This work was funded by a NSERC Discovery Grant (RGPIN/04246-2018 to G.Y.). G.Y. is a Canada Research Chair. S.K. was funded by Mitacs Globalink Graduate Fellowship and ACHRI Graduate Student Scholarship.

All animal use was supervised by the Animal Care Committee at the University of Calgary. CD1 mice used for the experiment were purchased from commercial vendor.
1. Preparation of solutions
NOTE To prevent RNA degradation, spray workbench and all equipment with RNase decontamination solution. RNase-free tips are used for the experiment. All solutions are prepared in RNase-free water.
<ol>
	<li>Prepare cycloheximide stock solution (100 mg/mL) in DMSO and store at -20 &#176;C.</li>
	<li>Prepare 2.2 M sucrose stock solution by adding 75.3 g of sucrose to RNase-free water and topping up the volume to 100 mL (for ~16 gradient preparation). The solution can be kept at -20 &#176;C for long-term storage.</li>
	<li>Prepare 10x salt solution (1 M NaCl; 200 mM Tris-HCl, pH 7.5; 50 mM MgCl2).</li>
	<li>Prepare 60% (w/v) sucrose chase solution, containing 30 g of sucrose, 5 mL of 10x salt solution and top the volume up to 50 mL with RNase-free water.</li>
	<li>Optionally, add a speck of bromophenol blue or approximately 5 &#181;L of 1% bromophenol blue in RNase-free water to the chase solution. 1.6. Store solutions at 4 &#176;C.</li>
</ol>
2. Preparation of sucrose gradient
NOTE: Accuracy in preparation of sucrose gradients is critical in obtaining consistent and reproducible results.
<ol>
	<li>To prepare six 10-50% sucrose gradients, dilute 2.2 M sucrose solution as in Table 1.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sucrose solution</td>
			<td>10%</td>
			<td>20%</td>
			<td>30%</td>
			<td>40%</td>
			<td>50%</td>
		</tr>
		<tr>
			<td>2.2 M sucrose</td>
			<td>2 mL</td>
			<td>4 mL</td>
			<td>6 mL</td>
			<td>8 mL</td>
			<td>10 mL</td>
		</tr>
		<tr>
			<td>10X salt solution</td>
			<td>1.5 mL</td>
			<td>1.5 mL</td>
			<td>1.5 mL</td>
			<td>1.5 mL</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td>Cycloheximide</td>
			<td>15 &#181;L</td>
			<td>15 &#181;L</td>
			<td>15 &#181;L</td>
			<td>15 &#181;L</td>
			<td>15 &#181;L</td>
		</tr>
		<tr>
			<td>Water</td>
			<td>11.5 mL</td>
			<td>9.5 mL</td>
			<td>7.5 mL</td>
			<td>5.5 mL</td>
			<td>3.5 mL</td>
		</tr>
		<tr>
			<td>Total volume</td>
			<td>15 mL</td>
			<td>15 mL</td>
			<td>15 mL</td>
			<td>15 mL</td>
			<td>15 mL</td>
		</tr>
	</tbody>
</table>
Table 1: Sucrose dilutions for preparations of sucrose gradients.
NOTE: Always prepare sucrose gradients in multiples of two to balance weight during ultracentrifugation.
<ol start="2">
	<li>Wipe the metal blunt end needle with RNase decontamination solution. Briefly rinse the ultracentrifuge tubes, tubing and the syringe using RNase-free water. Air-dry the tubes such that no water drop remains in the tubes as any remaining water will alter the sucrose concentration.</li>
	<li>Place the metal needle on the clamp holder and the centrifuge tube on the motorized stage. To prepare gradients consistently, a home-assembled system was used that contains a motorized stage to hold the ultracentrifuge tube and a syringe pump to inject sucrose solutions (Figure 1, see step 9 for details).</li>
	<li>Fill a 30 mL syringe with ~16 mL of 10% sucrose (enough for six gradients), place it on the syringe pump and connect it to the needle. Make sure that there are no air bubbles in the syringe, tubing, and needle. Wipe the tip off the needle to remove any residual solution.</li>
	<li>Move the motorized stage up such that the tip of the needle touches the centre of the tube at the bottom.</li>
	<li>Set the syringe pump at a flow rate of 2 mL/min for a volume of 2.3 mL.</li>
	<li>After dispensing 2.3 mL of 10% sucrose solution, move down the motorized stage and repeat the process for all six gradients.</li>
	<li>Repeat steps 2.4-2.7 to add 20% sucrose solution to the bottom of the tubes, followed by 30%, 40% and 50% sucrose solutions similarly.</li>
	<li>After preparation of the gradient, seal the ultracentrifuge tube using a paraffin film.</li>
	<li>Leave the tubes overnight at 4 &#176;C such that the different sucrose layers diffuse together to give a continuous gradient.</li>
</ol>
3. Tissue dissection
NOTE: Pregnant mice were euthanized by cervical dislocation preceded by anesthesia with 5% isoflurane.
<ol>
	<li>Collect CD1 mouse embryos at embryonic day 12, or other timepoints as needed, and place embryos in a &#216; 10 cm plate containing ice-cold Hank&#39;s Balanced Salt Solution (HBSS) on ice to retain cell viability.</li>
	<li>Under the dissection scope, transfer one embryo to a &#216; 6 cm plate containing ice-cold HBSS.</li>
	<li>Use 21-23 G needles to fix the position of the head by penetrating through eyes at an approximately 45&#176; angle and apply force to make sure the needles are fixed on the plate (Figure 2A).</li>
	<li>Use No.5 forceps to remove skin and skull, from the middle to the sides.</li>
	<li>Use forceps to cut the olfactory bulbs and remove the meninges to expose the cortical tissues.3.6.Use curved forceps to cut the cortical tissues into 2-3 mL of neurobasal medium on ice (Figure 2B). Pool cortical tissues from different embryos as needed. Tissues from 8-10 embryos usually give ~200 &#181;g total RNA.</li>
</ol>
4. Cell lysis
<ol>
	<li>On the day of tissue dissection, prepare fresh cell lysis buffer as described in Table 2.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Solution</td>
			<td>Final concentration</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>Tris-HCl (pH 7.5)</td>
			<td>20 mM</td>
			<td>100 &#181;L</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>100 mM</td>
			<td>250 &#181;L</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>5 mM</td>
			<td>25 &#181;L</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>1% (v/v)</td>
			<td>500 &#181;L</td>
		</tr>
		<tr>
			<td>Sodium deoxycholate</td>
			<td>0.5% (w/v)</td>
			<td>500 &#181;L</td>
		</tr>
		<tr>
			<td>Dithiothreitol (DTT)</td>
			<td>1 mM</td>
			<td>5 &#181;L</td>
		</tr>
		<tr>
			<td>Cycloheximide</td>
			<td>100 &#181;g/mL</td>
			<td>5 &#181;L</td>
		</tr>
		<tr>
			<td>RNase free water</td>
			<td></td>
			<td>Top up to 5 mL</td>
		</tr>
		<tr>
			<td>Total</td>
			<td>5 mL</td>
			<td>5 mL</td>
		</tr>
	</tbody>
</table>
Table 2: Preparation of&#160;polysome lysis buffer.
NOTE: Supplement lysis buffer with protease and phosphatase inhibitors.
<ol start="2">
	<li>Add cycloheximide to the neurobasal medium with dissected tissues to a final concentration of 100 &#181;g/mL and incubate at 37 &#176;C for 10 min. Cycloheximide blocks translation elongation and, therefore, prevents ribosome run-off 15.</li>
	<li>Centrifuge at 500 x g for 5 min at 4 &#176;C and discard the supernatant.</li>
	<li>Wash the tissues twice with ice-cold phosphate-buffered saline (PBS) supplemented with 100 &#181;g/mL cycloheximide.</li>
	<li>Add 500 &#181;L of cell lysis buffer supplemented with 4 &#181;L of RNase inhibitor. Pipette up and down to resuspend the tissue in lysis buffer. Use an insulin needle to gently lyse the tissue.</li>
	<li>Incubate the tissues on ice for 10 min with brief vortexing every 2-3 min. 
	NOTE: To prevent tissue degradation, ensure that all steps of tissue lysis are carried out on ice.</li>
	<li>Centrifuge at 2,000 x g for 5 min at 4 &#176;C.</li>
	<li>Transfer the supernatant to a new centrifuge tube on ice.</li>
	<li>Centrifuge at ~13,000 x g for 5 min at 4 &#176;C.</li>
	<li>Transfer the supernatant to a new centrifuge tube on ice.</li>
	<li>Measure the RNA concentration in the tissue lysate using a UV-Vis spectrophotometer. 
	NOTE: If multiple samples are included in the experiment, dilute samples to the same concentration with extra cell lysis buffer to minimize variation.</li>
</ol>
5. Sample loading and ultracentrifugation
<ol>
	<li>Pre-cool the ultracentrifugation rotor and swing buckets at 4 &#176;C, and set the temperature of the ultracentrifuge to 4 &#176;C.</li>
	<li>Keep 20 &#181;L of the tissue lysate as total RNA input.</li>
	<li>Load samples (50-300 &#181;g RNA with an equal volume) on the top of the sucrose gradients by slowly dispensing the lysate to the walls of the ultracentrifuge tubes.</li>
	<li>Gently place the ultracentrifuge tubes in the swing bucket. Ensure that all diametrically opposite buckets are balanced.</li>
	<li>Load the swing buckets on the rotor. Set the ultracentrifuge to 190,000 x g (~39,000 rpm) at 4 &#176;C for 90 min.</li>
	<li>Gently place the gradients on ice after centrifugation.</li>
</ol>
6. Fractionation and sample collection
NOTE: A home-assembled fractionating, recording and collecting system is used for the analysis and collecting samples from the gradients (Figure 3, see Device components).
<ol>
	<li>Place an empty ultracentrifuge tube on the tube piercer and gently penetrate the tube with the needle from the bottom.</li>
	<li>Switch on the UV monitor, and open the digital signal recording software.</li>
	<li>Fill a 30 mL syringe with 25 mL of 60% sucrose chase solution. Gently press the syringe such that the chase solution fills up the empty tube and go through the 254 nm UV monitor to set the baseline for detection.</li>
	<li>Press auto-zero on the UV-monitor to register a baseline for detection. Press play on the software to begin recording.</li>
	<li>Once the system establishes a baseline, pause recording on the software and retract the chase solution. Ensure that no residual chase solution remains in the system. 
	NOTE: Rinse the system with RNase-free water to remove residual sucrose solutions remaining from the previous run.</li>
	<li>Load one sample tube to the tube piercer. Gently penetrate the tube with the needle from the bottom.</li>
	<li>Begin recording on the software. Start the syringe pump (flow rate of 1 mL/min for a volume of 25 mL) and the fraction collector to collect polysome fractions. Set fractionator settings to 30 s. 
	NOTE: Before each run, ensure no air bubbles in the syringe or the tubing by gently pressing the syringe to let the chase solution flow continuously through the needle.</li>
	<li>Collect the fractions into 1.5 mL tubes (500 &#181;L each) using a fraction collector.</li>
	<li>The fractions can be processed immediately or stored at -80 &#176;C.</li>
	<li>After fraction collection, rinse the system with RNase-free water to remove any remnant sucrose.</li>
</ol>
7. Extraction of RNA
<ol>
	<li>To each fraction, add 10 ng of luciferase mRNA spike-in control.</li>
	<li>Add three volumes of guanidium hydrocholride based commercial RNA isolation reagent to each fraction.</li>
	<li>Vortex briefly for 15 s.</li>
	<li>Extract RNA using an RNA extraction kit compatible with the solution used in 7.2.</li>
</ol>
8. Reverse transcription and real-time PCR
<ol>
	<li>Measure the concentration of the RNA using UV-Vis spectrophotometer.</li>
	<li>Subject RNA to reverse transcription using a cDNA synthesis kit, according to manufacturer&#39;s protocol.</li>
	<li>Use quantitative real-time PCR (qPCR) to examine the polysomal distribution of gapdh mRNA as an example. qPCR was performed using a qPCR detection system and a qPCR mastermix reagent, with the following cycling conditions: 95 &#176;C for 10 min, then 40 cycles of 95 &#176;C for 15 s, 60 &#176;C for 30 s, 72 &#176;C for 40 s, followed by 95 &#176;C for 60 s.</li>
	<li>Obtain Threshold cycle (Ct) values from the amplification plots and used to calculate fold change using &#916;&#916;Ct method.</li>
</ol>
9. Sucrose gradient making system assembly
NOTE: Follow the steps to assemble each component (as described in Table 3) of the sucrose gradient maker (Figure 1).
<ol>
	<li>Mount two vertical brackets (A2) on the base breadboard (A1).</li>
	<li>Use two slim right-angle brackets (B2) to mount the linear stage actuator to the base breadboard (A1).</li>
	<li>Use the setscrew on the &#216;12.7 mm aluminum post (C2) to fix it on the tube holder base breadboard (C1) as a stand for the tube holder.</li>
	<li>Assemble the right-angle &#216;1/2&#34; to &#216;6 mm post clamp (C3) with the post (C2) and the mini-series optical post (C4).</li>
	<li>Use the setscrew on the optical post (C4) to connect a small V-clamp (C5) as the tube holder.</li>
	<li>Put a centrifuge tube in the tube holder and adjust the angle to make it vertical. Mark the position of the tube and mount the pedestal post holder (C6) on the base breadboard (C1) to support the tube.</li>
	<li>On the other side of the breadboard (A1), use the setscrew of the &#216;12.7 mm aluminum post (D1) to fix it and connect a mini-series optical post (D3) using a right-angle &#216;1/2&#34; to &#216;6 mm post clamp (D2).</li>
	<li>Connect the miniature V-clamp (D4) with blunt-end needle (D5) to the post (D3). Adjust the angle of the clamp to set the needle vertical, and adjust the length of post to ensure the needle meets the centrifuge tube.</li>
	<li>Connect the actuator (B1) to the stepper motor driver (E1) and use the UNO R3 controller board and the joystick module from the UNO starter kit (E2) to control the actuator. A power adaptor (e.g., 9-24 V AC/DC adjustable power adaptor) can be used to drive the motor separately.</li>
	<li>Place the syringe pump (F) next to the gradient making station and connect the syringe with the needle.</li>
	<li>Control the tube holder stage up and down using the joystick.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Item</td>
		</tr>
		<tr>
			<td>B1</td>
			<td>Linear stage actuator</td>
		</tr>
		<tr>
			<td>E1</td>
			<td>Stepper motor driver</td>
		</tr>
		<tr>
			<td>E2</td>
			<td>UNO project super starter kit</td>
		</tr>
		<tr>
			<td>A1</td>
			<td>Breadboard</td>
		</tr>
		<tr>
			<td>A2</td>
			<td>Vertical bracket</td>
		</tr>
		<tr>
			<td>B2</td>
			<td>Slim right-angle bracket</td>
		</tr>
		<tr>
			<td>C1</td>
			<td>Mini-series breadboard</td>
		</tr>
		<tr>
			<td>C5</td>
			<td>Small V-clamp</td>
		</tr>
		<tr>
			<td>D4</td>
			<td>Miniature V-clamp</td>
		</tr>
		<tr>
			<td>C2</td>
			<td>&#216;12.7 mm aluminum post</td>
		</tr>
		<tr>
			<td>C4, D3</td>
			<td>Mini-series optical post</td>
		</tr>
		<tr>
			<td>D1</td>
			<td>&#216;12.7 mm aluminum post</td>
		</tr>
		<tr>
			<td>C3, D2</td>
			<td>Right-angle &#216;1/2&#34; to &#216;6 mm post clamp</td>
		</tr>
		<tr>
			<td>C6</td>
			<td>Mini-series pedestal post holder base</td>
		</tr>
		<tr>
			<td>D5</td>
			<td>Blunt end needle</td>
		</tr>
		<tr>
			<td>F</td>
			<td>Syringe pump</td>
		</tr>
	</tbody>
</table>
Table 3: Gradient making system components.
10. Fractionating and detecting system assembly (Figure 2).
<ol>
	<li>Use a regular round jaw burette clamp to mount the optics module of the UV monitor on the tube piercer. Attach one end of the tubing (1 cm long and 0.56 mm internal diameter) to the connecter of the tube piercer and the other end to the Fluid In port of the optics module. Connect the Fluid Out port to the fractionator.</li>
	<li>Connect the optics module with the control unit of the UV monitor according the manual of the manufacturer.</li>
	<li>Use a breakout cable to connect the signal output socket to the digital converter at the ground and analog input connections, according to the manufacturer&#39;s manual.</li>
	<li>Connect the digital converter to a laptop using a regular USB cable, and record the converted digital signals using data acquisition software.</li>
</ol>

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The authors declare no competing interests.

Polysome profiling is a commonly used and powerful technique to assess the translational status at both single gene and genome-wide levels14 . In this report, we present a protocol of polysome profiling using a home-assembled platform and its application to analyze the developing mouse cortex. This cost-effective platform is easy to assemble and generate robust, reproducible sucrose gradients and polysome profiling with high sensitivity.
It is worthy to note that the pr...

During the development of the mammalian brain, neural stem cells proliferate and differentiate to generate neurons and glia1,2 . The perturbation of this process can lead to alterations in brain structure and function, as seen in many neurodevelopmental disorders3,4. The proper behavior of neural stem cells requires the orchestrated expression of specific genes5. While the epigenetic and transcriptional control of these genes has been intensively studied, recent findings suggest that gene regulation at other levels also contributes to the coordination of neural stem cell proliferation and differentiation6,7,8,9,10. Thus, addressing the translational control programs will greatly advance our understanding of the mechanisms underlying neural stem cell fate decision and brain development.
Three main techniques with different strengths have been widely applied to assess the translational status of mRNA, including ribosome profiling, translating ribosome affinity purification (TRAP) and polysome profiling. Ribosome profiling uses RNA sequencing to determine ribosome-protected mRNA fragments, allowing the global analysis of the number and location of translating ribosomes on each transcript to indirectly infer the translation rate by comparing it to transcript abundance11. TRAP takes advantage of epitope-tagged ribosomal proteins to capture ribosome-bound mRNAs12. Given that the tagged ribosomal proteins can be expressed in specific cell types using genetic approaches, TRAP allows the analysis of translation in a cell type-specific manner. In comparison, polysome profiling, which uses sucrose density gradient fractionation to separate free and poorly-translated portion (lighter monosomes) from those being actively translated by ribosomes (heavier polysomes), provides a direct measurement of ribosome density on mRNA13. One advantage this technique offers is its versatility to study the translation of specific mRNA of interest as well as genome-wide translatome analysis14.
In this paper, we describe a detailed protocol of polysome profiling to analyze the developing mouse cerebral cortex. We use a home-assembled system to prepare sucrose density gradients and collect fractions for downstream applications. The protocol presented here can be adapted easily to analyze other types of tissues and organisms.

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			<td>1.5 mL RNA free microtubes</td>
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			<td>MCT-150-C</td>
			<td></td>
		</tr>
		<tr>
			<td>10 cm dish</td>
			<td>Greiner-Bio</td>
			<td>664160</td>
			<td></td>
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			<td>1M MgCl2</td>
			<td>Invitrogen</td>
			<td>AM9530G</td>
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		<tr>
			<td>21-23G needle</td>
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			<td>305193</td>
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			<td>2M KCl</td>
			<td>Invitrogen</td>
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			<td>30 mL syringe</td>
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			<td>302832</td>
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			<td>Blunt end needle</td>
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		<tr>
			<td>Breadboard</td>
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			<td>MB2530/M</td>
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			<td>Bromophenol blue</td>
			<td>Sigma</td>
			<td>115-39-9</td>
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		<tr>
			<td>CD1 mouse</td>
			<td></td>
			<td>Charles River Laboratory</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved tip forceps</td>
			<td>Sigma</td>
			<td>#Z168785</td>
			<td></td>
		</tr>
		<tr>
			<td>Cycloheximide</td>
			<td></td>
			<td>Sigma</td>
			<td>66-81-9</td>
		</tr>
		<tr>
			<td>Data acquisition software TracerDAQ</td>
			<td>Measurement Computing</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Digital converter</td>
			<td></td>
			<td>Measurement Computing</td>
			<td>USB-1208LS</td>
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		<tr>
			<td>Direct-zol RNA miniprep kit</td>
			<td>Zymo</td>
			<td>R2070</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithiothreitol (DTT)</td>
			<td>Bio-basic</td>
			<td>12-03-3483</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Bioshop</td>
			<td>67-68-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont No.5 forceps</td>
			<td>Sigma</td>
			<td>#F6521</td>
			<td></td>
		</tr>
		<tr>
			<td>Fraction collector</td>
			<td>Bio-Rad</td>
			<td>Model 2110</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Wisent</td>
			<td>311-513-CL</td>
			<td></td>
		</tr>
		<tr>
			<td>Linear stage actuator</td>
			<td>Rattmmotor</td>
			<td>CBX1605-100A</td>
			<td></td>
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		<tr>
			<td>Luciferase control RNA</td>
			<td>Promega</td>
			<td>L4561</td>
			<td></td>
		</tr>
		<tr>
			<td>Maxima first strand cDNA synthesis kit</td>
			<td>Themo Fisher</td>
			<td>M1681</td>
			<td></td>
		</tr>
		<tr>
			<td>Miniature V-clamp</td>
			<td>Thorlabs</td>
			<td>VH1/M</td>
			<td></td>
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		<tr>
			<td>Mini-series breadboard</td>
			<td>Thorlabs</td>
			<td>MSB7515/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini-series optical post</td>
			<td>Thorlabs</td>
			<td>MS2R/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini-series pedestal post holder base</td>
			<td>Thorlabs</td>
			<td>MBA1</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Bio-basic</td>
			<td>7647-14-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurobasal media</td>
			<td>Gibco</td>
			<td>21103-049</td>
			<td></td>
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		<tr>
			<td>&#216;12.7 mm aluminum post</td>
			<td>Thorlabs</td>
			<td>TRA150/M</td>
			<td></td>
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		<tr>
			<td>Parafilm</td>
			<td>Bemis</td>
			<td>PM992</td>
			<td></td>
		</tr>
		<tr>
			<td>PerfeCTa SYBR green fastmix</td>
			<td>Quanta Bio</td>
			<td>CA101414-274</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS)</td>
			<td>Wisent</td>
			<td>311-010-CL</td>
			<td></td>
		</tr>
		<tr>
			<td>Puromycin</td>
			<td>Bioshop</td>
			<td>58-58-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Right-angle clamp</td>
			<td>Thorlabs</td>
			<td>RA90/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Right-angle &#216;1/2&#34; to &#216;6 mm post clamp</td>
			<td>Thorlabs</td>
			<td>RA90TR/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Rnase AWAY</td>
			<td>Molecular BioProducts</td>
			<td>7002</td>
			<td></td>
		</tr>
		<tr>
			<td>RNase free tips</td>
			<td>Frogga Bio</td>
			<td>FT10, FT200, FT1000</td>
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			<td>RNase free water</td>
			<td>Wisent</td>
			<td>809-115-CL</td>
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			<td>RNasin</td>
			<td>Promega</td>
			<td>N2111</td>
			<td></td>
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		<tr>
			<td>Slim right-angle bracket</td>
			<td>Thorlabs</td>
			<td>AB90B/M</td>
			<td></td>
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			<td>Small V-clamp</td>
			<td>Thorlabs</td>
			<td>VC1/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium deoxycholate</td>
			<td>Sigma</td>
			<td>302-95-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Stepper motor driver</td>
			<td>SongHe</td>
			<td>TB6600</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Bioshop</td>
			<td>57501</td>
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		<tr>
			<td>SW 41 Ti rotor</td>
			<td>Beckman Coulter</td>
			<td>331362</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td>Harvard Apparatus</td>
			<td>70-4500</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td>Harvard Apparatus</td>
			<td>70-4500</td>
			<td></td>
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		<tr>
			<td>Triton-X-100</td>
			<td>Bio-basic</td>
			<td>9002-93-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Trizol</td>
			<td>Thermofisher Scientific</td>
			<td>15596018</td>
			<td></td>
		</tr>
		<tr>
			<td>Tube piercer</td>
			<td>Brandel</td>
			<td>BR-184</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge</td>
			<td></td>
			<td>Beckman Coulter</td>
			<td>L8-70M</td>
		</tr>
		<tr>
			<td>Ultracentrifuge tubes</td>
			<td>Beckman Coulter</td>
			<td>331372</td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure 1M Tris-HCl pH 7.5</td>
			<td>Invitrogen</td>
			<td>15567-027</td>
			<td></td>
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		<tr>
			<td>UNO project super starter kit</td>
			<td>Elegoo</td>
			<td>EL-KIT-003</td>
			<td></td>
		</tr>
		<tr>
			<td>UV monitor</td>
			<td>Bio-Rad</td>
			<td>EM-1 Econo</td>
			<td></td>
		</tr>
		<tr>
			<td>Vertical bracket</td>
			<td>Thorlabs</td>
			<td>VB01A/M</td>
			<td></td>
		</tr>
	</tbody>
</table>

analysis of translation in the developing mouse brain using polysome profiling

The proper development of the mammalian brain relies on a fine balance of neural stem cell proliferation and differentiation into different neural cell types. This balance is tightly controlled by gene expression that is fine-tuned at multiple levels, including transcription, post-transcription and translation. In this regard, a growing body of evidence highlights a critical role of translational regulation in coordinating neural stem cell fate decisions. Polysome fractionation is a powerful tool for the assessment of mRNA translational status at both global and individual gene levels. Here, we present an in-house polysome profiling pipeline to assess translational efficiency in cells from the developing mouse cerebral cortex. We describe the protocols for sucrose gradient preparation, tissue lysis, ultracentrifugation and fractionation-based analysis of mRNA translational status.

As a demonstration, the cortical lysate containing 75 &#181;g RNA (pooled from 8 embryos) was separated by the sucrose gradient into 12 fractions. Peaks of UV absorbance at 254 nm identified fractions containing the 40S subunit, 60S subunit, 80S monosome and polysomes (Figure 4A). Analysis of fractions by western blot for the large ribosomal subunit, Rpl10 showed its presence in the 60S subunit (fraction 3), monosome (fraction 4) and polysomes (fractions 5-12) (Figure 4B...

Watch this Scientific Journal Video about Analysis of Translation in the Developing Mouse Brain using Polysome Profiling at JoVE.com

Analysis of Translation in the Developing Mouse Brain using Polysome Profiling

The development of the mammalian brain requires proper control of gene expression at the level of translation. Here, we describe a polysome profiling system with an easy-to-assemble sucrose gradient-making and fractionation platform to assess the translational status of mRNAs in the developing brain.

The proper development of the mammalian brain relies on a fine balance of neural stem cell proliferation and differentiation into different neural cell ...

analysis-translation-developing-mouse-brain-using-polysome

Research

JoVE Journal

Neuroscience

4.9K Views.  University of Calgary. The development of the mammalian brain requires proper control of gene expression at the level of translation. Here, we describe a polysome profiling system with an easy-to-assemble sucrose gradient-making and fractionation platform to assess the translational status of mRNAs in the developing brain.

Video: Analysis of Translation in the Developing Mouse Brain using Polysome Profiling

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

Single-cell Profiling of Developing and Mature Retinal Neurons

Highly specialized, but exceedingly small populations of cells play important roles in many tissues. The identification of cell-type specific markers and gene expression programs for extremely rare cell subsets has been a challenge using standard whole-tissue approaches. Gene expression profiling of individual cells allows for unprecedented access to cell types that comprise only a small percentage of the total tissue1-7. In addition, this technique can be used to examine the gene expression programs that are transiently expressed in small numbers of cells during dynamic developmental transitions8.
This issue of cellular diversity arises repeatedly in the central nervous system (CNS) where neuronal connections can occur between quite diverse cells9. The exact number of distinct cell types is not precisely known, but it has been estimated that there may be as many as 1000 different types in the cortex itself10. The function(s) of complex neural circuits may rely on some of the rare neuronal types and the genes they express. By identifying new markers and helping to molecularly classify different neurons, the single-cell approach is particularly useful in the analysis of cell types in the nervous system. It may also help to elucidate mechanisms of neural development by identifying differentially expressed genes and gene pathways during early stages of neuronal progenitor development.
As a simple, easily accessed tissue with considerable neuronal diversity, the vertebrate retina is an excellent model system for studying the processes of cellular development, neuronal differentiation and neuronal diversification. However, as in other parts of the CNS, this cellular diversity can present a problem for determining the genetic pathways that drive retinal progenitors to adopt a specific cell fate, especially given that rod photoreceptors make up the majority of the total retinal cell population11. Here we report a method for the identification of the transcripts expressed in single retinal cells (Figure 1). The single-cell profiling technique allows for the assessment of the amount of heterogeneity present within different cellular populations of the retina2,4,5,12. In addition, this method has revealed a host of new candidate genes that may play role(s) in the cell fate decision-making processes that occur in subsets of retinal progenitor cells8. With some simple adjustments to the protocol, this technique can be utilized for many different tissues and cell types.

A method for the isolation of single retinal cells and subsequent amplification of their cDNAs is described. Single-cell transcriptomics reveals the degree of cellular heterogeneity present in a tissue and uncovers new marker genes for rare cell populations. The accompanying protocol can be adjusted to suit many different cell types.

Highly specialized, but exceedingly small populations of cells play important roles in many tissues. The identification of cell-type specific markers and ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Genetic modification of specific regions of the developing mammalian brain is a very powerful experimental approach. However, generating novel mouse mutants is often frustratingly slow. It has been shown that access to the mouse brain developing in utero with reasonable post-operatory survival is possible. Still, results with this procedure have been reported almost exclusively for the most superficial and easily accessible part of the developing brain, i.e. the cortex. The thalamus, a narrower and more medial region, has proven more difficult to target. Transfection into deeper nuclei, especially those of the hypothalamus, is perhaps the most challenging and therefore very few results have been reported. Here we demonstrate a procedure to target the entire hypothalamic neuroepithelium or part of it (hypothalamic regions) for transfection through electroporation. The keys to our approach are longer narcosis times, injection in the third ventricle, and appropriate kind and positioning of the electrodes. Additionally, we show results of targeting and subsequent histological analysis of the most recessed hypothalamic nucleus, the mammillary body.

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Despite the functional and medical importance of the hypothalamus, in utero genetic manipulation of its development has rarely been attempted. We show a detailed procedure for in utero electroporation into the mouse hypothalamus and show representative results of total and partial (regional) hypothalamic transfection.

Genetic  modification of specific regions of the developing mammalian brain is a very  powerful experimental approach. However, generating novel mouse ...

Assessing Cell Cycle Progression of Neural Stem and Progenitor Cells in the Mouse Developing Brain after Genotoxic Stress

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a pseudostratified epithelium lining the lateral ventricles of the embryonic brain. Genotoxic stresses, such as ionizing radiation, have highly deleterious effects on the developing brain related to the high sensitivity of NSPC. Elucidation of the cellular and molecular mechanisms involved depends on the characterization of the DNA damage response of these particular types of cells, which requires an accurate method to determine NSPC progression through the cell cycle in the damaged tissue. Here is shown a method based on successive intraperitoneal injections of EdU and BrdU in pregnant mice and further detection of these two thymidine analogues in coronal sections of the embryonic brain. EdU and BrdU are both incorporated in DNA of replicating cells during S phase and are detected by two different techniques (azide or a specific antibody, respectively), which facilitate their simultaneous detection. EdU and BrdU staining are then determined for each NSPC nucleus in function of its distance from the ventricular margin in a standard region of the dorsal telencephalon. Thus this dual labeling technique allows distinguishing cells that progressed through the cell cycle from those that have activated a cell cycle checkpoint leading to cell cycle arrest in response to DNA damage.
An example of experiment is presented, in which EdU was injected before irradiation and BrdU immediately after and analyzes performed within the 4 hr&#160;following irradiation. This protocol provides an accurate analysis of the acute DNA damage response of NSPC in function of the phase of the cell cycle at which they have been irradiated. This method is easily transposable to many other systems in order to determine the impact of a particular treatment on cell cycle progression in living tissues.

Administration of two analogs of thymidine, EdU and BrdU, in pregnant mice allows the analysis of cell cycle progression in neural and progenitor cells in the embryonic mouse brain. This method is useful to determine the effects of genotoxic stress, including ionizing radiation, during brain development.

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

Modelling Zika Virus Infection of the Developing Human Brain In Vitro Using Stem Cell Derived Cerebral Organoids

The recent emergence of Zika virus (ZIKV) in susceptible populations has led to an abrupt increase in microcephaly and other neurodevelopmental conditions in newborn infants. While mosquitos are the main route of viral transmission, it has also been shown to spread via sexual contact and vertical mother-to-fetus transmission. In this latter case of transmission, due to the unique viral tropism of ZIKV, the virus is believed to predominantly target the neural progenitor cells (NPCs) of the developing brain.
Here a method for modeling ZIKV infection, and the resulting microcephaly, that occur when human cerebral organoids are exposed to live ZIKV is described. The organoids display high levels of virus within their neural progenitor population, and exhibit severe cell death and microcephaly over time. This three-dimensional cerebral organoid model allows researchers to conduct species-matched experiments to observe and potentially intervene with ZIKV infection of the developing human brain. The model provides improved relevance over standard two-dimensional methods, and contains human-specific cellular architecture and protein expression that are not possible in animal models.

Modelling Zika Virus Infection of the Developing Human Brain In Vitro Using Stem Cell Derived Cerebral Organoids

This protocol describes a technique used to model Zika virus infection of the developing human brain. Using wildtype or engineered stem cell lines, researchers may use this technique to uncover the various mechanisms or treatments that may affect early brain infection and resulting microcephaly in Zika virus-infected embryos.

The recent emergence of Zika virus (ZIKV) in susceptible populations has led to an abrupt increase in microcephaly and other neurodevelopmental conditions ...

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 ...

Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent development of thin, flexible ECoG electrodes has enabled conduction of stable recordings of large-scale cortical activity. We have developed a whole-cortical ECoG array for the common marmoset. The array continuously covers almost the entire lateral surface of cortical hemisphere, from the occipital pole to the temporal and frontal poles, and it captures whole-cortical neural activity in one shot. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain. Marmosets have two advantages regarding ECoG recordings, one being the homologous organization of anatomical structures in humans and macaques, including frontal, parietal, and temporal complexes. The other advantage is that the marmoset brain is lissencephalic and contains a large number of complexes, which are more difficult to access in macaques with ECoG, that are exposed to the brain surface.These features allow direct access to most cortical areas beneath the surface of the brain. This system provides an opportunity to investigate global cortical information processing with high resolutions at a sub-millisecond order in time and millimeter order in space.

We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent ...