We acknowledge NIH COBRE funding under number P20GM130456. We thank the UK Light Microscopy Core, which is supported by the Vice President for Research, for assistance in this work. Strains of S. mutans and E. faecalis were provided by Dr. Natalia Korotkova.&#160;The first mention of the play-on-words, Conpokal, to describe the combination of confocal microscopy and atomic force microscopy is attributed to Dr. Brad Berron, Chemical and Materials Engineering, at the University of Kentucky, in discussion with Dr. Martha Grady (corresponding author) in anticipation of the arrival of a seminar speaker Dr. Jonathan Pham, who joined the faculty in Chemical and Materials Engineering at the University of Kentucky in the Fall of 2017.

The authors declare that they have no competing financial interests.

Conpokal is an advanced, effective technique for collecting high-quality images and in situ mechanical properties of live biomaterials in a liquid environment. The ability to collect exceptional morphology and topography images combined with live-sample mechanical properties extends past typical electron and light microscopy techniques. Separately, a confocal microscope provides brightfield, epifluorescence, and laser scanning confocal capabilities to achieve high-quality, detailed fluorescent images of samples. An AFM provides mechanical characterization, but without auxiliary optics it becomes difficult to navigate the sample. With the two systems combined, a user can collect both images and mechanical properties of the exact same cell during the experiment, which is a great advantage over two separate instruments. The purpose of this manuscript is to inform and guide users on the extensive capabilities of the combined Conpokal system for the future of biology, engineering, and health. The protocol involves preparation of instruments, culture media, dishes, selection of microorganisms, AFM procedure, confocal procedure, and clean up. The most critical steps for a successful live, in-solution, Conpokal session are sample immobilization, judicious tip selection, and live stain execution. The discussion section for this manuscript will elaborate on culture recommendations, troubleshooting tips and operational guidelines, and future work for the Conpokal technique. Culture recommendations will cover guidance on sample culture, immobilization, and staining. AFM, confocal, and Conpokal tips and guidelines will discuss tip selection and calibration, resolution, limitations, and near simultaneous operation. Future work includes the outlook and potential for prospective research pathways.
The protocol covers culture, probing, and imaging of both live cell and fixed bacteria samples. One challenge present when conducting AFM in liquid stems from cantilever movement obstruction due to sample hindrance and hydrodynamic drag. If the sample is not well adhered to the substrate, there is potential for fouling of the cantilever by sections of the sample floating in the liquid. In that case, measurements are compromised since they are a supposition of elastic adherence and micromechanical properties of the sample. The HEK cells were not treated with fixative, however, S. mutans and E. faecalis require additional immobilization, similar to other prokaryotic cells. The bacteria chosen showed active movement which hindered cell probing and imaging, therefore, the samples were gently, chemically fixed to promote immobilization. There are alternatives to chemical fixation, such as filter membranes that physically trap individual cells in pores75.
Tip selection is also a critical component to the setup and operation of the AFM. When mechanical properties based on deformation of the biomaterial are sought, one must consider cantilever stiffness and material stiffness compatibility, which is a non-trivial task. The main goal is to best match the stiffness of the material with the stiffness of the selected cantilever. If the cantilever is much softer than the sample, it will be subject to too much deflection. If the cantilever is stiffer than the sample, then the AFM detector may be unable to capture such small deflections. It is recommended that when selecting a suitable AFM cantilever, to choose based on experimental application. To collect detailed images in intermittent contact mode, AFM tips within a range of 0.1 - 0.3 N/m for stiffness and tip radius within 5 nm &#8211; 100 nm are effective for live cell imaging. Small conical tips are recommended. Sharp tips provide a small contact area and ability to indent further aiding in collecting small details and features in sample morphology76. However, sharp tips can also be problematic as they are prone to puncturing samples when the settings (e.g., set point, pixel time, approach velocity) require too much indentation or the indentation is too fast. To avoid high-strain rate effects, local strain hardening near a sharp tip, or simply to control the contact area and approach speed, many choose to use force spectroscopy with a colloidal tip to measure an elastic modulus.
AFM tips within a range of 0.01 - 1.0 N/m for stiffness and tip radius within 1 &#8211; 5 &#181;m are recommended to measure the elastic moduli of living cells. Large tip sizes, typically glass colloids, provide a known contact area and are unlikely to puncture the cell while in contact. Rectangular cantilevers are preferred over triangular because during calibration, the contact-free method can be used for rectangular geometry as compared to contact-based calibration for triangular geometries. Also, due to the delicate nature of AFM tips, it is recommended that the operator implements tweezers with rubber tips to reduce the potential for damaging the delicate AFM chip or mounting block. Other parameters to keep in mind include the approach velocity of the AFM tip and the amount of indentation into the sample. A good guideline is to keep indentation to about 10% of the thickness (or height) of the sample and choose a velocity that precludes potential hydrodynamic resistance experienced by the cantilever38.
Intricate experimental instruments typically come with roadblocks that require troubleshooting in the instrument set up, calibration, and operation. Crashing the AFM tip or cantilever into the sample or sample substrate is a common mistake for new users. To avoid this issue, the protocol suggests backing the cantilever out 2000 &#181;m. This step ensures the tip will not come into contact with the bottom of the dish if the previous user forgot to back out the tip when cleaning up after experimentation. However, the 2000 &#181;m distance was chosen for the selected dish holder used in this protocol. A different range, larger or smaller, may need to be selected depending on the dish holder style being used. During alignment of the laser and detector, the protocol mentions adjustment of a mirror knob to maximize the sum signal. A mirror adjustment knob may not be present on all AFMs. If present, however, one way of manipulating the instrument to troubleshoot a low sum signal is to adjust the mirror knob, used to account for the medium in which scanning will take place; either liquid (e.g., water, media, PBS) or air. Due to the difference in index of refraction for light through air and through liquid, the mirror knob may need to be adjusted. The maximum bending sensitivity of the AFM cantilever will be at the location of the AFM tip, therefore, the laser light, which returns the bending position through its placement on a photodiode, must be located at the location of the AFM tip. Depending on the AFM tip chosen, the sum signal value could vary from 0.3 &#8211; 3.0 V. A backside coating on the AFM cantilever such as Cr-Au or Al will increase the sum signal and the sensitivity of the measurement.
Tip geometry is pertinent when completing tip calibration during AFM operation. Good agreement has been observed between non-contact and contact calibration. If the chosen AFM cantilever is not rectangular, contact calibration will need to be performed. Keep in mind that the medium in which the tip is calibrated must be the same as the sample medium. If those fluids differ, the user must recalibrate. When using the AFM tips in liquid, the frequency measured by the system should be one-fourth to one-third of that of the natural resonance frequency denoted by the manufacturer. A good way to check that the system calibrated the tip correctly is to verify values within the thermal noise file that is generated. Ensure this file saves to the appropriate folder. If the system has trouble calibrating or non-probable values come out, recalibrate, or adjust the laser position slightly then recalibrate.
Another cause of poor sum signal may be due to AFM cantilever alignment. When the AFM chip is mounted into the glass block, it is essential that the tip of the cantilever remains within the small laser window (smooth glass area). If the chip is too far forward, the angle at which the cantilever naturally rests may cause an issue with the reflection off the cantilever, missing the photodetector and resulting in poor sum signal. If the chip is too far back, the laser will be unable to reflect off the back of the cantilever, causing poor sum signal. For these reasons, the mounted AFM chip may need to be adjusted. Additionally, another issue that may be encountered occurs with the height of the sample. The AFM instrument used in this protocol has a maximum piezo z range (height) of 15 &#181;m. If a user finds that the software is unable to collect the height data and force maps in a particular pixel, a black box will appear indicating that the system is out of range (Figure 2C). One way to trouble shoot this issue is to set the piezo height to a lower value, such as 2 or 3 &#181;m so most of the 15 &#181;m range is committed to mapping the anticipated height of the cell. This technique should, in most cell- or bacteria-related experiments, fix the issue associated with the z range.
Experimentalists that require an extended z range for tall samples with heights greater than 10 &#8211; 15 &#181;m may need to pursue an additional module on the AFM. AFM manufacturers have this option available at additional costs for most systems. By extending the z range, the experimentalist has the availability to scan samples that are considered tall at the micro-scale with little issues for out of range values or AFM piezo motor modification. Although these modules cost extra, some, depending on the manufacturer, can offer additional height, up to 100 &#181;m in the z direction. Confocal is still possible with taller samples if the user has a long working distance, high magnification objective or is willing to use an air objective, perhaps a 20x or 40x. By lowering microscope objective magnification, the working distance increases, gaining distance to view the apex of a taller sample. This modification to a lower magnification objective will sacrifice resolution. In the Conpokal setup referred to in this manuscript, the 60x TIRF (total internal reflection fluorescence) objective has a working distance of near 100 &#181;m past the coverslip of the glass-bottomed sample dish.
In regard to the confocal microscope referred to in this manuscript, a few important stipulations are discussed. The confocal system used for the production of the figures in this manuscript implemented a 60x TIRF oil objective with a numerical aperture of 1.49. Laser&#160;lines at 405 nm, 488 nm and 561 nm excitation wavelengths were used for live cell sample imaging, shown in Figure 3 and Figure 4. The diffraction limit of the confocal microscope can be determined by using the Abbe Resolution equation, Abbe Resolution(x,y) = &#955;/2NA, where &#955; is the excitation wavelength for Alexa 488, at 488 nm, and NA is the numerical aperture for the confocal condenser, which is 0.3. Therefore, an axial resolution of 272 nm is determined. For epifluorescence imaging, two cases are considered to determine the resolution where the pinhole is set to one airy unit (AU) and 0.5 AU. In the latter case, the pinhole is closed down such that significant light loss occurs, but the resolution increases. The confocal software calculates lateral native and axial native resolutions at 170 nm and 290 nm for the pinhole at 0.5 AU, and 200 nm and 370 nm for the pinhole at 1 AU, respectively. Spherical aberrations introduced in the system can be accounted for through a deconvolution process to increase contrast and resolution in microscope images. Due to the diffraction limitations that are inherent with confocal microscopes, the confocal image of the bacteria colony in Figure 6 lacks the matching resolution for the detail seen in the AFM scan of bacteria in Figure 5. AFM provides access to nanoscale features and detail that is difficult to capture with a confocal microscope. However, depending on the required fluorescence resolution needed, Figure 5 and Figure 6 demonstrate the applicability of the Conpokal technique to microbes in addition to eukaryotic cells.
An advantage of using a confocal microscope allows the operator to collect 3D images of specific regions in a sample with sharpened detail. These images correlate with the AFM by viewing the surface that was probed via the AFM image and a confocal scan of the same region. Since the microscope is inverted, an epifluorescence image collects light information from the opposite side of the sample that was probed. The pinhole within the confocal system helps limit to a single plane from a certain distance while filtering out light that comes from the rest of the sample or even the room. Essentially, the pinhole helps isolate the light coming back from the single plane of interest in the sample. Typically, this plane of interest must contain strong fluorophore markers because, for inverted confocal systems, single molecule detection would be limited to higher-resolution confocal microscopes. Epifluorescence illumination is less desirable due to the fact that in epifluorescence imaging mode, any light from the sample that is reflected into the objective is collected and used to generate the image, therefore, impossible to isolate a single plane. Confocal techniques provide a more isolated single plane image of the sample feature in question because of the pinhole77. If, for example, the apex of a eukaryotic cell is probed by AFM, that same surface can be isolated with the laser scanning confocal capabilities of the microscope, rather than with epifluorescence imaging mode. It is recommended to monitor the health/shape of the cells during data acquisition via imaging simultaneously in differential interference contrast mode with the aid of the transmitted light detector and the 488 nm laser line. When capturing a z stack of the sample, for the procedure detailed above, only the gain of the detector is adjusted. Any morphological change in the cells during the measurement, which is not necessarily visible in the fluorescent channels, indicates that artifacts are introduced into the measurement.
Ideal spacing between planes can be obtained by following the software recommendation for the shortest wavelength used in the imaging technique. The native resolution and the signal to noise ratio in the image volumes can be effectively improved by employing deconvolution algorithms available in the image processing module of the acquisition software. However, performing fluorescent microscopy, selection of specific stains and dyes is vital to avoid early on-set bleaching or crosstalk from overlapping excitation/emission spectra. On occasion, a user may experience a malfunction in confocal light generation. If a user experiences a lack of light emission or malfunctioning laser lines, one way to troubleshoot is to reset the system, typically done by restarting the operating software. If the issue persists, the transmitted light detector may have failed to move into or out of place within the optical path of the light microscope. Resetting the position of the transmitter detector may help to alleviate light collection or laser imaging issues.
The focus of the Conpokal instrument is to provide users with the ability to collect optical and force-based information on live biomaterials in a liquid environment, simultaneously and on the same cell or feature. This work explicitly describes how to perform these experiments in liquid, a natural home for many biomaterials, although, dry experiments can still be performed using the instrumentation. With samples prepared in Petri dishes, the dish height is a limitation. Due to the configuration of the glass block that holds the AFM cantilever, the side walls of the dishes must be under 10 mm in height; if the dish is too tall, the instrument will not be able to lower the AFM tip to the surface of the sample or the substrate.
Although there is a limitation for sample size, there is no limitation with the instrument or software capabilities with regard to a time delay. Simultaneous confocal and AFM is possible with the right factors in place. The limitation that contributes to its near simultaneous capability refers to the noise that is generated when performing certain confocal microscopy functions and atomic force microscopy functions simultaneously. The vibrations from the motors that move the microscope objective during collection of a z stack will be added to the signal from the AFM probe tip during its motion. The noise will be enhanced by an oil objective, as the motors move up and down in the z direction to illuminate sequential planes in the sample. Therefore, the recommended protocol includes sequential collection of AFM scans and confocal z stack images. Simultaneous CLSM and AFM would require stationary imaging with the confocal, however, with the current technique, the delay time for the two instruments could be as short as tens of seconds. The actual time to switch from AFM operation to confocal imaging was around 2 &#8211; 4 minutes for the images collected in Figure 2A and Figure 4B. This value was determined by subtracting the two image collection time periods from the total time of 33 minutes, which includes the time to begin and complete the AFM scan, switch instrument modes to activate confocal imaging, and begin and complete the confocal image z stack.
The future of Conpokal aims to explore new structure-function relationships in addition to keen insight of single cell processes. For example, experimentation of in situ drug treatments on cell or bacterial samples to determine the effects of cell elasticity would be an advancement to the fields of biomaterials, biology, and biomechanics. Treatment therapy into the sample dish while imaging and probing would provide knowledge of how the sample responds to therapeutics in a live and carefully controlled environment, over time. Incorporating a new drug or environmental challenge would broaden the understanding of how the cytoskeleton or organelle location affects locomotion, topology, stiffness, etc. Another potential advancement of Conpokal is the ability to have complete environmental control of the system. The current Conpokal mentioned in this protocol is housed inside of an acoustic enclosure designed to reduce noise from inside of the laboratory. Advancement of this housing would provide the ability to test within, perhaps, one or a combination of factors, not limited to those such as a sterile environment, temperature-controlled, or even variable-gravity. As it stands, the Conpokal method provides an effective and useful approach for characterizing live, in-liquid biomaterials, but the future of the technique will only advance these capabilities further.

Micro- or nano-scale mechanical evaluation tools are often employed to uncover the deformation characteristics on the single cell or microorganism level, while separate high resolution microscopy tools are employed to visualize subcellular processes and structural features. Conpokal, is the combination of laser scanning confocal microscopy and atomic force microscopy (AFM) into a single instrumental platform. The Conpokal technique was first implemented in a non-biological polymer system, where the goal was to determine the adhesion, elasticity, and surface tension of hard surfaces in contact with soft surfaces. Confocal images provided a visual rendering of how the polymer deforms, adheres, and releases from the hard probe1,2. From polymers to biological samples, this technique provides the opportunity for near simultaneous live cell or microorganism confocal imaging and AFM.
Changes in cell elasticity have been implicated in the pathogenesis of many human diseases3 including vascular disorders4, malaria5,6, sickle cell anemia7, arthritis8, asthma9, and cancer6,10,11,12,13,14,15. Common techniques to measure the mechanics of cells include the use of magnetic beads16,17, optical tweezers18,19, micropipette aspiration8,20,21,22, and AFM11,23,24,25,26. Since the application of AFM to living cells, it has readily been adapted to characterize cell topography27,28 as well as mechanical properties11,24,29,30,31 with nanoscale precision. AFM has been further adapted for microrheology32,33, frequency modulation34,35, and creep25,36,37 experiments to study the viscoelastic properties of various cell lines. The use of an appropriate nanocontact model is critical to the extraction of quantitative elasticity values from AFM-produced force indentation measurements1,2. AFM studies that investigate the influence of cytoskeletal drugs on cell elasticity have shown the elastic modulus to be highly affected38. Based on the elastic modulus measurements, many researchers have shown that cancerous cells are softer than their non-transformed counterparts11,38,39. Increased deformability likely plays a prominent role in the ability of cancer cells to metastasize and infiltrate tissues40. Such behavior is regulated by modifications in the cytoskeletal organization of cells38,41,42. In addition to cancer, another class of mechanical-dependent diseases are respiratory illnesses. For example, acute respiratory stress syndrome affects more and more humans each year. It has been shown that when patients are put on mechanical ventilation, with high concentrations of oxygen, their condition can worsen43. In a series of studies observing alveolar epithelial macrophages with AFM, scientists found that the addition of an oxygen-rich environment increased the stiffness of cells due to actin formation; a prime example of the impact that AFM has on our detailed understanding of atmospheric environmental influence on respiratory function at the cellular level and, ultimately, human healing and health44,45,46. Epithelial cell stiffness during migration toward a wound can also be assessed via AFM and that a variation in elastic modulus occurs to signal future cell spreading47,48. Therefore, there is a practical need to measure cell mechanics quantitatively to understand how diseased cells differ from, and interact with, healthy ones.
AFM is also used to study adhesive properties and surface structure. An atomic force microscope has a variety of modes to collect information about a sample using contact mechanics. For living biological samples, two of the primary objectives are to obtain quantitative mechanical property values (e.g., elastic moduli, adhesive forces) as well as map surface height. These modes include tapping mode (also known as intermittent contact), which maintains a constant cantilever amplitude intermittently contacting the sample surface and contact mode, which raises or lowers the cantilever to maintain a constant deflection49. Adhesion maps can be collected using force mapping on the AFM system. The cantilever indents the sample to a certain force and is then retracted. The force resisting retraction is measured by the detector to generate a force-displacement graph. The adhesion force is the maximum force measured during retraction. Surface morphology provides a detailed view of the sample on the micro- or nano-level. The cantilever completes a raster scan across the sample based on the indentation and deflection captured by the motion of the cantilever at each pixel, revealing micro- and nano-sized features. With AFM, the size of small features such as peptidoglycan structure50, polymer chain alignment51, flagella52, lamellipodia53, and filipodia54 can be measured. While AFM&#8217;s power lies in force-displacement curves and non-optical topological images, confocal microscopy offers detailed imaging of fluorescently labeled samples.
Confocal laser scanning microscopy (CLSM) is a dominant technique for imaging live cells, fixed cells, and tissues55,56,57. CLSM has been employed for biological imaging since 1955 i.e., since its inception58,59. While the working principle of confocal microscopes (i.e., the rejection of out of focus light through a pinhole) has remained largely the same, the technical implementation has become very diverse56,57,58,60. Confocal imaging can be implemented via multipoint scanning, as in a spinning disc system61, point scanning of a single laser beam, as in line scanning62, or imaging the point spread function with subsequent pixel reassignments via a multi-element detector57. Recent advances that expand the utility of confocal imaging include laser scanning capability, high speed resonance scanning, and high sensitivity detectors63,64,65. The advantage that all confocal systems have over widefield microscopes is the ability to perform optical sectioning in the z axis with greater detail, especially in thick samples. Widefield microscopes using camera-based detection collect light that is emitted from the focal plane and, simultaneously, light emitted from everywhere in the illumination path. By closing the pinhole, at the cost of reducing the signal, both the axial and lateral resolution can be increased66. In CLSM, images are built up pixel by pixel through emission signal collection as an excitation laser is scanned across an x-y field of view. Collection of several x-y planes along the z axis of the sample can then be used to reconstruct the 3-dimensional architecture of the sample57,67. Confocal microscopy in conjunction with the introduction of fluorescent proteins, has revolutionized our understanding of cell biology68. For example, it has become an essential tool in neuroscience69. The CLSM is regularly used to observe cleared tissue, and to obtain comprehensive images of immune stained brain and neuronal network architecture70. This application has resulted in new insights into synaptic contacts and communication among neurons and glial cells in the brain71. From brain cells to vesicles, fluorescent staining protocols support inspection and capture of cell functions via confocal microscopy for advancements in therapeutic techniques72,73.
Although they are impressive and versatile tools individually, the combination of confocal and atomic force microscopy (Conpokal) allows researchers to uniquely correlate topographical and micromechanical cellular/tissue properties with the observation of various cellular organelles and their respective dynamics. Paired instrumentation allows users to, essentially, &#8220;see&#8221; what they are probing; a huge advantage over one technique on its own. With Conpokal, force mapping through AFM is overlaid on confocal images to correlate cell stiffness, adhesion, or morphology to the cytoskeletal structure within the sample, for example. The overall goal of the Conpokal method is to provide a platform for researchers to study living samples in a concise, effective manner, providing both image and material property data in a single platform. In this work, we demonstrate the operation of Conpokal including proper sample selection and preparation, instrument setup, cantilever calibration, and provide guidelines to successful troubleshooting. After the protocol, we provide representative results that include successful bacteria and cell AFM height mapping, bacteria and cell AFM modulus mapping, and confocal imaging of multi-labeled cells, through same-cell confocal and AFM. We conclude with a discussion of the Conpokal procedure critical steps, troubleshooting recommendations, technique limitations, significance, and future outlook for the method.

<table><tbody><tr><td>Acoustic Enclosure</td><td>JPK-Bruker</td><td>Acoustic Enclosure</td><td>Chamber used to mitigate noise</td></tr><tr><td>BioTracker 488 Green Microtubule Cytoskeleton Dye</td><td>Millipore Sigma</td><td>SCT142</td><td>Used to label microtubules in mammalian cells</td></tr><tr><td>CP-qp-CONT-BSG AFM tips</td><td>NanoAndMore</td><td>CP-qp-CONT-BSG-B</td><td>Collodial AFM tips good for modulus measurements on biological samples</td></tr><tr><td>Dil Stain (1,1&#039;-Dioctadecyl-3,3,3&#039;,3&#039;-Tetramethylindocarbocyanine Perchlorate (&#039;DiI&#039;; DiIC18(3)))</td><td>ThermoFischer Scientific</td><td>D282</td><td>Used to label plasma membrane of mammalian cells</td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium</td><td>VWR</td><td>VWRL0100-0500</td><td>Component of cell culture media</td></tr><tr><td>Dulbecco&#039;s Phosphate-Buffered Saline</td><td>ThermoFisher Scientific</td><td>A1285601</td><td>Used to simulate biological environment</td></tr><tr><td>Fetal Bovine Serum</td><td>Fischer Scientific</td><td>45000-736</td><td>Component of cell culture media</td></tr><tr><td>Fisher BioReagents Microbiology Media Additives: Yeast Extract</td><td>Fischer Scientific</td><td>BP1422-500</td><td>Component of bacterial culture medium</td></tr><tr><td>FluoroDish Cell Culture Dish</td><td>World Precision Instruments (WPI)</td><td>FD35-100</td><td>Glass-bottomed dishes whose side walls are less than 10 mm, necessary for full function of Nanowizard 4a AFM</td></tr><tr><td>Hoechst 33342 nucleic acid stain</td><td>ThermoFischer Scientific</td><td>62249</td><td>Used to label nucleus of mammalian cells, a good nuclear marker for live cells</td></tr><tr><td>Human embryonic kidney 293 cells</td><td>ATCC Global Bioresource Center</td><td>CRL-1573</td><td>Mammalian cells used in protocol</td></tr><tr><td>Matrigel</td><td>VWR</td><td>47743-716</td><td>Coating in cell culture dish</td></tr><tr><td>Model: PFQNM-LC-A-CAL AFM tips</td><td>Bruker</td><td>PFQNM-LC-A-CAL</td><td>AFM tips that work well for samples that have large heights (such as eukaryotic cells)</td></tr><tr><td>NanoWizard 4a</td><td>JPK-Bruker</td><td>NanoWizard 4a</td><td>AFM</td></tr><tr><td>Nikon A1 Laser Scanning Confocal Microscope</td><td>Nikon</td><td>A1 HD25</td><td>Microscope used for brightfield, epi-fluorescence, and confocal imaging</td></tr><tr><td>PENICILLIN/STREPTOMYCIN</td><td>VWR</td><td>16777-164</td><td>Component of cell culture media</td></tr><tr><td>PetriDishHeater</td><td>JPK-Bruker</td><td>PetriDishHeater</td><td>Maintains desired Petri dish sample temperature</td></tr><tr><td>qp-BioAC-Cl AFM tips</td><td>Nanosensors</td><td>qp-BioAC-Cl-10</td><td>Three different AFM cantilevers of varying stiffness</td></tr><tr><td>Replaceable Carbon Fiber Tip Tweezers</td><td>Ted Pella Inc.</td><td>5051</td><td>Used to carefully place and remove delicate AFM tips into the glass mounting block</td></tr><tr><td>Todd Hewitt Broth Bacto</td><td>BD DIFCO Bacterius Ltd</td><td>Bacto-249240</td><td>Component of bacterial culture medium</td></tr><tr><td>Vancomycin, BODIPY FL Conjugate</td><td>Thermo Fisher Scientific</td><td>V34850</td><td>Used to fluorescently label bacterial cell wall</td></tr></tbody></table>

near simultaneous laser scanning confocal and atomic force microscopy (conpokal) on live cells

Techniques available for micro- and nano-scale mechanical characterization have exploded in the last few decades. From further development of the scanning and transmission electron microscope, to the invention of atomic force microscopy, and advances in fluorescent imaging, there have been substantial gains in technologies that enable the study of small materials. Conpokal is a portmanteau that combines confocal microscopy with atomic force microscopy (AFM), where a probe &#8220;pokes&#8221; the surface. Although each technique is extremely effective for the qualitative and/or quantitative image collection on their own, Conpokal provides the capability to test with blended fluorescence imaging and mechanical characterization. Designed for near simultaneous confocal imaging and atomic force probing, Conpokal facilitates experimentation on live microbiological samples. The added insight from paired instrumentation provides co-localization of measured mechanical properties (e.g., elastic modulus, adhesion, surface roughness) by AFM with subcellular components or activity observable through confocal microscopy. This work provides a step by step protocol for the operation of laser scanning confocal and atomic force microscopy, simultaneously, to achieve same cell, same region, confocal imaging, and mechanical characterization.

The Conpokal technique is schematically represented in Figure 1A and an image of the setup is shown in Figure 1B. In order to accurately co-locate biological structures via confocal microscopy with the same structures in AFM, the alignment of the two systems during installation is critical. Select reputable confocal and AFM manufacturers that are familiar with each other&#8217;s spatial requirements. Additionally, successful implementation of the technique relies upon good staining procedures, immobilization of the biological structure, and the appropriate choice of AFM tip. The height of living cells often presents a challenge for AFM imaging. An AFM scan over a living HEK cell is shown in Figure 2A. The height for this particular HEK cell was around 10 &#956;m, demonstrated by the line scan in green (Figure 2B). An AFM tip with either zero offset (tip is at the very end of the cantilever) or the tip height greater than the cell height (e.g., tip height &#8805; 10 &#956;m) will produce a highly resolved image of individual cells. An example of a poor scan due to improper AFM tip choice is included in Figure 2C. In this image, black pixels appeared at the apex of the cell indicating that the AFM piezo is out of range due to a large cell height. The end of the AFM cantilever appeared in the image (a rounded square) because of tip offset combined with insufficient tip height compared to cell height. These artifacts in the AFM image indicate a different AFM tip should be chosen to image the cell.
Efficient labeling in fluorescence staining along with the correct probe for live cell imaging are required for this method. We performed a three-color confocal image shown in Figure 3A with a nuclear stain (fluorescing blue), a microtubule stain (fluorescing green), and a lipophilic membrane stain (fluorescing red). These live cell probes were chosen because of their limited spectral overlap and their compatibility with standard filter cubes. We have also included the brightfield optical image in Figure 3B.
From AFM, analysis of the tip indentations along with a nanomechanical model, produces a modulus map of the surface. An example of a modulus map constructed using a Hertz model fit and a parabolic profile (radius of 8 nm) for the tip shape is shown overlaid onto the 3D reconstruction of the cell shape in Figure 4A (which is the same two cells shown in Figure 3). The corresponding 3D projection of the laser confocal z stack is shown in Figure 4B.
The method is also appropriate for smaller biological structures, including single bacteria whose micro and nanoscopic features are difficult to resolve using light microscopy. An on-going study utilizes the Conpokal technique to explore mechanical mechanisms within the cell wall that influence antibiotic resistance in Streptococcus mutans.74 Manipulation of the cell wall components resulted in changes in morphology and antibiotic resistance. Conpokal facilitates live, in-solution, data collection (e.g., surface morphology, elastic modulus, adhesion strength, and surface roughness) on same-cell samples to couple mechanical properties with the presence or absence of cell wall components. Figure 5A,B include an AFM scan and measured modulus map of a Streptococcus mutans bacterium. Better resolution was achieved at this scale with AFM than with traditional confocal microscopy.
Figure 6 contains a confocal image of a colony of Enterococcus faecalis stained with a green fluorescent cell structure stain, which attaches to the cell wall. Figure 5 and Figure 6 demonstrate the applicability of the Conpokal technique to microbes in addition to eukaryotic cells.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61433/61433fig01.jpg" /> 
Figure 1: The Conpokal setup. 
(A) Schematic of Conpokal technique. (B) Image of the instrument setup. <a href="https://www.jove.com/files/ftp_upload/61433/61433fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61433/61433fig02.jpg" /> 
Figure 2: Atomic force microscopy of living cells. 
(A) AFM image of two HEK cells. (B) Height profile (green line in A) of an HEK cell indicating the cell height is quite large at 10 &#956;m. (C) Poor AFM image of an astrocyte cell where the AFM piezo is out range at the apex of the cell and there is evidence of reverse imaging of the cantilever end. <a href="https://www.jove.com/files/ftp_upload/61433/61433fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61433/61433fig03.jpg" /> 
Figure 3: Confocal and brightfield microscopy of living cells. 
(A) Laser scanning confocal microscopy of living HEK cells stained with a nuclear stain (fluorescing blue), a microtubule stain (fluorescing green) and a lipophilic membrane stain (fluorescing red). (B) Corresponding brightfield optical image. Scale bars are 10 &#956;m. <a href="https://www.jove.com/files/ftp_upload/61433/61433fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61433/61433fig04.jpg" /> 
Figure 4: Comparison of 3D renderings of AFM and confocal microscopy of living HEK cells. 
(A) Modulus map constructed using a Hertz model fit and a parabolic profile (radius of 8 nm) for the tip shape overlaid onto the 3D reconstruction of the cell from AFM. (B) The corresponding 3D projection of the laser confocal z stack with a nuclear stain (fluorescing blue), a microtubule stain (fluorescing green) and a lipophilic membrane stain (fluorescing red). <a href="https://www.jove.com/files/ftp_upload/61433/61433fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61433/61433fig05.jpg" /> 
Figure 5: AFM of bacteria sample. 
(A) AFM scan and (B) measured modulus map of a Streptococcus mutans bacterium. <a href="https://www.jove.com/files/ftp_upload/61433/61433fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61433/61433fig06.jpg" /> 
Figure 6: Confocal microscopy of bacteria sample. 
Confocal image of a colony of Enterococcus faecalis with green fluorescent membrane stain. Scale bar is 5 &#956;m. <a href="https://www.jove.com/files/ftp_upload/61433/61433fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy Conpokal on Live Cells at JoVE.com

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy Conpokal on Live Cells

Presented here is the protocol of the Conpokal technique, which combines confocal and atomic force microscopy into a single instrument platform. Conpokal provides same cell, same region, near simultaneous confocal imaging and mechanical characterization of live biological samples.

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy (Conpokal) on Live Cells

Techniques available for micro- and nano-scale mechanical characterization have exploded in the last few decades. From further development of the scanning ...

near-simultaneous-laser-scanning-confocal-atomic-force-microscopy

Research

JoVE Journal

Bioengineering

6.8K Views.  University of Kentucky. Conpokal combineert confocale microscopie met atoomkrachtmicroscopie met behulp van een sonde om het monsteroppervlak te porren. Hoewel beide technieken individueel effectief zijn, vergemakkelijkt Conpokal fluorescentiecolokalisatie met mechanische karakterisering. Het belangrijkste voordeel van de Conpokal techniek, is de bijna gelijktijdige dual-microscopie.Confocal onderzoekt cytoskeletale en andere cellulaire processen voor en na AFM indringende, die gebied specifieke mechanische e...

Video: Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy Conpokal on Live Cells

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

Atomic Force Microscopy of Red-Light Photoreceptors Using PeakForce Quantitative Nanomechanical Property Mapping

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be measured to ~10 pN by a laser and sectored detector, which can be converted to image topography. Amplitude modulation or &#8220;tapping mode&#8221; AFM involves the probe making intermittent contact with the surface while oscillating at its resonant frequency to produce an image. Used in conjunction with a fluid cell, tapping-mode AFM enables the imaging of biological macromolecules such as proteins in physiologically relevant conditions. Tapping-mode AFM requires manual tuning of the probe and frequent adjustments of a multitude of scanning parameters which can be challenging for inexperienced users. To obtain high-quality images, these adjustments are the most time consuming.
PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) produces an image by measuring a force response curve for every point of contact with the sample. With ScanAsyst software, PF-QNM can be automated. This software adjusts the set-point, drive frequency, scan rate, gains, and other important scanning parameters automatically for a given sample. Not only does this process protect both fragile probes and samples, it significantly reduces the time required to obtain high resolution images. PF-QNM is compatible for AFM imaging in fluid; therefore, it has extensive application for imaging biologically relevant materials.
The method presented in this paper describes the application of PF-QNM to obtain images of a bacterial red-light photoreceptor, RpBphP3 (P3), from photosynthetic R. palustris in its light-adapted state. Using this method, individual protein dimers of P3 and aggregates of dimers have been observed on a mica surface in the presence of an imaging buffer. With appropriate adjustments to surface and/or solution concentration, this method may be generally applied to other biologically relevant macromolecules and soft materials.

A method for investigating the structure of a protein photoreceptor using atomic force microscopy (AFM) is described in this paper. PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) reveals intact protein dimers on a mica surface.

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be ...

Engineering

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

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

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

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

Live Cell Response to Mechanical Stimulation Studied by Integrated Optical and Atomic Force Microscopy

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to investigate cells behavior at sub-cellular level with high spatial and temporal resolution was developed. Thus, an atomic force microscope (AFM) was integrated with total internal reflection fluorescence (TIRF) microscopy and fast-spinning disk (FSD) confocal microscopy. The integrated system is broadly applicable across a wide range of molecular dynamic studies in any adherent live cells, allowing direct optical imaging of cell responses to mechanical stimulation in real-time. Significant rearrangement of the actin filaments and focal adhesions was shown due to local mechanical stimulation at the apical cell surface that induced changes into the cellular structure throughout the cell body. These innovative techniques will provide new information for understanding live cell restructuring and dynamics in response to mechanical force. A detailed protocol and a representative data set that show live cell response to mechanical stimulation are presented.

This paper aims to instruct the reader in the operation of an integrated atomic force-optical imaging microscope for mechanical stimulation of live cells in culture. A step-by-step protocol is presented. A representative data set that shows live cell response to mechanical stimulation is presented.

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to ...

Biology

Functionalization of Atomic Force Microscope Cantilevers with Single-T Cells or Single-Particle for Immunological Single-Cell Force Spectroscopy

Atomic force microscopy based single cell force spectroscopy (AFM-SCFS) is a powerful tool for studying biophysical properties of living cells. This technique allows for probing interaction strengths and dynamics on a live cell membrane, including those between cells, receptor and ligands, and alongside many other variations. It also works as a mechanism to deliver a physical or biochemical stimulus on single cells in a spatiotemporally controlled manner, thus allowing specific cell activation and subsequent cellular events to be monitored in real-time when combined with live-cell fluorescence imaging. The key step in those AFM-SCFS measurements is AFM-cantilever functionalization, or in other words, attaching a subject of interest to the cantilever. Here, we present methods to modify AFM cantilevers with a single T cell and a single polystyrene bead respectively for immunological studies. The former involves a biocompatible glue that couples single T cells to the tip of a flat cantilever in a solution, while the latter relies on an epoxy glue for single bead adhesion in the air environment. Two immunological applications associated with each cantilever modification are provided as well. The methods described here can be easily adapted to different cell types and solid particles.

We present a protocol to functionalize atomic force microscope (AFM) cantilevers with a single T cell and bead particle for immunological studies. Procedures to probe single-pair T cell-dendritic cell binding by AFM and to monitor the real-time cellular response of macrophages to a single solid particle by AFM with fluorescence imaging are shown.

Atomic force microscopy based single cell force spectroscopy (AFM-SCFS) is a powerful tool for studying biophysical properties of living cells. This ...

Immunology and Infection

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Multicellular Spheroid Formation for FLIM-Based Metabolic and Hypoxia Analysis

Production and Multi-Parameter Live Cell Fluorescence Lifetime Imaging Microscopy (FLIM) of Multicellular Spheroids

Multicellular tumor spheroids are a popular 3D tissue microaggregate&#160;model for reproducing tumor microenvironment, testing and optimizing drug therapies&#160;and using bio- and nanosensors in a 3D context. Their ease of production, predictable size, growth, and observed nutrient and metabolite gradients are important to recapitulate the 3D niche-like&#160;cell microenvironment. However, spheroid heterogeneity and variability of their production methods can influence overall cell metabolism, viability, and drug response. This makes it difficult to choose the most appropriate&#160;methodology, considering the requirements in size, variability, needs of biofabrication, and use as in vitro 3D tissue models in stem and cancer cell biology. In particular, spheroid production can influence their compatibility with quantitative live microscopies, such as optical metabolic imaging, fluorescence lifetime imaging microscopy (FLIM), monitoring of spheroid&#160;hypoxia with nanosensors, or viability. Here, a number of conventional spheroid formation protocols are presented, highlighting their compatibility with the live widefield, confocal, and two-photon microscopies. The follow-up imaging to analysis&#160;pipeline with multiplexed autofluorescence FLIM and, using various types of cancer and stem cell spheroids, is also presented.

Author Spotlight: Standardizing Spheroid Formation Methods for Metabolic and Oxygenation Analysis Using Fluorescence Lifetime Imaging Microscopy

Here, we describe different multicellular spheroid formation methods to perform follow-up multi-parameter live cell microscopy. Using fluorescence lifetime imaging microscopy (FLIM), cellular autofluorescence, staining dyes, and nanoparticles, the approach for analysis of cell metabolism, hypoxia, and cell death in live three-dimensional (3D) cancer and stem cell-derived spheroids is demonstrated.

Multicellular tumor spheroids are a popular 3D tissue microaggregate&#160;model for reproducing tumor microenvironment, testing and optimizing drug ...

Simultaneous Live Imaging of Multiple Insect Embryos in Sample Chamber-Based Light Sheet Fluorescence Microscopes

Light sheet-based fluorescence microscopy offers efficient solutions to study complex processes on multiple biologically relevant scales. Sample chamber-based setups, which are specifically designed to preserve the three-dimensional integrity of the specimen and usually feature sample rotation, are the best choice in developmental biology. For instance, they have been used to document the entire embryonic morphogenesis of the fruit fly Drosophila melanogaster and the red flour beetle Tribolium castaneum. However, many available live imaging protocols provide only experimental frameworks for single embryos. Especially for comparative studies, such approaches are inconvenient, since sequentially imaged specimens are affected by ambient variance. Further, this limits the number of specimens that can be assayed within a given time. We provide an experimental framework for simultaneous live imaging that increases the throughput in sample chamber-based setups and thus ensures&#160;similar ambient conditions for all specimens. Firstly, we provide a calibration guideline for light sheet fluorescence microscopes. Secondly, we propose a mounting method for multiple embryos that is compatible with sample rotation. Thirdly, we provide exemplary three-dimensional live imaging datasets of Drosophila, for which we juxtapose three transgenic lines with fluorescently labeled nuclei, as well as of Tribolium, for which we compare the performance of three transgenic sublines that carry the same transgene, but at different genomic locations. Our protocol is specifically designed for comparative studies as it pro-actively addresses ambient variance, which is always present in sequential live imaging. This is especially important for quantitative analyses and characterization of aberrational phenotypes, which result e.g., from knockout experiments. Further, it increases the overall throughput, which is highly convenient when access to light sheet&#160;fluorescence microscopes is limited. Finally, the proposed mounting method can be adapted for other insect species and further model organisms, e.g., zebrafish, with basically no optimization effort.

Light sheet-based fluorescence microscopy is the most valuable tool in developmental biology. A major issue in comparative studies is ambient variance. Our protocol describes an experimental framework for simultaneous live imaging of multiple specimens and, therefore, addresses this issue pro-actively.

Light sheet-based fluorescence microscopy offers efficient solutions to study complex processes on multiple biologically relevant scales. Sample ...

Live-cell Imaging of Single-Cell Arrays (LISCA) - a Versatile Technique to Quantify Cellular Kinetics

Live-cell Imaging of Single-Cell Arrays (LISCA) is a versatile method to collect time courses of fluorescence signals from individual cells in high throughput. In general, the acquisition of single-cell time courses from cultured cells is hampered by cell motility and diversity of cell shapes. Adhesive micro-arrays standardize single-cell conditions and facilitate image analysis. LISCA combines single-cell microarrays with scanning time-lapse microscopy and automated image processing. Here, we describe the experimental steps of taking single-cell fluorescence time courses in a LISCA format. We transfect cells adherent to a micropatterned array using mRNA encoding for enhanced green fluorescent protein (eGFP) and monitor the eGFP expression kinetics of hundreds of cells in parallel via scanning time-lapse microscopy. The image data stacks are automatically processed by newly developed software that integrates fluorescence intensity over selected cell contours to generate single-cell fluorescence time courses. We demonstrate that eGFP expression time courses after mRNA transfection are well described by a simple kinetic translation model that reveals expression and degradation rates of mRNA. Further applications of LISCA for event time correlations of multiple markers in the context of signaling apoptosis are discussed.

Live-cell Imaging of Single-Cell Arrays LISCA - a Versatile Technique to Quantify Cellular Kinetics

We present a method for the acquisition of fluorescence reporter time courses from single cells using micropatterned arrays. The protocol describes the preparation of single-cell arrays, the setup and operation of live-cell scanning time-lapse microscopy and an open-source image analysis tool for automated preselection, visual control and tracking of cell-integrated fluorescence time courses per adhesion site.

Live-cell Imaging of Single-Cell Arrays (LISCA) is a versatile method to collect time courses of fluorescence signals from individual cells in high ...

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy (Conpokal) on Live Cells

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