Life Science

18 Mar 2022

Investigation of lipid bilayer formation by AFM

Introduction Cell membranes are highly complex structures in which a variety of cellular activities occur in response to both specific and non-specific extracellular signals [1,2]. Extracellular signals transmitted by membrane receptors regulate gene expression and regulate cell proliferation. Although the lipid bilayer constitutes the major structural component of the cell membrane, extracellular matrix (ECM) proteins coat the lipid membrane and provide a reactive surface for other cells and biomaterials to come into contact with [3,4,5]. Therefore, understanding the cell membrane is the first step in the study of cells, the fundamental building block of life itself. In this app note, we present a fundamental study on the lipid bilayer, which is the basic structural component of a cell membrane. As a first step, a defect-free lipid bilayer formation is essential. To form a continuous film, lipids are formed into vesicles using the extrusion method. These vesicles adsorb onto the solid support, where neighboring vesicles will coalesce. When a critical size is reached, the adsorbed vesicles rupture upon forming discs of bilayers that will eventually merge into a film. This bilayer formation strongly depends on the interaction between two vesicles and between a vesicle and the solid support, which is commonly silicon dioxide due to its hydrophilicity. (Fig.1). To investigate the lipid bilayer, a few measurement techniques are adopted such as the quartz crystal microbalance (QCM), which is a nanogram sensitive weighing balance that utilizes the piezoelectric properties of quartz crystal as the working principle [6], fluorescent recovery after photobleaching (FRAP), which utilizes the recovery of fluorescence intensity in an area of the membrane where the fluorophores are bleached [7]. The atomic force microscopy (AFM) [8] is a good candidate for investigating the lipid bilayer. It has the advantage of being able to measure the surface of the sample on a nanometer scale as well as in liquid conditions. In addition, it is an excellent technique for observing the lipid bilayer from various viewpoints because it can measure not only the image of the surface but also the physical and mechanical properties of the sample. In this app note, a fundamental study of the lipid bilayer formation is conducted using AFM. The formation of the lipid bilayer was confirmed using Non-contact and Contact mode and mechanical properties of the lipid bilayer are investigated by force distance spectroscopy by AFM. Fig. 1. Structure of lipid, micelle and, vesicle Materials and Methods The extrusion method was used to prepare vesicles consisting of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC). To create a vesicle with a 50±5 nm diameter, the precursor solution is passing through 100-nm, followed by 30-nm size of filter membranes several times using an extruder. This size-controlled vesicle solution was used to prepare intact vesicle and lipid bilayers on SiO2 substrates [9]. The Contact and Non-contact mode was applied for vesicles or lipid bilayer surface images and force distance spectroscopy was conducted for measuring physical information of lipid bilayer formation. A sharp, silicon nitride AFM probe (BL-AC40TS, 110 kHz resonance frequency, and a 0.09 N/m spring constant) was used for Non-contact mode imaging and force distance spectroscopy and relatively hard AFM probe (BL-AC160TS, 300 kHz resonance frequency, and a 26 N/m spring constant) was used for scratching the lipid bilayer formation in Contact mode. All AFM measurements were performed in liquid (10 mM Tris buffer solution [pH 7.5] with 150 mM NaCl). Lipid bilayer images by AFM As shown in Fig. 2, vesicle adsorption on the SiO2 substrate and lipid bilayer formation by ruptured vesicle were imaged on the nanoscale. The SiO2 substrate shows a very flat and uniform surface (Rpv (peak to valley): 2.38 nm) from which the rougher vesicles are easily distinguished, confirming the expected diameter of 50 to 60 nm. Although the lipid bilayer could also be distinguished from the substrate by an increased surface, direct thickness measurement and the detailed shape of the lipid heads (hydrophilic top part of the lipid molecules) were not accessible due to its very flexible and soft structure. Nevertheless, we confirmed the bilayer formation and measure its thickness by intentionally scraping an area with a silicon tip to remove the bilayer locally. For that, the surface of the lipid bilayer was imaged in Non-contact mode, and then a smaller area within the previously imaged area was “imaged” in Contact mode with high loading force. After that, the larger area was imaged again in Non-contact mode and compared with the first image. As can be seen in Fig. 3, in the area scraped in contact mode, the lipid bilayer was locally removed, exposing the bare substrate. The line profile of this hole reveals a depth of about 6 nm, which matches the thickness of the lipid bilayer very well.   Figure 2. AFM images of the solid substrate (SiO2), vesicles, and lipid bilayer together with 3D images and line profiles.   Figure 3. Confirming the lipid bilayer formation by physically scratching. Physical information of lipid bilayer by force distance spectroscopy Moreover, we confirmed the bilayer formation using force distance spectroscopy. By measuring the nanomechanical properties locally, it is possible to distinguish if a specific layer is covered with a bilayer. For that, a force-distance curve is recorded by monitoring the tip-sample interaction via the vertical deflection of the cantilever versus its Z movement. The bilayer is not deformed without any physical contact between sample and cantilever tip (Fig. 4a). Once the tip comes into contact with the sample, the bilayer is elastically deformed up to a certain force (Fig. 4b). When the force is further increases, the membrane is plastically deformed, and the cantilever penetrates the bilayer (Fig. 4c). The force needed to pierce the bilayer is called the breakthrough force and is reflected by a jump in the vertical deflection during a force distance curve.   Figure 4. Schematic view of the lipid bilayer study by force distance spectroscopy     Figure 5. Force distance spectroscopy result of both lipid bilayer region and non-lipid bilayer region (red points). In fully covered lipid bilayer, relatively flat line profile and breakthrough force was confirmed and in non lipid bilayer region, there was no breakthrough force. Conclusion In this app note, we confirmed vesicles and lipid bilayer formation on a solid SiO2 substrate. The surface of vesicles and lipid bilayers was imaged using Non-contact, revealing a larger surface roughness even on uniform bilayers compared to the uncovered SiO2 substrate. Moreover, we applied contact mode to mechanically scrape the bilayer and thus create a defined hole that allowed to confirm the expected bilayer thickness. Furthermore, we studied the mechanical properties of lipid bilayers using the force distance spectroscopy mode and demonstrated the occurrence of the breakthrough force as a reliable indicator for bilayer formation. Research always starts with the most basics. And the lipid bilayer is the most basic component of the cell that is the source of life. From the study of the lipid bilayer, we may find inspiration for how cells are protected, communicated with, external or internal, and furthermore, cell differentiation and proliferation. AFM, which can measure the surface of a material in nanometers, is a very versatile technique for lipid bilayer research. In addition, since physical and mechanical information as well as image of the surface can be measured together, it will be possible to study the lipid bilayer from various perspectives.

3 Jan 2020

Probiotic bacteria topography and nanomechanical properties analysis using atomic force microscopy

Gabriela Mendoza, Byong Kim and Keibock Lee, Park systems, INC., Santa Clara, CA, USA Introduction Probiotics are living microorganisms that provide numerous health benefits to the host when supplied in adequate amount as well as improve the intestinal microflora of the host. Probiotics are mainly administered to humans combined with food and are widely used in food products. From in vivo and in vitro studies of host intestinal epithelial or immune cell responses to probiotic strains, it has been reported that adhesion of probiotic strains to the intestinal surface and the subsequent colonization of the human gastrointestinal tract are important prerequisites for probiotic action [1]. This study looks into the topography and adhesion properties of those microorganisms to determine the shape and form that adheres well to the colon by means of using commercially available probiotics and the PinPoint nanomechanical mode of an atomic force microscopy (AFM) instrument from Park Systems. Experimental To isolate probiotic bacteria, 1 mL samples of a commercial product were put into microcentrifuge tubes and centrifuged (BioLab centrifuge) for 15 min at 10000 rpm to separate the biomass pellet. The supernatant was discarded and the biomass was recovered in a test tube. Serial dilutions (1 mL of biomass in 9 mL of sterile saline solution) from 10-1 to 10-5 were performed [2]. From the 10-5 dilutions, 0.1 mL drops were placed onto a silicon wafer chip (Ted Pella, Inc. 16007) and allowed to air dry. For bacteria characterization, a Park NX10 AFM from Park Systems was used; measurements were performed in air using an AC160TS cantilever. PinPoint™ Nanomechanical Mode from Park Systems can be used to study surface topography as well as determine quantitative nanomechanical properties (i.e., modulus, adhesion force, adhesion energy) [3]. In this study, topography, adhesion energy and adhesion force images were acquired using the PinPoint Nanomechanical Mode. Images obtained were 256 x 256 pixels and 2 μm x 2 μm sizes.   Figure 1. a) Adhesion energy, b) adhesion force and c) height images taken with AC160TS cantilever. d) Corresponding overlaid topography, adhesion energy and force line profiles. Result Figures 1a, b and c are the adhesion energy, adhesion force and topography images, respectively, of isolated Streptococcus chains from a 10-5 dilution. Figure 1d shows the corresponding overlaid topography, adhesion energy and force line profiles. The adhesion energy (Figure 1a) and adhesion force (Figure 1b) behaved in a similar manner; smaller values were observed for the bacteria compared with the substrate where they were attached. The adhesion energy on the silicon wafer chip was 400 aJ, but just 200 aJ on the probiotic bacteria. Similar results were obtained for the adhesion force (Figure 1d); 20 nN for the bacteria but 45 nN on the silicon wafer chip and clearly indicates that less force was required on the bacteria. The isolated bacteria in air is observed to be relatively less adhesive than that of the underlying Si substrate. Bacterial adhesion is based on physical interactions between two or more surfaces. There were three different surfaces identified in this study: the cantilever tip (surface 1), the bacteria (surface 2) and the silicon wafer (surface 3). All three surfaces have different physicochemical properties: surface area, surface energy, surface roughness size and shape, which affect the adhesion force and energy. Conclusion The test results show that probiotic bacteria under study were less adhesive to the Si cantilever tip compared to the Si substrate. This phenomenon can be correlated with the adhesion of probiotic strains to the site of action, an important prerequisite for probiotic action. We found from nanomechanical measurements that the adhesion force and adhesion energy are affected by the physicochemical characteristics from each substrate. Therefore, adhesion force and adhesion energy values were lower when the measurement was on the bacteria compared to that done on the silicon wafer. References [1] Saarela M et al.,Journal of Biotechnology. 2000; (84) 197-215. [2] Mendoza M AG et al., Revista Mexicana de Ingeniería Química. 2017; 16(1): 159-168. [3] Park Systems’s PinPoint™ Nanomehcanical mode https://www.parksystems.com/index.php/en/parkafm- modes/nanomechanical-modes?i=0  

7 Apr 2019

Mechanical properties of live and fixed cells measured by atomic force microscopy and scanning ion conductance microscopy

Jake Kim, Moses Lee and Cathy Lee Park Systems Corp., Suwon, Korea     Abstract During the process of cell sample preservation, cell fixation plays an essential role for a wide range of biological assays. To determine the importance of this procedure, a comparative analysis was performed between live and fixed cells using Scanning Probe Microscopy (SPM) techniques such as Atomic Force Microscopy (AFM) and Scanning Ion Conductance Microscopy (SICM) for mechanical property measurements and surface fluctuation measurements, respectively. Results from these techniques provide a broad understanding of how cell fixation affects the mechanical properties of cells, and offer new opportunities for the establishment of cell fixation protocols.   Introduction One of the primary objectives of cell fixation for in-vitro studies is preservation of the cell or cellular components in a life-like state by conserving essential chemical and physical characteristics of the cells. Cell fixation is also useful in immunostaining by allowing antibodies to access intracellular structures [1]. Comparing live and fixed cells, the fixed cell maintains a more uniform architecture over the entire cell surface [2]. However, there is no systematic assessment or correlation between changes in mechanical properties of live and fixed cells. By observing various states of cell fixation and the living cells,there is an opportunity to optimize cell fixation protocols, and gain valuable knowledge in aiding the process of cell fixation. Many fixation agents are available for cross-linking cell membranes and cytoplasmic proteins, with paraformaldehyde (PFA) as the most widely used for cell and tissue samples [1-3]. PFA works by covalently cross-linking molecules, bonding them together and creating an insoluble meshwork that alters the mechanical properties of the cell surface. In order to investigate the mechanical properties of live and fixed cell structures fixed with PFA, Atomic Force Microscopy (AFM) and Scanning Ion Conductance Microscopy (SICM) were used to measure the elastic modulus and surface fluctuation, respectively [3-6]. Experimental set-up Cell sample The cell sample used consisted of mouse fibroblast L929 cells (ATCC, USA) cultured in Dulbecco’s modified eagle medium (DMEM; Invitrogen Life Technique, USA) complemented with 10 % fetal bovine serum (Thermo Fisher Scientific, US) together with 1 % penicillin/streptomycin (Invitrogen Life Technique, USA) at 37 °C in a humidified atmosphere with a 5 % CO2 concentration.Cells (with densities of 1 x 104 / mL) were placed on a 35mm diameter cell culture petri dish (NUNC, Denmark),washed three times with phosphate buffered saline (PBS, Sigma-Aldrich, US) and treated with a 4 % PFA solution for 5 min. The fixed cell samples were rinsed with PBS three times prior to conducting AFM and SICM experiments. Set-up of AFM and SICM The setup for cell imaging consisted of the Park Systems scanning probe microscope (Park NX-Bio, Park Systems, Korea) equipped with an inverted optical microscope (Nikon Corp., Japan) specifically designed for biological applications. The AFM can gather the mechanical properties of a sample as well as cell surface information of soft material samples using SICM. All experiments performed on live cells were carried out in a live cell chamber. The chamber conditions of the live cell were regulated to 37 °C, 5 % CO2 and 95 % humidity and had the necessary environment to sustain living cell cultures. AFM measurements for Young’s modulus of cells The AFM was used to obtain force curve measurements to estimate the Young’s modulus of cells. A commercial cantilever (BL-AC40TS, Olympus, Japan) with a nominal spring constant of < 0.09 N/m was used. Utilizing such a cantilever with a small spring constant allows for a relatively large deflection with the application of a small force and allows for reliable data collection of the cell surface structure. Spring constant calibration of the AFM cantilever was performed using the thermal vibration method. Measurements were performed using 50 forces curves containing 512 data points each (Figure 1a). To analyze the force curves, a Hertz model was implemented using the Park Systems imaging analysis program (Park XEI, Park Systems). The shape of the AFM tip was assumed to be a four-sided pyramid with half cone angle α. As a result, the force (F) applied on the cantilever is expressed as: Where E is the Young’s modulus, ν is the Poisson’s ratio and δ is the indentation (depth). Poisson’s ratio was set to 0.5 and α was set to 35°. The AFM scan rate was set to 1μm/s and the maximum loading force was 8nN. SICM measurement for cell imaging and fluctuation analysis SICM utilizes an ion current that flows between an electrode placed inside a nano-pipette and an external electrode located in a bath solution (Figure 1b). The ion current provides a feedback signal that regulates a constant tip-to-sample distance and allows the nano-pipette to scan topographical information. Despite having low lateral resolution (~30 nm) [7], SICM can display useful topographical measurements without the need to apply mechanical forces onto the sample surface. Figure 2 shows the height images of the live and fixed L929 cell. At first glance, there is no difference between two cases; however, the surface of the fixed cell is slightly rougher due to cross-linking of cell membrane proteins. To perform SICM imaging and ion current-distance (I-D) curve experiments, 80 nm inner diameter nano-pipettes were fabricated from borosilicate capillaries (inner diameter 0.6mm, outer diameter 1.0 mm, World Precision Instruments,SSA) using a CO2 -laser pipette puller (Sutter Instruments, USA). Results and Discussion Young’s modulus of live and fixed cells To determine the stiffness of live and fixed cell surfaces, force spectroscopy measurements were obtained from live cells, fixed cells as well as a solid substrate and those force-distance curves are shown in Figure 3a. Fixed cells show a significantly steeper force curve slope compared to that of live cells. Additionally, the force required for surface indentation is also greater for fixed cells than for live cells. Table 1 presents the average Young’s modulus values which indicated that the stiffness of fixed cells (77.95 kPa) was greater than live cells (8.11 kPa). These AFM cell stiffness measurements suggested that the acting filamentous structures strongly affected the stiffness. Another notable mention is that PFA treatment affects the cross-linking of cell surface proteins, including F-actin filaments. In addition, it was noted that cell stiffening corresponds to protein cross linking [8]. More specifically, the PFA fixation process can be assumed to have a direct link to an increase of cell stiffness depending on the number of randomly distributed cross-linking sites available on the cell surface [9]. Surface fluctuations of live and fixed cells To obtain cell surface fluctuations, the I-D curves were obtained using SICM measurements on live cells, fixed cells and a solid substrate. I-D curves exhibit the steepest slope for solid substrates (petri dish) while untreated they possess a broader slope. For PFA-treated cells, I-D curves occur amidst the two. Subsequently it could be concluded that the live cells demonstrated more activity compared to fixed cells. The PFA treatment process results in smaller surface fluctuations for fixed cells which results in cross-linking of proteins between the membrane and cytoplasmic proteins. Conclusion A fundamental mechanical difference has been demonstrated by comparing live cells and cells fixed with PFA. Atomic force microscopy and scanning ion conductance microscopy measurements revealed a definite transition in surface fluctuation and elastic modulus of cells when exposed to PFA. Proceeding complete fixation with PFA, cell surface fluctuation decreased as compared to a live cell while the Young’s modulus increased by five-fold. These findings provide key insights into how cells react to chemical treatment with PFA. In addition to the traditional knowledge of the chemical influence of PFA on cells, this study has uncovered the effect of PFA on the mechanical properties of the cell’s surface. Cell membranes are assumed to be flexible and variable; however, in certain situations(chemical treatment in particular), transformations occur on biological and morphological levels. Such observations provide a strong motive for further study on cell surface fluctuations as a crucial requirement for the understanding of cell functions regarding cell dynamics. Scanning probe microscopy techniques, specifically the AFM and SICM, clearly serve as vital tools in the application of quantitative studies on both fixed and live cells. Fig 1. Optical images of L929 cell for AFM (a) and SICM (b). Each probe was positioned at the apex of the single cell.   Fig 2. Live (a),(c) and 4% PFA treated (b),(d) cell surface imaging by SICM.   Fig 3. Average force-distance curves (a), and ion-current-distance curves (b) of the solid substrate (black), 4% PFA treated (fixed) cell (red) and live cell (blue).   Table 1. Young’s modulus value.     References 1. Lanier, L. and N. Warner, Paraformaldehyde fixation of hematopoietic cells for quantitative flow cytometry (FACS) analysis. Journal of immunological methods, 1981. 47(1): p. 25-30. 2. Yamane, Y., et al., Quantitative analyses of topography and elasticity of living and fixed astrocytes. Journal of electron microscopy, 2000. 49(3): p. 463-471. 3. Binnig, G., C.F. Quate, and C. Gerber, Atomic force microscope. Physical review letters, 1986. 56(9): p. 930. 4. Korchev, Y.E., et al., Scanning ion conductance microscopy of living cells. Biophysical journal, 1997. 73(2): p. 653. 5. Cappella, B. and G. Dietler, Force-distance curves by atomic force microscopy. Surface science reports, 1999. 34(1): p. 1-104. 6. Mizutani, Y., et al., Nanoscale fluctuations on epithelial cell surfaces investigated by scanning ion conductance microscopy. Applied Physics Letters, 2013. 102(17): p. 173703. 7. Rheinlaender, J., et al., Comparison of scanning ion conductance microscopy with atomic force microscopy for cell imaging. Langmuir, 2010. 27(2): p. 697-704. 8. Hopwood D. Theoretical and practical aspects of glutaraldehyde fixation. InFixation in histochemistry, Springer, Boston, MA. 1973: p. 47-83. 9. Tanaka KA, et al., Membrane molecules mobile even after chemical fixation. Nature Methods. 2010 Nov;7(11):865.  

16 Oct 2018

Tooth Whitening Study using PinPoint Nanomechanical Mode of Park AFM

Alvin Lee, John Paul Pineda, Byong Kim, and Keibock Lee Park Systems Inc., Santa Clara, CA USA Introduction In recent years, the importance of mechanical properties measurement has become evident in various applications, ranging from understanding and screening of integrated circuit failure mechanisms of microelectronics, to diagnosis of diseases in the field of medicine, and to the development of restorative materials for human teeth [1-3]. Some samples when pressed with too much force can lead to surface deformation and change initssurface characteristics which may result to inaccurate assessment of measurements. Moreover, since the sizes and features of interest in functional materials are continuously shrinking downto nanoscale level,and given that material failures start from atomic lattice structures, conventional techniques used in materials characterization are no longer adequatein identifying defects and abnormalities on material surfaces asthey only allow up to millimeter range measurements. For these reasons, a nextgeneration characterization tools that is benign and allow nanoscale measurements is greatly needed.Recently, a new operational AFM mode developed by Park Systems called PinPoint ™ Nanomechanical mode [4] offers researchers an innovative solution to these problems. This technique operates by allowing the tip to move in approach-retract manner with only few nano-Newton force ensuring a frictionless operation thateliminates the lateral force due to continuous tip-sample contact and preserves tip and sample condition. At each point of the image, force-distance curve is acquired and used to calculate the mechanical characteristics of the sample being measured. During the data acquisition, the XY scanner stops, and the contact time is controlled to give enough time for the scanner to acquire precise and accurate data with nanoscale resolution. Every material structure deteriorates over time when kept in natural setting. Such changes originate from a point defect atthe nanoscale level. This also applies to the teeth whitening process. Everyone who whitens teeth desires for the teeth to remain whitened without redoing it too often. Thus, to realize this, one must know how the topological and mechanical changes take place on the surface of teeth at the nanoscale level. In this study, PinPoint Nanomechanical mode was used to acquire topographical and mechanical data of a tooth sample to understand the effect of tooth whitening strips on the tooth’s mechanical and topographical properties. The results show that tooth whitening strips lead to a decrease in tooth roughness and an increase in modulus and adhesion. This newly developed technique by Park Systems will enable researchers to investigate the topographical and mechanical properties variations ofdifferent varieties of samplesatnanoscale level. Experimental A tooth sample was analyzed using a Park NX10 AFM [5] under ambient air condition. PinPoint Nanomechanical mode with AC160TS tip (nominal spring constant k = 26 N/m and resonance frequency f = 300 kHz) [6]was used in the measurement to acquire topography and mechanical signals. Before undergoing AFM scanning, the tooth was treated by placing it in deionized water and left to sit for 30 minutes followed by placing in 50% by volume isopropyl alcohol for 2 minutes, then rinsed again in deionized water. Compressed air spray was used to dry off the tooth. This was in attempt to remove “debris” and “dirt” from the teeth that may have accumulate on to the teeth over the time.Figure 1 shows the setup of the experiment. A carbon tape and super glue were used to stabilize the sample during imaging. Tooth was placed onto a magnetic disk and stuck to double sided carbon tape. To be sure that the tooth was secured and would not move while taking measurements, a toothpick was placed to hold the tooth upward so that an area of the tooth would be leveled for examination. Superglue was used for extra stability.     Fig. 1.Experimental set up of the tooth sample. Tooth underneath Park NX10 AFM system (Left). Tooth with AC160-TS tip landed onto sample (Right) To validate the acquired modulus values, a referencesample was used. The sample was a PS-LDPE blend film (polystyrene and polyolefin elastomer). The scanparameters were setsuch thatthe modulus measurements were measuredto be ~2.0 GPa for the PS matrix, and ~0.1 GPa for the circular features that appeared in the modulus imagesthat are blended into the matrix which were in agreement with the manufacturers’ description of the PS-LDPE sample [7].Figure 2 shows the 15x15 μm acquired modulusimages of PS-LDPE reference sample. The images were analyzed using Park XEI software developed by Park Systemswhich mapped the acquired signals to a color table. The darker regions (color black in Fig. 2 Left)with circular features represent the areas with lower modulus values while the lighter regions (color brown in Fig. 2 Right) with flat surface known to be the PS matrix are the areas with higher modulus. All areas with purple color mean that these areas are excluded in calculating the mean modulus value. The calculated elastic modulus mean of the LDPE copolymer region is 0.116 GPa while for the PS polystyrene flat region is 2.039 GPa. Fig. 2.15x15 μm image of PS-LDPE reference sample. Image with polystyrene flat region excluded in the elastic modulus mean calculation (Left). Image with copolymer region excluded in the elastic modulus mean calculation (Right). After the reference sample measurements were taken, the tooth was placed under the Park NX10 AFM with the same tip used for reference sample measurements. For statistical relevance, three measurements were taken at three different teeth surfaces of the same teeth sample that could be found using the optical microscope built in to the Park AFM. Measurements were taken in areas of the tooth no farther than 500 microns apart so that roughly the same area arebeing imaged consistently. These measurements were performed using the same scan parameters as were used for the PS-LDPE reference sample. These parameters were utilized because under those conditions, the AFM recorded accurate modulus values. To whiten the tooth sample, whitening strips were used [8]. One strip was applied onto the tooth surface and left to sit for the recommended 1-hour period. After this, the tooth was rinsed in deionized water and dried off with compressed air spray. A total of 3 whitening strips were used as the final whitening. Results and Discussion The representative topographical and mechanical images acquired in the experiment were presented on Table1. The images were also analyzed using the Park XEI software. The quantitative results of elastic modulus are shown in Table 2. The experimental values of the elastic modulus before and after the whitening process were compared on the previous studies [2].The tooth is composed of three major layers, known as the enamel, dentin, and cementum. These layers are found in the respective order: enamel at the surface, dentin underneath the enamel, and cementum underneath the dentin.Based on the existing studies, the elastic modulus of the top layer of the teeth was found to be 1.3382 ± 0.3079 GPa. The measured values of the elastic modulus before were1.072 GPa and it increases up to 1.571 GPa after the whitening process.Same trend was observed on the adhesion energy values wherein the measured adhesion energy before was found to be 0.196fJ and it increases to 1.608fJ after the whitening process. On the other hand, the surface roughness was observed to be in decreasing trend wherein the surface roughness before was characterized to be 201.0 nm while the value for surface roughness after the application of the whitening strips was 38.33 nm. Tooth whitening via whitening strips is known to workby reacting the hydroxide chemicals on the strip with the staining molecules known as chromogens on the surface of the tooth. This reaction is known as oxidation. The oxidation causes the chromogen molecules to break down when gaining electrons from the hydroxide, causing it to split into molecules that do not reflect light and can be removed from the tooth later.As for whitening treatment, expectations based on previous studies were that the hardness would fall and the roughness would increase with more whitening applied [9-10]. This experiment however found that roughness decreased and modulus increased. A possible explanation wouldbe that the hydroxide oxidation caused the surface to be smoothed out as staining chromogen molecules would be removed from the tooth, while modulus increased because the chromogens had a lower modulus than the enamel itself. Table 1. A table of before and after whitening treatment was applied. The same 15 by 15 micron scan region was used for the before-images, as well as the after-images. These images used were representative of the change caused by whitening treatment, as the data values coincided closely with the mean.Scale bars: 3 μm. Table 2. Mean values of before whitening and the third and final whitening. Conclusion The topography and mechanical data of the tooth sample was successfully acquired by the Park NX10 AFM using PinPoint Nanomechanical mode.Based on the data acquired through this testing,PinPoint Nanomechanical mode found that its method of characterizingmechanical properties of teeth match and agree with mechanical properties found in other methodssuch as macro and microindentation. It was also found that the tooth roughness decreases and the elastic modulus as well as the adhesion properties increase after it had undergone whitening process. Overall, PinPoint Nanomechanical Mode introduced by Park Systems is an ideal approach to characterize and quantify mechanical properties at nanoscale, with faster scanning speed and maximized tip life due to its frictionless operation. References 1.J. Vella, et al., Mechanical properties and fracture toughness of organo-silicate glass (OSG) low-k dielectric thin films for microelectronic applications, International Journal of Fracture 119/120: 487–499, 2003. 2.M. Oyen, et al., A practical guide for analysis of nanoindentation data, J Mech Behav Biomed Mater. 2009 Aug;2(4):396-407. doi: 10.1016/j.jmbbm.2008.10.002. Epub 2008 Oct 15. 3.Y.Zhang, et al.,Review of research on the mechanical properties of the human tooth, Published online 2014 Apr 18. doi: 10.1038/ijos.2014.21. 4.https://www.parksystems.com/index.php/company/news/press-release/450-nanome chanical-mode-to-characterize-nano-mechanical 5.https://www.parksystems.com/index.php/products/small-sample-afm/park-nx10/overview 6.http://probe.olympus-global.com/en/product/omcl_ac160ts_r3/ 7.https://www.brukerafmprobes.com/a-3724-ps-ldpe-12m.aspx 8.https://crest.com/en-us/products/whitestrips/crest-3d-white-monthly-whitening-boo st-whitestrips-teeth-whitening-kit 9.K. Chun, et al., Comparison of mechanical property and role between enamel and dentin in the human teeth, Published online 2014 Feb 6. doi: 10.1177/1758736014520809 10.C. Pinto, et al., Peroxide bleaching agent effects on enamel surface microhardness, roughness and morphology, Braz. oral res. vol.18 no.4 São Paulo Oct./Dec. 2004.

31 Jan 2017

Imaging of Plasmids in Liquid using True Non-Contact Mode Atomic Force Microscopy

Deoxyribonucleic acid (DNA) is one of the most significant discoveries of the last century, primarily because it revolutionized many fields including genetics, molecular biology, medical research, agriculture, forensic science, and many others. DNA is a biological molecule that acts as the storage of genetic information essential for bodily function, growth, and the reproduction of all living organisms. This information is located in the nucleus of every cell in an organism and can be transmitted not only from one cell to another but also from parents to offspring. Hence, knowledge on the structure of DNA is very valuable in understanding how the transfer of genetic information occurs.

27 Dec 2016

Hair Damage From Sunlight Radiation Characterized Using Atomic Force Microscopy

Products for sun protection such as hats, sunscreen, and clothing are commonly used to prevent skin irritation and damage, but few products for protecting hair exist. Through researching the radiation effects of sunlight on hair, it is to be determined whether or not this is a concern worth looking into, especially for those with less photostable, fair-colored hair. With the Park NX20 Atomic Force Microscope (AFM) and its user-friendly Park SmartScan software’s Auto mode, images of hair can be produced to compare its nanoscale topography before and after prolonged exposure to sunlight.

10 Jul 2016

Collagen Fibrils Imaging in Air and in Liquid Using Atomic Force Microscope-Based Fast Nanomechanical Mode

Collagen is a protein that provides structure in various connective tissues in animals and can be found in ligaments, tendons, and skin. The characterization of collagen's mechanical properties at nanoscale can potentially reveal significant insights into the causes of macroscale phenomena such as the elasticity of skin and its degradation as we age. One tool that has been used to acquire nanoscale data of collagen is the atomic force microscope (AFM).

7 Jul 2016

Cellulose Nanowhiskers Topography & Young’s Modulus Imaging Using Atomic Force Microscopy

Carbon nanotubes (CNTs) are a popular sample for various nanotechnology studies. However, they are very difficult to synthesize en masse. As a result, researchers have started looking at alternatives with similar properties. One of the candidates to replace CNTs in some nanocomposite materials is cellulose nanowhiskers (CNWs), structures that can be readily produced from plant sources.

21 Jul 2015

Park PinPoint™ Mode For Cell Biology

By Brian Choi, Bio-application Scientist at Park Systems PinPoint Mode Enables Surface Nanomechanical Property Imaging In addition to nano-scale three dimensional imaging, nano-mechanical property measurement is one the most popular functions of atomic force microscopy (AFM). The basic AFM mode for nano-mechanical property measurement is force-distance (FD) spectroscopy. FD spectroscopy measures the physical properties of the sample surface, such as its hardness, adhesion, etc. by using nano-size cantilever tip to push the sample in the range of nN-scale. Nano-mechanical property characterization at the nN-scale is possible only with the AFM therefore it is widely used in material science, physics, chemistry, etc. for various nano level research. FD spectroscopy is also popular in biological research because it is the only technique available which can physically measure the single molecular binding force. Recently, new application of FD spectroscopy is being developed which is the study of comparing physical properties of a single cell or a tissue with various functions or diseases to make a diagnosis. In order to analyze the physical properties of a single cell in accordance with the biological functions, FD spectroscopy should be aligned exactly with the topographical data. In general, the topography of a sample is acquired first and FD spectroscopy is performed on the specific position using topographical data as a reference map. AC AFM mode (dynamic or tapping mode) is generally used to minimize the AFM artifact from sample-tip interaction in biological sample imaging. However, it is very difficult to avoid the positional errors correlating topographical data and FD spectroscopy data because the soft and sticky nature of the biological samples introduce unavoidable various AFM artifacts even during AC mode. The PinPoint mode was newly developed to compensate the positional errors between topographical and FD data. Figure 1. The PinPoint mode operation principle and the scheme.   The PinPoint mode does this by its capability to simultaneously acquire topographical data and the FD data at each pixel, as shown in Fig 1. There is a significant difference from the conventional AFM imaging of the raster scan with Z scanner feedback. In the PinPoint mode operation, the cantilever approaches at each pixel in the entire scan area (for example 256x256 pixel = 65536 points) to acquire the height of the sample surface to generate 3D topographical image of the sample and FD spectroscopy information at the same time. By lifting the cantilever tip perpendicularly from the sample surface at every pixel, it provides extra clearance space when the cantilever tip moves to the next pixel position during the PinPoint mode as shown in Fig. 2. Even if there is an adhesive interaction between the sample surface and the cantilever tip at the previous point (Pixel a) that connection is completely removed before arrival at the next pixel point (Pixel a+1), hence an accurate topographical information is obtained and the tip-sample interaction information at each pixel can be saved. Figure 2. Raster vs. PinPoint Mode Scanning   This is the biggest advantage of the PinPoint mode, in that simultaneous acquisition of height and FD data guarantees the accuracy of correlating between two types of information. PinPoint mode operation can be controlled by users using preset parameters shown on the left column of Fig. 3. Figure 3. The user interface for the PinPoint mode operation in the NXP software and its control parameters.   The preset parameters include such as the control height, the stiffness threshold for topographical and FD data, the approach and retract time, the lift height, the XY pixel to pixel move time, and the pre-approach delay for determining a scanning speed, etc. The important preset parameters to be determined carefully are the stiffness threshold and the control height. The AFM cantilever presses on the sample surface repeatedly with certain amount of force determined by the stiffness threshold. In order to obtain accurate FD spectroscopy data, the stiffness threshold should be determined carefully. The control height determines the retraction height during movement between pixels after cantilever pushes away from the sample surface. The control height is the most delicate parameter to control because it greatly depends on the type of sample surface characteristic. Generally speaking, smaller number can be used for smaller step height and bigger number for bigger step height. However, in the case of a sample with sticky surface the control height should be appropriately increased in order to clearly disconnect the tip-sample interaction before moving to the next pixel, even if the step height is small. The three physical properties which can beobtained simultaneously through PinPointmode are the surface morphology, elasticitymap, and adhesion force map. a. Surface Morphology (Height) The height information constituting the morphology of the sample surface is determined when the designated stiffness threshold is reached. In other words, the Z detector records how much Z scanner is extended when the certain cantilever deflection, determined by preset stiffness threshold, is reached. The level of cantilever deflection can be accurately designated in the NXP software from Park Systems for optimized surface morphology. b. Elasticity (Stiffness) The elasticity (stiffness) at the certain position of sample surface can be obtained by the same way in the FD spectroscopy. The stiffness can be obtained from the slope of FD curve from contact point to deflection threshold generated during the PinPoint mode operation. Applying Hertz model, designed for soft and elastic samples, user can calculate elasticity (Young's modulus) from the slop, too. In order to calculate Young's modulus using Hertz model equation, FD curve should be converted to tip-sample separation curve to determine the deformation depth from the sample surface. (Fig. 4) For the accurate measurement of the Young's modulus, the accurate positioning to determine the contact point with cantilever tip is very important. The data processing algorithm of NXP automatically generate a Young's modulus image map from the PinPoint mode operation. Figure 4. Force-Distance curve generated by the PinPoint mode c. Adhesion In addition to the elasticity map, the adhesion map also can be generated by using PinPoint mode imaging. The adhesion image is generated based on the maximum adhesion force value. The total adhesion energy at each pixel also can be obtained by measuring green area in Fig. 5. Figure 5. The calculation of the adhesion energy Application As shown in Fig. 6, the elasticity and the adhesion map of a single MRC5 cell was obtained one 3D topographical image at the same time on the NX-Bio using PinPoint mode. The coloring of each pixel representing the elasticity and the adhesion map was determined from the calculation generated by PinPoint FD curve at each pixel. 3D topography shows exact positional match with the FD spectroscopy maps. The higher elastic property can be noted on the nucleus compared to its surrounding areas. The absence of AFM artifacts, such as scratch or streak marks which can be seen frequently in imaging a cell or biological samples in liquid, is an additional benefit of PinPoint mode. Figure 6. The calculation of the adhesion energy  

19 Dec 2013

Targeted Patch Clamping with Scanning Ion Conductance Microscopy

Myung Hoon Choi, Goo Eun Jung (Research Product Management, Research & Development of Park Systems, Seoul, Korea) Targeted patch clamping (TPC)1 combines patch clamping with ion conductance microscopy (SICM)2 to guide the pipette to a specific patch clamping position. Patch clamping is vital to the study of excitable cells such as neurons, cardiomyocites, and muscle fiber in electrophysiology because the technique allows the researcher to examine single or multiple ion channels in those cells. But patch clamping cannot be applied to small cells or submicron-size structures on the cell surface, due to the resolution limit of the incorporated optical microscope, which is used to maneuver the patch clamp’s pipette near the cell surface. Because SICM can identify the cell surface at the submicron scale using the same patch clamping pipette, TPC overcomes the optical resolution hurdle, and greatly improves the applicability of the patch clamp and its accuracy beyond the classical technique. In this paper, live rat ventricular cardiomyocyte cells were examined with targeted patch clamping using a Park SICM. Ion channel signals were successfully recorded at a chosen Z groove location on the ventricular cardiomyocyte.   Operational Principle of Targeted Patch Clamping The main idea of TPC is to perform patch clamping on small cellular structures using  SICM for submicron-scale detection. The SICM utilizes the same pipette and electric circuit used with traditional patch clamping to detect the cell topography using Park XEP control software in the SICM imaging mode, high resolution cell topography  is acquired with feedback control. Once an interesting structure for patch clamping is identified via SICM imaging, the SICM Z scanner positions the pipette on the structure. A giga-ohm seal is formed by applying suction. Ion channel recording can then be performed as in conventional patch clamping. Because the pipette approaches the cell surface in the vertical direction with nanometer scale precision, TPCimproves the probability of making a  giga-ohm seal compared with the slanted approach used in conventional patch clamping3. Figure 1 Illustration of the operational steps for targeted patch clamping: (a) Park SICM acquires the topography of the cardiomycyte cell surface. (b) The nano-pipette is positioned on the chosen region (z groove) by the XY scanner of the SICM. (c) The nano-pipette approaches and forms a giga-ohm seal to record the channel opening current signal.   Instrumentation for Targeted Patch Clamping To implement TPC, the SICM is integrated with a patch clamp system by sharing the pipette and its ionic current circuit. SICM receives the ionic current signal for imaging from the patch clamp’s ion current detector (pre-amp.). The ionic current flows through the nano-pipette, it is detected by the patch clamp’s amplifier, and then it is divided into two: one for the SICM cell imaging (feedback control) and the other for the electrophysiological recording.   Figure 2 Configuration of targeted patch clamping:  (a) XE-Bio SICM integrated with the Axon Axopatch 200B. (b) A schematic diagram of patch clamp connection with XE-Bio SICM.   Samples and Methods Live rat ventricular cardiomyocyte cells were isolated as previously described4. Cells were allowed to attach to polystyrene cell culture dishes filled with bath solutions composed of 120 NaCl, 5.4 KCl, 5 MgSO4, 0.2 CaCl2, 5 Na-Pyryvate, 5.5 Glucose, 20 Taurine, 10 Hepes, and 29 Manitol, adjusted to PH 7.4 with NaOH. Pipette filling solution was the same as the bath solution. A live cell image was acquired in SICM-ARS mode5. The pipette used for the SICM was pulled from borosilicate glass (O.D.,1.0 mm, I.D., 0.58 mm, length 90 mm; Warner Instruments, US) using a CO2-laser-based micropipette puller (P-2000, Sutter Instruments, Novato, US). The inner diameter of the pipette was about 500 nm. Figure 3 (a) Optical image (top inset) and SICM topography of a live rat ventricular cardiomyocyte. The small red circle indicates a Z groove structure where patch clamping was  performed. (b) Current transient (3.45 pA) of cell attached, recording at voltage of ±0 mV. (c) The conductance measurement of the recorded channel. The conductance is 44 pS, and was estimated to be the inward-rectifier K channel.   Ion Channel Recording of Targeted Patch Clamping Figure 3 shows an example of targeted patch clamping using SICM. SICM topography illustrates typical Z grooves clearly, with details. At the position on the Z groove structure (red circle) selected from the SICM topography image, we examined the attached cell recording and successfully measured a transient current of about 3.45 pA at a voltage of ±0 mV.  This result indicates that Park XE-Bio SICM is well-suited for both precise targeting and patch clamping on the cell membrane. The SICM is especially useful for research on small cells such as sperm cells, subcellular structures such as microvilli6, and even opaque cells like intact tissues, trachea, and brain slices.   References 1J. Gorelik, G. Yuchun et al. Biophys. J. 83, 3296 (2002). 2P. K. Hansma, B. Drake, O. Marti, S. A. Gould, C. B. Prater,  Science 243, 641 (1989). 3O.P. Hamill, A. Marthy, E. Neher, B. Sakmann, F.J. Sigworth,  J. Physiol. 391, 85 (1981) 4S.E. Harding, G. Vescovo, M. Kirby, S.M. Jones, J. Gurden, and P.A. Poole-Wilson,  J.Mol. Cell. Cardiol. 20. 635 (1988). 5C. Myunghoon,  XE-Bio Application Note 1, Park Systems (2012) 6G. Yuchun, G. Julia, et al. FASEB J. 16, 748 (2002)