Name: peering at brain polysomes with atomic force microscopy
Uploaded: 2016-03-16T15:00:00.000+00:00
Duration: 8 min 49 s
Description: Watch this Scientific Journal Video about Peering at Brain Polysomes with Atomic Force Microscopy at JoVE.com

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&amp;nbsp;</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&amp;nbsp;</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

Universidad San Sebastian

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

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On the end of the microscope's cantilever is a nano-scale probe, which traverses image areas ranging from nanometers to micrometers, measuring the elevation of macromolecules resting on the substrate surface at any given point. Electrostatic forces cause proteins, lipids, and nucleic acids to loosely attach to the substrate in random orientations and permit imaging. The generated data resemble a topographical map, where the macromolecules resolve as three-dimensional particles of discrete sizes (Figure 1) 1,2. Tapping mode AFM involves the repeated oscillation of the cantilever, which permits imaging of relatively soft biomaterials such as DNA and proteins. One of the notable benefits of AFM over other nanoscale microscopy techniques is its relative adaptability to visualize individual proteins and macromolecular complexes in aqueous buffers, including near-physiologic buffered conditions, in real-time, and without staining or coating the sample to be imaged. 
The method presented here describes the imaging of DNA and an immunoadsorbed transcription factor (i.e. the glucocorticoid receptor, GR) in buffered solution (Figure 2). Immunoadsorbed proteins and protein complexes can be separated from the immunoadsorbing antibody-bead pellet by competition with the antibody epitope and then imaged (Figure 2A). This allows for biochemical manipulation of the biomolecules of interest prior to imaging. Once purified, DNA and proteins can be mixed and the resultant interacting complex can be imaged as well. Binding of DNA to mica requires a divalent cation 3,such as Ni2+ or Mg2+, which can be added to sample buffers yet maintain protein activity. Using a similar approach, AFM has been utilized to visualize individual enzymes, including RNA polymerase 4 and a repair enzyme 5, bound to individual DNA strands. These experiments provide significant insight into the protein-protein and DNA-protein biophysical interactions taking place at the molecular level. Imaging individual macromolecular particles with AFM can be useful for determining particle homogeneity and for identifying the physical arrangement of constituent components of the imaged particles. While the present method was developed for visualization of GR-chaperone protein complexes 1,2 and DNA strands to which the GR can bind, it can be applied broadly to imaging DNA and protein samples from a variety of sources.

A tapping mode atomic force microscope (AFM) method for the visualization of plasmid DNA, cytoplasmic proteins, and DNA-protein complexes is described. The method includes alternate approaches for preparing samples for AFM imaging following biochemical manipulation. DNA containing specific protein interacting regions are observed in near-physiologic buffer conditions.

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On ...

Bioengineering

Atomic Force Microscopy of Red-Light Photoreceptors Using PeakForce Quantitative Nanomechanical Property Mapping

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be measured to ~10 pN by a laser and sectored detector, which can be converted to image topography. Amplitude modulation or &#8220;tapping mode&#8221; AFM involves the probe making intermittent contact with the surface while oscillating at its resonant frequency to produce an image. Used in conjunction with a fluid cell, tapping-mode AFM enables the imaging of biological macromolecules such as proteins in physiologically relevant conditions. Tapping-mode AFM requires manual tuning of the probe and frequent adjustments of a multitude of scanning parameters which can be challenging for inexperienced users. To obtain high-quality images, these adjustments are the most time consuming.
PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) produces an image by measuring a force response curve for every point of contact with the sample. With ScanAsyst software, PF-QNM can be automated. This software adjusts the set-point, drive frequency, scan rate, gains, and other important scanning parameters automatically for a given sample. Not only does this process protect both fragile probes and samples, it significantly reduces the time required to obtain high resolution images. PF-QNM is compatible for AFM imaging in fluid; therefore, it has extensive application for imaging biologically relevant materials.
The method presented in this paper describes the application of PF-QNM to obtain images of a bacterial red-light photoreceptor, RpBphP3 (P3), from photosynthetic R. palustris in its light-adapted state. Using this method, individual protein dimers of P3 and aggregates of dimers have been observed on a mica surface in the presence of an imaging buffer. With appropriate adjustments to surface and/or solution concentration, this method may be generally applied to other biologically relevant macromolecules and soft materials.

A method for investigating the structure of a protein photoreceptor using atomic force microscopy (AFM) is described in this paper. PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) reveals intact protein dimers on a mica surface.

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be ...

Engineering

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. Atomic force microscopy (AFM) can address this problem by enabling stretching and relaxation of single protein molecules, which gives direct evidence of specific unfolding and refolding characteristics. AFM-based single-molecule force-spectroscopy (AFM-SMFS) provides a means to consistently measure high-energy conformations in proteins that are not possible in traditional bulk (biochemical) measurements. Although numerous papers were published to show principles of AFM-SMFS, it is not easy to conduct SMFS experiments due to a lack of an exhaustively complete protocol. In this study, we briefly illustrate the principles of AFM and extensively detail the protocols, procedures, and data analysis as a guideline to achieve good results from SMFS experiments. We demonstrate representative SMFS results of single protein mechanical unfolding measurements and we provide troubleshooting strategies for some commonly encountered problems.

We describe the detailed procedures and strategies to measure the mechanical properties and mechanical unfolding pathways of single protein molecules using an atomic force microscope. We also show representative results as a reference for selection and justification of good single protein molecule recordings.

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. ...

Biochemistry

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Sub-nanometer Resolution Imaging with Amplitude-modulation Atomic Force Microscopy in Liquid

Atomic force microscopy (AFM) has become a well-established technique for nanoscale imaging of samples in air and in liquid. Recent studies have shown that when operated in amplitude-modulation (tapping) mode, atomic or molecular-level resolution images can be achieved over a wide range of soft and hard samples in liquid. In these situations, small oscillation amplitudes (SAM-AFM) enhance the resolution by exploiting the solvated liquid at the surface of the sample. Although the technique has been successfully applied across fields as diverse as materials science, biology and biophysics and surface chemistry, obtaining high-resolution images in liquid can still remain challenging for novice users. This is partly due to the large number of variables to control and optimize such as the choice of cantilever, the sample preparation, and the correct manipulation of the imaging parameters. Here, we present a protocol for achieving high-resolution images of hard and soft samples in fluid using SAM-AFM on a commercial instrument. Our goal is to provide a step-by-step practical guide to achieving high-resolution images, including the cleaning and preparation of the apparatus and the sample, the choice of cantilever and optimization of the imaging parameters. For each step, we explain the scientific rationale behind our choices to facilitate the adaptation of the methodology to every user's specific system.

We present a method for achieving sub-nanometer resolution images with amplitude-modulation (tapping mode) atomic force microscopy in liquid. The method is demonstrated on commercial atomic force microscopes. We explain the rationale behind our choices of parameters and suggest strategies for resolution optimization.

Atomic force microscopy (AFM) has become a well-established technique for nanoscale imaging of samples in air and in liquid. Recent studies have shown ...

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record forces curves on 1000 cells in 4 hours automatically. To maintain a 4 hour analysis time, the number of force curves per cell is reduced to 9 or 16. The method combines a Jython based program and a strategy for assembling cells on defined patterns. The program, implemented on a commercial Bio-AFM, can center the tip on the first cell of the array and then move, automatically, from cell to cell while recording force curves on each cell. Using this methodology, it is possible to access the biophysical parameters of the cells such as their rigidity, their adhesive properties, etc. With the automation and the large number of cells analyzed, one can access the behavior of the cell population. This is a breakthrough in the Bio-AFM field where data have, so far, been recorded on only a few tens of cells.

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

This protocol aims to automate AFM measurements on hundreds of microbial cells. First, microbes are immobilized into PDMS stamp microstructures and then force spectroscopy measurements are performed automatically on hundreds of immobilized cells.

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record ...

Static Atomic Force Microscopy to Visualize and Characterize Assembled Nucleosomes

Source: Stormberg, T., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/58820">Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging.</a> J. Vis. Exp.(2019)
In this video, we demonstrate static atomic force microscopy to visualize assembled nucleosomes. This high-resolution imaging technique allows the study of the nucleosome structure.

Source: Stormberg, T., et al. Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging. J. Vis. Exp.(2019)
In this video, ...

JoVE Encyclopedia of Experiments

Biological Techniques

Microscopy Techniques

Atomic Force Microscopy

Atomic force microscopy (AFM) is a type of scanning probe microscopy that can analyze topographic details of various specimens like ceramics, glass, polymers, and biological samples. AFM offers over 1000 times more resolution than the optical imaging system. Images generated from AFM are three-dimensional surface profiles, offering an advantage over the flat, two-dimensional images from other imaging techniques.
The AFM Probe
The probe is regarded as the heart of any AFM setup and comprises the cantilever and tip assembly. Probes are the most commonly replaced part on this type of microscope because&#160; the constant interaction with the samples wears down the tip. Therefore, the choice of material for the probe depends on the properties of the sample. Silicon probes, used to analyze hard samples, are stiffer and sharper than silicon nitride probes, which are better suited to scan softer samples. These sharp tips are produced using electrochemical etching or carbon nanotubes for higher accuracy analysis.
Imaging Modes of AFM
In AFM, surface topography is studied using the interaction between the probe tip and the sample surface. There are two main imaging modes &#8212; a static mode, also referred to as the contact mode, and a dynamic mode.
In the static or contact mode, the tip of the probe is in continuous contact with the sample surface. As the tip drags over the surface, repulsive forces between the sample and the tip result in the cantilever bending, which is recorded. The entire specimen surface is scanned back and forth in both x- and y-axes, called raster scanning, while the vertical movement of the cantilever records the z-axis, thus generating a 3D image.
In the dynamic mode, the probe oscillates just above the sample surface, coming close to, but not touching the surface. Attractive and repulsive forces determine the variation in distance between the tip and the sample, affecting the amplitude of cantilever oscillation. This feedback is recorded to construct the surface topography of the sample.

Atomic force microscopy (AFM) is a type of scanning probe microscopy that can analyze topographic details of various specimens like ceramics, glass, ...

Peering at Brain Polysomes with Atomic Force Microscopy

Summary

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Chapters in this video

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

Atomic Force Microscopy of Red-Light Photoreceptors Using PeakForce Quantitative Nanomechanical Property Mapping

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Sub-nanometer Resolution Imaging with Amplitude-modulation Atomic Force Microscopy in Liquid

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

Static Atomic Force Microscopy to Visualize and Characterize Assembled Nucleosomes

Atomic Force Microscopy