The authors sincerely thank the Natural Sciences and Engineering Research Council of Canada, the Ontario Ministry of Research and Innovation, and the Canada Foundation for Innovation for funding this study.

1. Preparation of Glassware and Cultures
<ol>
	<li>Before proceeding with this experiment, prepare 5% sheep&#8217;s blood agar (SBA) plates for growing MRSA. Incubate SBA plates for 24 hr at 37 &#176;C. After this time, single, well-isolated colonies should be present to be used for subsequent inoculations into liquid media. Store streaked SBA plates with single colonies at 4 &#176;C for 1 - 2 months. 
	NOTE: Sheep&#8217;s blood agar plates come in pre-made packs containing between 20 - 25 plates based on the manufacturer, and typically consist of a tryptic soy agar base with the addition of 5% w/v sheep&#8217;s blood.</li>
	<li>Make 500 ml of tryptic soy broth (TSB) supplemented with 1% glucose and 0.1% NaCl. After this, collect and clean 2 glass test tubes. If autoclavable lids are unavailable for test tubes, use aluminum foil as a cap. Autoclave TSB and test tubes along with a box of 1 ml pipette tips.</li>
	<li>Under a biosafety cabinet or near a Bunsen burner, pipette 5 ml of previously autoclaved TSB into autoclaved test tubes. Be careful to ensure that enough time has been given to let the TSB and test tubes cool to room temperature. From a previously streaked 5% SBA plate, inoculate a single colony into 6 ml TSB broth. One will contain MRSA, and the second test tube will be used as a negative control to ensure sterility.</li>
	<li>Take inoculated TSB test tubes and place them in a reciprocal incubator for 24 hr at 37 &#176;C and at 200 rpm. After this time microbial growth should be apparent in the MRSA test tube while no growth should have occurred in the control test tube.</li>
	<li>Take 1 ml from 24 hr culture and pipette into 6 ml of fresh TSB. Incubate at 37 &#176;C for 6 hr at 200 rpm.</li>
	<li>Pipette 1 ml of 6 hr sub-culture into a 1.5 ml microfuge tube. Centrifuge for 3 min at 850 x g. Remove supernatant and re-suspend pellet in 1 ml of deionized H2O. Centrifuge, repeat, and re-suspend.</li>
</ol>
2. Cleaning and Inoculation of Stainless Steel and Gold Substrates
<ol>
	<li>Take stainless steel and gold AFM sample disks (20 mm and 10 mm diameters respectively) and clean each side with 5 ml of deionized H2O. Hold sample disks in the dominant hand with forceps and use the subdominant hand to pipette 5 ml on each side of the sample disk. After washing both sides, place sample disks in a beaker containing 20 ml of deionized H2O followed by sonication for 1 min.</li>
	<li>After sonication use forceps to remove sample disks from the beaker. Place the disks so that they lean at a 60&#176; angle against the edge of an open Petri plate onto a paper towel. Use the lid of the Petri plate to cover the drying disks. Let disks dry fully before proceeding. The drying time can vary between 4 - 8 hr.</li>
	<li>Once dry, use forceps to place sample disks into a Petri plate.
	<ol>
		<li>Optionally, functionalize the surfaces of sample disks with any material (e.g., collagen, hyaluronan, silver nano-particles, etc.).</li>
		<li>For this experiment, use poly-L-lysine to coat the substrate surface. For poly-L-lysine functionalization, pipette 200 - 400 &#181;l of 0.1% poly-L-lysine (in H2O) onto sample disks, and let incubate for 1 hr at room temperature in a Petri dish.</li>
		<li>After 1 hr, use forceps to hold the sample disk, and wash with 1 ml of deionized H2O. Let dry on paper towels as described in step 2.2.</li>
	</ol>
	</li>
	<li>Take 2x washed re-suspended cells and plate 200 - 400 &#181;l onto sample disks inside a biosafety cabinet. If sample disks are coated with poly-L-lysine, incubate for 0.5 hr at room temperature.</li>
	<li>After incubation of cells on sample disks, wash sample disks gently with 1 ml of deionized H2O. Let disks dry overnight before imaging.</li>
</ol>
3. KPFM Imaging
NOTE: For this experiment, use an Agilent 5500 series ILM-AFM.
<ol>
	<li>To start AFM, turn on computer unit along with the AFM&#8217;s HEB and MAC III controller. Open AFM imaging software (e.g., Agilent&#8217;s PicoView). Be sure to initially select ACAFM or whichever the correct designation is on the AFM for imaging in intermittent-contact mode.</li>
	<li>Using AFM forceps in the dominant hand, and holding the spring-loaded clip holder open with the subdominant hand, place a KPFM cantilever into to head-piece. Be careful when handling and transferring the AFM head-piece to the AFM as this unit (at least on the 5500 ILM-AFM) contains the fragile piezo crystal unit that controls X, Y, and Z scanning directionalities. 
	NOTE: The KPFM cantilevers used in this case were Mikromasch conductive DPE (low-noise) cantilevers with average resonant frequencies of 80 kHz and spring constants of 2.7 N/m. Cantilevers with these resonant frequencies and spring constants were chosen due to their ease of use. Cantilevers with larger spring constants and higher resonant frequencies can also be used for imaging.</li>
	<li>Carefully load head-piece into the AFM and connect the appropriate wires (in this case two wires need to be plugged in: laser light and stage lift motor).</li>
	<li>Align the laser onto the tip of the cantilever. Some units contain a camera functionality which allows for easier alignment. If camera functionality is not present on AFM, align onto the cantilever tip as described next:
	<ol>
		<li>Rotate the front-to-back knob clockwise to move the laser overtop the cantilever chip. When the laser reaches the chip it will be blocked and will no longer be visible. Only turn the knob a few times.</li>
		<li>Rotate the front-to-back knob counterclockwise until the laser spot reappears. The laser is now on the chip&#8217;s edge.</li>
		<li>Rotate the left-to-right knob to position the laser on the cantilever. As the laser passes over the cantilever it will disappear and reappear in rapid succession. The laser spot should be visible in the frosted glass cover where the photodiode unit will later sit.</li>
		<li>Turn the front-to-back knob counterclockwise to move the spot down the cantilever towards the tip until the spot on the frosted glass disappears.</li>
		<li>Turn the front-to-back knob clockwise only slightly in order to position the laser so that it just sits on the cantilever tip. The laser spot will reappear on the frosted glass. 
		NOTE: This is the laser positioning method for diving-board cantilevers. For triangular cantilevers, the alignment process differs slightly.</li>
	</ol>
	</li>
	<li>Once the laser is aligned, hold the stage lift motor in the &#8220;open&#8221; position for 10 sec to ensure enough space between the cantilever and sample when attaching the sample to the AFM.</li>
	<li>Place sample on the KPFM sample stage. Ground the sample using a copper wire. Connect the appropriate wires from the AFM to the sample stage. Connect the stage to the AFM. In most cases the stage is held in place using magnets.</li>
	<li>In the AFM software, tune the cantilever and then begin approach of the cantilever tip to the sample. Ensure that settling time is set at no more than 10 msec, and that approach speeds do not exceed 2.0 &#181;m/sec in order to avoid tip damage.</li>
	<li>Once the cantilever tip is on the sample surface, select an imaging window size and ideal imaging area. Optimize I and P gains along with the cantilever set point so that topographical imaging is optimized.
	<ol>
		<li>Once topographical imaging is optimized, turn on KPFM module (either FM or AM mode; here, use FM mode). It should be noted that KPFM imaging is very challenging outside of 5 &#181;m x 5 &#181;m imaging window. Also, for optimal KPFM imaging, use a 512 x 512 resolution with slow scan speeds (0.02 - 0.05 lines/sec).</li>
	</ol>
	</li>
	<li>For FM-KPFM settings (not ACAFM settings), set the drive percent between 5 - 10%. Set the cantilever frequency between 1 - 5 kHz with a bandwidth of 2 kHz. Set I and P gains for the FM-KPFM settings to 0.3%.
	<ol>
		<li>Vary signal bandwidth and I and P gains between sample surfaces. To optimize SP imaging, further adjust signal bandwidth and I and P gains to obtain optimal images. The recommended range of I and P gains are from 0.22 - 0.35%.</li>
	</ol>
	</li>
	<li>Process and analyze collected KPFM images using post-image processing software to collect SP data.</li>
</ol>

<ol>
	<li>Seth, A. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+modeling+of+biofilm-infected+wounds:+a+review.">In vivo modeling of biofilm-infected wounds: a review.</a> J. Surg. Research. 178 (1), 330-338 (2012).</li><li>Wolcott, R. D., Ehrlich, G. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biofilms+and+chronic+infections.">Biofilms and chronic infections.</a> JAMA. 299 (22), 2682-2684 (2008).</li><li>Hoiby, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+clinical+impact+of+bacterial+biofilms.">The clinical impact of bacterial biofilms.</a> Micro. Infect. 3 (2), 55-65 (2011).</li><li>Zhang, W., Hughes, J., Chen, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impacts+of+hematite+nanoparticle+exposure+on+biomechanical,+adhesive,+and+surface+electrical+properties+of+E.+coli+cells.">Impacts of hematite nanoparticle exposure on biomechanical, adhesive, and surface electrical properties of E. coli cells.</a> Appl. Environ. Microbiol. 78 (11), 3905-3915 (2012).</li><li>Lorite, G. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surface+physicochemical+properties+at+the+micro+and+nano+length+scales:+role+on+bacterial+adhesion+and+Xylella+fastidiosa+biofilm+development.">Surface physicochemical properties at the micro and nano length scales: role on bacterial adhesion and Xylella fastidiosa biofilm development.</a> PLoS One. 8 (9), (2013).</li><li>Lee, I., Chung, E., Kweon, H., Yiacoumi, S., Tsouris, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+surface+potential+microscopy+of+spore+adhesion+on+surfaces.">Scanning surface potential microscopy of spore adhesion on surfaces.</a> Coll. Surf. Biointer. 92, 271-276 (2012).</li><li>Tsai, C., Hung, H., Liu, C., Chen, Y., Pan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+in+plasma+membrane+surface+potential+of+PC12+cells+as+measured+by+Kelvin+probe+force+microscopy.">Changes in plasma membrane surface potential of PC12 cells as measured by Kelvin probe force microscopy.</a> PLoS One. 7 (4), (2012).</li><li>Jucker, B. A., Harms, H., Zehnder, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adhesion+of+the+positively+charged+bacterium+Strenotrophomonas+(Xanthomonas)+maltophilia+70401+to+glass+and+Teflon.">Adhesion of the positively charged bacterium Strenotrophomonas (Xanthomonas) maltophilia 70401 to glass and Teflon.</a> J. Bacteriology. 178 (18), 5472-5479 (1996).</li><li>Soon, R. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Different+surface+charge+of+colistin-susceptible+and+-resistant+Acinetobacter+baumannii+cells+measured+with+zeta+potential+as+a+function+of+growth+phase+and+colistin+treatment.">Different surface charge of colistin-susceptible and -resistant Acinetobacter baumannii cells measured with zeta potential as a function of growth phase and colistin treatment.</a> J. Anti. Chemo. 66, 126-133 (2011).</li><li>Tariq, M., Bruijs, C., Kok, J., Krom, B. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Link+between+culture+zeta+potential+homogeneity+and+Ebp+in+Enterococcus+faecalis.">Link between culture zeta potential homogeneity and Ebp in Enterococcus faecalis.</a> Appl. Environ. Microbiol. 78 (7), 2282-2288 (2012).</li><li>Ayala-Torres, C., Hernandez, N., Galeano, A., Novoa-Aponte, L., Soto, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zeta+potential+as+a+measure+of+the+surface+charge+of+mycobacterial+cells.">Zeta potential as a measure of the surface charge of mycobacterial cells.</a> Ann Microbiol. , (2013).</li><li>Allison, D. P., Mortensen, N. P., Sullivan, C. J., Doktycz, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atomic+force+microscopy+of+biological+samples.">Atomic force microscopy of biological samples.</a> Nanomed. Nanobiotech. 2 (6), 613-634 (2010).</li><li>Lindsay, S. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+tunneling+microscopy+and+atomic+force+microscopy+studies+of+biomaterials+at+a+liquid-solid+interface.">Scanning tunneling microscopy and atomic force microscopy studies of biomaterials at a liquid-solid interface.</a> J. Vac. Sci. Technol. 11 (4), 808-815 (1993).</li><li>Moores, B., Hane, F., Eng, L., Leonenko, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kelvin+probe+force+microscopy+in+application+to+biomolecular+films:+frequency+modulation,+amplitude+modulation,+and+lift+mode.">Kelvin probe force microscopy in application to biomolecular films: frequency modulation, amplitude modulation, and lift mode.</a> Ultramicroscopy. 110 (6), 708-711 (2010).</li><li>Melitz, W., Shen, J., Kummel, A. C., Kelvin Lee, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=probe+force+microscopy+and+its+application.">probe force microscopy and its application.</a> Surf. Sci. Reports. 66 (1), 1-27 (2011).</li><li>Loppacher, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FM+demodulated+Kelvin+probe+force+microscopy+for+surface+photovoltage+tracking.">FM demodulated Kelvin probe force microscopy for surface photovoltage tracking.</a> Nanotechnology. 16 (3), (2005).</li><li>Domanski, A. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kelvin+probe+force+microscopy+in+nonpolar+liquids.">Kelvin probe force microscopy in nonpolar liquids.</a> Langmuir. 28 (39), 13892-13899 (2012).</li><li>Collins, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual+harmonic+Kelvin+probe+force+microscopy+at+the+graphene-liquid+interface.">Dual harmonic Kelvin probe force microscopy at the graphene-liquid interface.</a> Appl. Phys. Letters. 104 (13), 133103 (2014).</li><li>Kobayashi, N., Asakawa, H., Fukuma, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanoscale+potential+measurements+in+liquid+by+frequency+modulation+atomic+force+microscopy.">Nanoscale potential measurements in liquid by frequency modulation atomic force microscopy.</a> Rev. Sci. Instru. 81, (2010).</li><li>Collins, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Probing+charge+screening+dynamics+and+electrochemical+processes+at+the+solid-liquid+interface+with+electrochemical+force+microscopy.">Probing charge screening dynamics and electrochemical processes at the solid-liquid interface with electrochemical force microscopy.</a> Nature Comm. 5, 2871 (2014).</li><li>Pastar, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interactions+of+methicillin+resistant+Staphylococcus+aureus+USA300+and+Pseudomonas+aeruginosa+in+polymicrobial+wound+infection.">Interactions of methicillin resistant Staphylococcus aureus USA300 and Pseudomonas aeruginosa in polymicrobial wound infection.</a> PLoS One. 8 (2), (2013).</li><li>Brien, D. J., Gould, I. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Does+vancomycin+have+a+future+in+the+treatment+of+skin+infections.">Does vancomycin have a future in the treatment of skin infections.</a> Cur. Opin. Infec. Diseas. 27 (2), 146-154 (2014).</li><li>Wang, P., Kinraide, T. B., Zhou, D., Kopittke, P. M., Peijnenburg, W. J. G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasma+membrane+surface+potential:+dual+effects+upon+ion+uptake+and.">Plasma membrane surface potential: dual effects upon ion uptake and.</a> 155 (2), 808-820 (2011).</li><li>Gross, M., Cramton, S. E., Gotz, F., Peschel, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Key+role+of+teichoic+acid+net+charge+in+Staphylococcus+aureus+colonization+of+artificial+surfaces.">Key role of teichoic acid net charge in Staphylococcus aureus colonization of artificial surfaces.</a> Infect. Immun. 69 (5), 3423-3426 (2001).</li><li>Sinensky, A. M., Belcher, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+and+high-resolution+protein/DNA+nanoarray+analysis+using+Kelvin+probe+force+microscopy.">Label-free and high-resolution protein/DNA nanoarray analysis using Kelvin probe force microscopy.</a> Nat. Nanotechnol. 2, 653-659 (2007).</li><li>Park, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+recognition+of+biomolecular+interaction+via+kelvin+probe+force+microscopy.">Single-molecule recognition of biomolecular interaction via kelvin probe force microscopy.</a> ACS Nano. 5 (9), 6981-6990 (2011).</li></ol>

The authors have nothing to disclose.

KPFM was employed as a novel technique for obtaining surface electrical data. It has commonly been used as a method for examining charge distribution in chemistry and has only recently begun to be applied for the study of biological systems on the micro- and nano-scales. From the data collected we found that microbes did not appear to readily attach to clean stainless steel and gold surfaces, even after 3 hr of static incubation. Poly-L-lysine functionalized surfaces showed rapid microbial attachment after 30 min. Longer...

Biofilms produced on equipment surfaces and in cutaneous wounds present a problem for the medical industry as biofilms are recalcitrant to removal and can lead to increased rates of disease transmission and antimicrobial resistance. Attachment is the first step in biofilm formation and is the most critical step due to its reversibility1-3. Substrate surface characteristics play a crucial role on microbial attachment. Factors such as surface hardness, porosity, roughness, and hydrophobicity have been shown to effect microbial attachment; however, little research examining the role of substrate surface potential (SP) on microbial adhesion has been done4,5. Negatively charged surfaces prevent the attachment of Bacillus thuringiensis spores to mica, silicon, and gold6. Changes in cellular membrane potential are indicative of changes in cellular attachment and motility5,7. It has been observed that electrically homogeneous surfaces more readily promote microbial adhesion5. Characterizing the surface potential of bacteria at the nanoscale can provide a novel way to understand the adhesion kinetics of bacteria to various surfaces, and thereby can aid in the development of anti-biofouling strategies. Unlike other methods used for characterizing the electro kinetics of bacteria, such as electrophoretic light scattering, zeta potential, and isoelectric point determination, Kelvin probe force microscopy (KPFM) allows for the examination of individual cells instead of entire cultures8-11. This is beneficial when wanting to compare cell-cell or biofilm electrical characteristics with high accuracy and precision.
KPFM is a module of atomic force microscopy (AFM). AFM was developed as a direct result of the scanning tunneling microscope (STM)12. The first published images using STM were done by Gerd Binnig and Heinrich Rorhrer in 198212. Their invention was able to resolve atomic structures by raster scanning a sharp conductive tip over a conductive surface in air. The implications of achieving high-resolution images excited biologists who quickly attempted to use STM to image dried samples of DNA, proteins, and viruses12. STM can also be done in liquids using specialized probe tips13. This was shown by Lindsay et al., who used STM and AFM to image DNA molecules in 10 mM HClO4 and in water, on gold electrodes13. KPFM has been demonstrated in determining the Surface potentials of DNA and protein analysis25 and biomolecules interaction with ligands26.
KPFM operates by measuring the contact potential difference (CPD) between an AFM conductive cantilever tip and an ideally conductive sample (Figure 1, i)14,15. Samples do not necessarily have to be conductive (i.e., biological samples). Imaging can be carried out on mica, glass, and silicon surfaces (non-conductive) as long as the non-conductive surface is thin and there is an underlying conductive material6,7. The CPD is equivalent to the surface potential voltage and can be described as the difference in the work functions between the tip (&#966;tip) and sample (&#966;sample), divided by the negative electron charge (-e). When a conductive AFM tip is brought close to a sample surface (separated by distance d), an electrostatic force (Fes) is generated due to the difference in Fermi energies (Figure 1, ii, a)15. At this point, the vacuum energies of the tip and sample (Ev) are in equilibrium and aligned. Upon bringing the tip closer to the sample surface, the tip and sample surface come into electrical contact and act as parallel plate capacitors (Figure 1, ii, b)14,15. At the point, the Fermi energies of the tip and sample surface become aligned, reaching a steady-state equilibrium (Figure 1, ii, b). The tip and sample surface will be charged and a VCPD will form due to a difference in Ev&#8217;s and work functions. A Fes acts on the electrical contact area due to the formed VCPD. This force is subsequently nullified through the application of an external VDC to the tip that has the same magnitude as the formed VCPD (Figure 1, ii, c). This applied DC voltage eliminates surface charge in the area of electrical contact, and the amount of VDC necessary to eliminate the Fes of VCPD is equal to the difference in the work functions between the sample surface and tip15. It should be noted that the work function of the tip is known and it is provided by manufacturers. In all KPFM methods, an AC voltage (VAC, approximately 100 - 500 mV) is also applied to the tip in order to generate oscillating electrical forces between the tip and sample14. This provides better resolution when measuring changes in VCPD and/or Fes. In this regard, changes in the frequency or amplitude of electrical oscillation can be corrected by VDC, and surface potential maps can be generated. Data from specific areas of these maps can be further analyzed to provide electrical information about specific topographical features.
KPFM can be operated in three modes: (1) lift mode, (2) amplitude modulation (AM) mode, and (3) frequency modulation (FM) mode14,16. Lift mode was the initial incarnation of KPFM. Lift mode relies on a two-pass method in which an oscillated tip is dragged across the surface to obtain a topographical image. For the second pass the tip is raised a preset distance above the sample (10 - 100 nm) and scanned back across the same area. Due to this two-pass method, lift mode, compared to AM- and FM-KPFM, takes the longest time for image acquisition. Raising the tip away from the surface ensures that only long-range Fes is measured. Also, cross talk between topography and surface potential measurements is decoupled at the expense of increased lateral resolution and sensitivity.
AM-KPFM improves lateral resolution and sensitivity by using dual-frequencies to simultaneously measure sample topography and surface potential (one-pass scan)14. In AM mode, the cantilever is mechanically oscillated, generally 5% below its first resonant frequency (f0), and electrically oscillated (through a VAC) at its second resonant frequency (f1). Changes in the amplitude of f0 lead to the generation of topographical data, while changes in the amplitude of f1, due to changes in Fes and VCPD, give surface potential measurement data. f0 and f1 of the cantilever are separated by significant frequencies and energies that signal cross-talk is minimized14. The head electronics box (HEB) of the AFM separates out the two signals to give both topographical and surface potential data simultaneously in one scan. FM-KPFM improves resolution even further than AM-KPFM on biological surfaces14. FM-KPFM works differently than AM-KPFM in that it measures changes in electrostatic force gradients rather than electrostatic force (Fes)15. Like AM mode, FM mode utilizes dual-frequencies and a single-pass scanning mechanism to obtain topographical and surface potential data simultaneously14. In FM mode, the cantilever is mechanically oscillated at f0 and electrically oscillated at a low modified frequency (fmod, typically 1 - 5 kHz). Upon electrostatic interactions, f0 and fmod mix to produce side bands f0 &#177; fmod. These sidebands are very sensitive to changes in electrostatic force, and can be separated from f0 through the AFM&#8217;s HEB. Since FM-KPFM measures changes in electrostatic force gradients, the tips apex shape and its maintenance/integrity play a critical role in overall surface potential resolution14, 15. Surface potential resolution using AM and FM modes are in the range of 1 nm laterally14-16. It should be noted that KPFM imaging can be done in non-polar liquids, and more recently, has been shown to be done in low-ionic (&#60;10 mM) polar liquids (in open-loop KPFM modes which do not require a bias feedback, obviating the application of a DC bias) such as MilliQ water; however, KPFM imaging has yet to be done on live cells in polar solutions17-20. Additional challenges associated with SP imaging in liquid is that solutions commonly used for maintaining cells (i.e., phosphate buffered saline) have high concentrations of mobile ions, which would lead to Faradaic reactions, bias-induced charge dynamics, and ion diffusion/redistribution20. Thus, for this experiment, measurements were taken from dried and dead MRSA cells on poly-L-lysine functionalized stainless steel and gold surfaces under ambient conditions. Imaging can be carried out in air or vacuum conditions on biological samples that have been previously dried or immobilized onto surfaces20. Humid conditions have also been shown to affect KPFM imaging of surfaces6.
In this study, we employed FM-KPFM and AFM to examine the role of SP on the attachment of methicillin-resistant Staphylococcus aureus USA100 (MRSA) to poly-L-lysine functionalized stainless steel and gold surfaces. MRSA has recently garnered status as a multi-drug resistant (MDR) &#8220;superbug&#8221; due to its naturally selected resistance to many &#946;-lactam antibiotics and cephalosporins21. MRSA infections are now more challenging, difficult, and formidable to treat, leading to the use of harsher antimicrobials such as vancomycin or oxazolidinones which have higher toxicity levels in humans, and therefore cannot be used as long-term treatments22. Stainless steel was chosen due to its medical relevance and common use as a material in hypodermic needles, bedpans, door handles, sinks, etc. Gold was used as a comparative metal. FM-KPFM was utilized to examine if microbial membrane SP changes upon attachment to the substrates.

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			<td style="width:167px;">Comments/Description</td>
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			<td height="63" style="height:63px;width:216px;">5500ILM Atomic Force Microscope</td>
			<td style="width:71px;">Agilent Technologies</td>
			<td style="width:119px;">#N9435S</td>
			<td></td>
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			<td height="63" style="height:63px;width:216px;">AFM/STM Metal Specimen Discs</td>
			<td style="width:71px;">TED PELLA, INC.</td>
			<td style="width:119px;">#16219</td>
			<td>Stainless steel sample discs</td>
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			<td height="42" style="height:42px;width:216px;">DPE (Low-Noise) Conductive SPM Probes</td>
			<td style="width:71px;">Mikromasch</td>
			<td style="width:119px;">#HQ:DPE-XSC11</td>
			<td>There are 4 Pt-coated cantilevers per chip.&#160; We utilized cantilever B for experiments.</td>
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			<td height="63" style="height:63px;width:216px;">PELCO Gold Coated AFM/STM Metal Specimen Discs</td>
			<td style="width:71px;">TED PELLA, INC.</td>
			<td style="width:119px;">#16218-G</td>
			<td>Gold sample discs</td>
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			<td height="63" style="height:63px;width:216px;">PicoView Software</td>
			<td style="width:71px;">Agilent Technologies</td>
			<td style="width:119px;">#N9797B</td>
			<td>5500ILM Atomic Force Microscope imaging software</td>
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			<td height="63" style="height:63px;width:216px;">Pico Image Software (Pico Image Basic)</td>
			<td style="width:71px;">Agilent Technologies</td>
			<td style="width:119px;">#N9797AU-1FP</td>
			<td>5500 ILM Atomic Force Microscope post-image processing software</td>
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			<td height="42" style="height:42px;width:216px;">Scilogex D3024 High Speed Micro-Centrifuge</td>
			<td style="width:71px;">Thomas Scientific</td>
			<td style="width:119px;">#91201513</td>
			<td>Centrifuge used in cell-washing steps to separate cells (pellet) from media</td>
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			<td height="42" style="height:42px;width:216px;">Trypticase Soy Agar with 5% Sheep Blood</td>
			<td style="width:71px;">BD</td>
			<td style="width:119px;">#221261</td>
			<td>Pre-made plates</td>
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			<td height="21" style="height:21px;width:216px;">Tryptic Soy Broth</td>
			<td style="width:71px;">BD</td>
			<td style="width:119px;">#257107</td>
			<td>Comes as a dry powder.&#160; Instruction on how to make come on the container.</td>
		</tr>
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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.

The ability to measure SP using KPFM relies on the principle that both the sample surface and cantilever tip are conductive to some extent. Stainless steel and gold acted as conductive surfaces to which MRSA were attached. KPFM images were taken of 15 MRSA cells on both surfaces with 512 x 512 resolutions, and with scan areas ranging from 5 x 5 &#181;m to 10 x 10 &#181;m. Scanning was carried out with line speeds ranging from 0.02 lines/sec to 0.05 lines/sec. Therefore, with 512 data points collected per line and 512 lin...

Watch this Scientific Journal Video about Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy at JoVE.com

Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy

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. ...

surface-potential-measurement-bacteria-using-kelvin-probe-force

Research

JoVE Journal

Bioengineering

21.7K Görüntüleme.  University of Guelph. 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. 

Video: Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy

Measurement of Bioelectric Current with a Vibrating Probe

Electric fields, generated by active transport of ions, are present in many biological systems and often serve important functions in tissues and organs. For example, they play an important role in directing cell migration during wound healing. Here we describe the manufacture and use of ultrasensitive vibrating probes for measuring extracellular electric currents. The probe is an insulated, sharpened metal wire with a small platinum-black tip (30-35 &mu;m), which can detect ionic currents in the &mu;A/cm2 range in physiological saline. The probe is vibrated at about 200 Hz by a piezoelectric bender. In the presence of an ionic current, the probe detects a voltage difference between the extremes of its movement. A lock-in amplifier filters out extraneous noise by locking on to the probe's frequency of vibration. Data are recorded onto computer. The probe is calibrated at the start and end of experiments in appropriate saline, using a chamber which applies a current of exactly 1.5 &mu;A/cm2. We describe how to make the probes, set up the system and calibrate. We also demonstrate the technique of cornea measurement, and show some representative results from different specimens (cornea, skin, brain).

The manufacture, calibration and use of non-invasive vibrating probes to measure bioelectric current in various biological systems is described.

Electric fields, generated by active transport of ions, are present in many biological systems and often serve important functions in tissues and organs. ...

Measurement of Aggregate Cohesion by Tissue Surface Tensiometry

Rigorous measurement of intercellular binding energy can only be made using methods grounded in thermodynamic principles in systems at equilibrium. We have developed tissue surface tensiometry (TST) specifically to measure the surface free energy of interaction between cells. The biophysical concepts underlying TST have been previously described in detail1,2. The method is based on the observation that mutually cohesive cells, if maintained in shaking culture, will spontaneously assemble into clusters. Over time, these clusters will round up to form spheres. This rounding-up behavior mimics the behavior characteristic of liquid systems. Intercellular binding energy is measured by compressing spherical aggregates between parallel plates in a custom-designed tissue surface tensiometer. The same mathematical equation used to measure the surface tension of a liquid droplet is used to measure surface tension of 3D tissue-like spherical aggregates. The cellular equivalent of liquid surface tension is intercellular binding energy, or more generally, tissue cohesivity. Previous studies from our laboratory have shown that tissue surface tension (1) predicts how two groups of embryonic cells will interact with one another1-5, (2) can strongly influence the ability of tissues to interact with biomaterials6, (3) can be altered not only through direct manipulation of cadherin-based intercellular cohesion7, but also by manipulation of key ECM molecules such as FN8-11 and 4) correlates with invasive potential of lung cancer12, fibrosarcoma13, brain tumor14 and prostate tumor cell lines15. In this article we will describe the apparatus, detail the steps required to generate spheroids, to load the spheroids into the tensiometer chamber, to initiate aggregate compression, and to analyze and validate the tissue surface tension measurements generated.

We describe a method of measuring binding energy, expressible as tissue surface tension, between cells within 3D tissue-like aggregates. Differences in tissue surface tension have been demonstrated to correlate with invasiveness of lung, muscle, and brain tumors, and are fundamental determinants of establishing spatial relationships between different cell types.

Rigorous measurement of intercellular binding energy can only be made using methods grounded in thermodynamic principles in systems at equilibrium.  We ...

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

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

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

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

Biosensor for Detection of Antibiotic Resistant Staphylococcus Bacteria

A structurally transformed lytic bacteriophage having a broad host range of Staphylococcus aureus strains and a penicillin-binding protein (PBP 2a) antibody conjugated latex beads have been utilized to create a biosensor designed for discrimination of methicillin resistant (MRSA) and sensitive (MSSA) S. aureus species 1,2. The lytic phages have been converted into phage spheroids by contact with water-chloroform interface. Phage spheroid monolayers have been moved onto a biosensor surface by Langmuir-Blodgett (LB) technique 3. The created biosensors have been examined by a quartz crystal microbalance with dissipation tracking (QCM-D) to evaluate bacteria-phage interactions. Bacteria-spheroid interactions led to reduced resonance frequency and a rise in dissipation energy for both MRSA and MSSA strains. After the bacterial binding, these sensors have been further exposed to the penicillin-binding protein antibody latex beads. Sensors analyzed with MRSA responded to PBP 2a antibody beads; although sensors inspected with MSSA gave no response. This experimental distinction determines an unambiguous discrimination between methicillin resistant and sensitive S. aureus strains. Equally bound and unbound bacteriophages suppress bacterial growth on surfaces and in water suspensions. Once lytic phages are changed into spheroids, they retain their strong lytic activity and show high bacterial capture capability. The phage and phage spheroids can be utilized for testing and sterilization of antibiotic resistant microorganisms. Other applications may include use in bacteriophage therapy and antimicrobial surfaces.

Lytic phage biosensors and antibody beads are able to discriminate between methicillin resistant (MRSA) and sensitive staphylococcus bacteria. The phages were immobilized by a Langmuir-Blodgett method onto a surface of a quartz crystal microbalance sensor and worked as broad range staphylococcus probes. Antibody beads recognize MRSA.

A structurally transformed lytic bacteriophage having a broad host range of Staphylococcus aureus strains and a penicillin-binding protein (PBP 2a) ...

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, ...

A Novel Method for Localizing Reporter Fluorescent Beads Near the Cell Culture Surface for Traction Force Microscopy

PA gels have long been used as a platform to study cell traction forces due to ease of fabrication and the ability to tune their elastic properties. When the substrate is coated with an extracellular matrix protein, cells adhere to the gel and apply forces, causing the gel to deform. The deformation depends on the cell traction and the elastic properties of the gel. If the deformation field of the surface is known, surface traction can be calculated using elasticity theory. Gel deformation is commonly measured by embedding fluorescent marker beads uniformly into the gel. The probes displace as the gel deforms. The probes near the surface of the gel are tracked. The displacements reported by these probes are considered as surface displacements. Their depths from the surface are ignored. This assumption introduces error in traction force evaluations. For precise measurement of cell forces, it is critical for the location of the beads to be known. We have developed a technique that utilizes simple chemistry to confine fluorescent marker beads, 0.1 and 1 &#181;m in diameter, in PA gels, within 1.6 &#956;m of the surface. We coat a coverslip with poly-D-lysine (PDL) and fluorescent beads. PA gel solution is then sandwiched between the coverslip and an adherent surface. The fluorescent beads transfer to the gel solution during curing. After polymerization, the PA gel contains fluorescent beads on a plane close to the gel surface.

Traditional techniques for fabricating polyacrylamide (PA) gels containing fluorescent probes involve sandwiching a gel between an adherent surface and a glass slide. Here, we show that coating this slide with poly-D-lysine (PDL) and fluorescent probes localizes the probes to within 1.6 &#181;m from the gel surface.

PA gels have long been used as a platform to study cell traction forces due to ease of fabrication and the ability to tune their elastic properties. When ...

Measurement of Maximum Isometric Force Generated by Permeabilized Skeletal Muscle Fibers

Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle function at the level of the single muscle cell. Single muscle fiber studies are useful in both basic science and clinical studies. For basic studies, single muscle fiber contractility measurements allow investigation of fundamental mechanisms of force production, and analysis of muscle function in the context of genetic manipulations. Clinically, single muscle fiber studies provide useful insight into the impact of injury and disease on muscle function, and may be used to guide the understanding of muscular pathologies. In this video article we outline the steps required to prepare and isolate an individual skeletal muscle fiber segment, attach it to force-measuring apparatus, activate it to produce maximum isometric force, and estimate its cross-sectional area for the purpose of normalizing the force produced.

Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle function at the level of the single muscle cell. In this article we outline a valid and reliable technique to prepare and test permeabilized skeletal muscle fibers in vitro.

Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle ...

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 ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Application of Atomic Force Microscopy to Detect Early Osteoarthritis

Biomechanical properties of cells and tissues not only regulate their shape and function but are also crucial for maintaining their vitality. Changes in elasticity can propagate or trigger the onset of major diseases like cancer or osteoarthritis (OA). Atomic force microscopy (AFM) has emerged as a strong tool to qualitatively and quantitatively characterize the biomechanical properties of specific biological target structures on a microscopic scale, measuring forces in a range from as small as the piconewton to the micronewton. Biomechanical properties are of special importance in musculoskeletal tissues, which are subjected to high levels of strain. OA as a degenerative disease of the cartilage results in the disruption of the pericellular matrix (PCM) and the spatial rearrangement of the chondrocytes embedded in their extracellular matrix (ECM). Disruption in PCM and ECM has been associated with changes in the biomechanical properties of cartilage. In the present study we used AFM to quantify these changes in relation to the specific spatial pattern changes of the chondrocytes. With each pattern change, significant changes in elasticity were observed for both the PCM and ECM. Measuring the local elasticity thus allows for drawing direct conclusions about the degree of local tissue degeneration in OA.

We present a method to investigate early osteoarthritic changes at the cellular level in articular cartilage by using atomic force microscopy (AFM).

Biomechanical properties of cells and tissues not only regulate their shape and function but are also crucial for maintaining their vitality. Changes in ...