the day prior to protocol execution, where applicable.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64574/64574fig01.jpg" /> 
Figure 1: Benchtop setup. Prior to brain dissections,&#160;(A) Dounce glass homogenizers and (B) all buffers were chilled on ice. (C) Protease inhibitor stock solutions were thawed on ice.&#160;A second container of wet ice for centrifuge tubes, a Dewar of liquid nitrogen (not shown), and&#160;(D) a container of dry ice for short-term storage of the samples flash-frozen in liquid nitrogen were obtained.&#160;(E) Microcentrifuge tubes were pre-labeled for all samples, as four aliquots of each subcellular fraction sample per genotype or condition were collected during this procedure (time-saving tip: thoroughly label all the tubes the day before the experiment is performed). (F)&#160;An appropriate biohazard waste container, (G)&#160;70% ethanol, (H)&#160;surgical tools, and (I)&#160;an absorbent surface pad. The required centrifuge tubes and disposables were set aside for efficient access during protocol implementation (not shown). <a href="https://www.jove.com/files/ftp_upload/64574/64574fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Prepare the benchtop for surgery and collect the scissors and forceps required for brain excision (see Table of Materials). Pre-label 1.5 mL microcentrifuge tubes for mouse tail biopsies and four tubes per collected fraction, as outlined in Figure 2.</li>
	<li>Obtain two containers of wet ice, one container of dry ice, and a benchtop liquid nitrogen Dewar flask.</li>
	<li>Thaw phenylmethylsulfonyl fluoride (PMSF), pepstatin A, aprotinin, and leupeptin stock solutions on ice (see Table of Materials). Prepare the necessary buffers (Table 1). 
	NOTE: Sucrose solutions can be prepared in advance and stored at 4 &#176;C. However, protease inhibitors (thawed stocks and tablets) must be added fresh to all the buffers at the start of the experiment due to the instability of these reagents in aqueous solutions. Further, all the buffers must be prepared with detergent-free glassware and detergent-free water to enable the collection of intact synaptosomes.</li>
	<li>Chill all the buffers and glass Dounce homogenizers (see Table of Materials) on ice. Set the centrifuges to 4 &#176;C and chill the rotors to 4 &#176;C.</li>
	<li>Add 14 mL of Buffer A (Table 1) to a Dounce homogenizer on ice.</li>
</ol>
Table 1: Composition of the subcellular fractionation buffers. <a href="https://www.jove.com/files/ftp_upload/64574/64574_Table 1_revised.xlsx" target="_blank">Please click here to download this Table.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64574/64574fig02.jpg" /> 
Figure 2: Overview of the subcellular fractionation protocol. Summary schematic of the subcellular fractionation steps and collected samples. <a href="https://www.jove.com/files/ftp_upload/64574/64574fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Mouse brain excision
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64574/64574fig03.jpg" /> 
Figure 3: Craniofacial anatomy. (A)&#160;Dorsal view of a mouse skull with relevant cranial structures indicated. (B) Left lateral view of a mouse skull and brain with relevant cranial structures and anatomical directions indicated. The dashed lines represent the locations where incisions should be made. <a href="https://www.jove.com/files/ftp_upload/64574/64574fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Deeply anesthetize each mouse with 100% isofluorane in an anesthesia chamber located in a fume hood or biosafety cabinet using an open drop method34. Sacrifice each mouse by cervical spine dislocation followed swiftly by decapitation. Alternate between genotypes or experimental groups for each sacrifice and dissection12. Obtain tail biopsies after euthanasia by excising 2 mm of the distal tail tip with fine scissors. Store the tissue for genotyping.</li>
	<li>Spray the decapitated head with 70% ethanol to prevent hair from adhering to the tissue and surgical instruments during dissection.</li>
	<li>Insert fine scissors under the skin at the decapitation incision to a pericranial depth and make a midsagittal incision up to the internasal suture (Figure 3A) to retract the scalp from the skull.</li>
	<li>Working from the occipital area toward each temporal aspect, trim the fascia and muscle to expose the external surface of the skull beyond each external acoustic meatus (Figure 3B).</li>
	<li>Secure the scalp and rostral aspect of the skull with the non-dominant hand. With the other, insert fine scissors 2 mm into the caudal side of the foramen magnum, where the spinal cord is visible exiting. Make a midline incision until the scissors reach the internal surface of the intraparietal bone (Figure 3; dashed lines). 
	NOTE: During the initial incision, the scissors must be parallel to the spinal cord with pressure applied toward the internal surface of the skull to prevent damage to the brainstem and cerebellum.</li>
	<li>Change the angle of the scissors so the blades run parallel with the dorsal surface of the skull. Continue advancing the midsagittal incision rostrally through the parietal and frontal bones, using the sagittal and interfrontal sutures as a guide. Use constant upward pressure to avoid damage to the cortex. Terminate the incision just beyond the internasal suture (Figure 3A).</li>
	<li>Make a small perpendicular incision (~3 mm) to the nasal bone, rostral to the internasal suture, by placing the scissors perpendicular to the skull with each blade positioned at a nasal-premaxillary suture and making one even cut (Figure 3; dashed lines). 
	NOTE: This step will increase the ease of retracting the skull and will be critical for collecting the olfactory bulb if this area is of interest.</li>
	<li>While securing the rostral aspect, use one side of a pair of textured forceps to gently lift the skull up from the brain, then laterally and ventrally. Repeat along the midline as needed, then for the other hemisphere until the entire brain surface is exposed.</li>
	<li>Using curved forceps or a fine spatula, gently lift the rostral side of the brain. Cut the optic and cranial nerves to complete the excision from the skull.</li>
	<li>For each condition, collect five to eight mouse brains together into the chilled glass Dounce homogenizer containing 14 mL of Buffer A (Table 1).</li>
</ol>
3. Synaptosome preparation
NOTE: The schematics of this procedure are shown in Figure 4.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64574/64574fig04.jpg" /> 
Figure 4: Synaptosome preparation. Schematic of step 3, the generation of synaptosomes (P2&#39;). <a href="https://www.jove.com/files/ftp_upload/64574/64574fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Homogenize the brains using a glass Dounce homogenizer in 12 up-down passes at 500 rpm (total). Pause briefly at each downstroke to ensure thorough homogenization of the tissue. Homogenize preferentially in an ice bath to avoid warming and protein denaturation. Take 5 &#181;L aliquots for protein concentration determination by the bicinchoninic acid assay (BCA, see Table of Materials). Take 100 &#181;L of whole brain lysate aliquots for western blot (WB). For this and all subsequent samples (Figure 2), take two aliquots for BCA and two aliquots for WB. Flash-freeze all the collected aliquots in liquid nitrogen and store them at &#8722;80 &#176;C.</li>
	<li>Spin the total brain homogenate in a high-speed round bottom centrifuge tube (14 mL) (see Table of Materials) at 800 x g for 10 min at 4 &#176;C to obtain the supernatant (S1). Transfer S1 to a new centrifuge tube, leaving the pellet behind (P1), which contains intact cells and nuclei. Avoid pipetting up the fluffy, white, loose, superficial pellet. Take 2 x 5 &#181;L of S1 for BCA and 2 x 100 &#181;L of S1 for WB.</li>
	<li>Spin S1 at 9,000 x g for 15 min at 4 &#176;C to obtain the synaptosomal supernatant (S2) and crude synaptosome pellet (P2). Take 2 x 10 &#181;L of S2 for BCA and 2 x 500 &#181;L of S2 for WB. Discard the supernatant after obtaining aliquots and proceed to the next step with the pellet.</li>
	<li>Resuspend P2 in 3 mL of ice-cold Buffer A with protease inhibitors and centrifuge at 9,000 x g for 15 min at 4 &#176;C to obtain the supernatant (S2&#39;) and washed synaptosomes (P2&#39;). Discard the supernatant and keep the pellet.</li>
	<li>Resuspend P2&#39; in 3 mL of Buffer A. Avoid resuspending the dark red portion at the bottom of the pellet, which mainly contains mitochondria.&#160;Take 2 x 20 &#181;L of P2&#39; for BCA and 2 x 100 &#181;L of P2&#39; for WB. 
	NOTE: This can be achieved by gently pipette-mixing the edges and surface of the pellet to resuspend the white washed synaptosomes while directing the pipette tip away from the red center of the pellet.</li>
</ol>
4. Hypotonic lysis
NOTE: The schematics of this procedure are shown in Figure 5.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64574/64574fig05.jpg" /> 
Figure 5: Hypotonic lysis. Schematic of step 4, the hypotonic lysis of synaptosomes to generate the lysis supernatant (LS1) and synaptosomal membrane fractions (LP1). <a href="https://www.jove.com/files/ftp_upload/64574/64574fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>For the hypotonic lysis of washed synaptosomes, add 9 volumes of chilled Buffer B (Table 1) to resuspended P2&#39; (~27 mL). Homogenize the synaptosomes in a glass Dounce homogenizer (three up-down passes at 500 rpm).</li>
	<li>Transfer the samples to 50 mL capped conical centrifuge tubes. Rotate them on a tube revolver in a 4 &#176;C cold room for 15 min.</li>
	<li>Centrifuge lysed P2&#39; at 25,000 x g for 20 min at 4 &#176;C to obtain the lysis supernatant (LS1) and the lysis pellet containing synaptosomal membranes (LP1). Take 2 x 50 &#181;L of LS1 for BCA and 2 x 400 &#181;L of LS1 for WB. Transfer LS1 into a capped centrifuge tube for ultracentrifugation (see Table of Materials).</li>
</ol>
5. Synaptic vesicle isolation
NOTE: The schematics of this procedure are shown in Figure 6.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64574/64574fig06.jpg" /> 
Figure 6: Synaptic vesicle isolation and synaptic plasma membrane isolation. (A)&#160;Schematic of step 5, the isolation of synaptic cytosol (LS2) and synaptic vesicle (LP2) fractions, and (B) step 6, the generation of myelin (MF), synaptic plasma membrane (SPM), and mitochondrial (Mito.) fractions following the ultracentrifugation of sucrose gradients. <a href="https://www.jove.com/files/ftp_upload/64574/64574fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Centrifuge LS1 in a fixed angle ultracentrifuge rotor (see Table of Materials) at 100,000 x g for 60 min at 4 &#176;C to obtain synaptic cytosol supernatant (LS2) and synaptic vesicle pellet (LP2). LP2 will be small, translucent, and strongly adhered to the side of the centrifuge tube.</li>
	<li>Resuspend LP2 in 500 &#181;L of Buffer A. Using a 23 G needle and a 1 mL syringe, shear LP2 with gentle trituration. Take 2 x 10 &#181;L of LP2 for BCA and 2 x 250 &#181;L of LP2 for WB.</li>
	<li>Transfer LS2 (~30 mL) to centrifugal filter units with a 10 kDa cutoff (see Table of Materials). 
	NOTE: If proteins smaller than 10 kDa are of interest, 4 kDa cutoff centrifugal filter units are available but will result in longer spin times.</li>
	<li>Concentrate LS2 to approximately 0.5 mL by spinning at 5000 x g for up to 1 h at 4 &#176;C. Take 2 x 10 &#181;L of concentrated LS2 for BCA and 2 x 250 &#181;L of concentrated LS2 for WB. After starting the spin, proceed directly to step 6.1.</li>
</ol>
6. Synaptic plasma membrane isolation
<ol>
	<li>Resuspend LP1 (step 4.3) in 1 mL of Buffer B (Table 1). Take 2 x 10 &#181;L of LP1 for BCA and 2 x 50 &#181;L of LP1 for WB. Adjust the remaining LP1 to a final volume of 7.5 mL and a final sucrose concentration of 1.1 M with Buffer B and Buffer C (Table 1).</li>
	<li>Transfer 7.5 mL of resuspended LP1 into a 14 mL ultracentrifuge tube (see Table of Materials). Carefully overlay LP1 with 3.75 mL of Buffer D (Table 1), and then overlay with 1.25 mL of Buffer A (or a larger volume to fill just below the top of the centrifuge tube). Avoid pipetting down the side of the tube, which will disrupt the sucrose gradient interfaces. After overlaying each sucrose fraction, mark the top of the solution with a pen. Balance the tubes for ultracentrifugation by weight, not volume, with the dropwise addition of Buffer A to within 10 mg. Centrifuge at 48,000 x g for 2.5 h at 4 &#176;C in a swinging bucket ultracentrifuge rotor (see Table of Materials).</li>
	<li>Acquire images of the intact gradients following ultracentrifugation to document the distinctness of each sucrose interface and the success of fractionation.</li>
	<li>Carefully remove the superficial layer of 320 mM sucrose (Buffer A). Recover the myelin fraction (MF) at the 320 mM/855 mM sucrose interface in an 800 &#181;L volume. Recover the synaptic plasma membrane (SPM) fraction at the 855 mM/1.1 M sucrose interface in a 1,000 &#181;L volume. Pipette each fraction up from the wall of the tube in a circular manner to ensure the complete fraction is collected. Carefully aspirate off the remaining sucrose and recover the mitochondrial pellet (Mito.) by resuspending in 200 &#181;L of Buffer B. Take 2 x 100 &#181;L of MF for BCA and 2 x 10 &#181;L of Mito. for BCA; divide the remainder of MF and Mito. samples in half for WB.</li>
	<li>Dilute the SPM fraction with 2 volumes of Buffer B (~2 mL), and then centrifuge in a fixed angle rotor in a 3.5 mL centrifuge tube (see Table of Materials) at 25,000 x g for 20 min at 4 &#176;C. Discard the supernatant and resuspend the SPM pellet in Buffer A for a final volume of 250 &#181;L. Take 2 x 5 &#181;L of SPM for BCA and divide the remaining SPM in half for WB.</li>
	<li>Perform a BCA to determine the protein concentration of each sample, accounting for variable aliquot volume. 
	NOTE: For WB analysis, the suggested working protein concentration for all subcellular fractions is 2 &#181;g/&#181;L (or as high as achievable for LS1 and MF).</li>
</ol>

The authors have nothing to disclose.

In their seminal studies, Whittaker and colleagues used four morphological criteria to identify synaptosomes: (1) the structures have a sealed plasma membrane; (2) the structures contain SVs resembling those in nerve terminals and varicosities in situ in size and number; (3) the structures possess one or more small mitochondria; and (4) the presynaptic membrane is frequently adhered to&#160;a post-synaptic component11,12,13.&#160;Though the first two criteria generally apply to every isolation method, in the most recent protocols described in this article, not all resulting synaptosomes will have mitochondria and attached post-synaptic terminals. Approximately 60% of the synaptosomes will have mitochondria, and only up to 15% are estimated to have attached post-synaptic terminals37. If post-synaptic components are of particular interest, the use of an isotonic Krebs-like homogenization buffer and pressure filtration for enrichment are known to yield high concentrations of synaptosomes with post-synaptic terminals (also termed synaptoneurosomes)22,38.
The method of sacrificing the animal can impact the quality of synaptosomes and synaptic subfractions. Adult animals sacrificed using a euthanasia method that does not require anesthesia will result in the best fraction quality. Further, the brains should be freshly dissected, not frozen, and homogenized using a 1:10 ratio of homogenization buffer (weight/volume) for the most viable synaptic fractions22. The brain has a heterogeneous population of synapses that can be differentiated by the type of neurotransmitters they carry. Synaptosome formation is generally unaffected by synapse type or neurotransmitter content13. An exception is mossy fibers in the cerebellum, which are known to be disrupted in optimal conditions for obtaining synaptosomes from the rest of the brain39,40. Thus, removal of the cerebellum prior to brain homogenization is recommended if the exclusion of this region does not affect the experimental goal. If interested in isolating synaptosomes of a particular neurotransmitter character, areas of the brain that are enriched for neurons containing the neurotransmitter of interest can first be isolated. However, this approach will impose limitations on the final fraction yield, depending on the size of the region of interest (the age of animals is also, therefore, a consideration). There are immunochemical methods for the isolation of neurotransmitter-specific synaptosomes, but the viability and yield will be significantly compromised22. If assessing synaptosome metabolic viability is important, the measurement of neurotransmitter release41,42 or certain enzymatic assays43 can be employed.
Common contaminants in synaptosome preparations include microsomes, free mitochondria, SVs, and neuronal and glial membranes. Contamination can be reduced by increasing the number of washes at the P1 and P2 fractions22 and avoiding resuspension of the red mitochondrial pellet in subsequent steps. In experiments where metabolic viability and time are crucial, reducing the number of washes and using Ficoll or Percoll gradients over sucrose gradients will be helpful44,45,46. These methods also reduce contamination significantly. Whittaker&#39;s original protocol yielded high-quality SVs. Further optimization by Nagy et al.23, included in this method, produces SVs with remarkable homogeneity and purity without compromising significantly on yield36. If specific SV subtypes are of interest, such as glutamatergic (VGLUT-1-containing) or GABAergic (VGAT-1-containing) SVs, immunoisolation using specific antibodies can be performed47,48. Alternative methods are also available for isolating CCVs from synaptosomes, which, due to differential density, may not be present at the same interface as SVs obtained with this method20,49,50.
Overall, the present protocol to isolate synaptic components can be further optimized to obtain fractions with improved homogeneity and viability based on the quality and quantity of the source brain tissue and the experimental goals. For further troubleshooting details, one should refer to book chapters by Dunkley and Robinson22 and Ganzella et al.36.

Synapses are the basic computational units of the brain through which neurons communicate and exert diverse and exquisitely complex functions. Synapses are, thus, fundamental to the health of the brain1; synaptic dysfunction is implicated as a source or result of many disorders2. Synapses are constituted by pre- and post-synaptic terminals, extensions of two different neurons that are closely apposed and separated by a synaptic cleft traversed by synaptic adhesion molecules. Information flows from the pre- to post-synaptic compartment in the form of chemical messengers called neurotransmitters1. The molecular processes involved in neurotransmission are active areas of research3,4,5. Understanding the pathogenic processes within synaptic terminals and the response of synapses to pathology in other neuronal sub-compartments are crucial steps to addressing disorders of the brain1,2. Several methodological advancements, predominantly applied to murine models, have advanced this pursuit6. The isolation of synaptic fractions by differential centrifugation is one such paradigm-shifting method that has enabled the detailed evaluation of synaptic processes in health and disease.
The adult human brain consists of 80-90 billion neurons7,8. Among murine species, the rat brain contains approximately ~200 million neurons, while mice have ~70 million9,10. Each neuron forms thousands of specific synaptic connections with a network of highly polarized neurons intermingled with glial cells and dense vasculature. In such complex and heterogeneous tissue, it was once unthinkable to isolate and study synapses as an independent system. In the 1960s, Victor Whittaker, Catherine Hebb, and others made this possible by isolating intact synaptic terminals using subcellular fractionation11,12,13,14. In an attempt to isolate synaptic vesicles (SVs), they homogenized brains through liquid shear force in iso-osmotic (0.32 M) sucrose followed by ultracentrifugation. They obtained pinched-off, plasma membrane-enclosed, intact nerve terminals or varicosities, which they called nerve-ending particles (NEPs)11,13. As the structural and functional characteristics of the synapse were preserved in these structures, NEPs were later termed &#34;synaptosomes&#34; for congruence with other subcellular organelles13,15. It is worth noting that the work of Eduardo de Robertis and colleagues, who coined the term &#34;synaptic vesicle&#34;, overlapped with that of Whittaker and colleagues and contributed to the validation of &#34;synaptosome&#34; isolation and characterization16,17,18.
Synaptosomes are physiologically active structures that contain all the cellular and molecular properties required for the storage, release, and reuptake of neurotransmitters13,18. The preservation of key synaptic characteristics in vitro and freedom from non-synaptic components also contribute to the utility of this isolation method. Synaptosomes have contributed immensely to the understanding of the chemical and physiological properties of neurotransmission and are now being used to study synaptic molecular processes and their alterations in disease19,20,21,22,23. Synaptosomes are also the initial source material for isolating synaptic components such as SVs, clathrin-coated vesicles (CCVs), synaptic cytosol, synaptic plasma membrane, synaptic mitochondria, synaptic adhesion molecules, and other components of interest, which can facilitate the understanding of the molecular mechanisms of synaptic function18,19,20,24,25,26,27,28. These sub-synaptic components can be obtained by the osmotic lysis of synaptosomes and sucrose density gradient ultracentrifugation15,29. Although the original subcellular fractionation method by Whittaker&#39;s research group is known to be efficient in isolating quality synaptosomes and SVs13,30, recent optimizations enhance the purity of the subcellular fractions22,23,31,32. This article provides a highly detailed and accessible version of a classic protocol for the subcellular fractionation of murine brain tissue to isolate synaptosomes, SVs, and other sub-synaptic components.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mL TB Syringe</td>
			<td>BD</td>
			<td>309649</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL Eppendorf Tubes</td>
			<td>USA Scientific</td>
			<td>1415-2500</td>
			<td></td>
		</tr>
		<tr>
			<td>14 mL, Open-Top Thinwall Ultra-Clear Tube</td>
			<td>Beckman Coulter</td>
			<td>344060</td>
			<td>Compatible with SW 40 Ti</td>
		</tr>
		<tr>
			<td>23 Gauge Precision Glide Hypodermic Needle</td>
			<td>BD</td>
			<td>305145</td>
			<td></td>
		</tr>
		<tr>
			<td>26.3 mL, Polycarbonate Bottle with Cap Assembly</td>
			<td>Beckman Coulter</td>
			<td>355618</td>
			<td>Compatible with Ti70</td>
		</tr>
		<tr>
			<td>3.5 mL, Open-Top Thickwall Polypropylene Tube</td>
			<td>Beckman Coulter</td>
			<td>349623</td>
			<td>Compatible with TLA-100.3</td>
		</tr>
		<tr>
			<td>50 mL Falcon Tubes</td>
			<td>Fisher Scientific</td>
			<td>14-432-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Amicon Ultra-15 Centrifugal Filter Unit&#160;</td>
			<td>Millipore Sigma</td>
			<td>UFC901024</td>
			<td></td>
		</tr>
		<tr>
			<td>Aprotinin</td>
			<td>Sigma-Aldrich</td>
			<td>A6279</td>
			<td>1 mg/mL in diH2O</td>
		</tr>
		<tr>
			<td>Avanti J-26 XP Centrifuge</td>
			<td>Beckman Coulter</td>
			<td>B22984</td>
			<td>&#60;26,000 rpm</td>
		</tr>
		<tr>
			<td>Benchtop HDPE Dewar Flask</td>
			<td>Thermo Scientific</td>
			<td>5028U19</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J Mice</td>
			<td>The Jackson Labs</td>
			<td>000664</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge 5810R</td>
			<td>Eppendorf</td>
			<td>EP022628168</td>
			<td>&#60;14,000 rpm</td>
		</tr>
		<tr>
			<td>complete, Mini, EDTA-free Protease Inhibitor Cocktail Tablets</td>
			<td>Roche</td>
			<td>11873580001</td>
			<td>Add 1 tablet per 50 mL of solution</td>
		</tr>
		<tr>
			<td>Curved Forceps</td>
			<td>Fine Science Tools</td>
			<td>11273-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Surgical Scissors</td>
			<td>Fine Science Tools</td>
			<td>8r</td>
			<td></td>
		</tr>
		<tr>
			<td>Glas-Col Tissue Homogenizing System</td>
			<td>Cole-Parmer</td>
			<td>UX-04369-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Graefe Forceps</td>
			<td>Fine Science Tools</td>
			<td>11650-10</td>
			<td></td>
		</tr>
		<tr>
			<td>High-Speed Polycarbonate Round Bottom Centrifuge Tubes</td>
			<td>ThermoFisher</td>
			<td>3117-0500</td>
			<td>Compatible with JA20</td>
		</tr>
		<tr>
			<td>Isofluorane</td>
			<td>Henry Schein Animal Health</td>
			<td>NDC 11695-6776-2</td>
			<td></td>
		</tr>
		<tr>
			<td>JA-20 Rotor</td>
			<td>Beckman Coulter</td>
			<td>334831</td>
			<td></td>
		</tr>
		<tr>
			<td>Leupeptin</td>
			<td>American Bio</td>
			<td>AB01108</td>
			<td>1 mg/mL in diH2O</td>
		</tr>
		<tr>
			<td>N-[2-Hydroxyethyl] piperazine-N&#8217;-[2-ethanesulfonic acid] (HEPES)</td>
			<td>American Bio</td>
			<td>AB00892</td>
			<td></td>
		</tr>
		<tr>
			<td>Optima L-80 XP Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td></td>
			<td>&#60;100,000 rpm</td>
		</tr>
		<tr>
			<td>Optima TLX Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td></td>
			<td>&#60;120,000 rpm</td>
		</tr>
		<tr>
			<td>Pepstatin A</td>
			<td>Thermo Scientific</td>
			<td>78436</td>
			<td>1 mg/mL in DMSO</td>
		</tr>
		<tr>
			<td>Phenylmethylsulfonyl fluoride (PMSF)</td>
			<td>American Bio</td>
			<td>AB01620</td>
			<td></td>
		</tr>
		<tr>
			<td>Pierce BCA Protein Assay Kit</td>
			<td>Thermo Scientific</td>
			<td>23335</td>
			<td>For determination of protein concentration</td>
		</tr>
		<tr>
			<td>Pipette Tips</td>
			<td></td>
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			<td>Serological Pipettes</td>
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		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich</td>
			<td>S0389</td>
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			<td>Surgical Scissors</td>
			<td>Fine Science Tools</td>
			<td>14002-12</td>
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			<td>SW 40 Ti Swinging-Bucket Rotor</td>
			<td>Beckman Coulter</td>
			<td>331301</td>
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			<td>Teflon-Coated Pestle and Mortar Tissue Grinder</td>
			<td>Thomas Scientific</td>
			<td>3431D94</td>
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			<td>Ti70 Rotor</td>
			<td>Beckman Coulter</td>
			<td>337922</td>
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			<td>TLA-100.3 Rotor</td>
			<td>Beckman Coulter</td>
			<td>349490</td>
			<td></td>
		</tr>
		<tr>
			<td>Tube Revolver</td>
			<td>Dot Scientific</td>
			<td>DTR-02VS</td>
			<td></td>
		</tr>
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</table>

subcellular fractionation for the isolation of synaptic components from the murine brain

Synaptic terminals are the primary sites of neuronal communication. Synaptic dysfunction is a hallmark of many neuropsychiatric and neurological disorders. The characterization of synaptic sub-compartments by biochemical isolation is, therefore, a powerful method to elucidate the molecular bases of synaptic processes, both in health and disease. This protocol describes the isolation of synaptic terminals and synaptic sub-compartments from mouse brains by subcellular fractionation. First, sealed synaptic terminal structures, known as synaptosomes, are isolated following brain tissue homogenization. Synaptosomes are neuronal pre- and post-synaptic compartments with pinched-off and sealed membranes. These structures retain a metabolically active state and are valuable for studying synaptic structure and function. The synaptosomes are then subjected to hypotonic lysis and ultracentrifugation to obtain synaptic sub-compartments enriched for synaptic vesicles, synaptic cytosol, and synaptic plasma membrane. Fraction purity is confirmed by electron microscopy and biochemical enrichment analysis for proteins specific to sub-synaptic compartments. The presented method is a straightforward and valuable tool for studying the structural and functional characteristics of the synapse and the molecular etiology of various brain disorders.

The presented method results in 11 brain subcellular fractions that can be subjected to further purification and various forms of downstream analysis35,36. The gold standard method to assess the quality of synaptosomes, SVs23, and other components is electron microscopy (EM) (Figure 7). Quantitative immunoblotting for proteins that are present in specific subcellular fractions can also be performed to assess markers of fraction purity (Figure 8). For example, immunoblot analysis of fractions reveals the enrichment of N-cadherin (CDH2, UniProt name) in the synaptic plasma membrane fraction (SPM), &#945;-synuclein (SYUA) in the synaptic cytosol (LS2), synaptophysin (SYPH) in the synaptic vesicle fraction (LP2), and myelin basic protein (MBP) in the myelin fraction (MF) when compared to protein levels in the initial whole brain homogenate (Total) (Figure 8). Once fraction purity has been established (for example, note the absence of CDH2 in the LS2 fraction or the many-fold increase in SYPH in the LP2 fraction), quantitative immunoblotting can be used to determine the localization of proteins of interest or query differences in protein distribution between genotypes or treatments. Understanding the subcellular localization of synaptic proteins can enable the dissection of previously undescribed protein functions. Further, this method may elucidate trafficking defects or synaptic dysfunction in disease states, especially when paired with functional assays. For example, our team has used this method to identify a pool of enzymatically active palmitoyl protein thioesterase 1 that is enriched in the synaptic cytosol19.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/64574/64574fig07.jpg" /> 
Figure 7: Electron microscopy (EM) of synaptosomes. (A) Representative EM image of a synaptosome containing synaptic vesicles (arrow). (B)&#160;Representative EM image of a synaptosome with both pre- (arrow) and post-synaptic components (double arrow). (C) Representative EM image of a synaptosome containing synaptic vesicles and a mitochondrion (arrow) (scale bars = 100 nm). <a href="https://www.jove.com/files/ftp_upload/64574/64574fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/64574/64574fig08.jpg" /> 
Figure 8: Immunoblot analysis of subcellular fractions. (A) Markers of subcellular fraction purity (indicated with UniProt nomenclature) are appropriately localized compared to the whole brain homogenate (total): N-cadherin (CDH2) in the synaptic plasma membrane fraction (SPM), synaptophysin 1 (SYPH) and synaptojanin 1 (SYNJ1) in the synaptic vesicle enriched fraction (LP2), &#945;-synuclein (SYUA) in the synaptic cytosol (LS2), and myelin basic protein (MBP) in the myelin fraction (MF). (B) Immunoblot quantification analysis reveals the enrichment (fold-change from total) of fraction purity markers. Data are represented as mean &#177; standard deviation on a log10 scale. The dotted line indicates a 1.5-fold change (y = 0.176) (n = 3 replicate experiments with 8 wild-type mice; age = 2 months; n = 4-5 blots for SYPH, SYUA, MBP, with n = 3 plotted values previously published by Gorenberg et al.19; n = 5 for SYNJ1; n = 1 for CDH2). <a href="https://www.jove.com/files/ftp_upload/64574/64574fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Subcellular Fractionation for the Isolation of Synaptic Components from the Murine Brain at JoVE.com

Subcellular Fractionation for the Isolation of Synaptic Components from the Murine Brain

This protocol presents a robust, detailed method to obtain highly pure synaptosomes, synaptic vesicles, and other synaptic fractions from the mouse brain. This method enables the evaluation of synaptic processes, including the biochemical analysis of protein localization and function with compartmental resolution.

Synaptic terminals are the primary sites of neuronal communication. Synaptic dysfunction is a hallmark of many neuropsychiatric and neurological ...

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JoVE Journal

Neuroscience

5.5K Views.  Yale University. This protocol presents a robust, detailed method to obtain highly pure synaptosomes, synaptic vesicles, and other synaptic fractions from the mouse brain. This method enables the evaluation of synaptic processes, including the biochemical analysis of protein localization and function with compartmental resolution. 

Video: Subcellular Fractionation for the Isolation of Synaptic Components from the Murine Brain

Monitoring Changes in the Intracellular Calcium Concentration and Synaptic Efficacy in the Mollusc Aplysia

It has been suggested that changes in intracellular calcium mediate the induction of a number of important forms of synaptic plasticity (e.g., homosynaptic facilitation) 1. These hypotheses can be tested by simultaneously monitoring changes in intracellular calcium and alterations in synaptic efficacy. We demonstrate how this can be accomplished by combining calcium imaging with intracellular recording techniques. Our experiments are conducted in a buccal ganglion of the mollusc Aplysia californica. This preparation has a number of experimentally advantageous features: Ganglia can be easily removed from Aplysia and experiments use adult neurons that make normal synaptic connections and have a normal ion channel distribution. Due to the low metabolic rate of the animal and the relatively low temperatures (14-16 &deg;C) that are natural for Aplysia, preparations are stable for long periods of time. 
To detect changes in intracellular free calcium we will use the cell impermeant version of Calcium Orange 2 which is easily 'loaded' into a neuron via iontophoresis. When this long wavelength fluorescent dye binds to calcium, fluorescence intensity increases. Calcium Orange has fast kinetic properties 3 and, unlike ratiometric dyes (e.g., Fura 2), requires no filter wheel for imaging. It is fairly photo stable and less phototoxic than other dyes (e.g., fluo-3) 2,4. Like all non-ratiometric dyes, Calcium Orange indicates relative changes in calcium concentration. But, because it is not possible to account for changes in dye concentration due to loading and diffusion, it can not be calibrated to provide absolute calcium concentrations. 
An upright, fixed stage, compound microscope was used to image neurons with a CCD camera capable of recording around 30 frames per second. In Aplysia this temporal resolution is more than adequate to detect even a single spike induced alteration in the intracellular calcium concentration. Sharp electrodes are simultaneously used to induce and record synaptic transmission in identified pre- and postsynaptic neurons. At the conclusion of each trial, a custom script combines electrophysiology and imaging data. To ensure proper synchronization we use a light pulse from a LED mounted in the camera port of the microscope. Manipulation of presynaptic calcium levels (e.g. via intracellular EGTA injection) allows us to test specific hypotheses, concerning the role of intracellular calcium in mediating various forms of plasticity.

Monitoring Changes in the Intracellular Calcium Concentration and Synaptic Efficacy in the Mollusc Aplysia

We demonstrate how changes in the intracellular free calcium concentration and synaptic efficacy can be simultaneously monitored in a ganglion preparation of Aplysia. We image intracellular calcium using a fluorescent dye, Calcium Orange, and induce and monitor synaptic transmission with sharp (intracellular) electrodes.

It has been suggested that changes in intracellular calcium mediate the induction of a number of important forms of synaptic plasticity (e.g., ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

Isolation and Culture of Endothelial Cells from the Embryonic Forebrain

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two vascular networks of the embryonic forebrain, pial and periventricular, are spatially distinctive and have different origins and growth patterns. Endothelial cells from the pial and periventricular vascular networks have unique gene expression profiles and functions. Here we present a step-by-step protocol for isolation, culture, and verification of pure populations of endothelial cells from the periventricular vascular network (PVECs) of the embryonic forebrain (telencephalon). In this approach, telencephalon devoid of pial membrane obtained from embryonic day 15 mice is minced, digested with collagenase/dispase, and dispersed mechanically into a single cell suspension. PVECs are purified from cell suspension using positive selection with anti-CD-31/PECAM-1 antibody conjugated to MicroBeads using a strong magnetic separation method. Purified cells are cultured on collagen 1&#160;coated culture dishes in endothelial cell culture medium until they become confluent and further subcultured. PVECs obtained with this protocol exhibit cobblestone and spindle shaped phenotypes, as visualized by phase-contrast light microscopy and fluorescence microscopy. Purity of PVEC cultures was established with endothelial cell markers. In our hands, this method reliably and consistently yields pure populations of PVECs. This protocol will benefit studies aimed at gaining mechanistic insights into forebrain angiogenesis, understanding PVEC interactions, and cross-talks with neuronal cell types and holds tremendous potential for therapeutic angiogenesis.

This video demonstrates an easy and reliable strategy for preparation of pure cultures of endothelial cells from the embryonic forebrain within 10-12 days and will be useful for research focused on many aspects of cerebral angiogenesis.

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two ...

Isolation of Primary Murine Brain Microvascular Endothelial Cells

The blood-brain-barrier is ultrastructurally assembled by a monolayer of brain microvascular endothelial cells (BMEC) interconnected by a junctional complex of tight and adherens junctions. Together with other cell-types such as astrocytes or pericytes, they form the neurovascular unit (NVU), which specifically regulates the interchange of fluids, molecules and cells between the peripheral blood and the CNS. Through this complex and dynamic system BMECs are involved in various processes maintaining the homeostasis of the CNS. A dysfunction of the BBB is observed as an essential step in the pathogenesis of many severe CNS diseases. However, specific and targeted therapies are very limited, as the underlying mechanisms are still far from being understood. 
Animal and in vitro models have been extensively used to gain in-depth understanding of complex physiological and pathophysiological processes. By reduction and simplification it is possible to focus the investigation on the subject of interest and to exclude a variety of confounding factors. However, comparability and transferability are also reduced in model systems, which have to be taken into account for evaluation. The most common animal models are based on mice, among other reasons, mainly due to the constantly increasing possibilities of methodology. In vitro studies of isolated murine BMECs might enable an in-depth analysis of their properties and of the blood-brain-barrier under physiological and pathophysiological conditions. Further insights into the complex mechanisms at the BBB potentially provide the basis for new therapeutic strategies.
This protocol describes a method to isolate primary murine microvascular endothelial cells by a sequence of physical and chemical purification steps. Special considerations for purity and cultivation of MBMECs as well as quality control, potential applications and limitations are discussed.

Brain microvascular endothelial cells (BMEC) are interconnected by specific junctional proteins forming a highly regulated barrier separating blood and the central nervous system (CNS), the so-called blood-brain-barrier (BBB). The isolation of primary murine brain microvascular endothelial cells, as discussed in this protocol, enables detailed in vitro studies of the BBB.

The blood-brain-barrier is ultrastructurally assembled by a monolayer of brain microvascular endothelial cells (BMEC) interconnected by a junctional ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

Culturing In Vivo-like Murine Astrocytes Using the Fast, Simple, and Inexpensive AWESAM Protocol

The AWESAM (a low-cost easy stellate astrocyte method) protocol entails a fast, simple, and inexpensive way to generate large quantities of in vivo-like mouse and rat astrocyte monocultures: Brain cells can be isolated from different brain regions, and after a week of cell culture, non-astrocytic cells are shaken off by placing the culture dishes on a shaker for 6 h in the incubator. The remaining astrocytes are then passaged into new plates with an astrocyte-specific medium (termed NB+H). NB+H contains low concentrations of heparin-binding EGF-like growth factor (HBEGF), which is used in place of serum in medium. After growing in NB+H, AWESAM astrocytes have a stellate morphology and feature fine processes. Moreover, these astrocytes have more in vivo-like gene expression than astrocytes generated by previously published methods. Ca2+ imaging, vesicle dynamics, and other events close to the membrane can thus be studied in the fine astrocytic processes in vitro, e.g., using live cell confocal or TIRF microscopy. Notably, AWESAM astrocytes also exhibit spontaneous Ca2+ signaling similar to astrocytes in vivo.

Culturing In Vivo-like Murine Astrocytes Using the Fast, Simple, and Inexpensive AWESAM Protocol

The AWESAM protocol described here is optimal for culturing murine astrocytes in isolation from other brain cells in a fast, simple, and inexpensive manner. AWESAM astrocytes exhibit spontaneous Ca2+ signaling, morphology, and gene expression profiles similar to astrocytes in vivo.

The AWESAM (a low-cost easy stellate astrocyte method) protocol entails a fast, simple, and inexpensive way to generate large quantities of in vivo-like ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a protein, it is vital to also determine its distribution. Here, we describe a protocol employing immunofluorescence, confocal microscopy, and computer-based analysis to determine the distribution of the synaptic protein Mover (also called TPRGL or SVAP30). We compare the distribution of Mover to that of the synaptic vesicle protein synaptophysin, thereby determining the distribution of Mover in a quantitative manner relative to the abundance of synaptic vesicles. Notably, this method could potentially be implemented to allow for comparison of the distribution of proteins using different antibodies or microscopes or across different studies. Our method circumvents the inherent variability of immunofluorescent stainings by yielding a ratio rather than absolute fluorescence levels. Additionally, the method we describe enables the researcher to analyze the distribution of a protein on different levels: from whole brain slices to brain regions to different subregions in one brain area, such as the different layers of the hippocampus or sensory cortices. Mover is a vertebrate-specific protein that is associated with synaptic vesicles. With this method, we show that Mover is heterogeneously distributed across brain areas, with high levels in the ventral pallidum, the septal nuclei, and the amygdala, and also within single brain areas, such as the different layers of the hippocampus.

Here, we describe a quantitative approach to determining the distribution of a synaptic protein relative to a marker protein using immunofluorescence staining, confocal microscopy, and computer-based analysis.

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a ...

Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

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

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

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

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.