I am grateful to Dr. Masaru Fujimoto for his technical suggestions for VAEM. I am also grateful to Prof. Koh Iba and Dr. Mimi Hashimoto-Sugimoto for providing GFP-PATROL1 transgenic plants, and discussions about PATROL1. I thank Prof. Seiichiro Hasezawa for his continuing support of my work. This work was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI grant number 25711017.

1. Preparation of Seedlings
<ol>
	<li>Sterilize the seeds.
	<ol>
		<li>Prepare the sterilization solution by adding 500 &#181;l NaClO (available chlorine: 5.0%) and 1 &#181;l 10% Triton X-100 to 500 &#181;l sterile water.</li>
		<li>Place ca. 10 transgenic A. thaliana seeds carrying GFP-PATROL18 into a 1.5-ml tube.</li>
		<li>Add 1 ml 70% ethanol solution, and mix well by inverting five times. Leave for 1 min.</li>
		<li>Observe the seeds sink to the bottom of the tube. In a clean laminar flow cabinet, gently remove the 70% ethanol using a micropipette, and add 1 ml sterilization solution. Mix well by inverting five times and leave for 5 min.&#160;</li>
		<li>Wash the seeds. Still working under aseptic conditions on a clean bench, gently remove the solution using a micropipette, and add 1 ml sterile water. Repeat this five times. The sterilized seeds may be stored in sterile water at 4 &#176;C for 2 days.</li>
	</ol>
	</li>
	<li>Sow the sterilized seeds on a 0.5% gellan gum-solidified 1/2 Murashige and Skoog medium plate (pH 5.8)9. Tape the lid onto the plate using two layers of surgical tape.</li>
	<li>Incubate the plate in the dark at 4 &#176;C with a cold storage chamber, O/N.</li>
	<li>Transfer the plate into a growth chamber set at 23.5 &#176;C with a 12-hr/12-hr light-dark cycle using 100 &#181;mol m&#8722;2 sec&#8722;1 white lights, and incubate for 7 days. Seedlings with cotyledons of around 1 mm long can then be harvested.</li>
</ol>
2. Sky Drop Mounting of Cotyledon Specimens
NOTE: An important factor in specimen preparation for VAEM observation is avoiding the inclusion of air bubbles between the specimen and the cover glass. Bubbles greatly reduce the image quality of VAEM by causing differences in the refractive index. A simple method, which we have called &#8216;sky drop&#8217; mounting, can be used to avoid bubbles between the A. thaliana cotyledons and the cover glass. This should be done immediately prior to observation.
<ol>
	<li>Place 30 &#181;l of basal buffer [5 mM 2-(N-morpholino)-ethanesulfonic acid-Tris (MES-Tris) at pH 6.5, 50 mM KCl, 100 &#181;M CaCl2] on the center of a glass slide (size: 76&#215;26 mm, thickness: 1.0&#8211;1.2 mm).</li>
	<li>Remove a cotyledon from a 7-day-old seedling using dissecting scissors. Float the cotyledon with the observation side facing up on the basal buffer drop (Figure 1, step 1).</li>
	<li>Place 30 &#181;l of basal buffer on the center of a cover glass (size: 18&#215;18 mm, thickness: 0.12&#8211;0.17 mm) (Figure 1, step 2). Turn the cover glass upside down gently. Surface tension will prevent the buffer drop from falling off. Hold the cover glass with tweezers on one edge. Place the opposite edge on the glass slide so that the buffer drop is approximately above the cotyledon.</li>
	<li>With the edge of the cover glass still on the slide, and still holding the opposite edge with tweezers, adjust the position of the buffer drop under the cover glass so that it is directly above the cotyledon sample (Figure 1, step 3). Let go of the cover glass. Mount a drop on the sample resulting in a preparation without air bubbles.</li>
	<li>Wipe off excess buffer using lint-free tissues. Observe the preparation immediately (see step 3). 
	NOTE: There is no need to seal the samples.</li>
</ol>
3. VAEM Observation and Movie Acquisition
NOTE : The TIRF microscope system9 used in the present study is described as follows: An inverted microscope is equipped with a TIRF unit and a TIRF objective lens with a numerical aperture of 1.49. For computerized control of the laser entry angle, a control box is used. Green fluorescent protein (GFP) is excited with a 488 nm optically pumped semiconductor laser, and the fluorescence is detected through a 510&#8211;550 nm band-pass filter to prevent autofluorescence from chloroplasts. The measured maximum value of fiber output power is 13.0&#8211;13.5 mW. For detection, an electron multiplying charge-coupled device (EM-CCD) camera head system and a C-mount camera magnification change unit are used.
<ol>
	<li>Calibrate the laser centering and focusing according to the manufacturer&#8217;s instructions. 
	NOTE: This step is crucial for precise VAEM observations. It is strongly recommended that periodic checks of the laser path adjustment procedures, appropriate to your microscope, are carried out.
	<ol>
		<li>Determine the central position illuminated with an objective lens-free light path on the ceiling of the microscopy room. To mark the central position, put a colored circle seal on the ceiling (Figure 2, step 1, 2).</li>
		<li>Illuminate the ceiling with an objective lens (Figure 2, step 3, 4). Move the illuminated region to the center position (Figure 2, step 5). Focus the laser (Figure 2, step 6). Fine-tune the position of the focused laser to the center (Figure 2, step 7).</li>
	</ol>
	</li>
	<li>Set a specimen on the stage of the microscope and select cells for observation using bright field illumination.</li>
	<li>Check that the fluorescent protein can be observed in the cells, and set the z-axis position at the cell surface using epifluorescence illumination (Figure 3A).</li>
	<li>Perform VAEM observations.
	<ol>
		<li>Incline the entry angle of the laser beam gradually with the controller box. At the same time, monitor the live image carefully. Initially, the image will be blurred (Figure 3B).</li>
		<li>As the laser angle increases, the VAEM image will become less blurred, eventually producing a clear image (Figure 3C and D). At this point, stop increasing the laser angle. If the fluorescence signal is lost, decrease to a shallower angle.</li>
		<li>Fine-tune the laser entry angle to obtain a better image. Fine-tuning of the z-axis position may also improve the image. If necessary, adjust the optical parameters (laser power, wavelength and filter set) and image sensor parameters [image size, exposure time, gain, digitizer and electron-multiplying (EM) gain]. 
		NOTE: In the case of the TIRF microscope equipment described above, representative parameters are as follows. Magnification of lens just in front of the camera: 2X; Laser output: 1.0 mW; Image size: 512 &#215; 512 pixels; Exposure time: 100 msec; Gain: 5&#215;; Digitizer: 11 MHz; and EM gain: 100.</li>
	</ol>
	</li>
	<li>&#160;Acquire a movie as a multi-page TIFF image file using the commercial microscope software. Here, the movie is of GFP-labeled dots blinking on the surface of stomatal cells. 
	NOTE: Representative image acquisition conditions are as follows. Acquisition Mode: Stream to RAM; Number of frames: 600; and multi-page TIFF file size: ~302 MB.</li>
</ol>
4. Kymograph Analysis for Quantification of GFP-labeled Dot Residence Time Using Fiji Software
<ol>
	<li>Install Fiji (&#8216;Fiji is Just ImageJ&#8217;) software10 using the authors&#8217; instructions (http://fiji.sc/Fiji).</li>
	<li>Run Fiji and open the acquired multi-page TIFF file using the Fiji menu &#8220;File-Open&#8221;.</li>
	<li>Place a line on the site of interest, using the Fiji tool bar menu &#8220;Straight Line Selection Tool&#8221;. Optionally, use the &#8220;Segmented Line Selection Tool&#8221; or &#8220;Freehand Line Selection Tool&#8221;. In the results presented here, a straight line of 100 pixels (corresponding to 8.0 &#181;m) was placed on a subsidiary cell (Figure 4A).</li>
	<li>Make a kymograph image using the Fiji menu &#8220;Image-Stacks-Dynamic Reslice&#8221; (http://fiji.sc/Dynamic_Reslice) (Figure 4B). Optionally, &#34;Flip Vertically&#8221; and &#8220;Rotate 90 Degrees&#8221; in a check box window are also available (not used in the results presented here). The x- and y-axis in the kymograph image indicate the line position (100 pixels corresponding to 8.0 &#181;m) and observation time (600 pixels corresponding to 60 sec), respectively. If the kymograph image is not satisfactory, the position and type (straight, segmented and freehand) of the line on the multi-page image may be changed. As the line changes, the kymograph image dynamically changes. Save a kymograph image as a TIFF file using the Fiji menu &#8220;File-Save As-Tiff&#8221;.</li>
	<li>Measure the length of the blob in the orientation of time axis.
	<ol>
		<li>To perform noise reduction, apply a Gaussian filter to the kymograph images using the Fiji menu &#8220;Process-Filters-Gaussian Blur&#8221;. The &#8220;Sigma (Radius)&#8221; parameter is tunable (1 pixel radius is used for the presented results).</li>
		<li>Segment the signal regions by thresholding, using the Fiji menu &#8220;Image-Adjust-Threshold&#8221;. 
		NOTE: There are several thresholding algorithms to choose from. In presented results, the &#8220;Yen&#8221; algorithm was chosen (Figure 4C).</li>
		<li>Prepare to measure the time (y)-axis length of the blob using the menu &#8220;Analyze-Set Measurements&#8221;. Check the &#8220;Bounding rectangle&#8221; in the &#8220;Set Measurements&#8221; window. Ideally the area set should be as small as possible, to exclude noise. Here, the minimum area was set at 50 pixels. Measure the time (y)-axis length of the blob using the menu &#8220;Analyze-Analyze Particles&#8221;. To obtain the result values and prove credibility of the measurement regions, check the &#8220;Display results&#8221; and &#8220;Add to Manager&#8221; boxes.</li>
		<li>Values in the &#8220;Height&#8221; column are the time (y)-axis lengths for the measured blob (Figure 4D). Save the values using the Results table menu &#8220;File-Save As&#8221; and calculate and process the dataset of blob image durations using a statistical package and/or spreadsheet software (Figure 4E).</li>
	</ol>
	</li>
</ol>

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The author has nothing to disclose.

In this video article, protocols are given for monitoring and measuring the dynamic behavior of GFP-PATROL1 dots on the stomatal complex of Arabidopsis thaliana. As shown here, VAEM observation is a powerful tool for live imaging of plant cell surfaces. Under the experimental conditions used here for GFP-PATROL1 monitoring, there was very little fluorescence photobleaching in the sample used for 1 min of video capture, because the highly sensitive EM-CCD permits the use of a relatively weak excitation laser in V...

The plant cell surface, including the plasma membrane and its immediately adjacent cytoplasm, is the main region of a plant cell&#8217;s perception and integration of biotic and abiotic cues from the extracellular environment. In response to these cues, cell surface components including plasma membrane proteins and the cortical cytoskeleton undergo dynamic changes, on a time scale of seconds to minutes1-4. Thus, real-time and high-resolution imaging of fluorescent proteins on the cell surface can illuminate a plant&#8217;s responses to environmental cues at the molecular level.
Confocal laser scanning microscopy is a powerful tool for determination of fluorescently-tagged protein localization3, however, it is often difficult to monitor the real-time protein dynamics because of its relatively long capturing times. An emerging technique for real-time monitoring of proteins in the plant cell is variable angle epifluorescence microscopy (VAEM), which is an adaptation of equipment usually used for total internal reflection fluorescence (TIRF) microscopy. In TIRF microscopy, the fluorescence-excitation light source is an evanescent light field that is generated when the entry angle of the laser is shallow enough to totally internally reflect light at the glass&#8211;water interface. The penetration depth of the evanescent light field is around 100 nm. TIRF microscopy is an outstanding tool for single molecule imaging, such as the detection of exocytosis in animal cells5. However, evanescent light cannot reach plasma membranes or the cortical cytoplasm in plant cells, because they have thick cell walls. Recently, TIRF microscopy equipment has been adapted by plant cell biologists, observing that a laser, if angled slightly more deeply than when being used to induce total internal reflection phenomena, could excite the surface of plant cell samples, resulting in high-quality plant cell imaging6,7. The excitation illumination depth is varied by adjusting the entry angle of the laser; therefore, this technique is described as VAEM. This optical system is also called variable angle TIRF microscopy (VA-TIRFM) because there is a possibility that total reflection may take place at the cell wall-periplasm interface7, however, the term VAEM is used in this article, as per the first report in plants6.
The goal of this protocol is to demonstrate practical procedures for using VAEM to visualize fluorescently-tagged protein dynamics on plant cell surfaces. Additionally, an image analysis protocol to quantify the residence time (duration of presence) of molecules is described for VAEM movie analysis. GFP-PATROL1 dot blinking on stomatal complex cells in Arabidopsis thaliana cotyledons is used as an example. PATROL1 was identified by forward genetic approaches as a causal gene of a stomatal response defect mutant in A. thaliana8. PATROL1 is a plant homolog of MUNC-13, which is a priming factor in synapse vesicle exocytosis8. In response to environmental cues, such as light or humidity, it is thought that PATROL1 reversibly regulates the delivery of a proton pump to plasma membranes in the stomatal complex. Stomatal complexes each comprise a pair of guard cells8 and subsidiary cells9, and they require a proton pump for stomatal movement. In these cells, GFP-tagged PATROL1 localizes to dot-like structures that remain on the plasma membrane for less than 1 min9.

<table border="0" cellpadding="0" cellspacing="0" width="573">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Inverted microscope</td>
			<td style="width:71px;">Olympus</td>
			<td style="width:119px;">IX-73</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">TIRF unit</td>
			<td style="width:71px;">Olympus</td>
			<td style="width:119px;">IX3-RFAEVAW</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">TIRF objective lens&#160;</td>
			<td style="width:71px;">Olympus</td>
			<td style="width:119px;">UAPON 100 &#215; OTIRF&#160;</td>
			<td>NA = 1.49</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Laser angle control box</td>
			<td style="width:71px;">Chuo Seiki</td>
			<td style="width:119px;">QT-AK</td>
			<td></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">Optically pumped semiconductor laser</td>
			<td style="width:71px;">Coherent</td>
			<td style="width:119px;">SapphireTM LP USB 488-20 CDRH Laser</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">510&#8211;550 nm band-pass filter</td>
			<td style="width:71px;">Olympus</td>
			<td style="width:119px;">U-FBNA</td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">EM CCD camera</td>
			<td style="width:71px;">Hamamatsu Photonics</td>
			<td style="width:119px;">ImagEM C9100-13</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">C-mount camera magnification change unit&#160;</td>
			<td style="width:71px;">Olympus</td>
			<td style="width:119px;">U-TVCAC</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">MetaMorph software</td>
			<td style="width:71px;">Molecular Devices</td>
			<td style="width:119px;">MetaMorph version 7.7.11.0</td>
			<td></td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:216px;">TIRF microscopy manual</td>
			<td style="width:71px;">Olympus</td>
			<td style="width:119px;">AX7385</td>
			<td style="width:167px;">Instructions: Total Internal Reflection Illumination System (Printed in Japan on August 24, 2012)</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Immersion oil</td>
			<td style="width:71px;">Olympus</td>
			<td style="width:119px;">Immersion Oil Typr-F</td>
			<td>ne = 1.518 (23 degrees)</td>
		</tr>
	</tbody>
</table>

real-time imaging of plant cell surface dynamics with variable-angle epifluorescence microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

In this video article, the protocols for VAEM observation of GFP-PATROL1 in A. thaliana cotyledon stomatal complex cells are provided. Sky drop mounting is a simple preparation method that may help reduce the occurrence of air bubbles in VAEM preparations of A. thaliana cotyledons (Figure 1). Overtilting of the entry laser and/or z-positioning of specimens for VAEM will provide an unclear image. If that happens, it is recommended to start again from a position immediately above the samp...

Watch this Scientific Journal Video about Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy at JoVE.com

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

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Biology

8.9K Views.  The University of Tokyo.  המטרה של פרוטוקול זה היא להדגים כיצד לפקח דינמיקת fluorescently- מתויג חלבון על משטחי תא צמח עם מיקרוסקופיה epifluorescence זווית משתנה, מראה מהבהב נקודות של PATROL1 מתויג GFP, חלבון סחר קרום, בקליפת התא של המתחם הפיוניות ב thaliana ארבידופסיס. 

Video: Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed centrifugation. When properly planned and executed, an AUC sedimentation velocity or sedimentation equilibrium experiment can reveal a great deal about a protein in regards to size and shape, sample purity, sedimentation coefficient, oligomerization states and protein-protein interactions.
This technique, however, requires a rigorous level of technical attention. Sample cells hold a sectored center piece sandwiched between two window assemblies. They are sealed with a torque pressure of around 120-140 in/lbs. Reference buffer and sample are loaded into the centerpiece sectors and then after sealing, the cells are precisely aligned into a titanium rotor so that the optical detection systems scan both sample and reference buffer in the same radial path midline through each centerpiece sector while rotating at speeds of up to 60, 000 rpm and under very high vacuum
Not only is proper sample cell assembly critical, sample cell components are very expensive and must be properly cared for to ensure they are in optimum working condition in order to avoid leaks and breakage during experiments. Handle windows carefully, for even the slightest crack or scratch can lead to breakage in the centrifuge. The contact between centerpiece and windows must be as tight as possible; i.e. no Newton s rings should be visible after torque pressure is applied. Dust, lint, scratches and oils on either the windows or the centerpiece all compromise this contact and can very easily lead to leaking of solutions from one sector to another or leaking out of the centerpiece all together. Not only are precious samples lost, leaking of solutions during an experiment will cause an imbalance of pressure in the cell that often leads to broken windows and centerpieces. In addition, plug gaskets and housing plugs must be securely in place to avoid solutions being pulled out of the centerpiece sector through the loading holes by the high vacuum in the centrifuge chamber. Window liners and gaskets must be free of breaks and cracks that could cause movement resulting in broken windows.
This video will demonstrate our procedures of sample cell assembly, torque, loading and rotor alignment to help minimize component damage, solution leaking and breakage during the perfect AUC experiment.

The analytical ultracentrifuge (AUC) sample cell holds sample and reference buffer and during experiments and is exposed to high vacuum and rotor speeds up to 60,000 rpm. This video will demonstrate the rigorous attention to detail necessary for assembly, loading and alignment of this very important component of an AUC experiment.

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed ...

Avidity-based Extracellular Interaction Screening (AVEXIS) for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ensuring cohesion within multicellular organisms. Proteins predicted to form extracellular interactions are encoded by approximately a quarter of human genes1, but despite their importance and abundance, the majority of these proteins have no documented binding partner. Primarily, this is due to their biochemical intractability: membrane-embedded proteins are difficult to solubilise in their native conformation and contain structurally-important posttranslational modifications. Also, the interaction affinities between receptor proteins are often characterised by extremely low interaction strengths (half-lives &lt; 1 second) precluding their detection with many commonly-used high throughput methods2. 
Here, we describe an assay, AVEXIS (AVidity-based EXtracellular Interaction Screen) that overcomes these technical challenges enabling the detection of very weak protein interactions (t1/2 &le; 0.1 sec) with a low false positive rate3. The assay is usually implemented in a high throughput format to enable the systematic screening of many thousands of interactions in a convenient microtitre plate format (Fig. 1). It relies on the production of soluble recombinant protein libraries that contain the ectodomain fragments of cell surface receptors or secreted proteins within which to screen for interactions; therefore, this approach is suitable for type I, type II, GPI-linked cell surface receptors and secreted proteins but not for multipass membrane proteins such as ion channels or transporters.
The recombinant protein libraries are produced using a convenient and high-level mammalian expression system4, to ensure that important posttranslational modifications such as glycosylation and disulphide bonds are added. Expressed recombinant proteins are secreted into the medium and produced in two forms: a biotinylated bait which can be captured on a streptavidin-coated solid phase suitable for screening, and a pentamerised enzyme-tagged (&beta;-lactamase) prey. The bait and prey proteins are presented to each other in a binary fashion to detect direct interactions between them, similar to a conventional ELISA (Fig. 1). The pentamerisation of the proteins in the prey is achieved through a peptide sequence from the cartilage oligomeric matrix protein (COMP) and increases the local concentration of the ectodomains thereby providing significant avidity gains to enable even very transient interactions to be detected. By normalising the activities of both the bait and prey to predetermined levels prior to screening, we have shown that interactions having monomeric half-lives of 0.1 sec can be detected with low false positive rates3.

Avidity-based Extracellular Interaction Screening AVEXIS for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

AVEXIS is a high throughput protein interaction assay developed to systematically screen for novel extracellular receptor-ligand pairs involved in cellular recognition processes. It is specifically designed to detect transient protein interactions that are difficult to identify using other high throughput approaches.

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

Real Time Measurements of Membrane Protein:Receptor Interactions Using Surface Plasmon Resonance (SPR)

Protein-protein interactions are pivotal to most, if not all, physiological processes, and understanding the nature of such interactions is a central step in biological research. Surface Plasmon Resonance (SPR) is a sensitive detection technique for label-free study of bio-molecular interactions in real time. In a typical SPR experiment, one component (usually a protein, termed 'ligand') is immobilized onto a sensor chip surface, while the other (the 'analyte') is free in solution and is injected over the surface. Association and dissociation of the analyte from the ligand are measured and plotted in real time on a graph called a sensogram, from which pre-equilibrium and equilibrium data is derived. Being label-free, consuming low amounts of material, and providing pre-equilibrium kinetic data, often makes SPR the method of choice when studying dynamics of protein interactions. However, one has to keep in mind that due to the method's high sensitivity, the data obtained needs to be carefully analyzed, and supported by other biochemical methods. 
SPR is particularly suitable for studying membrane proteins since it consumes small amounts of purified material, and is compatible with lipids and detergents. This protocol describes an SPR experiment characterizing the kinetic properties of the interaction between a membrane protein (an ABC transporter) and a soluble protein (the transporter's cognate substrate binding protein).

Real Time Measurements of Membrane Protein:Receptor Interactions Using Surface Plasmon Resonance SPR

Surface Plasmon Resonance (SPR) is a label-free method for detecting bio-molecular interactions in real time. Herein, a protocol for a membrane protein:receptor interaction experiment is described, while discussing the pros and cons of the technique.

Protein-protein interactions are pivotal to most, if not all, physiological processes, and  understanding the nature of such interactions is a central ...

Real Time Analysis of Metabolic Profile in Ex Vivo Mouse Intestinal Crypt Organoid Cultures

The small intestinal mucosa exhibits a repetitive architecture organized into two fundamental structures: villi, projecting into the intestinal lumen and composed of mature enterocytes, goblet cells and enteroendocrine cells; and crypts, residing proximal to the submucosa and the muscularis, harboring adult stem and progenitor cells and mature Paneth cells, as well as stromal and immune cells of the crypt microenvironment. Until the last few years, in vitro studies of small intestine was limited to cell lines derived from either benign or malignant tumors, and did not represent the physiology of normal intestinal epithelia and the influence of the microenvironment in which they reside. Here, we demonstrate a method adapted from Sato et al. (2009) for culturing primary mouse intestinal crypt organoids derived from C57BL/6 mice. In addition, we present the use of crypt organoid cultures to assay the crypt metabolic profile in real time by measurement of basal oxygen consumption, glycolytic rate, ATP production and respiratory capacity. Organoids maintain properties defined by their source and retain aspects of their metabolic adaptation reflected by oxygen consumption and extracellular acidification rates. Real time metabolic studies in this crypt organoid culture system are a powerful tool to study crypt organoid energy metabolism, and how it can be modulated by nutritional and pharmacological factors.

Real Time Analysis of Metabolic Profile in Ex Vivo Mouse Intestinal Crypt Organoid Cultures

Small intestinal crypt organoids cultured ex vivo provide a tissue culture system that recapitulates growth of crypts dependent on stem cells and their niche. We established a method to assay the metabolic profile in real time in primary mouse crypt organoids. We found organoids maintain physiological properties defined by their source.

The small intestinal mucosa exhibits a repetitive architecture organized into two fundamental structures: villi, projecting into the intestinal lumen and ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the observation of the whole organ with a high spatial- as well as temporal resolution over prolonged periods of time, which may cause photo-toxic effects. This protocol shows a plant sample preparation method for light-sheet microscopy, which is characterized by mounting the plant vertically on the surface of a gel. The plant is mounted in such a way that the roots are submerged in a liquid medium while the leaves remain in the air. In order to ensure photosynthetic activity of the plant, a custom-made lighting system illuminates the leaves. To keep the roots in darkness the water surface is covered with sheets of black plastic foil. This method allows long-term imaging of plant organ development in standardized conditions.

This protocol shows a plant sample preparation method for light-sheet microscopy. The setup is characterized by mounting the plant vertically on the surface of a gel and letting it grow in controlled bright conditions. This allows long-term observation of plant organ development in standardized conditions.

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the ...

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

Confocal Microscopy Reveals Cell Surface Receptor Aggregation Through Image Correlation Spectroscopy

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of pre-clinical agents labeled with fluorescent probes. With recent advancements in antibody based cytotoxic drug delivery systems, understanding the alterations induced by these agents within the realm of receptor aggregation and internalization is of critical importance. This protocol leverages the well-established methodology of fluorescent immunocytochemistry and the open source FIJI distribution of ImageJ, with its inbuilt autocorrelation and image mathematical functions, to perform spatial image correlation spectroscopy (ICS). This protocol quantitates the fluorescent intensity of labeled receptors as a function of the beam area of the confocal microscope. This provides a quantitative measure of the state of target molecule aggregation on the cell surface. This methodology is focused on the characterization of static cells with potential to expand into temporal investigations of receptor aggregation. This protocol presents an accessible methodology to provide quantification of clustering events occurring at the cell surface, utilizing well established techniques and non-specialized imaging apparatus.

Antibodies that bind to target receptors on the cell surface can confer conformation and clustering alterations. These dynamic changes have implications for characterizing drug development in target cells. This protocol utilizes confocal microscopy and image correlation spectroscopy through ImageJ/FIJI to quantify the extent of receptor clustering on the cell surface.

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of ...