We thank members of the Alto laboratory for helpful discussions. We also thank The Molecular and Cellular Imaging Facility at UT Southwestern Medical Center, specifically Dr. Chris Gilpin, Tom Januszewski, and Laurie Mueller for technical expertise and advice. We also thank Dr. Xionan Dong for critical reading of the manuscript. This work is supported by NIH grant AI083359 to N.M.A.

1. Mammalian Cell Culture
<ol>
 <li>
 Normal Rat Kidney (NRK), immortalized cervical cancer (HeLa), or other appropriate mammalian cell lines are cultured at 37&deg;C, 5% CO2, on photoetched glass gridded coverslips affixed to live cell dishes (see Table 1 for any reagent or apparatus used in this protocol) in DMEM + 10% FBS and 1% Pen/Strep. Precautions should be taken to maintain a sterile environment when working with cultured mammalian cells.
 </li>
 <li>
 Depending on your experimental time-line (0-5 hours vs 1-2 days post injection) the confluency of cells should be adjusted to ~50% or ~25%, respectively. 
 </li>
</ol>
2. Microinjection
<ol>
 <li>
 Prepare desired protein, DNA, or small molecules in your choice of buffer, depending on its amenability to cellular culture, to a final volume of ~50 &mu;l. Inert fluorescent dyes such as Texas Red or Cascade Blue are included in order to mark the microinjected cells; choice of injection dye and fluorescent probe should have non-overlapping excitation and emission wavelengths. The concentration of your molecule of interest needed for microinjection should be determined empirically, and the fluorescent tracking dyes are commonly used at 0.5-1.0 mg/ml. Typically, rhodamine or another fluorophore is covalently attached to your molecule of interest using a commercial kit, however other methods of detection may be used depending on sensitivity. For more information, see Critical Considerations below.
 </li>
 <li>Filter the injectant (~50 &mu;l) through a 0.22 &mu;m centrifugal filter. Centrifugal filters are preferred over vacuum and/or syringe filtration which have associated dead volumes. </li>
 <li>To generate the microinjection needle, borosilicate glass pipettes are pulled using a Flaming-Brown Micropipette Puller following manufacturers guidelines. Alternatively, microinjection needles can be purchased from vendors including Eppendorf which makes femtotip microinjection needles. </li>
 <li> Carefully pipette 4 &mu;l of injectant into the unmolested end of the microinjection needle using Eppendorf 20 Femtotips. Ensure that no bubbles are present within the pulled microinjection needle. Load microinjection needle into the micromanipulator arm of the Eppendorf FemptoJet Injectman attached to an inverted microscope. For a good review of the microinjection procedure, the reader is directed to a previous Journal of Visualized Experiments article 8. </li>
 <li>Place live cell dish with gridded coverslip and cultured cells into the dish holder of the inverted Zeiss microscope. Find cells growing within a photoetched grid and note the grids alphanumeric identifier, this will be used in all subsequent steps. Ideally, locate cells at the center of the coverslip as subsequent steps will limit sample transfer at the edges of the coverslip. </li>
 <li>Manually lower injection arm into place over the coverslip, and using the micromanipulator joystick and controls, lower injection needle down until the tip gently touches the surface of the cultured cells and press &#x2018;Limit to prevent movement of needle beyond this point and thus breaking the fragile pulled borosilicate glass. All other aspects of cell microinjection are performed as per Eppendorf FemtoJet InjectMan user manual. The microinjection technique was recently used to understand individual secreted virulence proteins from bacteria 9. </li>
 <li>Microinject every other cell within the chosen grid. With cells grown at ~50% confluency, one can expect ~80 cells per 600 &mu;m2 grid (example shown in Fig. 3A). Therefore, upon fluorescence and electron microscopy inspection, there will be an n of ~30-40 experimental and control cells. </li>
 <li>Place live-cell dish back into mammalian cell culture incubator for desired time as dictated by the experimental setup. Alternatively, one can proceed directly from microinjection to time-lapse microscopy analysis to await the appearance of particular subcellular structures of interest prior to fixation. </li>
</ol>
3. Fluorescent Microscopy
<ol>
<li>If desired, perform live-cell time-lapse fluorescence measurements of microinjected cells, after which, replace the media with 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) and incubate for 10 min at room temperature. However, if live-cell measurements are not required, discard cell growth media and fix the cells with 5 ml 3.7% formaldehyde in 1xPBS at room temperature for 10 minutes. Discard fixative and replace with 5 ml 1xPBS, proceed to step 3.2. Note: it is critical not to add glutaraldehyde until after fluorescence microscopy is complete.</li>
<li>Place the live-cell dish with photoetched gridded coverslip in a fluorescent microscope (confocal is not required for this step) and collect 5-10 bright field and fluorescent images of the appropriate grid to document the arrangement of the cells (Fig. 4A). Identify those cells that have been microinjected using the excitation and emission wavelengths of the inert injection dye (Fig. 4B). It is important to obtain images at different magnifications (e.g. 10x and 65x) in order to facilitate identification of microinjected cells in subsequent steps involving electron microscopy. </li>
<li> Using a confocal fluorescent microscope, collect a Z-stack of microinjected cells within the grid identified in step 2.5 above. High magnification (100x) is needed in order to correlate fluorescent images with subcellular structures that will be identified in electron microscopy. </li>
<li>The coverslips are removed from the bottom of the live cell dishes with commercially available coverslip removal fluid and placed cell-side up in a virgin cell culture dish containing 5 ml 1xPBS. The coverslip is then processed for electron microscopy as detailed below. </li>
</ol>
4. Transmission Electron Microscopy
<ol>
<li>Replace 1xPBS buffer with 2 ml 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) and incubate at room temperature for 10 minutes. Longer incubation, while not necessary, will aid in maintenance of subcellular structures. Note: formaldehyde alone is not a sufficient fixative for EM analysis.</li>
<li>Remove fixative and stain cells with the addition of 2 ml 1% osmium tetroxide in 0.1 M cacodylate buffer for 30 min followed by three washes with cacodylate buffer and finally replace all buffer with ddH2O. </li>
<li>Following standard EM protocols, dehydrate the sample in a graded (70% to 100%) series of ethanol. Prepare Epon resin and place coverslip cell-side down on top of an Epon resin-filled tube. Allow polymerization for 24 hr at 60&deg;C. By placing the photoetched gridded coverslip cell-side down onto the Epon resin, both the cells of interest as well as the grid pattern is transferred to the Epon resin during polymerization (example shown in Fig. 3B). </li>
<li>Remove coverslips from Epon resin by quickly immersing into liquid nitrogen and/or boiling water. Inspect the surface of the resin under a binocular dissection scope to locate the grid of interest (note: the grid patter will transfer as a mirror image). Using a sterile razor blade or scalpel, trim the Epon block down such that only the grid of interest is at the apex of the Epon resin block.</li>
<li>Using an ultramicrotome, acquire serial sections of the trimmed block at the desired thickness (~70 nm thickness for conventional TEM and ~250 nm thickness for tomographic TEM). Contrast the sections with uranyl acetate and lead citrate following standard protocols. </li>
</ol>
5. Correlative Light and Electron Microscopy
<ol>
 <li>Relocate the cells of interest in the electron microscope using images obtained in earlier steps. Once microinjected cells of interest have been located, use the high-resolution confocal fluorescent images to identify regions within the cell that were of interest. Image these at high resolution using TEM. Use landmarks such as the cell periphery and the nuclear envelope to determine which fluorescent z-stack image corresponds to the EM slice.
 </li>
 <li>
 Using the imaging program of your choice (e.g. Image J, Adobe Photoshop, etc.), reduce the opacity of the fluorescent image to 50% and overlay it onto the TEM image with which it correlates. Align the images by adjusting the scale (remember to constrain proportions) and the rotation of the confocal image to match the TEM image. Fine adjustments to contrast, sharpness, etc. in the fluorescent image may aid in the alignment. Note: we have found that heterochromatin, the nuclear envelope, and the cell periphery represent an excellent landmarks to aid in the orientation of the two images.
 </li>
</ol>





6. Critical Considerations
<ol>
 <li>To greatly aid in locating the injected small molecule, a fluorescent tag such as Rhodamine or Fluorescein can be covalently attached using commercially available kits. Additional information is gained by ectopically expressing a fluorescent subcellular localization marker protein prior to microinjection. Hence, choice of fluorophores and inert microinjection tracking dye is important.</li>
 <li>Choose cells for microinjection that are adhering close to the center of the coverslip and avoid choosing a grid close to the edge. This will present the greatest probability that the cells of interest are transferred to the Epon resin used for electron microscopy. </li>
 <li>Confluency of cells is important, as a fully confluent plate will make identification of microinjected cells in the electron microscope very difficult. Rather, seed cells such that they reach ~50% confluency on the day of a short experiment, or ~25% confluency on the day of a longer (1-2 day) experiment. Use the position and shape of the cells as well as the &#x2018;empty space as landmarks to orient the electron microscopy image. </li>
 <li>Confocal optical slice thickness should be tuned to the needs of the experiment and should therefore be empirically determined 10. </li>
 <li>The fixatives formaldehyde and glutaraldehyde should be of the highest grade available. Formaldehyde will fix the cells, yet preserve the fluorescence properties of both proteins and small fluorescent molecules (such as Rhodamine or FITC). However, it will not preserve the structures to permit high EM resolution. Glutaraldehyde preserves membranes and subcellular structures to a high degree for EM analysis, but will significantly hinder fluorescence. Therefore, the two-step fixation is critical for both fluorescence and EM microscopy. The formaldehyde fixation step can be omitted if performing live-cell measurements, as final fixation can be performed by glutaraldehyde treatment.</li>
</ol>
7. Representative Results
This procedure affords the ability to deliver a molecule of interest directly into the eukaryotic cellular cytoplasm and monitor its localization and/or effects on cellular and organeller architecture from millimeter to multi-nanometer resolution. A schematic illustration of the experimental setup and procedure is shown in Figure 2. This technique relies on microinjection of cells grown on a laser etched glass gridded coverslip (Fig. 3A), which after confocal microscopy, is able to transfer both the cells of interest as well as the grid pattern to the Epon resin (Fig. 3B).
A typical microinjection CLEM procedure yields fluorescence images and electron micrographs, that when correlated, permit both subcellular localization and ultrastructural analysis. Cells grown on a photoetched glass gridded coverslip are microinjected with a molecule of interest after which bright field images are obtained (Fig. 4A). An inert tracking dye is incorporated to distinguish microinjected cells from non-microinjected cells (Fig. 4B). Confocal z-stacks are obtained using a fluorophore or other identifier attached to the microinjected small molecule of interest (Fig. 4C). The sample is fixed with glutaraldehyde and processed for electron microscopy. Electron micrographs are obtained and overlaid onto the cells to which they correspond (Fig. 4D). Areas harboring fluorescent signals are inspected in greater detail at higher magnification (Fig. 4E). 
<img src="/files/ftp_upload/3650/3650fig1.jpg" alt="Figure 1"/> 
Figure 1. Timeline of microinjection CLEM procedure. Depending on the experimental setup, microinjection and fluorescence measurements can be performed on the same day. Preparing the sample for electron microscopy is most time consuming due to long incubations steps and will last 2-3 days. TEM analysis and correlating fluorescence signals with electron microscopy images can be performed in one day.
 <img src="/files/ftp_upload/3650/3650fig2.jpg" alt="Figure 2"/> 
 Figure 2. General illustration of the microinjection CLEM procedure. Step 1: Locate cells growing on laser etched glass gridded coverslip and microinject molecule of interest. Step 2: Capture both bright field images and confocal fluorescent z-stacks of microinjected cells. Step 3: Prepare coverslip for EM analysis and embed in Epon resin. Using transferred grid pattern, trim block such that cells of interest are at the apex. Step 4: Cut serial sections with ultramicrotome. Step 5: Correlate fluorescent images with EM images.
<img src="/files/ftp_upload/3650/3650fig3.jpg" alt="Figure 3"/> 
Figure 3. Grid pattern facilitates identification of cells of interest. Panel A depicts the microinjection of cells grown to ~50% confluency; note the grid identifier aK. Scale bar is 100 &mu;m. Panel B shows the transferred grid pattern from the coverslip onto the polymerized Epon resin block that will be sectioned serially after the grid of interest is identified in a dissection microscope. Panel C shows the transferred grid pattern in preparation for sectioning. Note that the transferred grid patter is in reverse on the resin block; scale bar is 1200 &mu;m. 
<img src="/files/ftp_upload/3650/3650fig4.jpg" alt="Figure 4"/> 
Figure 4. Representative results from microinjection correlative light and electron microscopy. Panel A shows the bright field micrograph of NRK cells growing on a laser etched glass gridded coverslip; arrows indicate two cells that are visualized by CLEM in panels C-D. Panel B shows the fluorescence of the inert tracking dye, in this case Cascade Blue, used to identify cells that were microinjected. Panel C shows the overlay of bright field and the fluorescence signature of a Rhodamine-labeled small molecule. The two cells shown here are indicated by arrows in panels A &amp; B. Panel D represents CLEM of bright field, fluorescent and electron microscopy; arrow head indicates fluorescent puncta imaged at higher magnification in Panel E. Scale bars are as follows: Panels A &amp; B, 100 &mu;m; C &amp; D, 50 &mu;m; E, 100 nm.

<ol>
	<li>Schmolze, D. B., Standley, C., Fogarty, K. E., Fischer, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+microscopy+techniques.">Advances in microscopy techniques.</a> Arch. Pathol. Lab Med. 135, 255-263 (2011).</li><li>Giepmans, B. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bridging+fluorescence+microscopy+and+electron+microscopy.">Bridging fluorescence microscopy and electron microscopy.</a> Histochem. Cell Biol. 130, 211-217 (2008).</li><li>Mayhew, T. M., Muhlfeld, C., Vanhecke, D., Ochs, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+review+of+recent+methods+for+efficiently+quantifying+immunogold+and+other+nanoparticles+using+TEM+sections+through+cells,+tissues+and+organs.">A review of recent methods for efficiently quantifying immunogold and other nanoparticles using TEM sections through cells, tissues and organs.</a> Ann. Anat. 191, 153-170 (2009).</li><li>Razi, M., Tooze, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+light+and+electron+microscopy.">Correlative light and electron microscopy.</a> Methods Enzymol. 452, 261-275 (2009).</li><li>Svitkina, T. M., Borisy, G. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+light+and+electron+microscopy+of+the+cytoskeleton+of+cultured+cells.">Correlative light and electron microscopy of the cytoskeleton of cultured cells.</a> Methods Enzymol. 298, 570-592 (1998).</li><li>van Weering, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+membrane+traffic+at+high+resolution.">Intracellular membrane traffic at high resolution.</a> Methods Cell Biol. 96, 619-648 (2010).</li><li>Polishchuk, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+light-electron+microscopy+reveals+the+tubular-saccular+ultrastructure+of+carriers+operating+between+Golgi+apparatus+and+plasma+membrane.">Correlative light-electron microscopy reveals the tubular-saccular ultrastructure of carriers operating between Golgi apparatus and plasma membrane.</a> J. Cell Biol. 148, 45-58 (2000).</li><li>Lim, J., Danuser, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+Cell+Imaging+of+F-actin+Dynamics+via+Fluorescent+Speckle+Microscopy+(FSM).">Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM).</a> J. Vis. Exp. (30), e1325 (2009).</li><li>Selyunin, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+assembly+of+a+GTPase-kinase+signalling+complex+by+a+bacterial+catalytic+scaffold.">The assembly of a GTPase-kinase signalling complex by a bacterial catalytic scaffold.</a> Nature. 469, 107-111 (2011).</li><li>Semwogerere, D., Weeks, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Confocal Microscopy in Encyclopedia of Biomaterials and Biomedical Engineering. , (2005).</li></ol>

No conflicts of interest are declared.

The method presented here enables the direct delivery of purified proteins, nucleic acids, or small molecules to the eukaryotic cytoplasm and affords ultra high-resolution analysis through the correlation of fluorescent and electron microscopy. The use of this method is simple yet robust and can be performed in-house with many existing core facilities and/or appropriately equipped electron microscopy labs. The technique is also amenable to live-cell, allowing the user to monitor the dynamics of fluorescently tagged molec...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Photoetched Coverslip and Live-cell Plate</td>
 <td>MatTek</td>
 <td>P35G-2-14-CGRD</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Texas Red</td>
 <td>Invitrogen</td>
 <td>D3329</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Cascade Blue</td>
 <td>Invitrogen</td>
 <td>D7132</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Rhodamine Labeling Kit</td>
 <td>Thermo Scientific</td>
 <td>53002</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>0.22 &mu;m centrifugal filtration unit</td>
 <td>Millipore</td>
 <td>UFC30GV00</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Borosilicate Glass Pipettes</td>
 <td>Sutter Instruments</td>
 <td>BF100-50-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micropipette Puller</td>
 <td>Sutter Instruments</td>
 <td>Model P-97</td>
 <td>Use program #4 after performing ramp test</td>
 </tr>
 <tr>
 <td>FemtoJet Microinjection System</td>
 <td>Eppendorf</td>
 <td>Injectman NI2</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Coverslip Removal Fluid</td>
 <td>MatTek</td>
 <td>PDCF OS 30</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Technai Transmission Electron Microscope</td>
 <td>FEI</td>
 <td>Technai G2 BioTWIN</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Ultramicrotombe</td>
 <td>Leica</td>
 <td>EM UC6</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

correlative light and electron microscopy (clem) as a tool to visualize microinjected molecules and their eukaryotic sub-cellular targets

The eukaryotic cell relies on complex, highly regulated, and functionally distinct membrane bound compartments that preserve a biochemical polarity necessary for proper cellular function. Understanding how the enzymes, proteins, and cytoskeletal components govern and maintain this biochemical segregation is therefore of paramount importance. The use of fluorescently tagged molecules to localize to and/or perturb subcellular compartments has yielded a wealth of knowledge and advanced our understanding of cellular regulation. Imaging techniques such as fluorescent and confocal microscopy make ascertaining the position of a fluorescently tagged small molecule relatively straightforward, however the resolution of very small structures is limited 1.
On the other hand, electron microscopy has revealed details of subcellular morphology at very high resolution, but its static nature makes it difficult to measure highly dynamic processes with precision 2,3. Thus, the combination of light microscopy with electron microscopy of the same sample, termed Correlative Light and Electron Microscopy (CLEM) 4,5, affords the dual advantages of ultrafast fluorescent imaging with the high-resolution of electron microscopy 6. This powerful technique has been implemented to study many aspects of cell biology 5,7. Since its inception, this procedure has increased our ability to distinguish subcellular architectures and morphologies at high resolution. 
Here, we present a streamlined method for performing rapid microinjection followed by CLEM (Fig. 1). The microinjection CLEM procedure can be used to introduce specific quantities of small molecules and/or proteins directly into the eukaryotic cell cytoplasm and study the effects from millimeter to multi-nanometer resolution (Fig. 2). The technique is based on microinjecting cells grown on laser etched glass gridded coverslips affixed to the bottom of live cell dishes and imaging with both confocal fluorescent and electron microscopy. Localization of the cell(s) of interest is facilitated by the grid pattern, which is easily transferred, along with the cells of interest, to the Epon resin used for immobilization of samples and sectioning prior to electron microscopy analysis (Fig. 3). Overlay of fluorescent and EM images allows the user to determine the subcellular localization as well as any morphological and/or ultrastructural changes induced by the microinjected molecule of interest (Fig. 4). This technique is amenable to time points ranging from &le;5 s up to several hours, depending on the nature of the microinjected sample.

Watch this Scientific Journal Video about Correlative Light and Electron Microscopy CLEM as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets at JoVE.com

Correlative Light and Electron Microscopy CLEM as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets

The CLEM technique has been adapted to analyze ultrastructural morphology of membranes, organelles, and subcellular structures affected by microinjected molecules. This method combines the powerful techniques of micromanipulation/microinjection, confocal fluorescent microscopy, and electron microscopy to allow millimeter to multi-nanometer resolution. This technique is amenable to a wide variety of applications.

Correlative Light and Electron Microscopy (CLEM) as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets


The eukaryotic cell relies on complex, highly regulated, and functionally distinct membrane bound compartments that preserve a biochemical polarity ...

correlative-light-electron-microscopy-clem-as-tool-to-visualize

Research

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Biology

19.1K 次观看.  University of Texas Southwestern Medical Center. CLEM技术已被用于分析超微结构形态的细胞膜，细胞器，通过显微注射的分子和亚细胞结构的影响。这种方法结合了强大的技术，的显微/显微注射，共聚焦荧光显微镜，电子显微镜，允许毫米多纳米的分辨率。此技术是适合于各种各样的应用。

视频: Correlative Light and Electron Microscopy CLEM as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets

Electroantennographic Bioassay as a Screening Tool for Host Plant Volatiles

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host plant.1,2 When a host plant is an important agronomic commodity feeding damage by insect pests can inflict serious economic losses to growers. Accordingly, kairomones can be used as attractants to lure or confuse these insects and, thus, offer an environmentally friendly alternative to pesticides for insect control.3 Unfortunately, plants can emit a vast number volatiles with varying compositions and ratios of emissions dependent upon the phenology of the commodity or the time of day. This makes identification of biologically active components or blends of volatile components an arduous process. To help identify the bioactive components of host plant volatile emissions we employ the laboratory-based screening bioassay electroantennography (EAG). EAG is an effective tool to evaluate and record electrophysiologically the olfactory responses of an insect via their antennal receptors. The EAG screening process can help reduce the number of volatiles tested to identify promising bioactive components. However, EAG bioassays only provide information about activation of receptors. It does not provide information about the type of insect behavior the compound elicits; which could be as an attractant, repellent or other type of behavioral response. Volatiles eliciting a significant response by EAG, relative to an appropriate positive control, are typically taken on to further testing of behavioral responses of the insect pest. The experimental design presented will detail the methodology employed to screen almond-based host plant volatiles4,5 by measurement of the electrophysiological antennal responses of an adult insect pest navel orangeworm (Amyelois transitella) to single components and simple blends of components via EAG bioassay. The method utilizes two excised antennae placed across a "fork" electrode holder. The protocol demonstrated here presents a rapid, high-throughput standardized method for screening volatiles. Each volatile is at a set, constant amount as to standardize the stimulus level and thus allow antennal responses to be indicative of the relative chemoreceptivity. The negative control helps eliminate the electrophysiological response to both residual solvent and mechanical force of the puff. The positive control (in this instance acetophenone) is a single compound that has elicited a consistent response from male and female navel orangeworm (NOW) moth. An additional semiochemical standard that provides consistent response and is used for bioassay studies with the male NOW moth is (Z,Z)-11,13-hexdecadienal, an aldehyde component from the female-produced sex pheromone.6-8

A method to rapidly screen host plant volatiles by measurement of the electrophysiological response of adult navel orangeworm (Amyelois transitella) antennae to single components and blends via electroantennographic analysis is demonstrated.

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host ...

Using SecM Arrest Sequence as a Tool to Isolate Ribosome Bound Polypeptides

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway that polypeptide chain follows during co-translational folding to achieve its functional form is still an enigma. In order to understand this process and to determine the exact conformation of the co-translational folding intermediates, it is essential to develop techniques that allow the isolation of RNCs carrying nascent chains of predetermined sizes to allow their further structural analysis.
SecM (secretion monitor) is a 170 amino acid E. coli protein that regulates expression of the downstream SecA (secretion driving) ATPase in the secM-secA operon6. Nakatogawa and Ito originally found that a 17 amino acid long sequence (150-FSTPVWISQAQGIRAGP-166) in the C-terminal region of the SecM protein is sufficient and necessary to cause stalling of SecM elongation at Gly165, thereby producing peptidyl-glycyl-tRNA stably bound to the ribosomal P-site7-9. More importantly, it was found that this 17 amino acid long sequence can be fused to the C-terminus of virtually any full-length and/or truncated protein thus allowing the production of RNCs carrying nascent chains of predetermined sizes7. Thus, when fused or inserted into the target protein, SecM stalling sequence produces arrest of the polypeptide chain elongation and generates stable RNCs both in vivo in E. coli cells and in vitro in a cell-free system. Sucrose gradient centrifugation is further utilized to isolate RNCs.
The isolated RNCs can be used to analyze structural and functional features of the co-translational folding intermediates. Recently, this technique has been successfully used to gain insights into the structure of several ribosome bound nascent chains10,11. Here we describe the isolation of bovine Gamma-B Crystallin RNCs fused to SecM and generated in an in vitro translation system.

We describe here a technique that is now routinely used to isolate stably bound ribosome nascent chain complexes (RNCs). This technique takes advantage of the discovery that a 17 amino acid long SecM "arrest sequence" can halt translation elongation in a prokaryotic (E. coli) system, when inserted into (or fused to the C-terminus) of virtually any protein.

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Correlative Confocal and 3D Electron Microscopy of a Specific Sensory Cell

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused throughout tissues made of diverse cell types, such as enteroendocrine cells of the intestinal epithelium. These gastrointestinal sensors of food and bacteria have been difficult to study because they are dispersed among other epithelial cells at a ratio of 1:1,000. Recently, transgenic reporter mice have been generated to identify enteroendocrine cells by means of fluorescence. One of those is the peptide YY-GFP mouse. Using this mouse, we developed a method to correlate confocal and serial block-face scanning electron microscopy. We named the method cocem3D and applied it to identify a specific enteroendocrine cell in tissue and unveil the cell&#8217;s ultrastructure in 3D. The resolution of cocem3D is sufficient to identify organelles as small as secretory vesicles and to distinguish cell membranes for volume rendering. Cocem3D can be easily adapted to study the 3D ultrastructure of other specific cell types in their native tissue.

Here, we introduce a method, cocem3D, to unveil the ultrastructure of a specific cell in its native tissue by bridging confocal and serial block-face scanning electron microscopy.

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy (SEM)

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to cells, tissues, organelles, and even whole organisms. This protocol focuses on two chemical drying methods, hexamethyldisilazane (HMDS) and t-butyl alcohol (TBA), and their application to imaging of both prokaryotic and eukaryotic organisms using SEM. In this article, we describe how to fix, wash, dehydrate, dry, mount, sputter coat, and image three types of organisms: cyanobacteria (Toxifilum mysidocida, Golenkina sp., and an unknown sp.), two euglenoids from the genus Monomorphina (M. aenigmatica and M. pseudopyrum), and the fruit fly (Drosophila melanogaster). The purpose of this protocol is to describe a fast, inexpensive, and simple method to obtain detailed information about the structure, size, and surface characteristics of specimens that can be broadly applied to a large range of organisms for morphological assessment. Successful completion of this protocol will allow others to use SEM to visualize samples by applying these techniques to their system.

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy SEM

SEM analysis is an effective method to aid in species identification or phenotypic discrimination. This protocol describes the methods for examining specific morphological details of three representative types of organisms and would be broadly applicable to examining features of many organismal and tissue types.

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

Ultrastructural Localization of Endogenous LC3 by On-Section Correlative Light-Electron Microscopy

The visualization of autophagic organelles at the ultrastructural level by electron microscopy (EM) is essential to establish their identity and reveal details that are important for understanding the autophagic process. However, EM methods often lack molecular information, obstructing the correlation of ultrastructural information obtained by EM to fluorescence microscopy-based localization of specific autophagy proteins. Furthermore, the rarity of autophagosomes in unaltered cellular conditions hampers investigation by EM, which requires high magnification, and hence provides a limited field of view.
In answer to both challenges, an on-section correlative light-electron microscopy (CLEM) method based on fluorescent labeling was applied to correlate a common autophagosomal marker, LC3, to EM ultrastructure. The method was used to rapidly screen cells in fluorescence microscopy for LC3 labeling in combination with other relevant markers. Subsequently, the underlying ultrastructural features of selected LC3-labeled spots were identified by CLEM. The method was applied to starved cells without adding inhibitors of lysosomal acidification.
In these conditions, LC3 was found predominantly on autophagosomes and rarely in autolysosomes, in which LC3 is rapidly degraded. These data show both the feasibility and sensitivity of this approach, demonstrating that CLEM can be used to provide ultrastructural insights on LC3-mediated autophagy in native conditions-without drug treatments or genetic alterations. Overall, this method presents a valuable tool for ultrastructural localization studies of autophagy proteins and other scarce antigens by bridging light microscopy to EM data.

Here, we present a protocol for optimized on-section correlative light-electron microscopy based on endogenous, fluorescent labeling as a tool to investigate the localization of rare proteins in relation to cellular ultrastructure. The power of this approach is demonstrated by ultrastructural localization of endogenous LC3 in starved cells without Bafilomycin treatment.

The visualization of autophagic organelles at the ultrastructural level by electron microscopy (EM) is essential to establish their identity and reveal ...

Novel Protocol for Simultaneous Fluorescence and Ultrastructure Preservation

Epon Post Embedding Correlative Light and Electron Microscopy

Correlative light and electron microscopy (CLEM) is a comprehensive microscopy that combines the localization information provided by fluorescence microscopy (FM) and the context of cellular ultrastructure acquired by electron microscopy (EM). CLEM is a trade-off between fluorescence and ultrastructure, and usually, ultrastructure compromises fluorescence. Compared with other hydrophilic embedding resins, such as glycidyl methacrylate, HM20, or K4M, Epon is superior in ultrastructure preservation and sectioning properties. Previously, we had demonstrated that mEosEM can survive osmium tetroxide fixation and Epon embedding. Using mEosEM, we achieved, for the first time, Epon post embedding CLEM, which maintains the fluorescence and the ultrastructure simultaneously. Here, we provide step-by-step details about the EM sample preparation, the FM imaging, the EM imaging, and the image alignment. We also improve the procedures for identifying the same cell imaged by FM imaging during the EM imaging and detail the registration between the FM and EM images. We believe one can easily achieve Epon post embedding correlative light and electron microscopy following this new protocol in traditional EM facilities.

Author Spotlight: Advancements in Correlative Light and Electron Microscopy with Fluorescent Protein Preservation

We present a detailed protocol for Epon post-embedding correlative light and electron microscopy using a fluorescent protein called mScarlet. This method can maintain the fluorescence and the ultrastructure simultaneously. This technique is amenable to a wide variety of biological applications.