We thank Dr. Till B&#246;cking for the original single molecule preparation protocol. 
This work was supported by NSF grant 1121972(SS).

1. Chemistry Preparation
<ol>
	<li>Preparation and storage of PLL-g-PEG-Biotin and avidin:
	<ol>
		<li>Dissolve the polymer PLL-g-PEG-biotin in PBS buffer according to manufacturer&#39;s directions at a concentration of 1.0 mg/ml.</li>
		<li>Aliquot according to coverslip size (see Table 1) and store at -80 &#176;C for up to one year.</li>
	</ol>
	</li>
	<li>Preparation and storage of Avidin/NeutrAvidin
	<ol>
		<li>Dissolve the unlabeled Avidin/NeutrAvidin in PBS buffer according to the manufacturer&#39;s directions at a concentration of 1.0 mg/ml.</li>
		<li>Aliquot according to coverslip size (see Table&#160;1) and store at -20 &#176;C for up to 3 years.</li>
	</ol>
	</li>
	<li>Virus sample preparation: Prepare virus samples according to experimental need and lab protocol prior to this assay. Utilize step 1.3.1 for method 5A, or 1.3.2 for method 5B.
	<ol>
		<li>Prepare virus samples for method 5A in advance of stage 2. Utilize prepared viral samples within 1-2 days of preparation, or aliquot according to cover slip size (see Table&#160;1) then flash freeze in liquid nitrogen and store at -80 &#176;C for up to one year. When thawed, use immediately.</li>
		<li>Prepare virus samples for method 5B similar to 1.3.1. Thaw aliquots of virus in advance of 5B and store at 4 &#176;C no more than 24 hr&#160;prior to mixing with antibodies.</li>
	</ol>
	</li>
	<li>Prepare appropriate biotinylated antibodies for the experiment.</li>
</ol>
2. Coverslip Cleaning in Preparation for Chemical Treatment
<ol>
	<li>Place coverslips in an appropriately sized Teflon coverslip rack that holds them in a vertical orientation and submerge them in a beaker filled with filtered (0.2 &#956;m) ethyl alcohol (190 proof).</li>
	<li>Sonicate the coverslips in the filtered ethyl alcohol (190 proof) for 30 min.</li>
	<li>Remove coverslips from alcohol and rinse coverslips extensively with ultrapure water.</li>
	<li>Place rinsed coverslips in a separate clean Teflon rack and beaker filled with 1 M NaOH.</li>
	<li>Sonicate the coverslips in the 1 M NaOH for 30 min.</li>
	<li>Remove coverslips from NaOH and rinse them extensively with ultrapure water.</li>
	<li>Dry the rinsed coverslips with a nitrogen stream, and place them in a clean, dry Teflon rack.</li>
	<li>Inspect the coverslips for any visible film or particles which remain on the coverslip. If any visible material remains, the coverslip has not been properly cleaned and rinsed. If not properly cleaned repeat steps 2.1-2.7.</li>
	<li>Clean the coverslips with oxygen-plasma for 2 min. This step is optional but recommended.</li>
</ol>
3. Formation of the PLL-g-PEG Thin Film Layer
<ol>
	<li>Immediately after drying in the nitrogen stream (or the optional oxygen-plasma), place one coverslip horizontally in a sterile Petri dish. Pipet in the center of the coverslip an appropriate amount of thawed PBS buffered PLL-g-PEG-biotin (see Table&#160;1).</li>
	<li>Place another cleaned coverslip atop the fluid in a sandwich method. Note that this method consists of having two coverslips incubate their chemical films by orienting their chemically treated &#34;active faces&#34; towards each other, separated only by the current chemical layer. Make sure there is sufficient fluid such that the volume between the two coverslips is completely filled with fluid, but that no fluid leaks out from between the coverslips (see&#160;Table&#160;1). The &#34;active face&#34; henceforth designates coverslip faces which have been in contact with the fluid in the sandwich configuration.</li>
	<li>Place the lid of the Petri dish over the coverslip sandwich and incubate at RT for 45-60 min.</li>
	<li>Remove lid of Petri dish and pick up the sandwich with a pair of tweezers. Be careful not to pinch out excess fluid. Use the thumb and index finger to pinch the sandwich lightly and slide the coverslips in opposite directions horizontally with respect to one another, then separate the two coverslips from each other without touching the active faces.</li>
	<li>Rinse both active faces with ultrapure water by pipetting 25 ml of ultrapure water over the active face 2-4x, and then dry the coverslips with a nitrogen stream. Be sure to note which coverslip faces are active.</li>
	<li>Utilize the coverslips immediately, or store O/N&#160;with the active face up in a sterile Petri dish placed in a dust free low humidity environment.</li>
</ol>
4. Avidin Binding Enhancement
<ol>
	<li>Place a PLL-g-PEG treated coverslip with active face up in a sterile Petri dish.</li>
	<li>Pipette an appropriate amount of thawed avidin in buffer (prepared in step 1.2) onto the middle of the active face.</li>
	<li>Place another coverslip (active face downward) atop the fluid in the sandwich method. Note the active face of the coverslips. It is important that the active faces be in contact with the fluid during the incubation (see step 3.2).</li>
	<li>Place the lid of the Petri dish over the coverslip sandwich and incubate at RT for 25-30 min.</li>
	<li>Remove the Petri dish lid and separate the two coverslips as described in step 3.4, making sure to keep track of and not touch the active faces.</li>
	<li>Rinse both active faces with ultrapure water by pipetting 25 ml of ultrapure water over the active face 2-4x and then dry the coverslips with a nitrogen stream. Note the active face of the coverslips. Use immediately for the next stage.</li>
</ol>
IMPORTANT NOTE: There are two distinct methods for completing sample preparation depending on the type of assay the experiment entails. The experimenter should proceed with stage 5A in which the virus is anchored to the glass before any additional antibody treatment required for super-resolution imaging is added. 5B describes an alternative method of labeling the virus in solution before anchoring it to the glass. The representative data in this manuscript is prepared using 5A. An appropriate method should be selected according to experimental assay type.
5A. Active Face Antibody Tether
<ol>
	<li>Immediately after drying with a nitrogen stream in stage 4.6, place an avidin treated coverslip active face up in a sterile Petri dish.</li>
	<li>Pipette an appropriate amount of thawed biotinylated antibody in buffer onto the middle of the active face (see Table&#160;1).</li>
	<li>Place another coverslip (active face downward) atop the fluid in a sandwich method. Note the active face of the coverslips, it is important the active faces are in contact with the fluid for incubation (see step 3.2.).</li>
	<li>Place the lid of the Petri dish over the coverslip sandwich and incubate at RT for 45-60 min.</li>
	<li>Remove the Petri dish lid and separate the two coverslips as described in step 3.4, making sure to keep track of and not touch the active faces.</li>
	<li>Rinse both active faces with PBS by pipetting 25 ml of PBS over the active face 2-4x, do not dry and use immediately. Be sure to keep track of which coverslip faces are active.</li>
	<li>Place an antibody treated coverslip active face up in a sterile Petri dish, and pipette an appropriate amount of thawed virus in buffer onto the middle of the active face.</li>
	<li>Place another coverslip (active face downward) atop the fluid in a sandwich method. Note the active face, as it is important the active faces are in contact with the fluid for incubation (see step 3.2.).</li>
	<li>Place the lid of the Petri dish over the coverslip sandwich and incubate at RT for 45-60 min.</li>
	<li>Remove the Petri dish lid and separate the two coverslips as described in step 3.4, making sure to keep track of and not touch the active faces.</li>
	<li>Rinse both active faces with PBS by pipetting 25 ml of PBS over the active face 2-4x. Do not dry the coverslips, instead place them in a Teflon rack and beaker filled with PBS immediately. Be sure to keep track of which coverslip faces are active.</li>
	<li>Utilize the prepared samples immediately or store in an appropriate buffer at 4 &#176;C for up to three days without significant loss of integrity. 
	 
	Important NOTE: If samples are to be used in an AFM assay, the preparation is complete and they can be utilized immediately or stored as described in step 5A.20 -&#160;5A.21. If samples are to be utilized in a super-resolution fluorescence imaging assay, proceed to the antibody labeling described in steps 5A.12 -&#160;5A.19.</li>
	<li>Place both virus treated coverslips active face up in a sterile Petri dish and pipet enough protein blocking buffer on the coverslips to bead enough fluid volume to cover the central 95% of the coverslip without flowing any blocking buffer off the coverslip (typically 100 - 200 &#956;l.) Incubate at RT for 60-90 min.</li>
	<li>Carefully remove blocking buffer by removing the coverslips one at a time from the Petri dish and pouring the fluid off of the coverslip into a waste beaker. Be careful not to drop the coverslip.</li>
	<li>Rinse both active faces with PBS by pipetting 25 ml of PBS over the active face 2-4x, do not dry and use immediately. Be sure to keep track of which coverslip faces are active.</li>
	<li>Place a single blocking buffer treated coverslip active face up in a sterile Petri dish, and pipette an appropriate amount of fluorescently labeled viral antibody in buffer onto the middle of the active face.</li>
	<li>Place another coverslip (active face downward) on top the fluid in a sandwich method. Note which face is active, it is important that the active faces are in contact with the fluid for incubation (see step 3.2).</li>
	<li>Place the lid of the Petri dish over the coverslip sandwich and incubate at RT for 30 min.</li>
	<li>Remove the Petri dish lid and separate the two coverslips as described in step 3.4, making sure to keep track of and not touch the active faces.</li>
	<li>Rinse both active faces with PBS by pipetting 25 ml of PBS over the active face 2-4x, do not dry and use immediately. Be sure to keep track of which coverslip faces are active.</li>
	<li>Do not dry the coverslips, instead place them in a Teflon rack within a beaker filled with PBS. Be sure to keep track of which coverslip faces are active.</li>
	<li>Utilize the prepared samples immediately or store in an appropriate buffer at 4 &#176;C for up to three days without significant loss of integrity.</li>
</ol>
5B. In Solution Antibody Attack
This method is an optional enhancement of the labeling strategy for the super-resolution fluorescent imaging. By applying the in solution attack, better antibody coating on the viral surface can be achieved.
<ol>
	<li>Mix antibody solutions in ratios according to experiments need. Example 1:4 ratio of 0.01 mg/ml biotinylated antibody to 0.01 mg/ml fluorescently labeled antibody will give a good binding affinity and allow for good envelope recovery in a super-resolution imaging assay.</li>
	<li>Mix equal portions of the antibody and virus solutions respectively prepared in (5B.1) and (1.3). Mix gently by pipetting up and down a few times in the aliquot. This resultant solution can be stored until needed for up to 24 hr&#160;at 4 &#176;C.</li>
	<li>Upon completion of step 4.6, place an avidin treated coverslip with active face up in a sterile Petri dish.</li>
	<li>Pipette an appropriate amount of virus with antibody solution (prepared in step 5B.2) onto the middle of the active face.</li>
	<li>Place another coverslip (active face downward) atop the fluid in a sandwich method. Note the active face, as it is important the active faces are in contact with the fluid for incubation (see step 3.2).</li>
	<li>Place the lid of the Petri dish over the coverslip sandwich and incubate at RT for 45-60 min.</li>
	<li>Remove the Petri dish lid and separate the two coverslips as described in step 3.4, making sure to keep track of and not touch the active faces.</li>
	<li>Rinse both active faces with PBS by pipetting 25 ml of PBS over the active face 2-4x. Do not dry the coverslips, instead place them in a Teflon rack and beaker filled with PBS immediately. Be sure to keep track of which coverslip faces are active.</li>
	<li>Utilize prepared samples immediately or store at 4 &#176;C for up to three days without significant loss of integrity.</li>
</ol>
6. Atomic Force Microscopy Material Property Measurement
<ol>
	<li>Turn on the AFM lasers and open the AFM software. Select the appropriate scanning mode for the desired assay from the opening dialog. All necessary windows should open by default; if not consult your user&#39;s manual in order to open the desired windows.</li>
	<li>Select a cantilever appropriate to the desired measurement and insert it into the cantilever housing according to manufacturer guidelines. There are a wide range of conditions and experiment types for which there are a variety of cantilevers. The experimenter should research cantilevers according to their need.</li>
	<li>Insert the cantilever housing into the AFM head according to manufacturer&#39;s guidelines.</li>
	<li>If the sample is to be used under wet conditions, remove the sample from the storage buffer and proceed to step 6.7.</li>
	<li>If the sample is to be used under ambient (dry) conditions, remove the sample from its storage buffer and rinse it briefly by dipping in a beaker of ultrapure water. Note: This will leave a thin hydration layer on the sample which may induce capillary attraction with cantilevers having a low spring constant.</li>
	<li>If the sample is to be utilized in ambient (dry) conditions, gently dry the sample with a nitrogen stream.</li>
	<li>Gently dry the back side of the coverslip with a dust free filter paper, making sure to not touch the active face of the sample.</li>
	<li>Place the sample in the appropriate AFM sample holder, then place the sample on the AFM stage.</li>
	<li>If the sample is to be imaged under wet conditions, pipette a small amount of buffer onto the center of the sample (~10 &#956;l).</li>
	<li>Place the AFM head over the sample.</li>
	<li>If the sample is wet, make sure that the cantilever and AFM tip penetrate the surface layer, and that there are no bubbles between the cantilever and cantilever housing.</li>
	<li>Align the AFM laser with the cantilever in accordance with manufacturer recommendations in order to obtain the optimal signal.</li>
	<li>Measure cantilever resonant frequency and set the scan frequency according to experiment type.</li>
	<li>Lower the AFM tip to the surface and optimize signal settings.</li>
	<li>Perform a 20 &#956;m square AFM scan in the mode of your choosing. Tapping (AC) mode was used in generating representative data. Identify virus candidates by approximate height, which can typically be seen as sharp spikes on the glass surface.</li>
	<li>Select a virus candidate and center the scan head above it.</li>
	<li>Perform a 250 nm square scan (or appropriate super-resolution scan) about the selected virus in order to obtain its topology and orientation.</li>
	<li>Select a point on the virus for the material property measurement (e.g. the center of a spherical virus for a Young&#39;s modulus measurement) and perform the measurement according to manufacturer&#39;s directions.</li>
</ol>
7. Super-resolution Imaging Method
<ol>
	<li>Open the super-resolution imaging software and calibrate the software according to manufacturer&#39;s guidelines.</li>
	<li>Calibrate the super-resolution imaging equipment in accordance with manufacturer&#39;s directions utilizing fluorescent beads.</li>
	<li>Assemble the sample imaging buffer from stocks just prior to imaging by combining 50 &#956;l&#160;of 1 M MEA and 100 &#956;l&#160;of 10x Gloxy to 850 &#956;l&#160;of stock buffer. Keep imaging buffer on ice for the duration of the experiment.</li>
	<li>Remove a prepared coverslip with sample from its storage buffer and rinse it 5x by dipping it in a beaker of ultrapure water.</li>
	<li>Dry the nonactive coverslip face utilizing a dust free filter paper. Make sure to not touch the active face.</li>
	<li>Insert into an imaging system sample holder and add 250 &#956;l&#160;of imaging buffer to the sample holder. It is reasonable to exchange the imaging buffer on the sample every 2 hr&#160;to maintain consistent results.</li>
	<li>Cover the sample holder with parafilm to decrease atmospheric interaction with the room environment.</li>
	<li>Place the sample holder into the imaging equipment for imaging.</li>
	<li>Bring the sample into focus utilizing the manufacturer&#39;s directions.</li>
	<li>Image the sample according to the super-resolution technique being employed. Be sure to adjust imaging conditions to optimize signal while decreasing simultaneous activations of photo-switchable fluorophores. 
	 
	IMPORTANT NOTE: Imaging conditions are sample dependent and vary according to experimental need. A typical experiment utilizing Alexa 647 efficiently initialized to their dark state in the imaging buffer can then be photo-activated through application of 405 nm UV light. Images are obtained by exciting a sparse subset of Alexa 647 molecules within a densely labeled sample and localizing each fluorophore with a precision limited primarily by the number of collected photons. This procedure is iterated repeatedly by the software until a desired number of acquisition cycles have happened. The experimenter should have this setting correlate with each fluorophore in the sample having been photo-bleached.</li>
	<li>When imaging is completed, shut down the imaging equipment according to manufacturer&#39;s directions. The software may be left open for data analysis.</li>
	<li>Data analysis is heavily dependent upon experiment; however, in all cases identify viruses by their full width at half maximum, which is compared to the known virus dimensions, making sure to take into consideration the additional size of fluorescent labels.</li>
	<li>For better visualization, view samples by utilizing either the Isosurface rendering option or the volumetric rendering option, both of which are typically found in super-resolution imaging software.</li>
</ol>

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The authors have no competing financial interests in this paper.

Single virion imaging with AFM and High-resolution fluorescence imaging can be used as an alternative methodology to CryoEM tomography. Each one of these methodologies has their specific strengths. For example AFM can be done on WT virions with no requirement on tagging the sample or the internal viral proteins. The location of viral structures is deconvolved from their contributions to the elastic properties of the virion.
Single virion super-resolution imaging in combination with the newly developed viral reverse genetic approaches becomes a powerful technique for localizing low copy number viral proteins20. Recombinant viruses with replacement of their proteins with proteins fused to fluorescent proteins can be made and purified using these reverse genetic approaches. Although super-resolution imaging has specificity in part due to the genetic tagging, it requires the use of the mutant viruses.
AFM and super-resolution imaging are complimentary methods that utilize very similar sample preparations. The methods outlined in this paper, which are adopted form earlier single molecule imaging assays, allow anchoring of single virions with little effects on the topology of the virions.

Charge based nonspecific interactions are routinely used for adhesion of virions in atomic force microscopy1,2. These techniques especially work well when used on nonenveloped virions with very stiff capsids3-6. Although these techniques are very effective in immobilizing the sample, they do not prevent nonspecific binding of proteins to the surface. The nonspecific binding can create a problem when attempting to image virions with AFM and super-resolution fluorescence techniques that require incubations of the sample with various antibodies. Here we outline a sample preparation method for specific immobilization of virions.
Poly(ethylene glycol) (PEG) grafted to poly(L-lysine) (PLL) and adsorbed onto a glass surface provides a significant block for the electrostatic interactions of proteins with glass7. Single molecule assays which require immobilization of single molecules on glass surfaces have taken advantage of this property and used it to create a specific PEG based single molecule immobilization technique8-10. This preparation was also used to immobilize clathrin cages onto the glass surface11 as well as creating a homogenous fibronectin coating for control cell adhesion 12.
We have adopted the sample preparation method based on PLL-g-PEG adsorption to glass surfaces from single molecule imaging methodologies and applied it to imaging single virions of vesicular stomatitis virus (VSV). These single virions were imaged using atomic force microscopy (AFM). Similar experiments were also performed on functionalized beads as a control. The virions were also imaged by super-resolution fluorescence microscopy were a VSV-G antibody labeled with Alexa 647 was used to create an image of the envelope of single virions. High resolution fluorescent imaging utilizes localization of single molecules to create an image13-15. fPALM bi-planar imaging allows localization of single molecules with 20 nm in plane and 50 nm resolution along the optical axis15,16.&#160;This bi-planar technique was used to image super-resolution fluorescent images present in this study. An alternative technique which has similar results is STORM13,17,18.
Both AFM and super-resolution fluorescent images of VSV anchored to the glass by the procedures outlined in this paper, showed specific binding of virions to the glass with minimum nonspecific interactions19. Here we present the sample preparation protocols for the AFM and super-resolution fluorescent imaging experiments. In brief: Clean glass coverslips are functionalized by adsorbing a mixture of PLL-g-PEG and PLL-g-PEG-biotin. It is typical for this thin film to be engineered to provide between 15-25% functionalized biotin on a coated surface. The coverslips are further incubated with tetrameric avidin. A biotinylated viral antibody is then used to create a unique binding site for the virions. Binding of the antibody can be done in two ways.
The first method is optimized for AFM however it remains suitable for super-resolution fluorescence imaging. In this method the avidin coated surface is treated with biotinylated antibody prior to viral adhesion. The immobilized virions are treated with Alexa 647 labeled antibody to cover the envelope and allow super-resolution fluorescence imaging of the virions.
The second method is to attack the virus in solution with the biotinylated antibody and Alexa 647 labeled antibody prior to adhering the viruses to the avidin treated surface; this method is optimized for super-resolution fluorescence imaging experiments in which recovery of the viral envelope is important. This method has the advantage of allowing uniform antibody coating on the viral surface. The biotinylated antibody may be mixed with Alexa 647 labeled viral antibodies in ratios that allow for sufficient adsorption of the virus to the avidin treated surface while maintaining a high concentration of fluorescent label on the exterior of the viral envelope. These Alexa 647 labeled viral antibodies are not biotinylated and their excess may be rinsed away in order to reduce background noise.

<table border="0" cellpadding="0" cellspacing="0" width="573">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="84">
			<td height="84" style="height:84px;width:216px;">Glass coverslips (fPALM)</td>
			<td style="width:71px;">Electron Microscopy Services</td>
			<td style="width:119px;">72225-01</td>
			<td style="width:167px;">25 mm Coverslips</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">PLL(20)-g[3.5]-PEG(2)/PEG(3.4)-Biotin (20%)&#160;</td>
			<td style="width:71px;">SuSoS&#160;</td>
			<td style="width:119px;">-</td>
			<td style="width:167px;">Stock: 0.5 mg/ml in PBS</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">NeutrAvidin biotin-binding protein</td>
			<td style="width:71px;">Invitrogen</td>
			<td style="width:119px;">A2666</td>
			<td style="width:167px;">Stock: 0.25 mg/ml in PBS</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:216px;">Alexa Fluor 647 goat anti-rabbit IgG (H+L)&#160;</td>
			<td style="width:71px;">Invitrogen</td>
			<td style="width:119px;">A21245</td>
			<td style="width:167px;">1:200 in 0.2 M KPO4, 0.15 M NaCl, 10% Glycol, pH 7.2 buffer</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:216px;">&#945;-VSV-G&#160;</td>
			<td style="width:71px;">Invitrogen</td>
			<td style="width:119px;">ab34774</td>
			<td style="width:167px;">1:200 in 0.2 M KPO4, 0.15 M NaCl, 10% Glycol, pH 7.2 buffer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gluox</td>
			<td style="width:71px;">Sigma</td>
			<td>G2133-250KU&#160;</td>
			<td>Glucose oxidase type seven from Aspergillius&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Catalase</td>
			<td style="width:71px;">Sigma</td>
			<td style="width:119px;">C40-100mg&#160;</td>
			<td>Catalase from Bovine liver&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MEA</td>
			<td style="width:71px;">Sigma</td>
			<td>30070-10G&#160;</td>
			<td>Cysteamine (MEA</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Stock Buffer</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>50 mM Tris-HCl (pH 8.0) + 10 mM NaCl + 10% glucose</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">NTE</td>
			<td style="width:71px;"></td>
			<td style="width:119px;"></td>
			<td>10 mM Tris pH 7.4, 100 mM NaCl, 66 mM EDTA</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Slide-A-Lyzer, mini dialysis unit</td>
			<td style="width:71px;">Thermo Scientific</td>
			<td style="width:119px;"></td>
			<td>10,000 MW</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Microscope Cover Glass: 35 CIRCLE #1</td>
			<td style="width:71px;">Fisherbrand</td>
			<td style="width:119px;">35 CIRCLE #1</td>
			<td style="width:167px;"></td>
		</tr>
	</tbody>
</table>

sample preparation for single virion atomic force microscopy and super-resolution fluorescence imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

Single virion imaging using AFM:
The sample preparation protocol outlined above was used in anchoring wild type virions to the glass surface. VSV virions are bullet shaped 180 nm in length and 80 nm in diameter. As there are a variety of virions for which this technique may be applied, the concept is also demonstrated here on biotinylated 36 nm beads as well. The resulting AFM experiments are shown in Figure 1. It is important to note that the VSV virions have a lower Young&#39;s modulus (100 MPa) compared to the nonenveloped viruses (GPa). The particular images of single virion VSV were obtained under tapping mode in ambient conditions with a stiff cantilever. The aim has been to deform the virus to the point that the extra protein density within the virus becomes visible as a bump within the AFM image. This method is used to detect the extra protein density within the virus cavity19.
Single virion super resolution imaging:
Alexa 647 labeled VSV-G antibodies were used to coat the envelope of individual VSV virions for super-resolution experiments using method 5A. To demonstrate the tethering density and the low unspecific binding, a large scan of the sample is shown in Figure 2A. The recovery of the viral envelope on a representative virion is shown in Figure 2B (blue isosurface).
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51366/51366fig1highres.jpg" src="/files/ftp_upload/51366/51366fig1.jpg" style="opacity: 0.9; font-size: 14.399999618530273px; line-height: 28px;" />
Figure 1.&#160;AFM imaging of beads and VSV on the functionalized PEGG surface.&#160;AFM scan of 36 nm biotinylated beads (A,&#160;B) and a VSV virion (C) anchored to the surface with a biotinylated VSVG antibody. AFM was done in AC air topography scan mode. The tip radius is &#60;25 nm and measurements were carried out under force modulation and light tapping. The tip had&#160;a force constant of 3 N/m and resonant frequency 75 kHz with&#160;uncertainty&#160;of 15 kHz. While the beads retained their height during the AFM scan, the VSV virion has a significantly smaller young&#8217;s modulus and is significantly deformed in height. In this image the virion was specifically imaged with a stiff cantilever in tapping mode which produced a small xy&#160;convolution (used to determine the tip vs blunt end of the virus) and the height difference between the tip and blunt end of the virus is used to detect extra protein density at the blunt end of the virus19.
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51366/51366fig2highres.jpg" src="/files/ftp_upload/51366/51366fig2.jpg" /> 
Figure 2. Fluorescence based imaging of VSV virions on the PEGG functionalized surface.&#160;A) recombinant VSV virions immobilized on the PEGG surface using a biotinylated anti VSVG antibody imaged in wide field fluorescence. B) High resolution fluorescence reconstruction of the envelope of VSV attached to the PEGG surface through a biotinylated anti VSVG antibody and decorated with Alexa 647 labeled anti-VSVG antibodies (Method 5A was used for creating these images). The blue surface is a 2D isosurface projection. Left shows a model of the bullet shaped virus.
<table border="0" cellpadding="0" cellspacing="0" width="212">
	<colgroup>
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="27">
			<td height="27" style="height:27px;width:119px;">Coverslip Size</td>
			<td style="width:93px;">Fluid</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">40 mm</td>
			<td>65 &#181;l</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">35 mm</td>
			<td>50 &#181;l</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">30 mm</td>
			<td>36 &#181;l</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">25 mm</td>
			<td>25 &#181;l</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">20 mm</td>
			<td>16 &#181;l</td>
		</tr>
	</tbody>
</table>
Table 1. Fluid aliquots according to coverslip size.

Watch this Scientific Journal Video about Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging at JoVE.com

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

sample-preparation-for-single-virion-atomic-force-microscopy-super

Research

JoVE Journal

Biology

9.4K Views.  University of Utah. The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Video: Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Test Samples for Optimizing STORM Super-Resolution Microscopy

STORM is a recently developed super-resolution microscopy technique with up to 10 times better resolution than standard fluorescence microscopy techniques. However, as the image is acquired in a very different way than normal, by building up an image molecule-by-molecule, there are some significant challenges for users in trying to optimize their image acquisition. In order to aid this process and gain more insight into how STORM works we present the preparation of 3 test samples and the methodology of acquiring and processing STORM super-resolution images with typical resolutions of between 30-50 nm. By combining the test samples with the use of the freely available rainSTORM processing software it is possible to obtain a great deal of information about image quality and resolution. Using these metrics it is then possible to optimize the imaging procedure from the optics, to sample preparation, dye choice, buffer conditions, and image acquisition settings. We also show examples of some common problems that result in poor image quality, such as lateral drift, where the sample moves during image acquisition and density related problems resulting in the 'mislocalization' phenomenon.

We describe the preparation of three test samples and how they can be used to optimize and assess the performance of STORM microscopes. Using these examples we show how to acquire raw data and then process it to acquire super-resolution images in cells of approximately 30-50 nm resolution.

STORM is a recently developed super-resolution microscopy technique with up to 10 times better resolution than standard fluorescence microscopy ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Open-source Single-particle Analysis for Super-resolution Microscopy with VirusMapper

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm allows for the routine imaging of nanoscale biological complexes and processes. This increase in resolution also means that methods popular in electron microscopy, such as single-particle analysis, can readily be applied to super-resolution fluorescence microscopy. By combining this analytical approach with super-resolution optical imaging, it becomes possible to take advantage of the molecule-specific labeling capacity of fluorescence microscopy to generate structural maps of molecular elements within a metastable structure. To this end, we have developed a novel algorithm &#8212; VirusMapper &#8212; packaged as an easy-to-use, high-performance, and high-throughput ImageJ plugin. This article presents an in-depth guide to this software, showcasing its ability to uncover novel structural features in biological molecular complexes. Here, we present how to assemble compatible data and provide a step-by-step protocol on how to use this algorithm to apply single-particle analysis to super-resolution images.

This manuscript uses the Fiji-based open-source software package VirusMapper to apply single-particle analysis to super-resolution microscopy images in order to generate precise models of nanoscale structure.

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...