This research was supported by the AXonomIX research project financed by the Provincia Autonoma di Trento, Italy.

The practices used to obtain the mouse tissues were approved by the Body for the Protection of Animals (OPBA) of the University of Trento (Italy), protocol no. 04-2015, as per art.31 Legislative Decree no. 26/2014. All mice were maintained at the Model Organism Facility of the Centre for Integrative Biology (CIBIO), University of Trento, Italy.
Caution: To avoid any RNA degradation of the samples, prepare all buffers using DEPC-treated water for minimizing RNase contamination.
1. Preparation of Polysomes from Whole Brains
<ol>
	<li>Gathering brain tissues (15 min)
	<ol>
		<li>Euthanize wild-type mouse strain C57BL/6 with CO2 asphyxiation for 5 min36. Carefully dissect the brain out from the skull36, place the tissue into a 1.5 ml tube and immediately place it in liquid nitrogen. Store at -80 &#176;C until use.</li>
	</ol>
	</li>
	<li>Preparation of lysate (30 min)
	<ol>
		<li>Pulverize the whole brain tissue using a mortar and pestle under liquid nitrogen.</li>
		<li>Transfer about 25 mg powder to a cold microcentrifuge tube and immediately (to avoid the thawing of the pulverized tissue) add 0.8 ml of Lysis Buffer (see Table 1) and disrupt the cell by pipetting up and down 25 times quickly.</li>
		<li>Centrifuge the tube at 12,000 x g for 1 min at 4 &#176;C to pellet cellular debris.</li>
		<li>Transfer the supernatant to a new microcentrifuge tube and keep the tube on ice for 15 min.</li>
		<li>Centrifuge the tube at 12,000 x g for 5 min at 4 &#176;C to pellet nuclei and mitochondria.</li>
		<li>Transfer the supernatant to a new microcentrifuge tube.</li>
		<li>Store the supernatant at &#8211;80 &#176;C for a maximum of 6 months or use it immediately.</li>
	</ol>
	</li>
	<li>Sucrose gradient preparation and centrifugation (2 hr and 30 min)
	<ol>
		<li>Wash ultracentrifuge tubes extensively with RNase-free water (diethylpyrocarbonate treated water (DEPC-water) or commercial) and 3% H2O2/DEPC H2O solution.</li>
		<li>Put the tubes on ice and add 5.5 ml of cold 50% sucrose solution at the bottom of each tube (see Table 1 for sucrose solutions). Carefully add the 15% sucrose solution drop by drop, staying close to the interphase in order to preserve a sharp interphase, until the tube is completely filled. When the tube is completely filled, close it with a rubber stopper to avoid air bubble formation.</li>
		<li>In a cold room gently lay down the tubes horizontally and keep it in this position for 2 hr. After this time, slowly straighten the tubes back into the vertical position and put them on ice. The gradients are now ready to be used. Alternatively, prepare the 15-50% sucrose gradient using a conventional gradient former.</li>
		<li>In a cold room remove carefully 1.0 ml from the top of the gradient and overlay the sucrose drop by drop with the cytosolic lysate (i.e., the supernatant obtained in step 1.2).</li>
		<li>Carefully lower the tubes into the buckets of swinging bucket rotor. Centrifuge the gradients for 100 min at 180,000 x g at 4 &#176;C using an ultracentrifuge. &#160;&#160;</li>
		<li>After the centrifugation, leave the tubes in their buckets for 20 min at 4 &#176;C to let the gradients stabilize.</li>
	</ol>
	</li>
	<li>Sucrose gradient fractionation (2 hr)
	<ol>
		<li>Carefully remove one ultracentrifuge tube from the ultracentrifuge rotor and mount it on the collector device of a Density Gradient Fractionation System. Collect 1 ml fractions monitoring the absorbance at 260 nm with a UV/VIS detector (See Figure 1 upper panel). Keep the collected fractions on ice.</li>
		<li>Prepare aliquots of 30-40 &#956;l of the fractions of interest, keep them on ice before storing them at -80 &#176;C till use. Do not use aliquots or sucrose fractions that underwent more than two freezing-thawing cycles (see Figure 1 lower panel).</li>
	</ol>
	</li>
</ol>
2. Sample Preparation for Atomic Force Microscopy (3 hr)
<ol>
	<li>Using tape, peel off the mica sheets.</li>
	<li>Wash the mica sheets 3-4 times with DEPC-water and place it into a small Petri dish. Then dry the surface using air.</li>
	<li>Cover the mica with 200 &#956;l of 1 mM NiSO4 and incubate for 3 min at RT.</li>
	<li>Remove the nickel solution and then dry the surface using air. Carry out all future steps at 4 &#176;C by placing the Petri dish with the mica on ice.</li>
	<li>Thaw an aliquot obtained in 1.4.2 on ice and gently add all of the sample drop by drop on the mica. Using a 100-200 &#956;l tip, spread the sample on the entire surface of the mica. Incubate the sample on ice for 3 min.</li>
	<li>Cover the mica sheet drop by drop with 200 &#956;l cold Buffer-AFM (see Table 1) and incubate for 1 hr on ice.</li>
	<li>Imaging in liquid
	<ol>
		<li>Remove carefully the Buffer-AFM and wash the mica sheets 3-4 times with 200 &#956;l cold Buffer-AFM to remove excess sucrose. Then wash the mica sheets 3 times with cold Washing Solution (see Table 1), leaving the mica surface covered by some microliters of solution.</li>
		<li>Go to point 3 (Image acquisition).</li>
	</ol>
	</li>
	<li>Imaging in air
	<ol>
		<li>Carefully remove the Buffer-AFM and wash the mica 3-4 times with 200 &#956;l cold Buffer-AFM to remove the excess sucrose. Then, wash the mica 3 times with cold Washing Solution (see Table 1) and drain the excess water using paper.</li>
		<li>Leave the sample to dry under the chemical hood with the top of the Petri dish partially open. After 2 hr, close the Petri dish and store at RT. Measure the sample after 2-3 hr as they are stable for years.</li>
	</ol>
	</li>
</ol>
3. Image Acquisition (15 min per image after thermal stabilization)
Note: Polysomes immobilized on mica can be imaged in air or in liquid, using AC mode.
<ol>
	<li>Attach the mica to the sample holder using double-sided tape.</li>
	<li>Insert the sample holder in the AFM stage following manufacturer&#39;s directions. When imaging in liquid, if possible, try to maintain the sample at a temperature lower than 25 &#176;C to increase polysome stability in time.</li>
	<li>Select a cantilever suitable for AC imaging and mount it on the tip holder following manufacturer&#39;s directions. Here, use cantilevers with force constant between 2-20 N/m for air imaging and around 0.1 N/m for liquid imaging.</li>
	<li>Adjust the laser spot on the cantilever and zero the quadrant detector signals.</li>
	<li>Select an opportune driving frequency and drive the cantilever with an amplitude of 10-20 nm.</li>
	<li>Approach the sample until the tip engages the surface.</li>
	<li>Select a scan area of 2x2 &#956;m, acquire at least 512x512 pixel images (pixel width &#60; 4 nm), select a live background subtraction mode and a Z scale of 20-25 nm.</li>
	<li>Inspect the image looking for the presence of round objects characterized by the height between 10 and 15 nm when acquiring in air and 25 and 30 nm when in liquid and the width in the range 25-30 nm. Adjust the setpoint and feedback parameters until sharp objects are visualized. The background should appear relatively flat in good samples, with some 2-4 nm height objects (see Figure 2A and B).</li>
	<li>If the image looks good (as indicated at point 3.8), acquire several (at least ten) 2x2 micron scans at different sample areas.</li>
	<li>(optional). If necessary, acquire high resolution images of selected polysomes (see Figure 2C).</li>
	<li>Apply software corrections (plane subtraction and line-by-line correction algorithms) to correct the AFM images removing arbitrary tilt and drifting effects.</li>
</ol>
4. Data Analysis (30 min per image)
<ol>
	<li>Export images in ImageJ (optional: apply a scale factor) preferentially using a lossless compression format, e.g., the TIFF format (see Figure 3A).</li>
	<li>Use the ImageJ macro toolset RiboPick.ijm (to be copied in the ImageJ macro/toolset subdirectory) to count ribosomes in polysomes (see Figure 3B) and compute statistical properties of the sample (see Figure 3C).
	<ol>
		<li>Start loading an image using the &#60;Read image&#62; tool that initializes the program.</li>
		<li>Pick the ribosome centers with the standard ImageJ &#60;Point selection&#62; tool (shift + left click allows the multi-selection of ribosomes). Mark the selected ribosomes with the &#60;Mark ribosomes&#62; tool. Ribosome coordinates will appear in a custom text window.</li>
		<li>Add more ribosomes to the same polysome repeating the procedure indicated at the point 4.2.2.</li>
		<li>Remove wrongly marked ribosomes using the &#60;Undo last pick&#62; tool (starts removing from the last added ribosome to the first one &#8212; in the current polysome only).</li>
		<li>When the polysome is completed, use the &#60;New polysome&#62; tool. As the polysome number is added as an overlay to the image, the text window is updated.</li>
		<li>Use the &#60;Save results and close&#62; to write the ribosome coordinates file (the default units of the image are used) and a PNG image that summarizes the picked ribosomes and polysomes. The original image is closed without being saved.</li>
	</ol>
	</li>
</ol>

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The authors have no competing financial interests in this paper and nothing to disclose.

As the structure of DNA was proven to be of paramount importance to describe the process of transcription and as the organization of chromatin advanced our understanding of transcriptional control of gene expression, it is essential to analyze the organization and structure of polysomes to improve the truthful comprehension of translation and its regulation.
With the described protocol, an optimal density of polysomes gently immobilized on a flat surface, suitable for AFM imaging is obtained. Using these conditions with conveniently prepared samples, nearly 50 polysomes per hour can be acquired, ready for further analyses and characterization. Moreover, dried samples stored at RT have proven to be suitable for imaging after more than one year.
To successfully obtain polysome images with our protocol, there are some crucial steps: to use tissues that has been properly flash-frozen immediately after dissection and stored at -80 &#176;C to minimize RNA degradation; to use the lysis buffer containing cycloheximide and RNase inhibitors to avoid ribosome dissociation from mRNA; to perform all incubations on ice during the sample deposition on mica; and when preparing the sample for AFM imaging on air, to extensively wash the sample on the mica with buffers and RNase free water in the presence of cycloheximide. The last point is particularly important. If the image appears to contain smooth surfaces at approximately the right height but 100-1,000 nm wide, this is a clear indication of a poorly washed sample (often ribosomes/polysomes can be still observed as faint, embedded objects). A method to resolve this issue is to repeat the washing procedure, but this does not usually mitigate the issue on an already dried sample. In this case it is worth starting again with a new aliquot.
Given the resolution of AFM, this method for studying polysomes cannot resolve the structural details of ribosomal interfaces in polysomes. In fact, AFM is a complementary and simpler approach to cryo-EM, representing an effective and robust technique, to acquire and analyze thousands of polysomes, and better understand the overall ribosome organization in polysomes. Importantly, this protocol has the unique advantage of identifying filaments compatible with RNA27. This very same procedure can be used for any polysomal sample obtained from any biological tissue or cell line. More importantly it can also be successfully employed to obtain transcript-specific information using in vitro translation systems, as recently demonstrated by Lauria and co-workers35. This methodology will pave the way for several experiments, to gain understanding of the kinetics of polysome formation or changes in polysome organization under different cellular or tissue conditions.

The synthesis of protein is the most energy consuming process in cells1,2. Hence, it is not surprising that protein abundances are mainly controlled at the translational rather than transcriptional level3,4,5. The polysome is the fundamental macromolecular component that converts the mRNA information into functional proteinaceous readouts. Polysomes are thus far recognized as macromolecular complexes where several translational controls converge6-13. Despite hundreds of studies on ribosome structure14- 16, the detailed molecular insights into the dynamics of translation and the topology of polysomes has encountered a limited interest. As a consequence, the organization of the native polysomal ribonucleoprotein complex and its potential effect on translation are still rather obscure issues. Polysomes may hide still unknown ordered and functional organizations, potentially mirroring what nucleosomes and epigenetic controls have represented for the transcription field. Indeed, the investigation of this intriguing hypothesis requires additional studies and new technical approaches. In this line, structural techniques and atomic force microscopy may fruitfully collaborate to unravel new mechanisms for controlling gene expression, similarly to what was achieved for the nucleosome17,18.
Since the discovery of the ribosome14-16, its structure has been extensively characterized in prokaryotes19,20, yeast21 and more recently in human22, providing the molecular description of the mechanisms at the base of protein synthesis. Polysomes were initially recognized on the membrane of the endoplasmic reticulum, forming typical 2D geometrical organizations14. As mentioned before, polysome assembly has not been the object of constant interest as the ribosome structure. In the past, polysomes have been studied essentially by transmission EM-based techniques. Only recently, cryo-EM techniques enabled the 3D reconstruction of purified polysomes from in vitro translation systems23-26, human cellular lysates27 or in cells28. These techniques offered more refined information about the ribosome-ribosome organization in polysomes23, 26-28 and a preliminary molecular description of the contact surfaces of adjacent ribosomes in wheat germ polysomes24. Thus, cryo-EM tomography allows the disclosure of ribosome-ribosome interactions with molecular detail, but it is burdened by extensive post-processing and reconstruction analyses that require heavy computational resources for data handling. Moreover, to obtain the molecular detail of ribosome-ribosome interactions, a highly resolved map of the ribosome is required, and this kind of reference ribosome is available only for a few species. Importantly, cryo-EM tomography is not able to detect free RNA. Therefore, new techniques are required to fully understand the organization of the translation machinery.
Beside cryo-EM, AFM has been also employed as a useful tool for direct imaging of polysomes in lower eukaryotes29-33 and humans27. Compared to EM, AFM requires no sample fixation or labeling. In addition, measurements can be carried out in near physiological conditions and with the unique possibility of clearly identifying both ribosomes and naked RNA strands27. Imaging of single polysomes can be performed relatively quickly, obtaining thousands of images at nano-resolution with little post-processing effort compared to the extensive and heavy post-processing and reconstruction analyses required by cryo-EM microscopy. Consequently, AFM data handling and analysis does not need expensive workstations and high computing power. As such, this technique collects information on polysomal shapes, morphological characteristics (such as height, length and width), ribosome densities, the presence of free RNA and the number of ribosomes per polysome27 with a higher throughput than cryo-EM. In such way, AFM represents a powerful and complementary approach to EM techniques to portray polysomes27.
Here we present a complete pipeline from purification to data analysis where AFM is applied to image and analyze mouse brain polysomes. The proposed protocol focuses on purification issues and on the accurate deposition of polysomes on mica substrates that are used for AFM imaging. In addition to conventional particle analysis that can be easily performed with common software used by the AFM community, an ImageJ34 plugin, called RiboPick, is presented for counting the number of ribosomes per polysome27, 35.

<table>
	<tbody>
		<tr>
			<td>Cycloheximide</td>
			<td>Sigma</td>
			<td>01810</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>DNAseI</td>
			<td>Thermo Scientific</td>
			<td>89836</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>RiboLock RNAse Inhibitor</td>
			<td>Life technologies</td>
			<td>EO-0381</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>DEPC</td>
			<td>Sigma</td>
			<td>40718</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>Triton X100</td>
			<td>Sigma</td>
			<td>T8532</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>DTT</td>
			<td>Sigma</td>
			<td>43815</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>Sodium Deoxycholate</td>
			<td>Sigma</td>
			<td>D6750</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>Microcentrifuge&#160;</td>
			<td>Eppendorf</td>
			<td>5417R</td>
			<td>Prepararation of lysate</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma</td>
			<td>S5016</td>
			<td>Sucrose gradient preparation</td>
		</tr>
		<tr>
			<td>Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td>Optima LE-80K</td>
			<td>Sucrose gradient centrifugation</td>
		</tr>
		<tr>
			<td>Ultracentrifuge Rotor</td>
			<td>Beckman Coulter</td>
			<td>SW 41 Ti</td>
			<td>Sucrose gradient centrifugation</td>
		</tr>
		<tr>
			<td>Polyallomer tube</td>
			<td>Beckman Coulter</td>
			<td>331372</td>
			<td>Sucrose gradient centrifugation</td>
		</tr>
		<tr>
			<td>Density Gradient Fractionation System&#160;</td>
			<td>Teledyne Isco</td>
			<td>67-9000-176</td>
			<td>Sucrose gradient fractionation</td>
		</tr>
		<tr>
			<td>AFM</td>
			<td>Asylum Research</td>
			<td>Cypher</td>
			<td>Polysome visualization</td>
		</tr>
	</tbody>
</table>

peering at brain polysomes with atomic force microscopy

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, formed by ribosomes, mRNAs, several proteins and non-coding RNAs, represent integrated platforms where translational controls take place. However, while the ribosome has been widely studied, the organization of polysomes is still lacking comprehensive understanding. Thus much effort is required in order to elucidate polysome organization and any novel mechanism of translational control that may be embedded. Atomic force microscopy (AFM) is a type of scanning probe microscopy that allows the acquisition of 3D images at nanoscale resolution. Compared to electron microscopy (EM) techniques, one of the main advantages of AFM is that it can acquire thousands of images both in air and in solution, enabling the sample to be maintained under near physiological conditions without any need for staining and fixing procedures. Here, a detailed protocol for the accurate purification of polysomes from mouse brain and their deposition on mica substrates is described. This protocol enables polysome imaging in air and liquid with AFM and their reconstruction as three-dimensional objects. Complementary to cryo-electron microscopy (cryo-EM), the proposed method can be conveniently used for systematically analyzing polysomes and studying their organization.

Sucrose gradient polysomal profiling of whole mouse brain 
It is possible to purify polysomes from a cellular lysate by polysomal profiling, which separates macromolecules in accordance to their weight and size. With polysomal profiling, cytoplasmic lysates obtained from cultured cells or tissues, such as in this example, are loaded onto a linear sucrose gradient and processed by ultracentrifugation to separate free RNA from 40S and 60S subunits, 80S ribosomes and polysomes according to their sedimentation coefficients (Figure 1 upper panel). The absorbance of the sample at 254 nm and the fraction collection enable the isolation of the sucrose fraction enriched in ribosomes or polysomes with different number of ribosomes per transcript. The fractions can be collected, aliquoted for AFM imaging and stored at -80 &#176;C (Figure 1 lower panel).
Native polysomes from mouse brain observed by AFM in air reveal tight ribosome interactions
Image acquisition is performed using AC mode (also known as tapping mode). This modality is especially suitable for imaging soft samples (such as the polysomes) because the tip-sample interaction and the corresponding shear forces are greatly reduced. In this way both the tip and the sample are preserved. AFM imaging allows the acquisition of data at high resolution, with exact values that depend on various factors, such as the measuring conditions, the kind of the tip, and the properties of the sample. In the conditions described in this paper a lateral resolution of about 4-6 nm and a vertical resolution of 0.1-0.2 nm are attainable. After the deposition of sucrose aliquots on mica, AFM provides descriptions of single polysomes that appear as clusters of tightly packed ribosomes (Figure 2A, C and C). By cross section analysis (Figure 2D), the height of ribosomal peaks is around 14 nm in accordance to what was previously observed for human ribosomes in polysomes after air drying27. The center to center distance of ribosomes identified by the line profile is 23 nm, which is in agreement with the dimension of ribosomes in solution37, 38.
The number of ribosomes per polysome from a single fraction shows a mono-dispersed distribution
Figure 3A reports a typical AFM image obtained by acquiring a sample absorbed on mica from a fraction of the sucrose gradient. The inset shows a digital zoom of a single polysome, with a magnification factor suitable for the identification of ribosomes in the polysome. Panel B shows the same image after the ribosome identification with the RiboPick macro. Red circles (see the inset) mark the ribosomes, and a progressive number (cyan) is used to help the association of the ribosome coordinates with a specific polysome. The frequency distribution of the number of ribosomes per polysome was analyzed for fraction #10 (See Figure1, lower panel, corresponding to Medium Molecular Weight polysomes (MMW)), which clearly shows a single peak (Panel C, grey bars). The experimental distribution was fitted with a Gaussian curve (black line) that was centered at 5.8 ribosomes/polysome, with a standard deviation of 1.3 ribosomes/polysome.
<img alt="Figure 1" src="/files/ftp_upload/53851/53851fig1.jpg" /> 
Figure 1. Representative absorbance profile for sucrose gradient sedimentation of whole mouse brain. (upper panel) A polysomal cytoplasmic lysate was obtained from P27 mouse brain and loaded onto a linear sucrose gradient (15-50% sucrose [w/v]). It is possible to observe clear peaks corresponding to absorbance at 254 nm of RiboNucleoParticles (RNP), ribosomal subunits (40S and 60S), ribosomes (80S) and polysomes. To avoid repeated freeze and thaw cycles of samples before AFM deposition on mica, it is convenient to split into aliquots both ribosome fraction and polysomal fractions (lower panel) with different number of ribosomes per transcript (i.e., low-, medium-, and high-molecular-weight polysomes (LMW, MMW, and HMW, respectively)). The aliquots can be stored at -80 &#176;C for as long as two years.
<img alt="Figure 2" src="/files/ftp_upload/53851/53851fig2.jpg" /> 
Figure 2. Images of brain polysomes by Tapping-Mode Atomic Force Microscopy. (A) Example of AFM image of MMW brain polysomes after absorption on mica using a linear grey color scale. (B) The same image shown in (A) is depicted using a different Z-ranges for the color scale: 0-0.5 nm gray for the background, 0.5-2 nm, and 2-10 nm yellow for ribosomes. In this case a better visual contrast is obtained for both low and high Z-range objects. (C) Magnification of a typical MMW polysome with cross-section profile (dotted white line). (D) Height profile of two ribosomes defined by cross-section profile in (C) shows ribosome height of 14 nm. This value is compatible with what previously observed in AFM for human ribosomes in polysomes27.
<img alt="Figure 3" src="/files/ftp_upload/53851/53851fig3.jpg" /> 
Figure 3. Example of counting the number of ribosomes per polysome in MMW brain polysomes. (A) AFM image that represents the input of the ImageJ macro, RiboPick. (B) After manually picking each ribosomes per polysome, RiboPick marks in the image the position of each ribosome (red circles). (C) The obtained number of ribosomes per polysomes can be plotted as a distribution that can be fitted with a Gaussian curve. The number of polysomes considered in this example is 164, obtained from 9 independent images. The fitting with the Gaussian curve returns the mean value of the population of polysomes in the MMW fraction (#ribosomes/polysome 5.8&#177;1.3, R2= 0.99).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Buffer</td>
			<td>Composition</td>
			<td>Application</td>
		</tr>
		<tr>
			<td>Lysis Buffer</td>
			<td>10 mM Tris&#8211;HCl, pH 7.5</td>
			<td>Preparation of lysate (1.1)</td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM NaCl</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM MgCl2</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>1% Triton-X100</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>1% Na-deoxycholate</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>0.4 U/ml RNase Inhibitor&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>1 mM DTT</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>0.2 mg/ml cycloheximide</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>5 U/ml Dnase I&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>50% sucrose solution</td>
			<td>50% (w/v) sucrose in&#160;</td>
			<td>Sucrose gradient preparation (1.2)</td>
		</tr>
		<tr>
			<td></td>
			<td>100 mM NaCl</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM MgCl2</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM Tris/HCl pH 7.5&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>15% sucrose solution</td>
			<td>15% (w/v) sucrose in&#160;</td>
			<td>Sucrose gradient preparation (1.2)</td>
		</tr>
		<tr>
			<td></td>
			<td>100 mM NaCl</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM MgCl2</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM Tris/HCl pH 7.5&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nickel Solution</td>
			<td>1 mM NiSO4&#160;</td>
			<td>Sample preparation for AFM (2)</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Buffer-AFM</td>
			<td>100 mM NaCl</td>
			<td>Sample preparation for AFM (2)</td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM MgCl2</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>100 &#181;g/ml cycloheximide</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10 mM Hepes</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td colspan="2">3% (w/v) sucrose, pH = 7.4</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Washing Solution</td>
			<td>DEPC-Water</td>
			<td>Sample preparation for AFM (2)</td>
		</tr>
		<tr>
			<td></td>
			<td>100 &#181;g/ml cycloheximide</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1. Buffers.

Watch this Scientific Journal Video about Peering at Brain Polysomes with Atomic Force Microscopy at JoVE.com

Peering at Brain Polysomes with Atomic Force Microscopy

While ribosome structure has been extensively characterized, the organization of polysomes is still understudied. To overcome this lack of knowledge, we present here a detailed preparation protocol for accurate imaging of mammalian polysomes by atomic force microscopy (AFM) in air and liquid.

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, ...

peering-at-brain-polysomes-with-atomic-force-microscopy

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.3K Views.  Fondazione Bruno Kessler. While ribosome structure has been extensively characterized, the organization of polysomes is still understudied. To overcome this lack of knowledge, we present here a detailed preparation protocol for accurate imaging of mammalian polysomes by atomic force microscopy (AFM) in air and liquid. 

Video: Peering at Brain Polysomes with Atomic Force Microscopy

State-Dependency Effects on TMS:  A Look at Motive Phosphene Behavior

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability (either increasing or decreasing it) via the application of localized magnetic field pulses.1,2 Within the field of TMS, the term state dependency refers to the initial, baseline condition of the particular neural region targeted for stimulation. As can be inferred, the effects of TMS can (and do) vary according to this primary susceptibility and responsiveness of the targeted cortical area.3,4,5

In this experiment, we will examine this concept of state dependency through the elicitation and subjective experience of motive phosphenes. Phosphenes are visually perceived flashes of small lights triggered by electromagnetic pulses to the visual cortex. These small lights can assume varied characteristics depending upon which type of visual cortex is being stimulated. In this particular study, we will be targeting motive phosphenes as elicited through the stimulation of V1/V2 and the V5/MT+ complex visual regions.6

In this article, we examine the effects of visually relevant state dependency on TMS induced motive phosphenic presentations.

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate ...

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron microscope (FIB/SEM). The samples are fixed with aldehydes, heavy metal stained using osmium tetroxide and uranyl acetate. They are then dehydrated with alcohol and infiltrated with resin, which is then hardened. Using a light microscope and ultramicrotome with glass knives, a small block containing the region interest close to the surface is made. The block is then placed inside the FIB/SEM, and the ion beam used to roughly mill a vertical face along one side of the block, close to this region. Using backscattered electrons to image the underlying structures, a smaller face is then milled with a finer ion beam and the surface scrutinised more closely to determine the exact area of the face to be imaged and milled. The parameters of the microscope are then set so that the face is repeatedly milled and imaged so that serial images are collected through a volume of the block. The image stack will typically contain isotropic voxels with dimenions as small a 4 nm in each direction. This image quality in any imaging plane enables the user to analyse cell ultrastructure at any viewing angle within the image stack.

This protocol describes how resin embedded brain tissue can be prepared and imaged in the three dimensions in the focussed ion beam, scanning electron microscope.

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Experimental Demyelination and Remyelination of Murine Spinal Cord by Focal Injection of Lysolecithin

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost oligodendrocytes, myelin, axons, and neurons. Remyelination is an endogenous repair mechanism whereby new myelin is produced subsequent to proliferation, recruitment, and differentiation of oligodendrocyte precursor cells into myelin-forming oligodendrocytes, and is necessary to protect axons from further damage. Currently, all therapeutics for the treatment of multiple sclerosis target the aberrant immune component of the disease, which reduce inflammatory relapses but do not prevent progression to irreversible neurological decline. It is therefore imperative that remyelination-promoting strategies be developed which may delay disease progression and perhaps reverse neurological symptoms. Several animal models of demyelination exist, including experimental autoimmune encephalomyelitis and curprizone; however, there are limitations in their use for studying remyelination. A more robust approach is the focal injection of toxins into the central nervous system, including the detergent lysolecithin into the spinal cord white matter of rodents. In this protocol, we demonstrate that the surgical procedure involved in injecting lysolecithin into the ventral white matter of mice is fast, cost-effective, and requires no additional materials than those commercially available. This procedure is important not only for studying the normal events involved in the remyelination process, but also as a pre-clinical tool for screening candidate remyelination-promoting therapeutics.

Demyelinating diseases can be modeled in animals by focal application of lysolecithin into the CNS. A single injection of lysolecithin into mouse spinal cord produces a lesion that spontaneously repairs over time. The goal is to study factors involved in de- and remyelination, and to test agents for enhancing repair.

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost ...

Characterizing Multiscale Mechanical Properties of Brain Tissue Using Atomic Force Microscopy, Impact Indentation, and Rheometry

To design and engineer materials inspired by the properties of the brain, whether for mechanical simulants or for tissue regeneration studies, the brain tissue itself must be well characterized at various length and time scales. Like many biological tissues, brain tissue exhibits a complex, hierarchical structure. However, in contrast to most other tissues, brain is of very low mechanical stiffness, with Young&#39;s elastic moduli E on the order of 100s of Pa. This low stiffness can present challenges to experimental characterization of key mechanical properties. Here, we demonstrate several mechanical characterization techniques that have been adapted to measure the elastic and viscoelastic properties of hydrated, compliant biological materials such as brain tissue, at different length scales and loading rates. At the microscale, we conduct creep-compliance and force relaxation experiments using atomic force microscope-enabled indentation. At the mesoscale, we perform impact indentation experiments using a pendulum-based instrumented indenter. At the macroscale, we conduct parallel plate rheometry to quantify the frequency dependent shear elastic moduli. We also discuss the challenges and limitations associated with each method. Together these techniques enable an in-depth mechanical characterization of brain tissue that can be used to better understand the structure of brain and to engineer bio-inspired materials.

We present a set of techniques to characterize the viscoelastic mechanical properties of brain at the micro-, meso-, and macro-scales.

To design and engineer materials inspired by the properties of the brain, whether for mechanical simulants or for tissue regeneration studies, the brain ...

Automated Gait Analysis in Mice with Chronic Constriction Injury

The von Frey test is a classical method that has been widely used to examine the sensory function of neuropathic pain animals. However, it has some disadvantages such as subjective data and the requirement of a skilled, experienced experimenter. To date, a variety of modifications have improved the von Frey method, but it still has a few limitations. Recent reports have suggested that gait analysis produces more accurate and objective data from the neuropathic animals. This protocol demonstrates how to perform the automated gait analysis to determine the degree of neuropathic pain in mice. After several days of acclimation, the mice were allowed to walk freely on the glass floor to illuminate footprints. Then, quantification of the footprints and gait were performed through video clips with automatic analysis of various walking parameters, such as area of paw print, swing time, angle of paw, etc.
The main purpose of this study is to describe the methodology of automated gait analysis and briefly compare it with data from the classical sensory test using von Frey filament.

The precise assessment of pain response in a neuropathic animal model is critical to investigate the pathophysiology of pain diseases and develop new analgesics. We present a sensitive and objective method to determine the sensory function of the rodent hind paw by an automated gait analysis system.

The von Frey test is a classical method that has been widely used to examine the sensory function of neuropathic pain animals. However, it has some ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.