This work was funded by National Institutes of Health Grant GM086822. The authors would like to thank Drs. Alec Pakhomov &amp; Paul Wallace (Univ. of Washington Nanotechnology User Facility (NTUF)) and Andrea Slade (Bruker AXS) for their expert technical assistance. DNA AFM imaging was conducted at the Univ. of Washington NTUF, a member of the National Nanotechnology Infrastructure Network. The 2xGal4-2xGRE-luciferease plasmid was kindly provided by the laboratory of Dr. Keith Yamamoto (Univ. of California, San Francisco).

1. Preparing DNA and protein samples to be imaged free of contaminants
<ol>
	<li>Isolate biomolecules to be imaged and place in a suitable aqueous buffer. Proteins to be imaged may be purified using liquid chromatography 6, immunoadsorption followed by removal of antibody and pellet 1,2, or Profinity eXact purification and tag removal 7 (Figure 2A). Isolate DNA molecules to be imaged by miniprep purification (Figure 2B).</li>
	<li>Confirm composition and purity of protein samples to be imaged using SDS-PAGE and western blotting. For DNA, confirm sequence and purity by completing a restriction digest and agarose gel electrophoresis.</li>
	<li>Incubate samples in an AFM adsorption buffer to promote adherence of the sample to mica. For protein samples, 10 mM HEPES buffer may be used. Additional low-ionic strength salts or other cofactors may be added as well. For DNA samples, include a divalent cation (e.g. Mg2+ or Ni2+) to promote adsorption to the mica substrate. A suitable Mg2+-containing buffer for DNA is 10 mM Tris, pH 7.5, 10 mM NaCl, 2 mM MgCl2 3. An alternative Ni2+ buffer is 10 mM HEPES, pH 6.8, 10 mM NiCl2.</li>
	<li>Dilute the sample to a concentration of 5 &mu;g/ml in adsorption buffer. Gently mix. Store on ice while the microscope is being prepared.</li>
</ol>
2. Mounting AFM probe
<ol>
	<li>Place the liquid cell probe holder onto the corresponding cantilever installation docking station.</li>
	<li>Locate the long, thick cantilever (referred to as the 'B' cantilever) of the sharp nitride lever (SNL) probe to be used for imaging. Carefully transfer it to the liquid cell probe holder, ensuring that the tip remains upright. Technical note: Extreme care must be paid while the probe is in transit, as dropping it from any height may damage or destroy the cantilever or tip. A microlever SNL (MSNL) probe may be used as an alternative to the SNL probe.</li>
	<li>Remove the probe holder from the docking station and examine the cantilever under a light microscope to ensure it is intact and properly seated in the liquid cell probe holder.</li>
	<li>Carefully remove the AFM head from the instrument dovetail assembly by tightening the knurled head clamp screw. Technical note: Extreme care must be paid while manipulating the AFM head, as dropping it from even a short distance (e.g. if not seated correctly in the dovetail assembly) may cause substantial damage.</li>
	<li>Invert the AFM head and firmly push the liquid cell probe holder onto the four pins at the base of the AFM head. Carefully return the AFM head into the instrument dovetail assembly. Release the knurled head clamp screw to fasten the AFM head into the instrument.</li>
</ol>
3. Locating cantilever tip, aligning laser, and adjusting photodetector
<ol>
	<li>Using the AFM instrument software, locate the cantilever tip by moving the knobs of the onboard light microscope. Adjust the illumination of the microscope if necessary to improve tip identification. Bring the end of the cantilever tip into close proximity of the crosshairs displayed on the instrument software. </li>
	<li>Bring the cantilever tip into focus by adjusting the up and down arrows of the optics controller on the software. Use slow (S) or medium (M) speed while adjusting the optics.</li>
	<li>Align the laser to the cantilever tip using the laser adjustment knobs. Move the adjustment knobs until the red laser dot is inside the white illumination spot. Trace along the cantilever arm using a zig-zag pattern until the laser is positioned on the cantilever tip. When properly aligned, a strong reflection spot will appear in the front window of the AFM head.</li>
	<li>Center the photodetector by adjusting the photodetector knobs on the AFM head. This will align the laser dot in the crosshairs of the photodetector display of the instrument software. The observed signal sum should be approximately 4-6.</li>
</ol>
4. Positioning AFM head and tuning cantilever
<ol>
<li>Place a freshly cleaved sheet of mica attached to a metal AFM specimen disc on the magnetized sample holder atop the AFM holder plate. This setup will be used to position the AFM head prior to imaging the DNA or protein sample prepared in Step 1. 
<ol><li>Technical note: The magnetized sample holder and liquid cell probe holder are thicker than other sample fastening mechanisms and probe holders. If there is currently insufficient clearance between the mica-sample holder assembly and AFM head, move the AFM head up by selecting the withdraw icon on the instrument software several times.</li></ol></li>
<li>Rotate the AFM holder plate so the specimen disc is positioned for the initial imaging. Ensure the mica substrate is centered under the AFM head.</li>
<li>Move the AFM head down toward the surface of the mica-specimen disc using the focus surface control of the instrument software. Select the medium (M) speed on the Z motor, and continue towards the surface until the AFM head is approximately 2 mm above the mica. At this time, switch to using the slow (S) Z motor speed in order to avoid abrupt contact between the probe and the surface (referred to as crashing the tip). </li>
<li>Proceed to move the probe closer to the mica substrate surface until distinct features on the mica surface or the tip reflection are in focus. Toggle between surface and tip reflection focal planes using the "focus on:" icon from the instrument software. </li>
<li>Select the tune icon from the instrument software and tune the cantilever. The resonance frequency of the recommended SNL and MSNL cantilever probes is 20-60 kHz. Select a 5% peak offset. Technical note: Adequate cantilever tuning is essential for adequate sample imaging.</li>
</ol>
5. Imaging mica sample surface
<ol>
	<li>Engage the probe onto the mica surface. Set initial scan size to 10 &mu;m and scan rate to 1 Hz. Integral gain and proportional gain can be initially set to 0.2 and 0.4, respectively. Adjust gains as needed in order to have trace and retrace scans roughly overlap. Decreasing the amplitude setpoint will increase the interaction between the probe and the mica and ensure engagement.</li>
	<li>Capture complete scan images of the 10 &mu;m field. Decrease scan size to 5, 1, and 0.5 &mu;m and capture complete images of each field. Adjust gains and scan rate as necessary. Capturing images of the mica surface provides a baseline for comparison to the DNA- and protein-containing samples.</li>
	<li>Disengage the probe with a single click on the disengage icon on the instrument software. </li>
</ol>
6. Imaging biomolecules of interest
<ol>
	<li>Gently mix the DNA or protein sample prepared in Step 1 and prepare to load it onto the imaged mica surface. Mix 5 &mu;l of the prepared sample with 45 &mu;l of fresh adsorption buffer (final sample concentration = 0.5 &mu;g/ml).</li>
	<li>Carefully add 50 &mu;l of the 0.5 &mu;g/ml biomolecule-containing solution directly onto the mica surface. Do this without moving the probe or AFM head, and ensure the sample solution (but not pipet tip) makes contact with the probe. </li>
	<li>Pause 5 minutes to allow the biomolecules in the sample to adhere to the mica surface. </li>
	<li>At the conclusion of 5 minutes, pipet an additional 50-100 &mu;l of adsorption buffer into the liquid cell probe holder being careful to not touch the probe, AFM head, or mica with the pipet tip. This will form a meniscus and allow for AFM tapping mode imaging in fluid. Figure 3 shows the final arrangement of the fluid cell, cantilever, sample, sample holder, and meniscus.</li>
	<li>Readjust the photodetector and realign the laser to the cantilever as necessary. Technical note: viewing the probe tip and laser alignment is complicated due to the diffraction pattern caused by adding sample buffer into the probe holder. Manually retune the cantilever, if needed.</li>
	<li>Re-engage the probe using the instrument software. Adjust vertical deflection, integral gain, and proportional gain as necessary. This will begin the process of re-imaging the section of mica scanned immediately prior to adding sample and capture images.</li>
	<li>Increase scan size (500 nm-10 &mu;m) to identify regions of interest. Zoom-in and reduce scan rate to 0.5 Hz to improve image resolution</li>
	<li>Once imaging is complete, disengage the probe by selecting withdraw icon on the instrument software several times. Ensure sufficient clearance between the AFM head and specimen disc before disassembly of the liquid cell or specimen disc. </li>
	<li>Thoroughly rinse the liquid cell probe holder and specimen disc with distilled water to prevent salt crystals from forming as residual adsorption buffer evaporates. Dry with compressed air.</li>
</ol>
7. Representative Results: 
Examples of AFM images are presented in Figure 4. Mica substrate (A) provides a molecularly flat surface onto which DNA and protein can adsorb. Imaging mica prior to biomolecule samples provides a negative control and assessment of imaging noise. It also provides a level of assurance that the cantilever is properly tuned and subsequent sample imaging will be successful. Double-stranded plasmid DNA (B, D), presumptively supercoiled, is readily identified by its asymmetric appearance and uniform depositing on the previously unremarkable mica substrate. Protein complexes of discreet particle sizes (C, E) are also uniquely distinguishable from the mica substrate. Particle size differences indicate sample heterogeneity, and can be useful to approximating protein complex stoichiometry or biochemical activity. The general diagonal shape and consistent orientation of the protein particles observed in Panel C is an imaging artifact, as proteins would expectedly be oriented on the mica substrate in a random fashion. Possible causes of the observed artifact is a physical tip abnormality or AFM imaging at too rapid of a scan rate. While tip convolution (illustrated in Figure 1B) prevents absolute length measurement calculations in the x and y axes, height measurement (z axis) and relative x and y measurements may be nonetheless useful for estimating biophysical properties of the imaged biomolecules.
<img src="/files/ftp_upload/3061/3061fig1.jpg" alt="Figure 1"/> Figure 1: Schematic presentation of tapping mode atomic force microscopy (AFM). A, the microscope. At the end of the cantilever is a sharp tip that oscillates up and down as it scans over the surface of a mica substrate to which biomolecular complexes adsorb. As the scanner moves in the x and y directions, a laser beam is reflected from the back of the cantilever onto a position-sensitive photodiode detector to map the vertical (z) distance the tip moves as it passes over the biomolecular complexes seated on the mica. B. Distortion of the image in the x and y directions caused by tip convolution. The nominal radius of a conventional silicon nitride tip is larger than the particles to be imaged, and the edge of the tip contacts the sample as it traverses the surface. Tip convolution results in the imaged particles appearing larger in the x and y directions, but not the z direction. This effect can be minimized by the use of tips with a smaller nominal tip radius.
<img src="/files/ftp_upload/3061/3061fig2.jpg" alt="Figure 2"/> Figure 2: Illustration of immunoadsorbed protein and DNA sample preparation. A, Release of immunoadsorbed GR&bull;hsp70 protein complex from the monoclonal antibody (mAb)-protein A-Sepharose (PAS) pellet. Incubating the immunopellet with a peptide containing the mAb epitope will facilitate the release of the GR&bull;hsp70 protein complex from the pellet, allowing the complex to be collected from the supernatant and visualized by AFM. B, Preparation of DNA for AFM imaging requires the use of a divalent cation adsorption buffer. The divalent cation increases the affinity of DNA for the mica substrate.
<img src="/files/ftp_upload/3061/3061fig3.jpg" alt="Figure 3"/> Figure 3: Final arrangement of the AFM head, liquid cell probe holder, sample, specimen holder, and buffer meniscus. A, Care must be taken when depositing sample and additional buffer onto the mica substrate to avoid physical contact between the pipet tip and AFM head, liquid call, and mica substrate. B, Magnification of A. The buffer meniscus spans from the specimen disc to the liquid cell probe holder. 
<img src="/files/ftp_upload/3061/3061fig4.jpg" alt="Figure 4"/> Figure 4: Atomic force micrographs of mica substrate (A), 2xGal4-2xGRE-luciferease plasmid DNA 8 (B, D), and GR&bull;hsp70 protein complexes (C, E) in buffered solution. Mica serves as a control image to compare with the visualizations of DNA and protein. GR&bull;hsp70 protein complexes were prepared by immunoadsorption of GR primed with hsp70 and then released from the antibody-bead pellet using a mAb competing antibody (illustrated in Figure 2A). DNA was prepared by conventional plasmid miniprep. Panels A, B, and C are of an equivalent magnification (scale bars = 200 nm), as are Panels D and E (scale bars = 40 nm).

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	<li>Murphy, P. J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualization+and+mechanism+of+assembly+of+a+glucocorticoid+receptor&#149;hsp70+complex+that+is+primed+for+subsequent+Hsp90-dependent+opening+of+the+steroid+binding+cleft.">Visualization and mechanism of assembly of a glucocorticoid receptor&#149;hsp70 complex that is primed for subsequent Hsp90-dependent opening of the steroid binding cleft.</a> J Biol Chem. 278, 34764-34773 (2003).</li><li>Harrell, J. 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No conflicts of interest declared.

AFM provides a unique microscopic technique capable of imaging individual uncoated biomolecules in aqueous and near-physiologic buffered solutions in real-time. This allows for the visualization of individual proteins and DNA molecules, as well as multiprotein complexes and protein-DNA complexes. Imaging macromolecular particles with AFM can be useful for assessing sample homogeneity and for identifying the physical arrangement of the constituent components of the observed particles. This approach of observing individual macromolecular particles can be a useful adjunct to conventional biochemical techniques, such as immunoprecipitation assays, polyacrylamide gel electrophoresis, and size exclusion column chromatography, which provide significant summative data representing the 103-1011 individual biomolecular complexes of interest present in a sample. 
Research into multiprotein complex stoichiometry, biomolecular interactions, and cofactor requirements can all be investigated using AFM. The method presented here can be adapted to accommodate specific biophysical questions of interest. For example, using a liquid cell probe holder capable of exchanging buffer, it is be possible to assay cofactor requirements for protein complex formation. DNA-protein assemblies can be formed and dissociated in near-real-time and even while being imaged. DNA can be generated with specific sequences of interest (e.g. a presumptive response elements or novel protein binding sequence), mixed with the hypothesized binding protein, and imaged to provide direct evidence of intermolecular interactions. 
Tapping mode AFM imaging is substantially less complicated if dry samples are utilized rather than samples in aqueous solutions, and linear DNA stands and closed-circle DNA plasmids have both been imaged in this way 3,9. The method presented here utilizes samples in buffered solutions in order to provide a more physiological imaging environment. Other noteworthy methods of imaging proteins should also be considered, including near field infrared microscopy 10. The complementary use of immunoadsorption in consort with AFM provides an opportunity to prepare a myriad of protein complexes, using well-established biochemical techniques, and then visualize them following release from the immunoadsorbing antibody-bead pellet. For example, immunoadsorbed GR has been assayed for its association with the molecular chaperone protein hsp70 1 as well as the dynein motor protein 2 using this approach in order to estimate complex size and stoichiometries. Particle size and biophysical features (e.g. rigidity) have been useful in ascertaining the identity of the biomolecules observed in AFM imaged samples 11,12. It is also possible to confirm biomolecule identity by the addition of an interacting biomolecule (e.g. a ligand or monoclonal antibody), that would bind to its target and cause a particle size increase if the target biomolecule is present.

<table border="1">
	<tbody>
		<tr>
			<th>Name of the reagent</th>
			<th>Company</th>
			<th>Catalogue number</th>
		</tr>
		<tr>
			<td>Dimension 3100</td>
			<td>Bruker</td>
			<td>Dimension 3100</td>
		</tr>
		<tr>
			<td>Dimension Fluid Cell</td>
			<td>Bruker</td>
			<td>DTFML-DD-HE</td>
		</tr>
		<tr>
			<td>Sharp nitride lever (SNL) silicon nitride AFM probe</td>
			<td>Bruker</td>
			<td>SNL-10</td>
		</tr>
		<tr>
			<td>Microlever sharp nitride lever (MSNL) silicon nitride AFM probe </td>
			<td>Bruker</td>
			<td>MSNL-10 </td>
		</tr>
		<tr>
			<td>NanoScope AFM instrument software</td>
			<td>Bruker</td>
			<td>004-132-000</td>
		</tr>
		<tr>
			<td>Metal AFM specimen discs</td>
			<td>Ted Pella</td>
			<td>16208</td>
		</tr>
		<tr>
			<td>Grade V1 Mica Discs, 12 mm</td>
			<td>Ted Pella</td>
			<td>50-12</td>
		</tr>
	</tbody>
</table>

visualization of recombinant dna and protein complexes using atomic force microscopy

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

Watch this Scientific Journal Video about Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy at JoVE.com

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

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

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

visualization-recombinant-dna-protein-complexes-using-atomic-force

Research

JoVE Journal

Bioengineering

22.4K visualizações.  Seattle University. Um modo de tocar microscópio de força atômica método (AFM) para a visualização de DNA plasmídeo, proteínas citoplasmáticas e proteína-DNA complexos é descrito. O método inclui abordagens alternativas para a preparação de amostras para AFM imagem após a manipulação bioquímica. DNA contendo regiões específicas interagindo proteínas são observados em condições quase fisiológica buffer.

Vídeo: Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

Bacterial Immobilization for Imaging by Atomic Force Microscopy

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface. It has the ability to image cells and biomolecules, in a liquid environment, without the need to chemically treat the sample. In order to accomplish this goal, the sample must sufficiently adhere to the mounting surface to prevent removal by forces exerted by the scanning AFM cantilever tip. In many instances, successful imaging depends on immobilization of the sample to the mounting surface. Optimally, immobilization should be minimally invasive to the sample such that metabolic processes and functional attributes are not compromised. By coating freshly cleaved mica surfaces with porcine (pig) gelatin, negatively charged bacteria can be immobilized on the surface and imaged in liquid by AFM. Immobilization of bacterial cells on gelatin-coated mica is most likely due to electrostatic interaction between the negatively charged bacteria and the positively charged gelatin. Several factors can interfere with bacterial immobilization, including chemical constituents of the liquid in which the bacteria are suspended, the incubation time of the bacteria on the gelatin coated mica, surface characteristics of the bacterial strain and the medium in which the bacteria are imaged. Overall, the use of gelatin-coated mica is found to be generally applicable for imaging microbial cells.

Live Gram-negative and Gram-positive bacteria can be immobilized on gelatin-coated mica and imaged in liquid using Atomic Force Microscopy (AFM).

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface.  It has the ability to image cells and biomolecules, in a liquid ...

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Quantifying the Mechanical Properties of the Endothelial Glycocalyx with Atomic Force Microscopy

Our understanding of the interaction of leukocytes and the vessel wall during leukocyte capture is limited by an incomplete understanding of the mechanical properties of the endothelial surface layer. It is known that adhesion molecules on leukocytes are distributed non-uniformly relative to surface topography 3, that topography limits adhesive bond formation with other surfaces 9, and that physiological contact forces (&asymp; 5.0 &minus; 10.0 pN per microvillus) can compress the microvilli to as little as a third of their resting length, increasing the accessibility of molecules to the opposing surface 3, 7. We consider the endothelium as a two-layered structure, the relatively rigid cell body, plus the glycocalyx, a soft protective sugar coating on the luminal surface 6. It has been shown that the glycocalyx can act as a barrier to reduce adhesion of leukocytes to the endothelial surface 4. In this report we begin to address the deformability of endothelial surfaces to understand how the endothelial mechanical stiffness might affect bond formation. Endothelial cells grown in static culture do not express a robust glycocalyx, but cells grown under physiological flow conditions begin to approximate the glycocalyx observed in vivo 2. The modulus of the endothelial cell body has been measured using atomic force microscopy (AFM) to be approximately 5 to 20 kPa 5. The thickness and structure of the glycocalyx have been studied using electron microscopy 8, and the modulus of the glycocalyx has been approximated using indirect methods, but to our knowledge, there have been no published reports of a direct measurement of the glycocalyx modulus in living cells. In this study, we present indentation experiments made with a novel AFM probe on cells that have been cultured in conditions to maximize their glycocalyx expression to make direct measurements of the modulus and thickness of the endothelial glycocalyx.

The mechanical characteristics of endothelial glycocalyx were measured by indentation using micron sized spheres on AFM cantilevers. Endothelial cells were cultured in a custom chamber under physiological flow conditions to induce glycocalyx expression. Data were analyzed using a thin film model to determine the glycocalyx thickness and modulus.

Our  understanding of the interaction of leukocytes and the vessel wall during  leukocyte capture is limited by an incomplete understanding of the ...

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

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

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

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

Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. Kelvin probe force microscopy (KPFM) is a module of atomic force microscopy (AFM) that measures the contact potential difference between surfaces at the nano-scale. The combination of KPFM with AFM allows for the simultaneous generation of surface potential and topographical maps of biological samples such as bacterial cells. Here, we employ KPFM to examine the effects of surface potential on microbial adhesion to medically relevant surfaces such as stainless steel and gold. Surface potential maps revealed differences in surface potential for microbial membranes on different material substrates. A step-height graph was generated to show the difference in surface potential at a boundary area between the substrate surface and microorganisms. Changes in cellular membrane surface potential have been linked with changes in cellular metabolism and motility. Therefore, KPFM represents a powerful tool that can be utilized to examine the changes of microbial membrane surface potential upon adhesion to various substrate surfaces. In this study, we demonstrate the procedure to characterize the surface potential of individual methicillin-resistant Staphylococcus aureus USA100 cells on stainless steel and gold using KPFM.

Here, we present a protocol explaining the use of Kelvin probe force microscopy as a tool for generating high resolution nano-scale surface potential maps. This tool was applied to assess the role of surface potential on the binding capacity of microorganisms to substrate surfaces.

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. ...

Novel Atomic Force Microscopy Based Biopanning for Isolation of Morphology Specific Reagents against TDP-43 Variants in Amyotrophic Lateral Sclerosis

Because protein variants play critical roles in many diseases including TDP-43 in Amyotrophic Lateral Sclerosis (ALS), alpha-synuclein in Parkinson&#8217;s disease and beta-amyloid and tau in Alzheimer&#8217;s disease, it is critically important to develop morphology specific reagents that can selectively target these disease-specific protein variants to study the role of these variants in disease pathology and for potential diagnostic and therapeutic applications. We have developed novel atomic force microscopy (AFM) based biopanning techniques that enable isolation of reagents that selectively recognize disease-specific protein variants. There are two key phases involved in the process, the negative and positive panning phases. During the negative panning phase, phages that are reactive to off-target antigens are eliminated through multiple rounds of subtractive panning utilizing a series of carefully selected off-target antigens. A key feature in the negative panning phase is utilizing AFM imaging to monitor the process and confirm that all undesired phage particles are removed. For the positive panning phase, the target antigen of interest is fixed on a mica surface and bound phages are eluted and screened to identify phages that selectively bind the target antigen. The target protein variant does not need to be purified providing the appropriate negative panning controls have been used. Even target protein variants that are only present at very low concentrations in complex biological material can be utilized in the positive panning step. Through application of this technology, we acquired antibodies to protein variants of TDP-43 that are selectively found in human ALS brain tissue. We expect that this protocol should be applicable to generating reagents that selectively bind protein variants present in a wide variety of different biological processes and diseases.

Using atomic force microscopy in combination with biopanning technology we created a negative and positive biopanning system to acquire antibodies against disease-specific protein variants present in any biological material, even at low concentrations. We were successful in obtaining antibodies to TDP-43 protein variants involved in Amyotrophic Lateral Sclerosis.

Because protein variants play critical roles in many diseases including TDP-43 in Amyotrophic Lateral Sclerosis (ALS), alpha-synuclein in ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

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

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

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

3D Magnetic Stem Cell Aggregation and Bioreactor Maturation for Cartilage Regeneration

Cartilage engineering remains a challenge due to the difficulties in creating an in vitro functional implant similar to the native tissue. An approach recently explored for the development of autologous replacements involves the differentiation of stem cells into chondrocytes. To initiate this chondrogenesis, a degree of compaction of the stem cells is required; hence, we demonstrated the feasibility of magnetically condensing cells, both within thick scaffolds and scaffold-free, using miniaturized magnetic field sources as cell attractors. This magnetic approach was also used to guide aggregate fusion and to build scaffold-free, organized, three-dimensional (3D) tissues several millimeters in size. In addition to having an enhanced size, the tissue formed by magnetic-driven fusion presented a significant increase in the expression of collagen II, and a similar trend was observed for aggrecan expression. As the native cartilage was subjected to forces that influenced its 3D structure, dynamic maturation was also performed. A bioreactor that provides mechanical stimuli was used to culture the magnetically seeded scaffolds over a 21-day period. Bioreactor maturation largely improved chondrogenesis into the cellularized scaffolds; the extracellular matrix obtained under these conditions was rich in collagen II and aggrecan. This work outlines the innovative potential of magnetic condensation of labeled stem cells and dynamic maturation in a bioreactor for improved chondrogenic differentiation, both scaffold-free and within polysaccharide scaffolds.

Chondrogenesis from stem cells requires fine tuning the culture conditions. Here, we present a magnetic approach for condensing cells, an essential step to initiate chondrogenesis. In addition, we show that dynamic maturation in a bioreactor applies mechanical stimulation to the cellular constructs and enhances cartilaginous extracellular matrix production.

Cartilage engineering remains a challenge due to the difficulties in creating an in vitro functional implant similar to the native tissue. An approach ...

Visualization of Germinosomes and the Inner Membrane in Bacillus subtilis Spores

The small size of spores and the relatively low abundance of germination proteins, cause difficulties in their microscopic analyses using epifluorescence microscopy. Super-resolution three-dimensional Structured Illumination Microscopy (3D-SIM) is a promising tool to overcome this hurdle and reveal the molecular details of the process of germination of Bacillus subtilis (B. subtilis) spores. Here, we describe the use of a modified SIMcheck (ImageJ)-assistant 3D imaging process and fluorescent reporter proteins for SIM microscopy of B. subtilis spores&#8217; germinosomes, cluster(s) of germination proteins. We also present a (standard)3D-SIM imaging procedure for FM4-64 staining of B. subtilis spore membranes. By using these procedures, we obtained unsurpassed resolution for germinosome localization and show that &#62;80% of B. subtilis KGB80 dormant spores obtained after sporulation on defined minimal MOPS medium have one or two GerD-GFP and GerKB-mCherry foci. Bright foci were also observed in FM4-64 stained spores&#8217; 3D-SIM images suggesting that inner membrane lipid domains of different fluidity likely exist. Further studies that use double labeling procedures with membrane dyes and germinosome reporter proteins to assess co-localization and thus get an optimal overview of the organization of Bacillus germination proteins in the inner spore membrane are possible.

Visualization of Germinosomes and the Inner Membrane in Bacillus subtilis Spores

Germinant receptor proteins cluster in &#8216;germinosomes&#8217; in the inner membrane of Bacillus subtilis spores. We describe a protocol using super resolution microscopy and fluorescent reporter proteins to visualize germinosomes. The protocol also identifies spore inner membrane domains that are preferentially stained with the membrane dye FM4-64.

The small size of spores and the relatively low abundance of germination proteins, cause difficulties in their microscopic analyses using epifluorescence ...

Fabrication and Implementation of a Reference-Free Traction Force Microscopy Platform

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their microenvironment. While many approaches exist for measuring cell-induced material strain, here we provide a methodology for monitoring strain with sub-micron resolution in a reference-free manner. Using a two-photon activated photolithographic patterning process, we demonstrate how to generate mechanically and bio-actively tunable synthetic substrates containing embedded arrays of fluorescent fiducial markers to easily measure three-dimensional (3D) material deformation profiles in response to surface tractions. Using these substrates, cell tension profiles can be mapped using a single 3D image stack of a cell of interest. Our goal with this methodology is to make traction force microscopy a more accessible and easier to implement tool for researchers studying cellular mechanotransduction processes, especially newcomers to the field.

This protocol provides instructions for implementing multiphoton lithography to fabricate three-dimensional arrays of fluorescent fiducial markers embedded in poly(ethylene glycol)-based hydrogels for use as reference-free, traction force microscopy platforms. Using these instructions, measurement of 3D material strain and calculation of cellular tractions is simplified to promote high-throughput traction force measurements.

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their ...