We thank Dr. Mario Borgnia from the cryo-EM core at the National Institute of Environmental Health Sciences and Dr. Joshua Strauss from the University of North Carolina at Chapel Hill for their mentorship and training in the cryo-EM grid preparation. We also thank Dr. Juliana Mello Da Fonseca Rezende for technical assistance in the initial stages of this project. We appreciate the key contribution and support of the late Dr. Samuel H. Wilson and his lab members, especially Dr. Rajendra Prasad and Dr. Joonas Jamsen for the purification of the genetically fused APE1-Pol&#946; complex. Research has been supported by the Intramural Research Program of the National Institutes of Health, National Institute of Environmental Health Sciences [grant numbers Z01ES050158, Z01ES050159, and K99ES031662-01].

1. Assemble nucleosome core particles via salt-gardient dialysis
NOTE: The preparation of nucleosome core particles using recombinant histone proteins for structural studies has been extensively described in detail by others14,15,16. Follow the purification of recombinant X. laevis histones and histone octamer assembly described by others14,15, and assemble the nucleosomal substrate as described below.
<ol>
	<li>Purchase three oligonucleotides (listed in Table 1) at the indicated scale: UND-197mer, 161mer, 35mer.
	<ol>
		<li>PAGE purify ultramers (197mer and 161mer) as described17.
		<ol>
			<li>Anneal equimolar amounts of oligonucleotides in 1x annealing buffer (10 mM Tris-Cl, pH 8.0, 50 mM NaCl). For example, mix 40 &#181;L of 161-mer [166.7 &#181;M]; 8 &#181;L of 35-mer [292.4 &#181;M]; 16.8 &#181;L of 197-mer comp strand [408.2 &#181;M]; 10 &#181;L of 10x anneal buffer and 10.9 &#181;L of dH2O. 
			NOTE: It is critical to control the annealing reaction with a temperature gradient. See Table 2 for the details on the temperature gradient.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Perform small-scale reconstitutions (scale by a factor of 1/57 of the example shown below for a final volume of 50 &#181;L) to determine the ratio of DNA to histone octamer (typical starting ratios of DNA:octamer are as follows: 1:0.9, 1:1.1, 1:1.2, 1:2; shown in Figure 2A); it is critical to determine this ratio empirically with small-scale reconstitutions.
	<ol>
		<li>After this ratio is determined, prepare a large-scale reconstitution. For example, mix 32.9 &#181;L of DNA-197 bp [99.4 &#181;M]; 1647.1 &#181;L of TE (1x); 1120 &#181;L of 5 M NaCl, and 62.3 &#181;L of WT histone octamer [2.56 &#181;M].</li>
	</ol>
	</li>
	<li>Make dialysis buffers: RBlow and RBhigh as described in Table 3 and Table 4, respectively; set up the reconstitutions as previously described14.
	<ol>
		<li>Transfer the dialysis tube into the beaker containing RBhigh, and allow for dialysis to proceed, with gentle stirring, for 16 h or until 2 L RBlow has been transferred into the waste beaker.</li>
	</ol>
	</li>
	<li>Transfer the dialysis tube into a 1 L beaker containing RB50mM buffer (Table 5) and allow the sample to dialyze for at least 3 h or overnight.
	<ol>
		<li>Evaluate reconstitution efficiency on a 6% polyacrylamide nondenaturing gel, ensuring there is less than 10% free DNA and a single NCP band; store NCPs at 4 &#176;C. See Figure 2A (lane 5) and Figure 2B for representative optimal reconstitution results.</li>
	</ol>
	</li>
</ol>
2. Prepare NCP-DNA repair factor complex (NCP-DRF)
<ol>
	<li>Purification by preparative-gel electrophoresis 
	NOTE: This method is the most labor-intensive (of the two described), and while it is effective at removing high molecular weight (HMW) species (Figure 6A) because of the high dilution factor, it can lead to some disassembly as shown in Figure 7A (NCP prep cell out lane), releasing DNA. However, up to 25% free DNA is still compatible with cryo-EM studies. Because of the high dilution, use this method to purify the NCP only, rather than the whole complex.
	<ol>
		<li>Prepare 70 mL of 6% nondenaturing polyacrylamide gel solution in 0.2x TBE, using polyacrylamide gel solution 37.5:1, acrylamide:bisacrylamide. Pour a cylindrical gel with outer radius of 28 mm and polymerize overnight.</li>
		<li>Assemble the preparative gel running apparatus, using a dialysis membrane with a 6-8 kDa MW cutoff. Use 0.25x TBE as the running buffer and 1x elution buffer (50 mM KCl, 10 mM HEPES, pH 7.5). Pre-run the cylindrical gel at a constant 12 W for 1 h, and collect fractions running the peristaltic pump at a flow rate of approximately 1 mL/min. 
		NOTE: If a UV detector is not available to identify fractions containing DNA, a screening experiment with 32P-NCP can be performed to identify the fractions of interest. Keeping all conditions the same, unlabeled NCPs can then be purified without a UV detector.</li>
		<li>Concentrate the 2.8 mL of NCP to 250 &#181;L using a centrifugal filter (MW cutoff 30 kDa), and load onto the preparative gel and electrophorese for a total of 6 h. At 2 h, when the xylene cyanol has ran out of the gel, start collecting 1.5 mL fractions. 
		NOTE: At 2.5 h, fractions will be clear of xylene cyanol; the DNA (197mer) elutes at approximately 3-3.5 h, and the NCP is expected to elute at 4.5-5 h mark.</li>
		<li>Analyze fractions of interest on a 6% nondenaturing polyacrylamide gel. Pool fractions containing the NCP, and immediately concentrate with a centrifugal filter (MW cutoff 30 kDa) to 1 mg/mL. The next day, evaluate the quality of the NCP on a 6% polyacrylamide nondenaturing gel before complexing with the DNA repair factor (DRF) of interest.</li>
		<li>Incubate the NCP and the DRF of interest on ice for 15 min using an optimal buffer for the specific complex. For example, mix 1,000 &#181;L of NCP [1.2 &#181;M]; 260 &#181;L of 5x binding buffer (250 mM HEPES, pH 8, 500 mM KCl, 25 mM MgCl2), 22 &#181;L of 3 M KCl, and 18 &#181;L of MBP- Pol &#946;-APE1 [338 &#181;M]. 
		NOTE: The temperature and ionic strength at which the complex is made may need optimization&#160;for different complexes.</li>
		<li>Add glutaraldehyde (freshly opened; EM grade) to a final concentration of 0.005%. Therefore, to this reaction, add 26.8 &#181;L&#160;of 0.25% glutaraldehyde with 13.2 &#181;L of dH2O. Mix well, and incubate at room temperature for 13 min.</li>
		<li>Quench with 1 M Tris-Cl, pH 7.5, to a final concentration of 20 mM Tris-Cl, pH 7.5. Concentrate to ~50 &#181;L and exchange the buffer with 1x freezing buffer: 50 mM KCl, 10 mM HEPES, pH 7.5 using a desalting column. 
		CAUTION: Glutaraldehyde can cause severe skin burns and eye damage; it is harmful if swallowed, and it is toxic if inhaled. Wear a lab coat, goggles, gloves, and handle glutaraldehyde in a fume hood and&#160;wear a mask. 
		NOTE: It is critical to perform the crosslinking reaction in a large volume with the NCP at this lower concentration. Previous attempts of crosslinking, even at this concentration of glutaraldehyde, but at a 10x higher concentrated NCP led to aggregation of the sample and over-crosslinking as indicated by SDS PAGE and size exclusion chromatography. These grids had no discernable particles with just clumps (Figure 5D,E).</li>
		<li>Determine the absorbances at OD280 and OD260 and concentrate further if needed to reach 1.3-3 mg/mL, based on OD280 (typical ratio of OD260/ OD280= 1.7-2 yields good particles). For this small volume of 50 &#181;L, the best method to reduce sample loss is to make a dialysis button with a lid of a 1.7 mL tube covered by a dialysis membrane (6-8 kDa MW cutoff), cutting the bottom of the tube and using the rim of the tube to seal the membrane on top of the lid.</li>
		<li>Place the dialysis button containing the sample on top of a polyethylene glycol bed, with the membrane facing down (check progress every 2 min). This method is preferred where the concentration needed is less than or equal to 2.5-fold to avoid diluting or losing the sample in the concentrator. It is critical to immediately prepare cryo-EM grids of the complex after this step.</li>
	</ol>
	</li>
	<li>Purification by size exclusion chromatography
	<ol>
		<li>The day before freezing, wash and equilibrate a size exclusion column with 60 mL of dH2O, followed by 80 mL of freezing buffer (50 mM KCl, 10 mM HEPES, pH 8) overnight at a rate of 0.4 mL/min. Using NCPs straight after reconstitution, prepare the same complex using 2.5x greater amounts as follows: mix 2,500 &#181;L of NCP [1.2 &#181;M]; 650 &#181;L of 5x binding buffer; 45 &#181;L of MBP-Pol &#946;-APE1 [338 &#181;M] and 55 &#181;L of 3M KCl.</li>
		<li>Incubate the mixture on ice without crosslinker for 15 min. Then, add glutaraldehyde to a final concentration of 0.005% as described in the preparative gel method. In this case, however, concentrate the complex in a pre-equilibrated (with freezing buffer) centrifugal filter to approximately 120 &#181;L.</li>
		<li>Immediately analyze the peak fractions and concentrate those fractions containing the histones and MBP-Pol &#946;-APE1 as indicated previously. See Figure 5A,B,C for successful results using the size exclusion method&#160;and Figure 7&#160;using both methods.</li>
	</ol>
	</li>
</ol>
3. Freeze nucleosomal complex
<ol>
	<li>After turning on the plunge freezer and filling the humidifier with 50 mL of dH2O, set the chamber temperature to 22 &#176;C and HR (humidity) to 98%. Place the ethane cup and liquid nitrogen cup (containing a labeled grid box) in their respective space holders.</li>
	<li>Cover the ethane cup with the ethane lid dispenser, and carefully pour liquid nitrogen (LN2) over it, while also filling the LN2 cup with LN2. When the LN2 level has stabilized and reaches 100%&#160;and a temperature of -180 &#176;C, carefully open the ethane valve and fill the ethane cup until it forms a bubble on the clear lid. Remove the ethane dispenser.</li>
	<li>Place two filter papers onto the blotting device and secure them with a metal ring. Go to the set up and use the following parameters for blotting: 0 pre-blot, 3 s blot, 0 post-blot; select A-plunge and click on OK.</li>
	<li>On the main screen, click on Load Forceps, and load them with a grid with the carbon/application side facing toward the left (prepared as before). Calibrate the forceps adjusting the Z-axis to ensure a solid blot. Click on Lower Chamber and apply 3 &#181;L of the complex (1.3-3 mg/mL; OD280). Click on Blot/A-plunge. This will rotate the grid to blot from the front and will plunge freeze it.</li>
	<li>Transfer and store the grid in the grid box in the LN2 chamber. When all four grids have been frozen and placed in the grid box, rotate the lid to neutral position, where all the grids are covered by the lid, and tighten the screw. Grids can be stored in LN2 until screening is initiated.</li>
</ol>
4. Screen grids
<ol>
	<li>Before loading the samples in the microscope, place the vitrified grids on a ring and secure them using a C-clip. Perform this clipping process under LN2 in a humidity-controlled room, to avoid ice contamination.</li>
	<li>When loading the microscope, insert the grids in a 12-slot cassette. Shuttle the cassette into a nanocab capsule and load it into the autoloader. The autoloader robotic mechanism initiates the process.</li>
	<li>Transfer the grid from the cassette to the microscope stage. Adjust the stage to eucentric height by wobbling the stage 10&#176; and simultaneously moving the Z-height until minimal planar shift is observed in the images.</li>
	<li>Once the eucentric height is reached, start imaging. First, obtain the atlas by taking a 3 x 3 montage image of the grid, in which each montage is taken at 62x magnification. Pick three squares of varying sizes; take the eucentric height, and then image at 210x magnification.</li>
	<li>Once a square is imaged, pick one hole each from the edge, center, and in between within the squares. Image each hole at 2600x magnification.</li>
	<li>Before taking this high magnification image, the autofocusing occurs at an offset of the imaging area. Take the high magnification image from the center of the hole at 36,000x magnification and at 7.1 s exposure time, 60 frames, 1.18-pixel size, and 3 &#181;m defocus. See representative images of screening in Figure 5, Figure 6, and&#160;Figure 7.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solvent+mediated+interactions+in+the+structure+of+the+nucleosome+core+particle+at+1.9+a+resolution.">Solvent mediated interactions in the structure of the nucleosome core particle at 1.9 a resolution.</a> Journal of Molecular Biology. 319 (5), 1097-1113 (2002).</li><li>Suto, R. K., Clarkson, M. J., Tremethick, D. 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The authors declare no competing interests.

A specific protocol for purifying the DNA repair factor will be dependent on the enzyme of interest. However, there are some general recommendations, including the use of recombinant methods for protein expression and purification18; if the protein of interest is too small (&#60;50 kDa), structure determination by cryo-EM had been nearly impossible until more recently through the use of fusion systems19, nanobody-binding scaffolds20, and optimizing imaging strategies21.
It is quite common to obtain poor quality grids in the initial stages. Initial attempts to determine NCP homogeneity and stability without any crosslinker, using uranyl acetate and carbon-coated copper grids for negative staining, showed stable, nicely dispersed NCPs with minimal aggregation. However, once this sample was vitrified (at 20x higher concentration) in cryo-EM grids, no particles were detected. Sample optimization to keep the NCPs from falling apart or aggregating was the biggest hurdle. Additionally, two methods of preparation are described separately; however, they are not mutually exclusive and can be used back-to-back by first purifying the NCP via a preparative gel, complexing it with the repair factor, and purifying the complex with a size exclusion column. This requires twice as much material for the initial step, and it is also more time-consuming, but helpful for difficult samples (Figure 6).
Finding a starting point for the concentration of crosslinker was difficult due to the lack of consistency in methods of crosslinking and the amounts of crosslinker used by others22,23. Attempts to replicate the less-controlled crosslinking reaction on the bench top described by Anderson et al. failed to give positive results, and the complex was overly crosslinked. This is likely due to the source of glutaraldehyde (freshly opened ampule compared to long-term stored glutaraldehyde). Because this level of detail is often omitted in non-methods, research articles, it is difficult to replicate the exact conditions as a starting point. Optimal results were obtained when the complex was crosslinked at a concentration of 1.2 &#181;M in 150 mM KCl ionic strength compared to crosslinking the complex at 10 &#181;M NCP, at the same glutaraldehyde concentration of 0.005% for 13 min at room temperature. The crosslinking efficiency is also likely dependent on the complex that is being formed as some chromatin-interacting factors may induce/require greater destabilization influencing where the crosslinking will occur and the overall effect on NCP stability. Therefore, while optimization of these conditions is inevitable, finding critical steps such as optimal NCP and salt concentration during the crosslinking reactions will be critical.
Another important parameter was the ratio of NCP:DRF that yielded enough molecules on the grid containing the DRF bound to the NCP. Indeed, this parameter has also been identified by others as one that is critical to optimize for structural studies16 given that even for cryo-EM homogeneity of the way the DRF binds to the NCP is important for the 3D map. Not optimizing this may yield unbound or heterogenous, non-specific binding. When a 1:1 molar ratio was used, yielding only 50% the NCP bound by the DRF (Figure 6), the 3D map did not show any additional density corresponding to the DRF. Therefore, it is important to determine the ratio that yields a single band at the same location on an EMSA and shows a single peak on the chromatogram of a sizing column, corresponding to the complex. Several parameters for freezing of the grids, including the blot time, added several rounds of optimization.
Taken together, this protocol provides detailed instructions for the first time on two different methods to prepare nucleosomal complexes for cryo-EM structural determination. The baseline parameters for the grid preparation provided can be used as a starting point. And while not a single method works equally well for all macromolecular complexes, focusing on optimizing the parameters described in this protocol will narrow down the strategy for tailored optimization of other nucleosomal complexes.

Eukaryotic DNA is organized and compacted by histone proteins, forming chromatin. The nucleosome core particle (NCP) constitutes the fundamental repeating unit of chromatin that regulates accessibility to DNA-binding proteins for DNA repair, transcription, and replication1. Although the first X-ray crystal structure of the NCP was first solved more than two decades ago2 and many more structures of the NCP have been published since3,4,5,6, DNA repair mechanisms in nucleosomal substrates have not yet been delineated. Uncovering the molecular details underlying DNA repair in chromatin will require structural characterization of the participating components to understand how local structural features of the NCP regulate DNA repair activities. This is particularly important in the context of base excision repair (BER) given that biochemical studies with BER enzymes suggest unique DNA repair mechanisms in nucleosomes that are dependent on enzyme-specific structural requirements for catalysis and the structural position of the DNA lesion within the nucleosome7,8,9,10,11,12,13. Given that BER is a vital DNA repair process, there is considerable interest to fill in these gaps while also establishing a starting point from which other technically feasible structural studies involving relevant nucleosomal complexes can be carried out.
Cryo-EM is rapidly becoming the method of choice for solving the three-dimensional (3D) structure of complexes whose large-scale preparation of homogeneous sample is challenging. Although, the design and purification of NCPs complexed with a DNA repair factor (NCP-DRF) will likely necessitate tailored optimization, the procedure presented here to generate and freeze a stable NCP-DRF complex provides details on how to optimize the sample and cryo-EM grid preparation. Two workflows (not mutually exclusive) shown in Figure 1, and the specific details in the protocol identify critical steps and provide strategies for optimizing these steps. This work will propel the chromatin and DNA repair field in a direction where complementing biochemical with structural studies becomes technically feasible to better understand the molecular mechanisms of nucleosomal DNA repair.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 M HEPES; pH 7.5</td>
			<td>Thermo Fisher Scientific</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>1 M MgCl2</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM9530G</td>
			<td></td>
		</tr>
		<tr>
			<td>10x TBE</td>
			<td>Bio-rad</td>
			<td>1610733</td>
			<td></td>
		</tr>
		<tr>
			<td>25% glutaraldehyde</td>
			<td>Fisher Scientific</td>
			<td>50-262-23</td>
			<td></td>
		</tr>
		<tr>
			<td>3 M KCl</td>
			<td>Thermo Fisher Scientific</td>
			<td>043398.K2</td>
			<td></td>
		</tr>
		<tr>
			<td>491 prep cell</td>
			<td>Bio-rad</td>
			<td>1702926</td>
			<td></td>
		</tr>
		<tr>
			<td>Amicon Ultra 15 centrifugal filter (MW cutoff 30 kDa)</td>
			<td>Millipore Sigma</td>
			<td>Z717185</td>
			<td></td>
		</tr>
		<tr>
			<td>Amicon Ultra 4 centrifugal filter (MW cutoff 30 kDa)</td>
			<td>Millipore Sigma</td>
			<td>UFC8030</td>
			<td></td>
		</tr>
		<tr>
			<td>AutoGrid Tweezers</td>
			<td>Ted Pella</td>
			<td>47000-600</td>
			<td></td>
		</tr>
		<tr>
			<td>Automatic Plunge Freezer</td>
			<td>Leica</td>
			<td>Leica EM GP</td>
			<td></td>
		</tr>
		<tr>
			<td>C-1000 touch thermocycler</td>
			<td>Bio-rad</td>
			<td>1851148</td>
			<td></td>
		</tr>
		<tr>
			<td>C-clips and rings</td>
			<td>Thermo Fisher</td>
			<td>6640--6640</td>
			<td></td>
		</tr>
		<tr>
			<td>Clipping station</td>
			<td>SubAngostrom</td>
			<td>SCT08</td>
			<td></td>
		</tr>
		<tr>
			<td>Dialysis Membrane (MW cufoff 6-8 kDa)</td>
			<td>Fisher Scientific</td>
			<td>15370752</td>
			<td></td>
		</tr>
		<tr>
			<td>Diamond Tweezers</td>
			<td>Techni-Pro</td>
			<td>758TW0010</td>
			<td></td>
		</tr>
		<tr>
			<td>dsDNA</td>
			<td>Integrated DNA techonologies</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>FEI Titan Krios</td>
			<td>Thermo Fisher</td>
			<td>KRIOSG4TEM</td>
			<td></td>
		</tr>
		<tr>
			<td>FPLC purification system</td>
			<td>AKTA Pure</td>
			<td>29018224</td>
			<td></td>
		</tr>
		<tr>
			<td>Fraction collector Model 2110</td>
			<td>Bio-rad</td>
			<td>7318122</td>
			<td></td>
		</tr>
		<tr>
			<td>Glow Discharge Cleaning System</td>
			<td>Ted Pella</td>
			<td>91000S</td>
			<td></td>
		</tr>
		<tr>
			<td>Grid Boxes</td>
			<td>SubAngostrom</td>
			<td>PB-E</td>
			<td></td>
		</tr>
		<tr>
			<td>Grid Storage Accessory Pack</td>
			<td>SubAngostrom</td>
			<td>GSAX</td>
			<td></td>
		</tr>
		<tr>
			<td>Liquid Ethane</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Liquid Nitrogen</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Minipuls 3 peristaltic two-head pump</td>
			<td>Gilson</td>
			<td>F155008</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanodrop</td>
			<td>Thermo Fisher Scientific</td>
			<td>ND-2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Novex 16%, Tricine, 1.0 mm, Mini Protein Gels</td>
			<td>Thermo Fisher Scientific</td>
			<td>EC6695BOX</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetman</td>
			<td>Gilson</td>
			<td>FA10002M</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips (VWR) Low Retention</td>
			<td>VWR</td>
			<td>76322-528</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyacrylamide gel solution (37.5:1)</td>
			<td>Bio-rad</td>
			<td>1610158</td>
			<td></td>
		</tr>
		<tr>
			<td>polyethylene glycol (PEG)</td>
			<td>Millipore Sigma</td>
			<td>P4338-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Pur-A-lyzer Maxi 3500</td>
			<td>Millipore Sigma</td>
			<td>PURX35050</td>
			<td></td>
		</tr>
		<tr>
			<td>Purified recombinant DNA repair factor</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>R 1.2/1.3 Cu 300 mesh Grids</td>
			<td>Quantifoil</td>
			<td>N1-C14nCu30-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant histone octamer</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring clipping tools</td>
			<td>SubAngostrom</td>
			<td>CSA-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Superdex 200 column 10/300</td>
			<td>Millipore Sigma</td>
			<td>GE28-9909-44</td>
			<td></td>
		</tr>
		<tr>
			<td>Transmission Electron Microscope</td>
			<td>Thermo Fisher</td>
			<td>Talos Arctica 200 kV</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers Assembly for FEI Vitrobot Mark IV-I</td>
			<td>Ted Pella</td>
			<td>47000-500</td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure Glycerol</td>
			<td>Thermo Fisher Scientific</td>
			<td>15514011</td>
			<td></td>
		</tr>
		<tr>
			<td>Vitrobot</td>
			<td>Thermo Fisher</td>
			<td>Mark IV System</td>
			<td></td>
		</tr>
		<tr>
			<td>Whatman Filter paper (55 mM)</td>
			<td>Cytiva</td>
			<td>1005-055</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylene cyanol</td>
			<td>Thermo Fisher Scientific</td>
			<td>440700500</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeba Micro Spin Desalting Columns, 7K MWCO, 75 &#181;L</td>
			<td>Thermo Fisher Scientific</td>
			<td>89877</td>
			<td></td>
		</tr>
	</tbody>
</table>

preparation of nucleosome core particles complexed with dna repair factors for cryo-electron microscopy structural determination

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of chromatin, show most DNA repair enzymes remove DNA damage at reduced rates as compared to free DNA. The molecular details on how base excision repair (BER) enzymes recognize and remove DNA damage in nucleosomes have not been elucidated. However, biochemical BER data of nucleosomal substrates suggest the nucleosome presents different structural barriers dependent on the location of the DNA lesion and the enzyme. This indicates the mechanisms employed by these enzymes to remove DNA damage in free DNA may be different than those employed in nucleosomes. Given that the majority of genomic DNA is assembled into nucleosomes, structural information of these complexes is needed. To date, the scientific community lacks detailed protocols to perform technically feasible structural studies of these complexes. Here, we provide two methods to prepare a complex of two genetically fused BER enzymes (Polymerase &#946; and AP Endonuclease1) bound to a single-nucleotide gap near the entry-exit of the nucleosome for cryo-electron microscopy (cryo-EM) structural determination. Both methods of sample preparation are compatible for vitrifying quality grids via plunge freezing. This protocol can be used as a starting point to prepare other nucleosomal complexes with different BER factors, pioneer transcription factors, and chromatin-modifying enzymes.

Properly assembled NCPs (Figure 2) were used to make a complex with a recombinant fusion protein of MBP-Pol&#946;-APE1 (Figure 3). To determine the ratio of NCP to MBP-Pol&#946;-APE1 to form a stable complex, we performed electrophoretic mobility shift assays (EMSA) (Figure 4), which showed a singly shifted band of the NCP with 5-fold molar excess of MBP-Pol&#946;-APE1. During the optimization of making this complex, crosslinking with glutaraldehyde was critical to prevent the NCP from falling apart. Initially, assembly of the complex of NCP-Pol&#946;-APE1 was performed in a smaller volume, at approximately 10 &#181;M NCP, using 0.005% final glutaraldehyde concentration. Under these conditions, the sample was overly crosslinked (Figure 5D-E) and resulted in aggregates without discernable individual complexes. Crosslinking at approximately 10-fold diluted NCPs (1.2 &#181;M NCP) resulted in significantly reduced aggregation and improved particle stability (Figure 5A-C). Figure 6 illustrates that both methods shown in Figure 1 can be combined with the preparative gel improving the quality of the NCPs when the NCP contains a significant amount of high molecular weight (HMW) aggregates (Figure 6A), followed by the purification of the complex via size exclusion. Although this showed great success in the preparation of the sample (Figure 5B-C) and grids, yielding stable particles (Figure 6D), the suboptimal formation of the complex (Figure 6A) yielded a 3D map of the NCP alone (data not shown). These results show that both methods (size exclusion and preparative gel) can be used independently to generate stable complexes (Figure 7A-C). Indeed, data have been collected from both grids and show almost identical 2D classes (Figure 7D) and 3D maps (Figure 7E) at approximately 3.2 &#197; resolution.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64061/64061fig01.jpg" /> 
Figure 1: Workflow of the two methods presented in this protocol to prepare and freeze an NCP-Pol&#946;-APE1 complex via 1) preparative gel and 2) size exclusion. Although not shown here, these methods can be combined (Figure 6), starting with (1) and purifying the concentrated complex from (1) using a sizing column. This will require doubling the starting material of NCPs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64061/64061fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64061/64061fig02.jpg" /> 
Figure 2: Representative NCP reconstitutions. (A) DNA was titrated with increasing amounts of histone octamer at ratios of 1:0.9, 1:1.1, 1:1.2, 1:2, DNA:octamer. (B) Duplicate large-scale assembled NCPs (1:2, DNA:octamer). Reconstitutions were electrophoresed in a 6% nondenaturing polyacrylamide gel, followed with sybr green I staining. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64061/64061fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64061/64061fig03.jpg" /> 
Figure 3: Schematic of genetically fused Pol&#946;-APE1 with MBP at its N-terminal. MBP-Pol&#946;-APE1 fusion protein was enzymatically characterized for both activities (data not shown). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64061/64061fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64061/64061fig04.jpg" /> 
Figure 4: MBP-Pol&#946;-APE1 binding to NCPs. Nucleosomal substrate (100 nM) was incubated with increasing amounts of MBP-Pol&#946;-APE1 for 15 min on ice. Bound and unbound NCP were separated in a 6% polyacrylamide nondenaturing gel, followed by sybr green I staining. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64061/64061fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64061/64061fig05.jpg" /> 
Figure 5: Analysis of successful and unsuccessful crosslinking results. The elution profile of the complex is shown in (A) with the peak of interest labeled. Fractions, denoted by &#34;F&#34;, collected from this elution were analyzed in a 16% tricine polyacrylamide gel and shown in (B). (C) Sample from (B) was frozen using holey carbon support grids and imaged using the 200 kV field emission cryo-transmission electron microscope. Corresponding unsuccessful experiments are shown in (D), (E), and (F). Note these unsuccessful experiments contain the fusion protein without MBP. HOc and HOD denote histone octamer concentrated and diluted, respectively; RT denotes room temperature; &#34;X&#34; corresponds to crosslinking with glutaraldehyde. Scale bars = 100 nm (C, F). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64061/64061fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64061/64061fig06.jpg" /> 
Figure 6: Preparation of NCP-Pol&#946;-APE1 complex using the two methods in tandem. (A) Unbound and bound (1:1, NCP- Pol&#946;-APE1) were analyzed in 6% polyacrylamide nondenaturing gel (note that at this ratio only 50% is bound). (B) Elution profile of NCP-Pol&#946;-APE1 complex from a sizing column. (C) Analysis of eluted fractions in a 16% tricine polyacrylamide gel; fractions were pooled and concentrated, as indicated. (D) Sample was frozen and imaged as described in Figure 5B. Data processing did not show an additional density corresponding to Pol&#946;-APE1 (not shown). &#34;X&#34; in lanes 5 and 6 denotes crosslinking with glutaraldehyde as indicated in the protocol. Scale bar = 100 nm (D). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64061/64061fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/64061/64061fig07.jpg" /> 
Figure 7: Analysis of NCP-MBP-Pol&#946;-APE1 complex. (A) Samples from the different steps in the methods illustrated in Figure 1 were electrophoresed in a 6% polyacrylamide nondenaturing gel. Notice that complex formation is reproducible by the two methods. (B,C) Representative grid holey carbon support grid images taken using a 300 kV transmission electron microscope where the red boxes highlight different orientations of the complex (some of which correspond to the 2D classes shown in (D). (E) Data processed from (C) show a 3D map at 3.2 &#197; resolution of the NCP-MBP-Pol&#946;-APE1 complex.). Scale bars = 100 nm (B, C). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64061/64061fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Oligonucleotide name</td>
			<td colspan="3">Sequence (5&#39;--&#62;3&#39;)</td>
			<td>Purification scale</td>
			<td>Method of purificaiton</td>
			<td>Number of vials ordered</td>
		</tr>
		<tr>
			<td>UND complementary strand</td>
			<td colspan="3">TGATGGACCCTATACGCGGCCGCCCTGGAGAAT 
			CCCGGTGCCGAGGCCGCTCAATTGGTCGTAGA 
			CAGCTCTAGCACCGCTTAAACGCACGTACGCGC 
			TGTCCCCCGCGTTTTAACCGCCAAGGGGATTAC 
			TCCCTAGTCTCCAGGCACGTGTCAGATATCAAC 
			ATCCTGTGCATGTATTGAACAGCGACCTTGCCG</td>
			<td>&#160;20 nmole</td>
			<td>PAGE purified</td>
			<td>6</td>
		</tr>
		<tr>
			<td>161mer</td>
			<td colspan="3">/5Phos/GATATCTGACACGTGCCTGGAGACTAGG 
			GAGTAATCCCCTTGGCGGTTAAAACGCGGGGG 
			ACAGCGCGTACGTGCGTTTAAGCGGTGCTAGA 
			GCTGTCTACGACCAATTGAGCGGCCTCGGCA 
			CCGGGATTCTCCAGGGCGGCCGCGTATAGGG 
			TCCATCA</td>
			<td>20 nmole</td>
			<td>PAGE purified</td>
			<td>4</td>
		</tr>
		<tr>
			<td>35mer</td>
			<td colspan="3">CGGCAAGGTCGCTGTTCAATACATGCACAGG 
			ATGT</td>
			<td>250 nmole</td>
			<td>HPLC</td>
			<td>1</td>
		</tr>
	</tbody>
</table>
Table 1: DNA substrate. Three oligonucleotides were annealed to generate the dsDNA substrate. The underlined sections are 25 bp of linker DNA flanking the 147 bp 601 DNA positioning sequence containing a single nucleotide gap.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Step</td>
			<td>Temperature (&#176;C)</td>
			<td>Time (min)/step</td>
		</tr>
		<tr>
			<td>1</td>
			<td>95</td>
			<td>10</td>
		</tr>
		<tr>
			<td>2-5</td>
			<td>90, 85, 80, 75</td>
			<td>5</td>
		</tr>
		<tr>
			<td>6-41</td>
			<td>69 decrease by 1</td>
			<td>3</td>
		</tr>
		<tr>
			<td>42</td>
			<td>4</td>
			<td>hold</td>
		</tr>
	</tbody>
</table>
Table 2: Annealing temperature gradient. Oligos were incubated in a thermocycler at these temperatures to generate the dsDNA substrate.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Components (stock)</td>
			<td>Volume/amount</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>3M KCl</td>
			<td>167 mL</td>
			<td>0.250 M</td>
		</tr>
		<tr>
			<td>1 M HEPES, pH 8.0</td>
			<td>20 mL</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>0.5 M EDTA</td>
			<td>4 mL</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>1 M DTT</td>
			<td>2 mL</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>CHAPS</td>
			<td>2 g</td>
			<td>1.6 mM</td>
		</tr>
		<tr>
			<td colspan="3">QS with dH2O to make 2L</td>
		</tr>
	</tbody>
</table>
Table 3: RBlow reconstitution buffer. Instructions to make 2 L as previously reported14, except 1.6 mM CHAPS was added.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Components (stock)</td>
			<td>Volume/amount</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>3M KCl</td>
			<td>267 mL</td>
			<td>2 M</td>
		</tr>
		<tr>
			<td>1 M HEPES, pH 8.0</td>
			<td>4 mL</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>0.5 M EDTA</td>
			<td>800 &#956;L</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>1 M DTT</td>
			<td>400 &#956;L</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>CHAPS</td>
			<td>0.4 g</td>
			<td>1.6 mM</td>
		</tr>
		<tr>
			<td colspan="3">QS with dH2O to make 400 mL</td>
		</tr>
	</tbody>
</table>
Table 4: RBhigh reconstitution buffer. Instructions to make 400 mL as previously reported14, except 1.6 mM CHAPS was added.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Components (stock)</td>
			<td>Volume</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>3M KCl</td>
			<td>16.7 mL</td>
			<td>50 mM</td>
		</tr>
		<tr>
			<td>1 M HEPES, pH 8.0</td>
			<td>10 mL</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>0.5 M EDTA</td>
			<td>2 mL</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>1 M DTT</td>
			<td>1 mL</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td colspan="3">QS with dH2O to make 1L</td>
		</tr>
	</tbody>
</table>
Table 5:&#160;RB50mM reconstitution buffer. Instructions to make 1 L of reconstitution buffer compatible for freezing.

Watch this Scientific Journal Video about Preparation of Nucleosome Core Particles Complexed with DNA Repair Factors for Cryo-Electron Microscopy Structural Determination at JoVE.com

Preparation of Nucleosome Core Particles Complexed with DNA Repair Factors for Cryo-Electron Microscopy Structural Determination

This protocol outlines in detail the preparation of nucleosomal complexes using two methods of sample preparation for freezing TEM grids.

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of ...

preparation-nucleosome-core-particles-complexed-with-dna-repair

Nanyang Technological University

Research

JoVE Journal

Biochemistry

1.7K Views.  National Institute of Environmental Health Sciences, National Institutes of Health. This protocol outlines in detail the preparation of nucleosomal complexes using two methods of sample preparation for freezing TEM grids.

Video: Preparation of Nucleosome Core Particles Complexed with DNA Repair Factors for Cryo-Electron Microscopy Structural Determination

Expression and Purification of the Human Lipid-sensitive Cation Channel TRPC3 for Structural Determination by Single-particle Cryo-electron Microscopy

Transient receptor potential channels (TRPCs) of the canonical TRP subfamily are nonselective cation channels that play an essential role in calcium homeostasis, particularly store-operated calcium entry, which is critical to maintaining proper function of synaptic vesicle release and intracellular signaling pathways. Accordingly, TRPC channels have been implicated in a variety of human diseases including cardiovascular disorders such as cardiac hypertrophy, neurodegenerative disorders such as Parkinson&#39;s disease, and neurologic disorders such as spinocerebellar ataxia. Therefore, TRPC channels represent a potential pharmacologic target in human diseases. However, the molecular mechanisms of gating in these channels are still unclear. The difficulty in obtaining large quantities of stable, homogeneous, and purified protein has been a limiting factor in structure determination studies, particularly for mammalian membrane proteins such as the TRPC ion channels. Here, we present a protocol for the large-scale expression of mammalian ion channel membrane proteins using a modified baculovirus gene transfer system and the purification of these proteins by affinity and size-exclusion chromatography. We further present a protocol to collect single-particle cryo-electron microscopy images from purified protein and to use these images to determine the protein structure. Structure determination is a powerful method for understanding the mechanisms of gating and function in ion channels.

This protocol describes techniques used to determine ion channel structures by cryo-electron microscopy, including a baculovirus system used to efficiently express genes in mammalian cells with minimum effort and toxicity, protein extraction, purification, and quality checking, sample grid preparation and screening, as well as data collection and processing.

Transient receptor potential channels (TRPCs) of the canonical TRP subfamily are nonselective cation channels that play an essential role in calcium ...

Enrichment of Native Lipoprotein Particles with microRNA and Subsequent Determination of Their Absolute/Relative microRNA Content and Their Cellular Transfer Rate

Lipoprotein particles are predominately transporters of lipids and cholesterol in the bloodstream. Furthermore, they contain small amounts of strands of noncoding microRNA (miRNA). In general, miRNA alters the protein expression profile due to interactions with messenger-RNA (mRNA). Thus, knowledge of the relative and absolute miRNA content of lipoprotein particles is essential to estimate the biological effect of cellular particle uptake. Here, a quantitative real-time polymerase chain reaction (qPCR)-based protocol is presented to determine the absolute miRNA content of lipoprotein particles&#8212;exemplified shown for native and miRNA-enriched lipoprotein particles. The relative miRNA content is quantified using multiwell microfluidic array cards. Furthermore, this protocol allows scientists to estimate the cellular miRNA and, thus, the lipoprotein particle uptake rate. A significant increase of the cellular miRNA level is observable when using high-density lipoprotein (HDL) particles artificially loaded with miRNA, whereas incubation with native HDL particles yields no significant effect due to their rather low miRNA content. In contrast, the cellular uptake of low-density lipoprotein (LDL) particles&#8212;neither with native miRNA nor artificially loaded with it&#8212;did not alter the cellular miRNA level.

Here, a quantitative real-time polymerase chain reaction-based protocol is presented for the determination of the native micro-RNA content (absolute/relative) of lipoprotein particles. In addition, a method for increasing the micro-RNA level, as well as a method for determining the cellular uptake rate of lipoprotein particles, is demonstrated.

Lipoprotein particles are predominately transporters of lipids and cholesterol in the bloodstream. Furthermore, they contain small amounts of strands of ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

Rapid Determination of Antibody-Antigen Affinity by Mass Photometry

Measurements of the specificity and affinity of antigen-antibody interactions are critically important for medical and research applications. In this protocol, we describe the implementation of a new single-molecule technique, mass photometry (MP), for this purpose. MP is a label- and immobilization-free technique that detects and quantifies molecular masses and populations of antibodies and antigen-antibody complexes on a single-molecule level. MP analyzes the antigen-antibody sample within minutes, allowing for the precise determination of the binding affinity and simultaneously providing information on the stoichiometry and the oligomeric state of the proteins. This is a simple and straightforward technique that requires only picomole quantities of protein and no expensive consumables. The same procedure can be used to study protein-protein binding for proteins with a molecular mass larger than 50 kDa. For multivalent protein interactions, the affinities of multiple binding sites can be obtained in a single measurement. However, the single-molecule mode of measurement and the lack of labeling imposes some experimental limitations. This method gives the best results when applied to measurements of sub-micromolar interaction affinities, antigens with a molecular mass of 20 kDa or larger, and relatively pure protein samples. We also describe the procedure for performing the required fitting and calculation steps using basic data analysis software.

We describe a single-molecule approach to antigen-antibody affinity measurements using mass photometry (MP). The MP-based protocol is fast, accurate, uses a very small amount of material, and does not require protein modification.

Measurements of the specificity and affinity of antigen-antibody interactions are critically important for medical and research applications. In this ...

Fast Grid Preparation for Time-Resolved Cryo-Electron Microscopy

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures and information on more challenging systems. Sample preparation for cryo-EM is undergoing a similar revolution with new approaches being developed to supersede the traditional blotting systems. These include the use of piezo-electric dispensers, pin printing and direct spraying. As a result of these developments, the speed of grid preparation is going from seconds to milliseconds, providing new opportunities, especially in the field of time-resolved cryo-EM where proteins and substrates can be rapidly mixed before plunge freezing, trapping short lived intermediate states. Here we describe, in detail, a standard protocol for making grids on our in-house time-resolved EM device both for standard fast grid preparation and also for time-resolved experiments. The protocol requires a minimum of about 50 &#181;L sample at concentrations of &#8805; 2 mg/mL for the preparation of 4 grids. The delay between sample application and freezing can be as low as 10 ms. One limitation is increased ice thickness at faster speeds and compared to the blotting method. We hope this protocol will aid others in designing their own grid making devices and those interested in designing time-resolved experiments.

Here, we provide a detailed protocol for the use of a rapid grid making device for both fast grid-making and for rapid mixing and freezing to conduct time-resolved experiments.

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures ...

Preparation of Sample Support Films in Transmission Electron Microscopy using a Support Floatation Block

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant bottleneck. Macromolecular samples are ideally imaged directly from random orientations in a thin layer of vitreous ice. However, many samples are refractory to this, and protein denaturation at the air-water interface is a common problem. To overcome such issues, support films-including amorphous carbon, graphene, and graphene oxide-can be applied to the grid to provide a surface which samples can populate, reducing the probability of particles experiencing the deleterious effects of the air-water interface. The application of these delicate supports to grids, however, requires careful handling to prevent breakage, airborne contamination, or extensive washing and cleaning steps. A recent report describes the development of an easy-to-use floatation block that facilitates wetted transfer of support films directly to the sample. Use of the block minimizes the number of manual handling steps required, preserving the physical integrity of the support film, and the time over which hydrophobic contamination can accrue, ensuring that a thin film of ice can still be generated. This paper provides step-by-step protocols for the preparation of carbon, graphene, and graphene oxide supports for EM studies.

Sample preparation for cryo-electron microscopy (cryo-EM) is a significant bottleneck in the structure determination workflow of this method. Here, we provide detailed methods for using an easy-to-use, three-dimensionally printed block for the preparation of support films to stabilize samples for transmission EM studies.

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant ...

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in situ. Owing to the strong interaction of an electron with the matter, high-resolution cryo-ET studies are limited to specimens with a thickness of less than 200 nm, which restricts the applicability of cryo-ET only to the peripheral regions of a cell. A complex workflow that comprises the preparation of thin cellular cross-sections by cryo-focused ion beam micromachining (cryo-FIBM) was introduced during the last decade to enable the acquisition of cryo-ET data from the interior of larger cells. We present a protocol for the preparation of cellular lamellae from a sample vitrified by plunge freezing utilizing Saccharomyces cerevisiae as a prototypical example of a eukaryotic cell with wide utilization in cellular and molecular biology research. We describe protocols for vitrification of S. cerevisiae into isolated patches of a few cells or a continuous monolayer of the cells on a TEM grid and provide a protocol for lamella preparation by cryo-FIB for these two samples.

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

We present a protocol for lamella preparation of plunge frozen biological specimens by cryo-focused ion beam micromachining for high-resolution structural studies of macromolecules in situ with cryo-electron tomography. The presented protocol provides guidelines for the preparation of high-quality lamellae with high reproducibility for structural characterization of macromolecules inside the&#160;Saccharomyces cerevisiae.

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in ...

Single Particle Cryo-Electron Microscopy: From Sample to Structure

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The overall process involves i) vitrifying the specimen in a thin film supported on a cryoEM grid; ii) screening the specimen to assess particle distribution and ice quality; iii) if the grid is suitable, collecting a single particle dataset for analysis; and iv) image processing to yield an EM density map. In this protocol, an overview for each of these steps is provided, with a focus on the variables which a user can modify during the workflow and the troubleshooting of common issues. With remote microscope operation becoming standard in many facilities, variations on imaging protocols to assist users in efficient operation and imaging when physical access to the microscope is limited will be described.

Structure determination of macromolecular complexes using cryoEM has become routine for certain classes of proteins and complexes. Here, this pipeline is summarized (sample preparation, screening, data acquisition and processing) and readers are directed towards further detailed resources and variables that may be altered in the case of more challenging specimens.

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The ...

Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules present in the cellular context and preserved in a near-native frozen-hydrated state. However, there are challenges associated with culturing and/or adhering cells onto TEM grids in a manner that is suitable for tomography while retaining the cells in their physiological state. Here, a detailed step-by-step protocol is presented on the use of micropatterning to direct and promote eukaryotic cell growth on TEM grids. During micropatterning, cell growth is directed by depositing extra-cellular matrix (ECM) proteins within specified patterns and positions on the foil of the TEM grid while the other areas remain coated with an anti-fouling layer. Flexibility in the choice of surface coating and pattern design makes micropatterning broadly applicable for a wide range of cell types. Micropatterning is useful for studies of structures within individual cells as well as more complex experimental systems such as host-pathogen interactions or differentiated multi-cellular communities. Micropatterning may also be integrated into many downstream whole-cell cryo-ET workflows, including correlative light and electron microscopy (cryo-CLEM) and focused-ion beam milling (cryo-FIB).

The goal of this protocol is to direct cell adhesion and growth to targeted areas of grids for cryo-electron microscopy. This is achieved by applying an anti-fouling layer that is ablated in user-specified patterns followed by deposition of extra-cellular matrix proteins in the patterned areas prior to cell seeding.

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules ...

A Robust Single-Particle Cryo-Electron Microscopy (cryo-EM) Processing Workflow with cryoSPARC, RELION, and Scipion

Recent advances in both instrumentation and image processing software have made single-particle cryo-electron microscopy (cryo-EM) the preferred method for structural biologists to determine high-resolution structures of a wide variety of macromolecules. Multiple software suites are available to new and expert users for image processing and structure calculation, which streamline the same basic workflow: movies acquired by the microscope detectors undergo correction for beam-induced motion and contrast transfer function (CTF) estimation. Next, particle images are selected and extracted from averaged movie frames for iterative 2D and 3D classification, followed by 3D reconstruction, refinement, and validation. Because various software packages employ different algorithms and require varying levels of expertise to operate, the 3D maps they generate often differ in quality and resolution. Thus, users regularly transfer data between a variety of programs for optimal results. This paper provides a guide for users to navigate a workflow across the popular software packages: cryoSPARC v3, RELION-3, and Scipion 3 to obtain a near-atomic resolution structure of the adeno-associated virus (AAV). We first detail an image processing pipeline with cryoSPARC v3, as its efficient algorithms and easy-to-use GUI allow users to quickly arrive at a 3D map. In the next step, we use PyEM and in-house scripts to convert and transfer particle coordinates from the best quality 3D reconstruction obtained in cryoSPARC v3 to RELION-3 and Scipion 3 and recalculate 3D maps. Finally, we outline steps for further refinement and validation of the resultant structures by integrating algorithms from RELION-3 and Scipion 3. In this article, we describe how to effectively utilize three processing platforms to create a single and robust workflow applicable to a variety of data sets for high-resolution structure determination.

A Robust Single-Particle Cryo-Electron Microscopy cryo-EM Processing Workflow with cryoSPARC, RELION, and Scipion

This article describes how to effectively utilize three cryo-EM processing platforms, i.e., cryoSPARC v3, RELION-3, and Scipion 3, to create a single and robust workflow applicable to a variety of single-particle data sets for high-resolution structure determination.

Recent advances in both instrumentation and image processing software have made single-particle cryo-electron microscopy (cryo-EM) the preferred method ...