communications Atomic Force Microscopy
Nanoscale Probing the Kinetics of Oriented Bacterial Cell Growth Using Atomic Force Microscopy Peipei Chen, Luping Xu, Jing Liu, Felix J. H. Hol, Juan E. Keymer, François Taddei, Dong Han,* and Ariel B. Lindner* The real-time observation of living cells is critical to understanding biological mechanisms. Despite rapid progress in bio-imaging technologies, it remains a significant challenge to dynamically image cellular processes in living cells with nanometer resolution. AFM,[1] as a versatile nano-technological instrument, provides the opportunity to study biological systems at the nanoscale. It can map biological surfaces under physiological conditions and thus offers significant advantages for imaging biological samples in their native state compared to other imaging techniques with comparable resolution.[2] Furthermore, AFM can be used as a forcespectroscopy tool to measure the nanoscale chemical and mechanical properties of cells.[3] Real-time AFM imaging of biological systems can further provide insight into dynamic events originating from physiological or pathological status.[4] Bacteria are easy to observe and manipulate in the laboratory and are considered an optimal model for understanding many basic biological phenomena.[5] The nanoscale imaging of living bacteria in an aqueous environment, howDr. P. P. Chen,[+] Dr. L. P. Xu,[+,†] Prof. F. Taddei, Prof. A. B. Lindner Institut National de la Santé et de la Recherche Medicale U1001; Faculty of Medicine Paris Descartes University 75014, Paris, France E-mail: [email protected] J. Liu, Prof. D. Han National Center for Nanoscience and Technology Beijing 100190, China E-mail: [email protected] Dr. F. J. H. Hol, Prof. J. E. Keymer [‡] Department of Bionanoscience Kavli Institute of Nanoscience Delft University of Technology Delft 2628 CJ, The Netherlands J. Liu University of Chinese Academy of Sciences Beijing 100049, China [+]
Peipei Chen and Luping Xu contributed equally to this work. Present address: Center for Nano and Micro Mechanics, Tsinghua University, 100084, Beijing, China [‡]Present address: Instituto de Ecología y Biodiversidad, Casilla 653, Santiago, Chile [†]
DOI: 10.1002/smll.201303724
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ever, remains extremely challenging.[6] Improvements in AFM have enabled the imaging of dynamic bacterial processes,[7] but these approaches are often limited to small areas (usually less than 2 µm on a single cell). However, many basic biological phenomena, such as bacterial division and aging, are related to multiple cells in a lineage context.[8] The AFM imaging of these processes requires a large scan size to monitor both the mother and daughter cells and is therefore technically challenging. One limitation hindering AFM use in such studies is the immobilization of growing and dividing cells on substrates to prevent the cells from being displaced by the considerable shear force during AFM tip scanning. Chemical methods using poly-L-lysine (PLL) to immobilize cells via weak charge attractions cannot effectively maintain the daughter cells on the surface.[9] Furthermore, PLL can influence bacterial physiology.[10] The method of mechanically trapping cells by porous membranes is limited to single spherical cells.[11] Microfluidic devices have been used to arrange growing cells and their offspring in linear channels (i.e., create linear colonies) without affecting their physiology.[12] Although the chips prevent the necessary direct contact between the AFM probe and the cells for AFM use, the concept of trapping cells in micro-channels for oriented growth has been proposed. Inspired by the concept of linearly growing colonies, we developed a novel device that facilitates the AFM nanoimaging of E. coli cells and their offspring growing in a linear channel. We nanofabricated a micro-channel array with an open-top structure in which E. coli cells are physically trapped but can grow freely. The cells in these channels grow and divide into linear colonies. Using this device, the growth kinetics of both mother and daughter cells were measured. We furthermore demonstrated that this device can be used to dynamically monitor the nano-mechanics of single cells. The kinetics of the linearly growing E. coli cells revealed that, under the imaging conditions, the cells elongated exponentially. Our findings furthermore suggested that the cell's average stiffness changed throughout the cell cycle. An array of parallel micro-channels was designed to arrange the growing E. coli cells in the same orientation. Each channel consisted of two layers: a micro-tube for cell growth and a submicron opening at the top of the tube that provided access for the AFM tip. Figure 1a depicts the fabrication process of the chip, which combined micro-/nano-fabrication and soft lithography. The structure was initially constructed
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Figure 1. Micro-/nano-fabrication of the structured substrate for cell loading. (a) Schematic representation of the methodology to produce the structured substrate. (b) SEM image of the micro-channels on the silicon mold in a cross-sectional view. Two parts, consisting of open-top structures and tube-like channels, can be clearly observed. Scale bar: 1 µm. (c) SEM image of the PDMS mold surface obtained by replication from the silicon mold. Scale bar: 2 µm. (d) SEM image of the structures on the PDMS film obtained by replication from the PDMS mold shown in (c). Scale bar: 2 µm.
on a silicon substrate, which was followed by polydimethylsiloxane (PDMS) replication. First, we used electron-beam (e-beam) lithography to expose parallel trenches on an e-beam sensitive resist of polymethylmethacrylate (PMMA) and then transferred this pattern to silicon by dry etching (Figure 1a-1 to 1a-4). In this process, the side-wall angle of the trench was anisotropically etched to 85° by fine tuning the mixture ratio of the etching gases, the substrate bias power, and the temperature in the reactive ion etcher (RIE) chamber.[13] Second, we deposited a fluorocarbon polymer over the trench walls by C4F8 (octafluorocyclobutane) plasma treatment. This deposited polymer acted as a passivation film (Figure 1a-5) to protect the trench-walls from lateral etching in the following step. Third, we applied the etching gas SF6 (sulfur hexafluoride) and O2 (oxygen) to yield isotropic etching and obtain tube-like channels (Figure 1a-6). After stripping off residues of the PMMA resist left on the silicon surface, a silicon mold containing micro-channels was obtained (Figure 1a-7). Scanning electron microscopy (SEM) images of the cross-sections of the channels (Figure 1b) show that each channel consisted of a tube-like structure approximately 1 µm in diameter, corresponding to the size of an E. coli cell, and an open-top channel 400 nm in height and 600 nm in width (more silicon-chip images and parameters in supporting information 1). After fabrication of the silicon mold, we made the first replica using PDMS. As Figure 1c illustrates, on the PDMS surface, we obtained an array of protuberances that were complementary to the silicon structure. small 2014, 10, No. 15, 3018–3025
Taking the first PDMS replica as the master, we made a second replica and obtained the final PDMS slide with the originally designed structures. The obtained PDMS slide displayed in Figure 1d was utilized as the patterned substrate for cell immobilization and oriented growth in the following AFM study. We investigated E. coli wild type strain MG1655 constitutively expressing the yellow fluorescence protein (YFP) by AFM. To load the cells into the channels, a PDMS block with a flow channel was first mounted on top of the microchannels (as shown in Figure 2a, top image). Then, two metal plates with screws were used to tightly clamp the two PDMS layers to maintain the cell medium during loading. After that, a concentrated cell culture with a cell density of 1010 cells/ml was injected into the flow channel. A centrifugation step assured cell loading into the micro-channels (Figure 2a, bottom image). As a control, we followed the cellular growth within the channels by fluorescence microscopy. In three hours of incubation, the cells grew and divided unhindered in the channels (Figure 2b). Furthermore, the three-hour timelapse fluorescence microscopy images indicated oriented growth of the cells within the channel (Figure 2c). The doubling time of the E. coli cells measured from fluorescence images was 45 min, in agreement with measurements in bulk culture by optical density. This demonstrates that the microchannels do not hinder cellular growth. Compared to previously described microfluidic chips for bacterial cell growth, our structures demonstrate significant advantages for AFM
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Figure 2. Cell loading. (a) Sketch of the loading method. Top: injection of E. coli cells and medium into the two-layer chip. Bottom: the cells can be loaded into the channels by centrifugation. (b) Fluorescence image of the cells in the PDMS channels after incubation. The green signal shows cells with YFP. Scale bar: 10 µm. (c) Time-lapse fluorescence images of cells dividing along a micro-channel. Scale bar: 2 µm.
studies: the micro-channels immobilize both the mother and daughter cells in the channels, and the cells undergo oriented growth in a linear colony. Interestingly, the open-top structure provided homogenous medium conditions along the channel; this chip comprised an “open” system (compared to a sealed microfluidic chip), which allowed the AFM probe to touch the cells and the growth medium to be changed over time. To enable AFM imaging on these unique channels, modifications to the AFM tip were necessary because the normal pyramid tip was too wide to reach the cells loaded in the 600 nm wide channels.[14] The AFM probe used here was a cantilever with a modified tip. A focused ion beam (FIB) was applied to sharpen the pyramidal Si3N4 tip to obtain an ultrasharp tip measuring 2.5 µm in length, only 20 nm in diameter at the tip point, and 300 nm at the tip base (Figure 3a and b). The E. coli cells were loaded into the PDMS channels as described above. The PDMS chip was then carefully released from the metal plates and attached to the bottom of a petri dish. We then washed the chip surface and added culture medium M9 with a low carbon source to the petri dish. This low nutrition condition and low temperature (e.g., room temperature instead of 37 °C) allowed for slow growth, which was necessary to visualize the division process in detail by AFM. Before AFM imaging, an inverted optical microscope was used to locate the cells in the channels (Figure 3a). Using the ultra-sharp AFM tip, topographic images of living E. coli cells (Figure 3c) were obtained. The top image in Figure 3c shows multiple dividing cells in one channel. Underneath, there are four single cells and a zoomed-in image with a resolution of 2 nm. All of the images in Figure 3c have fine imaging quality, shown by the smooth surfaces of the cells. These results demonstrate that the chip and the modified AFM tip facilitated the nanoscale probing of living E. coli cells.
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Figure 3. Chip setup and AFM imaging. (a) Scheme of the liquid AFM imaging system combined with inverted optical microscopy. The AFM tip is indicated by an arrow. (b) SEM image of the super-sharp AFM tip etched by FIB. Scale bar: 5 µm. (c) AFM images of single cells. Top: the cells dividing along a micro-channel. Scale bar: 1 µm. Bottom: five AFM images of single cells. The last image is a magnification of the fourth cell. Scale bars are 500 nm, 1 µm, 1 µm, 500 nm and 200 nm, respectively.
In the following study, we provide proof-of-concept experiments demonstrating the utility of the device as a tool to measure the dynamics of cell growth in a linear colony. Time-lapse AFM imaging was performed to monitor the growth and division processes of the E. coli cells loaded in the channel for more than 4 hours. Figure 4a consists of 16 topographic images showing a cell dividing into two daughter cells, then subsequently dividing into four cells. At 36 min (see Figure 4b), the cell membrane of the mother cell in the middle constricts at the division site (labeled by a black arrow). From 52 min to 96 min, the cell continues growing while the division site gradually contracts. During division in vivo, a circumferential protein ring (named the Z-ring) develops at the inner surface of the cell membrane at the division site, and this ring is believed to be an essential component of the bacterial division machinery targeted to the division site.[15] Here, AFM was used to visualize the 3D topography of the division site of the E. coli cell. The constriction at the division site was revealed by a depression in the cross-sectional curve (labeled by a black arrow) of the cell. Therefore, the time point of cell-division termination can be easily identified (see experimental section and supplementary information 2 for more details) using a cross-sectional profile of the division site. When the vertical distance between the top membrane surface and the division site (labeled as h in Figure S2a) equaled half the cell height, we considered the division to be complete. We measured the dynamic proportion of h/Height. At 107 min, this value suddenly ascend to 0.519 (shown in Figure S2), indicating division termination. At this moment, the AFM tip can protrude in between the two poles of the daughter cells and slightly separate them. The time point at 107 min, therefore, marked division termination in Figure 4a. At this time, the division site disappeared and the division was complete.
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Figure 4. Growth curves of E. coli cells based on time-series AFM images. (a) Time-lapse AFM images of E. coli growth over 274 min in M9 and at room temperature. Scale bar: 2 µm. (b) Zoomed-in images of cells obtained at 36, 219 and 237 minutes (black boxes in (a)). Scale bar: 500 nm. The black arrows show the constriction-structures in the cells’ division planes. The curves under each image are cross-sectional profiles along the long axis of each dividing cell. The division planes are indicated by arrows in these profiles. (c) Growth curves of the AFM-probed growing cells. The black dots correspond to the mother cell length measurements over time, and the blue and red dots represent its two daughter cells. Both the mother and daughter cell sizes increased exponentially with time.
To quantify the dynamic change, the cells’ lengths throughout the growth and division process described above were measured (supporting information 3). As shown in Figure 4c, the mother cell’s length was measured during the first 96 min. After the estimated point of division termination, at 107 min, the growth curves of the mother cell and the two daughter cells were plotted separately. The fitted growth curves demonstrate that both the mother and daughters in Figure 4a exponentially elongated under the imaging conditions. It is currently under debate whether cell elongation is linear or exponential during the life cycle of an E. coli cell.[16] This debate has persisted for decades primarily due to the lack of quantitative methods for measuring cell size with the required sensitivity. To distinguish an exponential pattern from a linear one, a resolution of under 6% in cell size is required.[17] Our results obtained by AFM have a resolution of 31 nm, which is only 3% of the cell size and well below this limit. Additionally, the cell increased in length during the AFM scan. The magnitude of this error depends on the speed of AFM scanning relative to cell growth. The scanning rate used to collect the images was 0.49 Hz, which represents a scanning speed of 19.5 nm/s along the Y axis of each image over 10 µm. The largest growth rate of the cells we measured was 0.39 nm/s. The scanning rate of AFM was thus approximately 50 times faster than the growth rate of the cells. Therefore, the error in the size measurement derived from the scanning time can be neglected. Thus, it can be concluded that this chip facilitates AFM measurements of cell growth with sub-diffraction-limited resolution. Examples of asymmetry in division as well as of variations in the cell growth rate, length-at-division, and time-to-divide that have previously been reported were also observed in this imaging experiment. First, the division site of E. coli is accurately positioned by specific protein machinery.[18] Here, the small 2014, 10, No. 15, 3018–3025
distance of the division site from the midcell at the beginning of cell constriction (at 36 min in Figure 4a) was 4.9% of the cell length. Second, the growth rates of the two daughter cells acquired from fitting the growth curves (Figure 4c) differed, leading to variations in the length-at-division with time. For example, the bottom cell (Figure 4a) was initially 300 nm longer than the upper cell (at 107 min), and this asymmetry increased to 550 nm before the second division (at 199 min). Finally, the future division site was visible in the longer cell 18 minutes before contraction could be observed in the shorter cell’s septation site (the division planes are labeled by black arrows in the zoomed-in images and in the crosssectional curves in Figure 4b at 219 min and 237 min). These observations demonstrate that the asymmetry of the mother and daughter cells loaded into the channels could be detected by AFM. With this on-chip AFM technique, the dynamic nanomechanical properties across E. coli cells can be investigated by Peak Force Tapping (PFT) AFM.[19] PFT is a relatively new AFM mode for studying the mechanical properties of cells at high speeds (similar to Tapping Mode[20]) and high resolution that has been used to explore the structural and physical properties of a variety of biological samples.[21] The imaging feedback is based on the peak force of the forcedistance curves. Because this force curve performance is repeated for each pixel of the image, the resolution of the mechanical images will be the same as that of a conventional height AFM image. Once the system is calibrated, the Young's modulus (YM) determined by the DMT (DerjaguinMuller-Toropov model)[22] fit can be directly displayed in the quantitative units of Pascals. Hence, this technique offers the possibility to map cellular mechanical properties at nanometer resolution. Figure 5a shows a time-series over 53 minutes of topographical AFM images (height image) of E. coli
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Figure 5. Time-lapse AFM topographic images and kinetics of mechanical measurements on E. coli cells. (a) Topographic images over 53 minutes. Scale bar: 500 nm. (b) Magnified, real-time Young’s modulus (YM) images over 53 minutes of the cell labeled by a black rectangle in (a). Scanning size: 0.74 µm×3.5 µm. Scale bar: 200 nm. (c) Cross-sections along the long axis of the cell showing the time progression of the YM variation. Each slice represents data extracted from one image in the full time series. (d) Histogram representations of the average distance between two stripes on 20 cells. The median distribution is 400 nm. The figure was obtained in M9 and at room temperature.
cells loaded in a micro-channel. Zoomed-in YM images with a resolution of 20 nm for the cell (labeled by the black box in Figure 5a) at each time point are displayed in Figure 5b, which depicts the logarithmic YM in pseudo color. The YM cross-section along the long axis of the cell in the full time series was further extracted and is shown in Figure 5c. In this figure, each slice represents the data extracted from each time point of Figure 5b, showing the time progression of the YM variation. The dynamic cell nano-mechanics can be quantified from this figure. First, many stripe-shaped areas with a dark blue or purple color are distributed perpendicularly to the cell's long axis (Figure 5b). The color scale reveals that these areas are more rigid than other parts of the cell. This feature can also be observed as a waved cross-sectional profile (Figure 5c). Recently, actin-like filaments forming a helical cytoskeleton under the inner membrane in E. coli cells were observed.[23] It was further shown that this cytoskeleton-like structure was sufficiently stiff to affect the overall mechanical rigidity of an E. coli cell, in much the same way that the actin cytoskeleton does in eukaryotes.[24] Therefore, we assumed that the observed oriented patterns on the single cells reflected the cytoskeletal arrangement underneath the cell membrane (supporting information 4). Presumably, when the tip interacts with the areas with a cytoskeleton support underneath, the YM information represents the mechanical integrity of the cell, which involves the cytoskeleton, cell wall, etc. In the other areas, there is lack of cytoskeleton information in the
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YM. To quantify this structural feature, the distances between the stripes distributed on 20 cells were measured. Figure 5d shows the histogram representation of the distance distribution, which reveals that the average distance between two stripes measured by AFM was approximately 400 nm. Another measurement is the average YM of the division site. It can be observed in Figure 5b that the division site had a much larger YM than the rest of the cell (see the image at 26 min and beyond). In the division process from 26 min to 53 min, the constriction-structure was rigidified (Figure 5c). We measured the average YM based on the data in this figure. At 53 min, the average YM of the division site was 1 MPa, while the average YM for the rest of the curve was 630.96 KPa. The YM of the division site was therefore 370 KPa larger than that of the rest of the cell. The rigidity of the division site presumably increased in order to facilitate the septation necessary for division. Finally, the average YM (excluding the division plane area) along the long axis throughout the division process (supporting information 5) was measured. It first increased from 288.40 KPa (5.46 log(Pa)) to 630.96 KPa (5.80 log(Pa)) and then decreased to 537.03 KPa (5.73 log(Pa)) at 0, 26 and 53 minutes, respectively. This YM variation implies that the average stiffness of an E. coli cell changes throughout the cell cycle. In summary, we developed a novel chip to facilitate the AFM imaging of oriented, growing E. coli cells. To immobilize the freely growing cells on a substrate and adapt the AFM tip
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to this system, technical improvements in the substrate fabrication and probe sharpening were achieved. These improvements extend AFM imaging applications to non-adherent living cells. These techniques can also be applied to trap other prokaryotic or eukaryotic (e.g., mammalian) cultured cells. And the tip sharpening will allow quantitative detection to be more accurate and credible. This novel design principle can be used and extended further to microfluidic channels of different internal dimensions and shapes. As a result of these technical improvements, the kinetics of oriented cell growth, including the growth pattern and the nano-mechanical properties, can be quantified at the single-cell level. These advances in micro-/nano-fabrication, AFM technology and their applications establish a powerful tool for exploring cellular processes in real time at the nanometer scale.
Experimental Section Fabrication of the Micro-Channel Array: Fabrication of the micro-channels was performed on a 100 mm diameter singlecrystal (100) silicon wafer (University Wafer, USA). The wafer were ultrasonically cleaned in fuming nitric acid (100% HNO3) for 10 min, rinsed in deionized (DI) water and spun dry. Then, the wafer surface was primed for resist adhesion using hexamethyldisilazane (HMDS) by spin-coating at 1000 rpm for 60 s, followed by baking for 90 s at 200 °C. After priming, a 1.2 µm layer of PMMA 950k positive electron-beam resist (MicroChem) was spin-coated onto the wafer at 2000 rpm for 60 s and baked at 175 °C for 10 min. Feature patterning with 9 individual channel-arrays arranged on the wafer was performed using a Vistec EBPG5000 Plus with an acceleration voltage of 100 kV and an aperture of 400 µm. Spot-size diameters of 56 nm were achieved by focusing the electron spots with currents of 129 nA. The doses were set to 1200 µC/cm2 with a 40 nm beam step size (BSS). The total machine time was approximately 3 h for the entire wafer. Following electron-beam exposure, development was conducted for 1 min in 1:3 MIBK (methyl isobutyl ketone):IPA (isopropyl alcohol), followed by a 30 s soak in IPA. The wafer was then rinsed in DI water and spun dry. The wafer was cut into 9 pieces containing trench-structures. The trench-structure layer was then dry-etched in a reactive-ion etcher (Adixen AMS100 I-speeder) with a mixture of 15 sccm SF6 (sulfur hexafluoride), 20 sccm C4F8 (octafluorocyclobutane) and 10 sccm CH4 (methane) diluted in 100 sccm He (helium). The source power was set to 2000 W, and the substrate bias was approximately −22 V. Under this etching condition, the obtained side-wall for this layer was almost perpendicular with the silicon surface (anisotropic etching). A chemically inert passivation layer was immediately deposited after anisotropic etching. Here, a 80 sccm C4F8 plasma was used to yield a fluorocarbon polymer on the structures for 10 s to protect the trench-walls from lateral etching in the following step. We next applied the etching gas SF6 and O2 to yield isotropic etching and obtain tube-like channels. Here, the etching gas was a mixture of 200 sccm SF6 and 120 sccm O2, the source power was 1200 W, and the substrate bias RF power was set to 20 W. In this etching process, the reactive ions first attacked the passivation layer on the bottoms of trenches. After the silicon surface under the passivation layer was exposed, a tube-like structure was obtained resulting from the isotropic small 2014, 10, No. 15, 3018–3025
ion path in the second etching process. Compared to anisotropic etching, this process is isotropic etching. Etching times of 22 s, 25 s, 27 s and 30 s were utilized to obtain variable tube diameters. After cleaning by HNO3, we obtained structures of open-top, tube-like channels. Table S1 in the supporting information lists all the structures on silicon we obtained with different sized microchannels. These silicon molds were applied for cells with diverse sizes. Figure S1a shows the zoomed-in cross-section of one microchannel with a size measurement. To transfer the patterns from the silicon chip to the PDMS surface, two PDMS replicas were necessary. A 10 µL volume of the hydrophobic chemical trimethylchlorosilane (TMCS) was dropped beside the silicon mold, sealed and maintained for 10 min. After hydrophobic treatment, 1:10 PDMS was poured on the chip and solidified by baking at 80°C for 2 h. The hydrophobic treatment to silicon and greatly decreased the interaction of the PDMS with the silicon. Besides, with the V-shaped structure (as marked by a black arrow in Figure S1b in the supporting information) at one end of the micro-channel array, it was therefore effortless to peel the PDMS slab from the silicon surface. The structure we obtained on the PDMS surface was the inverse of the silicon mold and was then used as the mold for the second PDMS replica. Then, a 2 min activation of the PDMS surface was conducted using oxygen plasma, followed by hydrophobic TMCS treatment for 10 h. In this process, each hydroxyl group on the silicon surface (a monolayer of hydroxyl groups was formed after plasma bombardment) reacted with a silane group of TMCS. After baking at 80°C for 2 h, the formation of a hydrophobic molecular film prevented adhesion between the two PDMS layers during peel off, and the second PDMS replication was easy to accomplish. Eventually, PDMS substrates containing surface micro-channels were obtained. E. coli Strain Information: E. coli K12 MG1655 fliC was prepared by standard lambda recombination-based replacement of the gene with a cat cassette that was further removed by FRT recombination. P1 transduction was used to insert the yellow fluorescence protein (YFP) gene under the control of a constitutive promoter (λPR) together with a chloramphenicol resistance gene (from MRR strain) at the intC locus. The fliC gene determines the flagella of E. coli. Thus, the fliC knockout strain has no flagella, which leads to their inability to swim in the medium. For the AFM investigation, 50 µL of cryo-preserved E. coli cells were recovered in 5 ml M9 medium (2 × 10−3M MgSO4, 1 × 10−4 M CaCl2, 3.4 × 10−4 g/mL Vitamin B1 with 0.4% w/v glucose) and cultured at 37 °C for 16 hours until reaching the stationary phase. The E. coli cells were then refreshed in M9 to reach the exponential phase (OD600 = 0.2). Cell Loading: To load the cells into the channels, the chip was covered with a PDMS block with a flow channel (500 µm in width, 20 µm in height and 1 cm in length) facing down into the channel array. Two metal plates with surrounding screws were utilized to tightly clamp two PDMS layers to maintain the cell medium in the flow channel in subsequent steps. A 1 mL volume of E. coli cell medium was concentrated by centrifuging at 2000 rpm for 5 min. Then, 100 µL of concentrated medium with a cell density of 1010 cells/ml was injected from one entrance of the channel. By centrifugation at 2000 rpm for 5 min under centrifugal force parallel with the chip surface, the E. coli cells were driven from the V-shaped entrance into the micro-channels. After centrifuging, the PDMS film was immediately released from the metal plates and then carefully put into a petri dish with the medium.
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Tip Sharpening: In air, the intrinsic resonant frequency of the probe was 97.42 kHz. We utilized a focused ion beam (FIB) (FEI Nova200 Nanolab) with an acceleration voltage of 30 kV to sharpen the probe. The ring-shaped (inside diameter 0.3 µm and outside diameter 1 µm) Gallium ion-beam with a current of 0.3 nA was used to etch the tip for 10 s. AFM Imaging System and Parameters: Time-series AFM images for the cell length measurements were acquired by an Agilent 5500 AFM (Agilent Corp.) in magnetic alternating current (MAC)mode placed on an inverted microscope (Nikon, TE2000). A Picoscan controller and a top MAC-Mode control-box were used to control the scanner and to acquire and convert the analog signals to digital signals. The software Picoview was used to command the controller and display the images. The MAC-mode AFM probe’s cantilever was coated with a magnetic material. A top magnetic field generated by a solenoid directly shook the cantilever. The probe was immersed in M9 medium to scan over the E. coli cells. In the imaging process, the resonant frequency of the AFM probe was 31.6 kHz. We set the ratio of the actual to raw amplitude of the probe, A/A0, to 0.83. The scanning speed was 0.49 Hz. The integral gain and proportional gain were 11% and 21%, respectively. The scanning range in the Z-axis was 3.3 µm. Speculation of Cell-Division Termination: The AFM 3D profile was used to determine this time point. As Figure S2 in the supporting information shows, AFM scanning allowed us to obtain a cross-section of an E. coli cell. In the division process of an E. coli cell, the division site gradually contracts. When the AFM tip scans the division site, the vertical depth of the tip required to contact the cell increases. At the time-point of cell-division termination, the two daughter cells are completely separated but remain physically in contact. At this time, the depth of the AFM tip is half of the height of the cell. We therefore considered the ratio of the vertical distance between the top membrane surface and the division site (labeled as h in Figure S2a) to the total cell height to be 0.5 when cell division is complete. The dynamic proportion of h/Height was measured. In Figure S2b, we plot the ratio in each image as a function of time. From 52 min to 96 min, the division site first appears and then constricts gradually. Correspondingly, the measured ratio increases from 0.066 to 0.327. At 107 min, this value suddenly increases to 0.519, and afterwards fluctuates between 0.4 and 0.5. Growth Curve Measurement: We measured the cell lengths in each image in the imaging sequence using the software ImageJ. The AFM image was first imported into the software. Then, lines were drawn on the image, and ImageJ gave the length result in pixels. We set the width of the drawing line (yellow line in Figure S3a of the supporting information) to 5, so the measured length was given by the average value of 5 lines. To remove imaging aberrations from scanning, a calibration along the cell's long axis for all the images was performed. We measured the distance between two particles on the edge of a channel (the distance shown by the black arrow in Figure S3a) in each image. The calibration factor for each image was acquired (listed in Table S2 of supporting information) using the normalization of each measurement by their average value (average distance was divided by respective measured value). In our measurements, the longest distance from one end of the cell to the other was considered to be the length of the cell. From the cross-sectional profile of this line (Figure S3b in supporting information), we determined the initial and terminal points
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of each measurement. Given all the cell calibration factors and the absolute length value of 1 pixel (31 nm), the final lengths were obtained (Table S2). Kinetic Nano-Mechanical Measurement: Our results were achieved by the Peak Force Tapping (PFT) mode (Bruker Dimension Icon SPM). In our imaging process, the probe was oscillated at 2 kHz. A deflection sensitivity of 37 nm/V was calculated by engaging the tip on a stiff glass slide and recording a force curve. The spring constant of the AFM probe calculated by “Thermal Tune” was 0.35 N/m. The tip radius measured from the SEM image was 10 nm. The scanning rate of each image was 0.65 Hz. We obtained Young's modulus images by entering these parameters into the software and employing a DMT fit.
Supporting Information Supporting Information is available from the Wiley Online Library or from the author.
Acknowledgements This work was supported by the Franco-Chinese cooperation program (PFCC) between INSERM and the Chinese Academy of Sciences (INSERM/CAS AO 2010–2011) and the AXA Foundation Chair. We acknowledge Dr. François-Xavier Pellay for his helpful discussion, Dr. Zhuangxiong Huang for advice on micro-fabrication, the TU Delft Nanofacility for micro-fabrication process support and Dr. Kaiwu Peng for FIB operation.
[1] G. Binning, C. F. Quate, C. Gerber, Phys. Rev. Lett. 1986, 56, 930. [2] J. H. Hoh, C. A. Schoenenberger, J. Cell Sci. 1994, 107, 1105. [3] a) P. Hinterdorfer, Y. F. Dufrene, Nat. Methods 2006, 3, 347; b) T. G. Kuznetsova, M. N. Starodubtseva, N. I. Yegorenkov, S. A. Chizhik, R. I. Zhdanov, Micron 2007, 38, 824; c) A. B. Mathur, A. M. Collinsworth, W. M. Reichert, W. E. Kraus, G. A. Truskey, J. Biomech. 2001, 34, 1545; d) D. J. Muller, Y. F. Dufrene, Nat. Nanotechnol. 2008, 3, 261; e) C. Rotsch, K. Jacobson, M. Radmacher, Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 921; f) A. Touhami, B. Nysten, Y. F. Dufrene, Langmuir 2003, 19, 4539. [4] G. E. Fantner, R. J. Barbero, D. S. Gray, A. M. Belcher, Nat. Nanotechnol. 2010, 5, 280. [5] K. Struhl, Nature 2002, 417, 22. [6] Y. F. Dufrene, Nat. Protoc. 2008, 3, 1132. [7] a) G. Francius, O. Domenech, M. P. Mingeot-Leclercq, Y. F. Dufrene, J. Bacteriol. 2008, 190, 7904; b) M. Plomp, T. J. Leighton, K. E. Wheeler, H. D. Hill, A. J. Malkin, Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 9644. [8] a) A. B. Lindner, R. Madden, A. Dernarez, E. J. Stewart, F. Taddei, Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 3076; b) M. Ni, A. L. Decrulle, F. Fontaine, A. Demarez, F. Taddei, A. B. Lindner, PLoS Genet. 2012, 8, 1003148; c) E. J. Stewart, R. Madden, G. Paul, F. Taddei, PLoS. Biol. 2005, 3, 295. [9] M. J. Doktycz, C. J. Sullivan, P. R. Hoyt, D. A. Pelletier, S. Wu, D. P. Allison, Ultramicroscopy 2003, 97, 209. [10] K. Colville, N. Tompkins, A. D. Rutenberg, M. H. Jericho, Langmuir 2010, 26, 2639.
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[11] a) Y. F. Dufrene, Nat. Rev. Microbiol. 2004, 2, 451; b) S. Kasas, A. Ikai, Biophys. J. 1995, 68, 1678. [12] a) N. Q. Balaban, J. Merrin, R. Chait, L. Kowalik, S. Leibler, Science 2004, 305, 1622; b) Y. Caspi, PLoS ONE 2014, 8, e83775; c) F. Gasset-Rosa, A.-S. Coquel, M. Moreno-del Álamo, P. Chen, X. Song, A. M. Serrano, M. E. Fernández-Tresguerres, S. Moreno-Díaz de la Espina, A. B. Lindner, R. Giraldo, Mol. Microbiol. 2014, 91, 1070; d) P. Wang, L. Robert, J. Pelletier, W. L. Dang, F. Taddei, A. Wright, S. Jun, Curr. Biol. 2010, 20, 1099. [13] Z. X. Huang, F. Pedaci, M. van Oene, M. J. Wiggin, N. H. Dekker, ACS Nano 2011, 5, 1418. [14] H. J. Dai, J. H. Hafner, A. G. Rinzler, D. T. Colbert, R. E. Smalley, Nature 1996, 384, 147. [15] a) S. Ben-Yehuda, R. Losick, Cell 2002, 109, 257; b) E. Bi, J. Lutkenhaus, Nature 1991, 354, 161; c) F. Guo, H. A. Tao, J. Buss, C. Coltharp, Z. Hensel, X. A. Jie, PLoS ONE 2010, 5, e12680. [16] S. Cooper, Theor. Biol. Med. Model. 2006, 3, 10. [17] A. Tzur, R. Kafri, V. S. LeBleu, G. Lahav, M. W. Kirschner, Science 2009, 325, 167. [18] J. M. Guberman, A. Fay, J. Dworkin, N. S. Wingreen, Z. Gitai, PLoS Comp. Biol. 2008, 4, e1000233.
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[19] a) A. Berquand, Bruker Application Note 2011, #135; b) B. Pittenger, Bruker Application Note 2012, #128. [20] Q. Zhong, D. Inniss, K. Kjoller, V. B. Elings, Surf. Sci. 1993, 290, L688. [21] a) J. Adamcik, A. Berquand, R. Mezzenga, Appl. Phys. Lett. 2011, 98, 193701; b) D. Alsteens, V. Dupres, S. Yunus, J. P. Latge, J. J. Heinisch, Y. F. Dufrene, Langmuir 2012, 28, 16738; c) C. Heu, A. Berquand, C. Elie-Caille, L. Nicod, J. Struct. Biol. 2012, 178, 1; d) I. Medalsy, U. Hensen, D. J. Muller, Angew. Chem. Int. Ed. 2011, 50, 12103; e) G. Pletikapic, A. Berquand, T. M. Radic, V. Svetlicic, J. Phycol. 2012, 48, 174; f) F. Rico, C. M. Su, S. Scheuring, Nano Lett. 2011, 11, 3983. [22] B. V. Derjaguin, V. M. Muller, Y. P. Toporov, Prog. Surf. Sci. 1994, 45, 131. [23] Y. L. Shih, T. Le, L. Rothfield, Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 7865. [24] S. Y. Wang, H. Arellano-Santoyo, P. A. Combs, J. W. Shaevitz, Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 9182.
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Nanoscale Probing the Kinetics of Oriented Bacterial Cell Growth Using Atomic Force Microscopy Peipei Chen, Luping Xu, Jing Liu, Felix J. H. Hol, Juan E. Keymer, François Taddei, Dong Han,* and Ariel B. Lindner* The real-time observation of living cells is critical to understanding biological mechanisms. Despite rapid progress in bio-imaging technologies, it remains a significant challenge to dynamically image cellular processes in living cells with nanometer resolution. AFM,[1] as a versatile nano-technological instrument, provides the opportunity to study biological systems at the nanoscale. It can map biological surfaces under physiological conditions and thus offers significant advantages for imaging biological samples in their native state compared to other imaging techniques with comparable resolution.[2] Furthermore, AFM can be used as a forcespectroscopy tool to measure the nanoscale chemical and mechanical properties of cells.[3] Real-time AFM imaging of biological systems can further provide insight into dynamic events originating from physiological or pathological status.[4] Bacteria are easy to observe and manipulate in the laboratory and are considered an optimal model for understanding many basic biological phenomena.[5] The nanoscale imaging of living bacteria in an aqueous environment, howDr. P. P. Chen,[+] Dr. L. P. Xu,[+,†] Prof. F. Taddei, Prof. A. B. Lindner Institut National de la Santé et de la Recherche Medicale U1001; Faculty of Medicine Paris Descartes University 75014, Paris, France E-mail: [email protected] J. Liu, Prof. D. Han National Center for Nanoscience and Technology Beijing 100190, China E-mail: [email protected] Dr. F. J. H. Hol, Prof. J. E. Keymer [‡] Department of Bionanoscience Kavli Institute of Nanoscience Delft University of Technology Delft 2628 CJ, The Netherlands J. Liu University of Chinese Academy of Sciences Beijing 100049, China [+]
Peipei Chen and Luping Xu contributed equally to this work. Present address: Center for Nano and Micro Mechanics, Tsinghua University, 100084, Beijing, China [‡]Present address: Instituto de Ecología y Biodiversidad, Casilla 653, Santiago, Chile [†]
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ever, remains extremely challenging.[6] Improvements in AFM have enabled the imaging of dynamic bacterial processes,[7] but these approaches are often limited to small areas (usually less than 2 µm on a single cell). However, many basic biological phenomena, such as bacterial division and aging, are related to multiple cells in a lineage context.[8] The AFM imaging of these processes requires a large scan size to monitor both the mother and daughter cells and is therefore technically challenging. One limitation hindering AFM use in such studies is the immobilization of growing and dividing cells on substrates to prevent the cells from being displaced by the considerable shear force during AFM tip scanning. Chemical methods using poly-L-lysine (PLL) to immobilize cells via weak charge attractions cannot effectively maintain the daughter cells on the surface.[9] Furthermore, PLL can influence bacterial physiology.[10] The method of mechanically trapping cells by porous membranes is limited to single spherical cells.[11] Microfluidic devices have been used to arrange growing cells and their offspring in linear channels (i.e., create linear colonies) without affecting their physiology.[12] Although the chips prevent the necessary direct contact between the AFM probe and the cells for AFM use, the concept of trapping cells in micro-channels for oriented growth has been proposed. Inspired by the concept of linearly growing colonies, we developed a novel device that facilitates the AFM nanoimaging of E. coli cells and their offspring growing in a linear channel. We nanofabricated a micro-channel array with an open-top structure in which E. coli cells are physically trapped but can grow freely. The cells in these channels grow and divide into linear colonies. Using this device, the growth kinetics of both mother and daughter cells were measured. We furthermore demonstrated that this device can be used to dynamically monitor the nano-mechanics of single cells. The kinetics of the linearly growing E. coli cells revealed that, under the imaging conditions, the cells elongated exponentially. Our findings furthermore suggested that the cell's average stiffness changed throughout the cell cycle. An array of parallel micro-channels was designed to arrange the growing E. coli cells in the same orientation. Each channel consisted of two layers: a micro-tube for cell growth and a submicron opening at the top of the tube that provided access for the AFM tip. Figure 1a depicts the fabrication process of the chip, which combined micro-/nano-fabrication and soft lithography. The structure was initially constructed
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Figure 1. Micro-/nano-fabrication of the structured substrate for cell loading. (a) Schematic representation of the methodology to produce the structured substrate. (b) SEM image of the micro-channels on the silicon mold in a cross-sectional view. Two parts, consisting of open-top structures and tube-like channels, can be clearly observed. Scale bar: 1 µm. (c) SEM image of the PDMS mold surface obtained by replication from the silicon mold. Scale bar: 2 µm. (d) SEM image of the structures on the PDMS film obtained by replication from the PDMS mold shown in (c). Scale bar: 2 µm.
on a silicon substrate, which was followed by polydimethylsiloxane (PDMS) replication. First, we used electron-beam (e-beam) lithography to expose parallel trenches on an e-beam sensitive resist of polymethylmethacrylate (PMMA) and then transferred this pattern to silicon by dry etching (Figure 1a-1 to 1a-4). In this process, the side-wall angle of the trench was anisotropically etched to 85° by fine tuning the mixture ratio of the etching gases, the substrate bias power, and the temperature in the reactive ion etcher (RIE) chamber.[13] Second, we deposited a fluorocarbon polymer over the trench walls by C4F8 (octafluorocyclobutane) plasma treatment. This deposited polymer acted as a passivation film (Figure 1a-5) to protect the trench-walls from lateral etching in the following step. Third, we applied the etching gas SF6 (sulfur hexafluoride) and O2 (oxygen) to yield isotropic etching and obtain tube-like channels (Figure 1a-6). After stripping off residues of the PMMA resist left on the silicon surface, a silicon mold containing micro-channels was obtained (Figure 1a-7). Scanning electron microscopy (SEM) images of the cross-sections of the channels (Figure 1b) show that each channel consisted of a tube-like structure approximately 1 µm in diameter, corresponding to the size of an E. coli cell, and an open-top channel 400 nm in height and 600 nm in width (more silicon-chip images and parameters in supporting information 1). After fabrication of the silicon mold, we made the first replica using PDMS. As Figure 1c illustrates, on the PDMS surface, we obtained an array of protuberances that were complementary to the silicon structure. small 2014, 10, No. 15, 3018–3025
Taking the first PDMS replica as the master, we made a second replica and obtained the final PDMS slide with the originally designed structures. The obtained PDMS slide displayed in Figure 1d was utilized as the patterned substrate for cell immobilization and oriented growth in the following AFM study. We investigated E. coli wild type strain MG1655 constitutively expressing the yellow fluorescence protein (YFP) by AFM. To load the cells into the channels, a PDMS block with a flow channel was first mounted on top of the microchannels (as shown in Figure 2a, top image). Then, two metal plates with screws were used to tightly clamp the two PDMS layers to maintain the cell medium during loading. After that, a concentrated cell culture with a cell density of 1010 cells/ml was injected into the flow channel. A centrifugation step assured cell loading into the micro-channels (Figure 2a, bottom image). As a control, we followed the cellular growth within the channels by fluorescence microscopy. In three hours of incubation, the cells grew and divided unhindered in the channels (Figure 2b). Furthermore, the three-hour timelapse fluorescence microscopy images indicated oriented growth of the cells within the channel (Figure 2c). The doubling time of the E. coli cells measured from fluorescence images was 45 min, in agreement with measurements in bulk culture by optical density. This demonstrates that the microchannels do not hinder cellular growth. Compared to previously described microfluidic chips for bacterial cell growth, our structures demonstrate significant advantages for AFM
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Figure 2. Cell loading. (a) Sketch of the loading method. Top: injection of E. coli cells and medium into the two-layer chip. Bottom: the cells can be loaded into the channels by centrifugation. (b) Fluorescence image of the cells in the PDMS channels after incubation. The green signal shows cells with YFP. Scale bar: 10 µm. (c) Time-lapse fluorescence images of cells dividing along a micro-channel. Scale bar: 2 µm.
studies: the micro-channels immobilize both the mother and daughter cells in the channels, and the cells undergo oriented growth in a linear colony. Interestingly, the open-top structure provided homogenous medium conditions along the channel; this chip comprised an “open” system (compared to a sealed microfluidic chip), which allowed the AFM probe to touch the cells and the growth medium to be changed over time. To enable AFM imaging on these unique channels, modifications to the AFM tip were necessary because the normal pyramid tip was too wide to reach the cells loaded in the 600 nm wide channels.[14] The AFM probe used here was a cantilever with a modified tip. A focused ion beam (FIB) was applied to sharpen the pyramidal Si3N4 tip to obtain an ultrasharp tip measuring 2.5 µm in length, only 20 nm in diameter at the tip point, and 300 nm at the tip base (Figure 3a and b). The E. coli cells were loaded into the PDMS channels as described above. The PDMS chip was then carefully released from the metal plates and attached to the bottom of a petri dish. We then washed the chip surface and added culture medium M9 with a low carbon source to the petri dish. This low nutrition condition and low temperature (e.g., room temperature instead of 37 °C) allowed for slow growth, which was necessary to visualize the division process in detail by AFM. Before AFM imaging, an inverted optical microscope was used to locate the cells in the channels (Figure 3a). Using the ultra-sharp AFM tip, topographic images of living E. coli cells (Figure 3c) were obtained. The top image in Figure 3c shows multiple dividing cells in one channel. Underneath, there are four single cells and a zoomed-in image with a resolution of 2 nm. All of the images in Figure 3c have fine imaging quality, shown by the smooth surfaces of the cells. These results demonstrate that the chip and the modified AFM tip facilitated the nanoscale probing of living E. coli cells.
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Figure 3. Chip setup and AFM imaging. (a) Scheme of the liquid AFM imaging system combined with inverted optical microscopy. The AFM tip is indicated by an arrow. (b) SEM image of the super-sharp AFM tip etched by FIB. Scale bar: 5 µm. (c) AFM images of single cells. Top: the cells dividing along a micro-channel. Scale bar: 1 µm. Bottom: five AFM images of single cells. The last image is a magnification of the fourth cell. Scale bars are 500 nm, 1 µm, 1 µm, 500 nm and 200 nm, respectively.
In the following study, we provide proof-of-concept experiments demonstrating the utility of the device as a tool to measure the dynamics of cell growth in a linear colony. Time-lapse AFM imaging was performed to monitor the growth and division processes of the E. coli cells loaded in the channel for more than 4 hours. Figure 4a consists of 16 topographic images showing a cell dividing into two daughter cells, then subsequently dividing into four cells. At 36 min (see Figure 4b), the cell membrane of the mother cell in the middle constricts at the division site (labeled by a black arrow). From 52 min to 96 min, the cell continues growing while the division site gradually contracts. During division in vivo, a circumferential protein ring (named the Z-ring) develops at the inner surface of the cell membrane at the division site, and this ring is believed to be an essential component of the bacterial division machinery targeted to the division site.[15] Here, AFM was used to visualize the 3D topography of the division site of the E. coli cell. The constriction at the division site was revealed by a depression in the cross-sectional curve (labeled by a black arrow) of the cell. Therefore, the time point of cell-division termination can be easily identified (see experimental section and supplementary information 2 for more details) using a cross-sectional profile of the division site. When the vertical distance between the top membrane surface and the division site (labeled as h in Figure S2a) equaled half the cell height, we considered the division to be complete. We measured the dynamic proportion of h/Height. At 107 min, this value suddenly ascend to 0.519 (shown in Figure S2), indicating division termination. At this moment, the AFM tip can protrude in between the two poles of the daughter cells and slightly separate them. The time point at 107 min, therefore, marked division termination in Figure 4a. At this time, the division site disappeared and the division was complete.
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Figure 4. Growth curves of E. coli cells based on time-series AFM images. (a) Time-lapse AFM images of E. coli growth over 274 min in M9 and at room temperature. Scale bar: 2 µm. (b) Zoomed-in images of cells obtained at 36, 219 and 237 minutes (black boxes in (a)). Scale bar: 500 nm. The black arrows show the constriction-structures in the cells’ division planes. The curves under each image are cross-sectional profiles along the long axis of each dividing cell. The division planes are indicated by arrows in these profiles. (c) Growth curves of the AFM-probed growing cells. The black dots correspond to the mother cell length measurements over time, and the blue and red dots represent its two daughter cells. Both the mother and daughter cell sizes increased exponentially with time.
To quantify the dynamic change, the cells’ lengths throughout the growth and division process described above were measured (supporting information 3). As shown in Figure 4c, the mother cell’s length was measured during the first 96 min. After the estimated point of division termination, at 107 min, the growth curves of the mother cell and the two daughter cells were plotted separately. The fitted growth curves demonstrate that both the mother and daughters in Figure 4a exponentially elongated under the imaging conditions. It is currently under debate whether cell elongation is linear or exponential during the life cycle of an E. coli cell.[16] This debate has persisted for decades primarily due to the lack of quantitative methods for measuring cell size with the required sensitivity. To distinguish an exponential pattern from a linear one, a resolution of under 6% in cell size is required.[17] Our results obtained by AFM have a resolution of 31 nm, which is only 3% of the cell size and well below this limit. Additionally, the cell increased in length during the AFM scan. The magnitude of this error depends on the speed of AFM scanning relative to cell growth. The scanning rate used to collect the images was 0.49 Hz, which represents a scanning speed of 19.5 nm/s along the Y axis of each image over 10 µm. The largest growth rate of the cells we measured was 0.39 nm/s. The scanning rate of AFM was thus approximately 50 times faster than the growth rate of the cells. Therefore, the error in the size measurement derived from the scanning time can be neglected. Thus, it can be concluded that this chip facilitates AFM measurements of cell growth with sub-diffraction-limited resolution. Examples of asymmetry in division as well as of variations in the cell growth rate, length-at-division, and time-to-divide that have previously been reported were also observed in this imaging experiment. First, the division site of E. coli is accurately positioned by specific protein machinery.[18] Here, the small 2014, 10, No. 15, 3018–3025
distance of the division site from the midcell at the beginning of cell constriction (at 36 min in Figure 4a) was 4.9% of the cell length. Second, the growth rates of the two daughter cells acquired from fitting the growth curves (Figure 4c) differed, leading to variations in the length-at-division with time. For example, the bottom cell (Figure 4a) was initially 300 nm longer than the upper cell (at 107 min), and this asymmetry increased to 550 nm before the second division (at 199 min). Finally, the future division site was visible in the longer cell 18 minutes before contraction could be observed in the shorter cell’s septation site (the division planes are labeled by black arrows in the zoomed-in images and in the crosssectional curves in Figure 4b at 219 min and 237 min). These observations demonstrate that the asymmetry of the mother and daughter cells loaded into the channels could be detected by AFM. With this on-chip AFM technique, the dynamic nanomechanical properties across E. coli cells can be investigated by Peak Force Tapping (PFT) AFM.[19] PFT is a relatively new AFM mode for studying the mechanical properties of cells at high speeds (similar to Tapping Mode[20]) and high resolution that has been used to explore the structural and physical properties of a variety of biological samples.[21] The imaging feedback is based on the peak force of the forcedistance curves. Because this force curve performance is repeated for each pixel of the image, the resolution of the mechanical images will be the same as that of a conventional height AFM image. Once the system is calibrated, the Young's modulus (YM) determined by the DMT (DerjaguinMuller-Toropov model)[22] fit can be directly displayed in the quantitative units of Pascals. Hence, this technique offers the possibility to map cellular mechanical properties at nanometer resolution. Figure 5a shows a time-series over 53 minutes of topographical AFM images (height image) of E. coli
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Figure 5. Time-lapse AFM topographic images and kinetics of mechanical measurements on E. coli cells. (a) Topographic images over 53 minutes. Scale bar: 500 nm. (b) Magnified, real-time Young’s modulus (YM) images over 53 minutes of the cell labeled by a black rectangle in (a). Scanning size: 0.74 µm×3.5 µm. Scale bar: 200 nm. (c) Cross-sections along the long axis of the cell showing the time progression of the YM variation. Each slice represents data extracted from one image in the full time series. (d) Histogram representations of the average distance between two stripes on 20 cells. The median distribution is 400 nm. The figure was obtained in M9 and at room temperature.
cells loaded in a micro-channel. Zoomed-in YM images with a resolution of 20 nm for the cell (labeled by the black box in Figure 5a) at each time point are displayed in Figure 5b, which depicts the logarithmic YM in pseudo color. The YM cross-section along the long axis of the cell in the full time series was further extracted and is shown in Figure 5c. In this figure, each slice represents the data extracted from each time point of Figure 5b, showing the time progression of the YM variation. The dynamic cell nano-mechanics can be quantified from this figure. First, many stripe-shaped areas with a dark blue or purple color are distributed perpendicularly to the cell's long axis (Figure 5b). The color scale reveals that these areas are more rigid than other parts of the cell. This feature can also be observed as a waved cross-sectional profile (Figure 5c). Recently, actin-like filaments forming a helical cytoskeleton under the inner membrane in E. coli cells were observed.[23] It was further shown that this cytoskeleton-like structure was sufficiently stiff to affect the overall mechanical rigidity of an E. coli cell, in much the same way that the actin cytoskeleton does in eukaryotes.[24] Therefore, we assumed that the observed oriented patterns on the single cells reflected the cytoskeletal arrangement underneath the cell membrane (supporting information 4). Presumably, when the tip interacts with the areas with a cytoskeleton support underneath, the YM information represents the mechanical integrity of the cell, which involves the cytoskeleton, cell wall, etc. In the other areas, there is lack of cytoskeleton information in the
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YM. To quantify this structural feature, the distances between the stripes distributed on 20 cells were measured. Figure 5d shows the histogram representation of the distance distribution, which reveals that the average distance between two stripes measured by AFM was approximately 400 nm. Another measurement is the average YM of the division site. It can be observed in Figure 5b that the division site had a much larger YM than the rest of the cell (see the image at 26 min and beyond). In the division process from 26 min to 53 min, the constriction-structure was rigidified (Figure 5c). We measured the average YM based on the data in this figure. At 53 min, the average YM of the division site was 1 MPa, while the average YM for the rest of the curve was 630.96 KPa. The YM of the division site was therefore 370 KPa larger than that of the rest of the cell. The rigidity of the division site presumably increased in order to facilitate the septation necessary for division. Finally, the average YM (excluding the division plane area) along the long axis throughout the division process (supporting information 5) was measured. It first increased from 288.40 KPa (5.46 log(Pa)) to 630.96 KPa (5.80 log(Pa)) and then decreased to 537.03 KPa (5.73 log(Pa)) at 0, 26 and 53 minutes, respectively. This YM variation implies that the average stiffness of an E. coli cell changes throughout the cell cycle. In summary, we developed a novel chip to facilitate the AFM imaging of oriented, growing E. coli cells. To immobilize the freely growing cells on a substrate and adapt the AFM tip
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to this system, technical improvements in the substrate fabrication and probe sharpening were achieved. These improvements extend AFM imaging applications to non-adherent living cells. These techniques can also be applied to trap other prokaryotic or eukaryotic (e.g., mammalian) cultured cells. And the tip sharpening will allow quantitative detection to be more accurate and credible. This novel design principle can be used and extended further to microfluidic channels of different internal dimensions and shapes. As a result of these technical improvements, the kinetics of oriented cell growth, including the growth pattern and the nano-mechanical properties, can be quantified at the single-cell level. These advances in micro-/nano-fabrication, AFM technology and their applications establish a powerful tool for exploring cellular processes in real time at the nanometer scale.
Experimental Section Fabrication of the Micro-Channel Array: Fabrication of the micro-channels was performed on a 100 mm diameter singlecrystal (100) silicon wafer (University Wafer, USA). The wafer were ultrasonically cleaned in fuming nitric acid (100% HNO3) for 10 min, rinsed in deionized (DI) water and spun dry. Then, the wafer surface was primed for resist adhesion using hexamethyldisilazane (HMDS) by spin-coating at 1000 rpm for 60 s, followed by baking for 90 s at 200 °C. After priming, a 1.2 µm layer of PMMA 950k positive electron-beam resist (MicroChem) was spin-coated onto the wafer at 2000 rpm for 60 s and baked at 175 °C for 10 min. Feature patterning with 9 individual channel-arrays arranged on the wafer was performed using a Vistec EBPG5000 Plus with an acceleration voltage of 100 kV and an aperture of 400 µm. Spot-size diameters of 56 nm were achieved by focusing the electron spots with currents of 129 nA. The doses were set to 1200 µC/cm2 with a 40 nm beam step size (BSS). The total machine time was approximately 3 h for the entire wafer. Following electron-beam exposure, development was conducted for 1 min in 1:3 MIBK (methyl isobutyl ketone):IPA (isopropyl alcohol), followed by a 30 s soak in IPA. The wafer was then rinsed in DI water and spun dry. The wafer was cut into 9 pieces containing trench-structures. The trench-structure layer was then dry-etched in a reactive-ion etcher (Adixen AMS100 I-speeder) with a mixture of 15 sccm SF6 (sulfur hexafluoride), 20 sccm C4F8 (octafluorocyclobutane) and 10 sccm CH4 (methane) diluted in 100 sccm He (helium). The source power was set to 2000 W, and the substrate bias was approximately −22 V. Under this etching condition, the obtained side-wall for this layer was almost perpendicular with the silicon surface (anisotropic etching). A chemically inert passivation layer was immediately deposited after anisotropic etching. Here, a 80 sccm C4F8 plasma was used to yield a fluorocarbon polymer on the structures for 10 s to protect the trench-walls from lateral etching in the following step. We next applied the etching gas SF6 and O2 to yield isotropic etching and obtain tube-like channels. Here, the etching gas was a mixture of 200 sccm SF6 and 120 sccm O2, the source power was 1200 W, and the substrate bias RF power was set to 20 W. In this etching process, the reactive ions first attacked the passivation layer on the bottoms of trenches. After the silicon surface under the passivation layer was exposed, a tube-like structure was obtained resulting from the isotropic small 2014, 10, No. 15, 3018–3025
ion path in the second etching process. Compared to anisotropic etching, this process is isotropic etching. Etching times of 22 s, 25 s, 27 s and 30 s were utilized to obtain variable tube diameters. After cleaning by HNO3, we obtained structures of open-top, tube-like channels. Table S1 in the supporting information lists all the structures on silicon we obtained with different sized microchannels. These silicon molds were applied for cells with diverse sizes. Figure S1a shows the zoomed-in cross-section of one microchannel with a size measurement. To transfer the patterns from the silicon chip to the PDMS surface, two PDMS replicas were necessary. A 10 µL volume of the hydrophobic chemical trimethylchlorosilane (TMCS) was dropped beside the silicon mold, sealed and maintained for 10 min. After hydrophobic treatment, 1:10 PDMS was poured on the chip and solidified by baking at 80°C for 2 h. The hydrophobic treatment to silicon and greatly decreased the interaction of the PDMS with the silicon. Besides, with the V-shaped structure (as marked by a black arrow in Figure S1b in the supporting information) at one end of the micro-channel array, it was therefore effortless to peel the PDMS slab from the silicon surface. The structure we obtained on the PDMS surface was the inverse of the silicon mold and was then used as the mold for the second PDMS replica. Then, a 2 min activation of the PDMS surface was conducted using oxygen plasma, followed by hydrophobic TMCS treatment for 10 h. In this process, each hydroxyl group on the silicon surface (a monolayer of hydroxyl groups was formed after plasma bombardment) reacted with a silane group of TMCS. After baking at 80°C for 2 h, the formation of a hydrophobic molecular film prevented adhesion between the two PDMS layers during peel off, and the second PDMS replication was easy to accomplish. Eventually, PDMS substrates containing surface micro-channels were obtained. E. coli Strain Information: E. coli K12 MG1655 fliC was prepared by standard lambda recombination-based replacement of the gene with a cat cassette that was further removed by FRT recombination. P1 transduction was used to insert the yellow fluorescence protein (YFP) gene under the control of a constitutive promoter (λPR) together with a chloramphenicol resistance gene (from MRR strain) at the intC locus. The fliC gene determines the flagella of E. coli. Thus, the fliC knockout strain has no flagella, which leads to their inability to swim in the medium. For the AFM investigation, 50 µL of cryo-preserved E. coli cells were recovered in 5 ml M9 medium (2 × 10−3M MgSO4, 1 × 10−4 M CaCl2, 3.4 × 10−4 g/mL Vitamin B1 with 0.4% w/v glucose) and cultured at 37 °C for 16 hours until reaching the stationary phase. The E. coli cells were then refreshed in M9 to reach the exponential phase (OD600 = 0.2). Cell Loading: To load the cells into the channels, the chip was covered with a PDMS block with a flow channel (500 µm in width, 20 µm in height and 1 cm in length) facing down into the channel array. Two metal plates with surrounding screws were utilized to tightly clamp two PDMS layers to maintain the cell medium in the flow channel in subsequent steps. A 1 mL volume of E. coli cell medium was concentrated by centrifuging at 2000 rpm for 5 min. Then, 100 µL of concentrated medium with a cell density of 1010 cells/ml was injected from one entrance of the channel. By centrifugation at 2000 rpm for 5 min under centrifugal force parallel with the chip surface, the E. coli cells were driven from the V-shaped entrance into the micro-channels. After centrifuging, the PDMS film was immediately released from the metal plates and then carefully put into a petri dish with the medium.
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Tip Sharpening: In air, the intrinsic resonant frequency of the probe was 97.42 kHz. We utilized a focused ion beam (FIB) (FEI Nova200 Nanolab) with an acceleration voltage of 30 kV to sharpen the probe. The ring-shaped (inside diameter 0.3 µm and outside diameter 1 µm) Gallium ion-beam with a current of 0.3 nA was used to etch the tip for 10 s. AFM Imaging System and Parameters: Time-series AFM images for the cell length measurements were acquired by an Agilent 5500 AFM (Agilent Corp.) in magnetic alternating current (MAC)mode placed on an inverted microscope (Nikon, TE2000). A Picoscan controller and a top MAC-Mode control-box were used to control the scanner and to acquire and convert the analog signals to digital signals. The software Picoview was used to command the controller and display the images. The MAC-mode AFM probe’s cantilever was coated with a magnetic material. A top magnetic field generated by a solenoid directly shook the cantilever. The probe was immersed in M9 medium to scan over the E. coli cells. In the imaging process, the resonant frequency of the AFM probe was 31.6 kHz. We set the ratio of the actual to raw amplitude of the probe, A/A0, to 0.83. The scanning speed was 0.49 Hz. The integral gain and proportional gain were 11% and 21%, respectively. The scanning range in the Z-axis was 3.3 µm. Speculation of Cell-Division Termination: The AFM 3D profile was used to determine this time point. As Figure S2 in the supporting information shows, AFM scanning allowed us to obtain a cross-section of an E. coli cell. In the division process of an E. coli cell, the division site gradually contracts. When the AFM tip scans the division site, the vertical depth of the tip required to contact the cell increases. At the time-point of cell-division termination, the two daughter cells are completely separated but remain physically in contact. At this time, the depth of the AFM tip is half of the height of the cell. We therefore considered the ratio of the vertical distance between the top membrane surface and the division site (labeled as h in Figure S2a) to the total cell height to be 0.5 when cell division is complete. The dynamic proportion of h/Height was measured. In Figure S2b, we plot the ratio in each image as a function of time. From 52 min to 96 min, the division site first appears and then constricts gradually. Correspondingly, the measured ratio increases from 0.066 to 0.327. At 107 min, this value suddenly increases to 0.519, and afterwards fluctuates between 0.4 and 0.5. Growth Curve Measurement: We measured the cell lengths in each image in the imaging sequence using the software ImageJ. The AFM image was first imported into the software. Then, lines were drawn on the image, and ImageJ gave the length result in pixels. We set the width of the drawing line (yellow line in Figure S3a of the supporting information) to 5, so the measured length was given by the average value of 5 lines. To remove imaging aberrations from scanning, a calibration along the cell's long axis for all the images was performed. We measured the distance between two particles on the edge of a channel (the distance shown by the black arrow in Figure S3a) in each image. The calibration factor for each image was acquired (listed in Table S2 of supporting information) using the normalization of each measurement by their average value (average distance was divided by respective measured value). In our measurements, the longest distance from one end of the cell to the other was considered to be the length of the cell. From the cross-sectional profile of this line (Figure S3b in supporting information), we determined the initial and terminal points
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of each measurement. Given all the cell calibration factors and the absolute length value of 1 pixel (31 nm), the final lengths were obtained (Table S2). Kinetic Nano-Mechanical Measurement: Our results were achieved by the Peak Force Tapping (PFT) mode (Bruker Dimension Icon SPM). In our imaging process, the probe was oscillated at 2 kHz. A deflection sensitivity of 37 nm/V was calculated by engaging the tip on a stiff glass slide and recording a force curve. The spring constant of the AFM probe calculated by “Thermal Tune” was 0.35 N/m. The tip radius measured from the SEM image was 10 nm. The scanning rate of each image was 0.65 Hz. We obtained Young's modulus images by entering these parameters into the software and employing a DMT fit.
Supporting Information Supporting Information is available from the Wiley Online Library or from the author.
Acknowledgements This work was supported by the Franco-Chinese cooperation program (PFCC) between INSERM and the Chinese Academy of Sciences (INSERM/CAS AO 2010–2011) and the AXA Foundation Chair. We acknowledge Dr. François-Xavier Pellay for his helpful discussion, Dr. Zhuangxiong Huang for advice on micro-fabrication, the TU Delft Nanofacility for micro-fabrication process support and Dr. Kaiwu Peng for FIB operation.
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Received: December 3, 2013 Revised: March 15, 2014 Published online: April 6, 2014
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