We are grateful to the members of the Stasevich lab for innumerable conversations that helped improve and develop this protocol. Specifically, Dr. Linda Forero and Dr. Phil Fox for advice on bead loading different cell types. We would like to sincerely thank Dr. Yoko Hayashi-Takanaka, Dr. Yuko Sato and Dr. Hiroshi Kimura for sharing their glass bead loading protocol. We are very grateful to Dr. Ashok Prasad and Dr. Diego Krapf for generously sharing their bead loading protocols for introducing inorganic particles into cells. We are grateful to Dr. Travis Sanders, Craig Marshall, and Dr. Thomas Santangelo for generously sharing their labeled RNA reagent. ALK, MNS,&#160;CAC, GG&#160;and TJS were supported by the National Institutes of Health (NIH) grant R35GM119728&#160;and the National Science Foundation (NSF) CAREER grant MCB-1845761, both&#160;to TJS. CAC was also supported by the&#160;NSF&#160;NRT award DGE-1450032.

1. Clean, sterilize, and dry glass beads to avoid clumping and ensure even spreading atop the cells.
<ol>
	<li>Sterilize approximately 5 mL of glass beads in sodium hydroxide (NaOH). Measure the beads in a 50 mL conical tube. Add 25 mL of 2 M NaOH and mix gently using a shaker or rotator for 2 h.</li>
	<li>Decant the NaOH, retaining as many beads as possible. If the beads are in suspension, spin down the tube of beads briefly in a centrifuge (1 min at ~1000 &#215; g, room temperature).</li>
	<li>Wash the beads thoroughly with cell culture-grade water until the pH is neutral (use a pH test strip on the eluent to confirm a neutral pH). Decant the wash water each time, as before.</li>
	<li>Wash the beads thoroughly with 100% ethanol 2-3x. Decant the ethanol each time, as before.</li>
	<li>Dry the beads. Sprinkle the beads to form a thin layer inside a sterile container (such as a 10 cm Petri dish). Leaving the container open, let the beads air dry in a biosafety cabinet overnight. Ensure that the beads are completely dry by tapping or gently shaking the container and checking that the beads have a sandy texture with no clumping or flaking.</li>
	<li>UV-sterilize the dry beads for 15 min.</li>
</ol>
2. Assemble the bead loader apparatus.
<ol>
	<li>Fasten a patch of mesh (polypropylene or equivalent material, 105 &#181;m openings to allow the beads to pass through) to cover the entire opening of the beads holding chamber with either tape or clamping the mesh between the male and female ends of a metal reusable imaging chamber (Figure 1A).</li>
	<li>UV-sterilize the apparatus for 15 min. Add the beads to the apparatus and seal it tightly with waxy film. 
	NOTE: It is essential that the beads are completely clean and dry at this step. They should be loose and look sandy with no clumps. If they do not appear so, re-wash and completely dry the beads.</li>
	<li>Store the apparatus in a sealed, dry container desiccated by silica gel or other desiccant medium. If the beads become damp, which will be apparent by bead clumping, thoroughly dry and sterilize the bead loader and replace with fresh beads. 
	&#8203;NOTE: All these precautions will prevent any mold or bacteria from growing on or around the beads within the bead loader. The bead loader apparatus can be made in different ways. See the details in the discussion.</li>
</ol>
3. Prepare glass-bottom chambers of adherent cells.
<ol>
	<li>Seed adherent mammalian cells onto a 35 mm glass-bottom chamber. Ensure that the cells are approximately 80% confluent at the time of bead loading. (See Table 1 for more information on various cell types and notes on the effectiveness of bead loading in different cell types.) 
	NOTE: Cells can be seeded in only the microwell in the center of the chamber to conserve how many cells are used.</li>
	<li>Incubate the cells under normal conditions until they are completely adherent to the glass. 
	&#8203;NOTE: It is essential that the cell density is high enough and that the cells are securely adhered to the glass. If these requirements are not met, cells will likely peel off during bead loading. The timeline between cell seeding and bead loading can be lengthened to ensure proper cell adhesion and confluency.</li>
</ol>
4. Bead loading cells
NOTE: If required, wash the cells briefly with phosphate-buffered saline (PBS) and then add 2 mL of the optimal medium. Incubate for at least 30 min.
<ol>
	<li>Make a solution of 3-8 &#181;L containing the desired plasmids, protein, and/or particles. Use ~1 &#181;g (0.1-1 pmol) of each type of plasmid and ~0.5 &#181;g (0.01 nmol) of protein, depending on experimental requirements. Use a low-retention tube for proteins so that they are not left behind on the tube walls. Bring the solution up to a minimum of 3 &#181;L with PBS, and adjust the solution volume to coat the entire area of cells to be loaded (i.e., the chamber&#39;s microwell, Figure 1B).</li>
	<li>Mix the solution thoroughly by pipetting up and down and/or flicking the tube. Briefly spin the solution down to the bottom of the tube in a tabletop microfuge.</li>
	<li>Transfer the bead loading solution and the chamber of cells into a tissue culture hood. Perform the remaining steps in the tissue culture hood using sterile technique.</li>
	<li>Remove the medium from the cells and temporarily store it in a sterile tube. Gently aspirate all medium from around the edges of the chamber, and tilt the chamber at approximately a 45&#176; angle and remove the remaining drop of media in the center microwell. During medium removal, make sure to avoid letting the pipette tip touch the glass, which may result in cell peeling and loss. Move quickly to the next step so that the cells are not dry for long.</li>
	<li>Gently pipette the bead loading solution onto the glass microwell in the center of the chamber. Optional: Incubate with gentle rocking for ~30 s without allowing the chamber to dry up completely.</li>
	<li>Gently disperse a monolayer of glass beads on top of the cells, preferably using a bead loading apparatus (Figure 1A). Ensure that the beads cover the cells in the glass-bottom microwell completely.</li>
	<li>Pinching the chamber with two fingers, tap it against the hood surface by lifting it ~2 inches and bringing it down firmly. Use a force approximately equivalent to dropping the dish from that height. Repeat for a total of ~10 taps. 
	NOTE: Ensure that the taps do not substantially peel the cells. Tapping can be optimized for the cell type. If cells load poorly, tap harder; however, if many cells peel off, tap more lightly.</li>
	<li>Gently add medium back into the chamber by pipetting slowly onto the plastic side of the chamber. Try to aspirate any floating beads without disturbing the cells. Add more pre-warmed media at this step if too much was removed. Incubate the cells for 0.5-2 h in the incubator.</li>
	<li>UV-sterilize the bead loader for 15 min before returning it to storage under desiccating conditions.</li>
	<li>Add dye (e.g., DAPI or HaloTag ligand stain, if required by the experiment) to the cells as per the manufacturer&#39;s recommended protocol.</li>
	<li>Wash the cells 3x with medium before imaging to remove the beads and excess loading components in solution. Avoid pipetting directly onto cells to keep them from peeling.</li>
</ol>
5. Imaging the bead-loaded cells
<ol>
	<li>Image the cells immediately or when required by the experiment. Use a microscope capable of capturing fluorescence (lasers or monochromatic light source). Ensure that the excitation wavelengths are appropriate for the chosen fluorophore or dye (e.g., 488 nm wavelength light for green fluorescence protein (GFP)). 
	NOTE: Bead-loaded proteins may be imaged once the cells have recovered (as soon as 30 min post loading for the cell lines described here). Plasmid expression takes &#8805;2 h depending on expression vector elements (Figure 1C, and further explanation in the discussion). Imaging of bead-loaded cells can be performed on any microscope equipped with the appropriate fluorescent sources associated with loaded probes, a camera capable of capturing fluorescence images, such as an electron-multiplying charge-coupled device (EMCCD) or scientific complementary metal-oxide semiconductor (sCMOS) camera, and an incubator to control temperature, humidity, and carbon dioxide. For a guide to fluorescence microscopy, refer to 27.</li>
</ol>

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The authors have no conflicts of interest to disclose.

The bead loading technique described here is a cost-effective and time-efficient method for introducing macromolecules and other particles into adherent cells. This versatile process can load protein (Figure 2A)15,16,26,27, a combination of protein and plasmids (Figure 2B,C)22,25, RNA (Figure 4C), 100 and 250 nm polystyrene beads (personal correspondence), synthetic dyes39&#160;or quantum dots34,40. Bead loading may have the capability to load other types of membrane-impermeable particles as well. Its most frequently used application is for loading antibodies or Fabs to target endogenous epitopes, such as post-translational modifications (PTMs), into live cells. Targets, such as PTMs, are often difficult to label in live cells without established PTM-specific, genetically encoded probes41,42. In contrast, bead loading can introduce multiple types of probes, reporters, or other molecular tools together into the same cell for monitoring multiple readouts simultaneously. We anticipate that bead loading will be a useful technique for loading a variety of macromolecules or particles.
A major advantage of bead loading is the low cost: each experiment costs less than 0.01 USD. A bead loader apparatus can be made easily using inexpensive materials costing in total ~$150, which is significantly less expensive than any other cell-loading method. The cost of a bead loader apparatus can be further reduced to under $10 by replacing the reusable metal chamber with a plastic one. For this, either drill a hole in a 35 mm chamber or remove the glass from a 35 mm glass-bottom chamber, then securely fasten the mesh in place with tape. In lieu of an apparatus, bead loading can even be performed using a wide-bore 1000 &#181;L pipette tip to scoop and sprinkle beads onto cells, although this variation makes it difficult to sprinkle a monolayer of beads onto cells (step 4.6).
Another benefit of bead loading is that cells can retain normal overall morphology, recover rapidly, and continue to grow and divide, at least for the U2OS, RPE1, and HeLa cells studied here and for the other cell lines studied elsewhere (Figure 3; Figure 4A,B; Supplemental Video 1; and Table 1)31. During bead loading, cells undergo physical stress and sometimes dislodge and peel (~5% of cells peel under optimal conditions, but greater cell loss can happen if bead loading is performed too forcefully or too many glass beads are loaded atop the cells, as depicted in Figure 3B). However, bead-loaded cells that remain attached to the coverslip usually appear healthy and can be imaged as soon as 30 min after bead loading (Figure 3A). We generally allow cells a 30-min recovery period but anticipate that imaging sooner post-bead loading is feasible.
A major drawback of this technique is that the cells need to be capable of withstanding minor physical stress during loading and remain securely adherent to the coverslip. Poorly/non-adherent cell lines or cells grown on coated plates (e.g., HEK and stem cells) often detach upon gentle tapping during bead loading. Further, experience has shown that primary neurons are too sensitive for bead loading.
Bead loading is best suited for single-cell or single-molecule experiments. In our experience, bead loading has a roughly 20-40% protein loading efficiency, and ~20% of bead-loaded cells also expressed a co-loaded plasmid (Figure 2A,B). Thus, bead loading plasmids may be less efficient for protein expression than bead loading purified proteins because plasmids must not only enter cells but also be expressed (which involves, among other things, nuclear import, transcription, and translation, each of which can lower expression efficiency). The low efficiency of bead-loaded plasmid expression can be circumvented by using alternative transfection protocols, such as lipofection, before bead loading proteins or probes16,27. Additionally, incubating cells in optimal media for 30 min before bead loading may assist plasmid expression. Due to low plasmid expression, bead loading has not often been used as an alternative to lipofection-based transfection in this laboratory. The only exception is when a purified protein, such as Fab, is to be co-loaded, in which case it is quite convenient to bead-load the protein and plasmid at the same time. Moreover, for cells that are unresponsive or intolerant to lipofection, bead loading may provide an alternate, albeit low-efficiency, method for transient plasmid expression.

Loading macromolecules into mammalian cells necessitates methodology that allows them to cross the cell&#39;s plasma membrane1. Several methods can introduce plasmids into mammalian cells through transfection, including liposomal transfection2 and diethylaminoethyl-dextran transfection3. However, methods for loading proteins or membrane-impermeable particles into cells are more limited.
Several techniques have bypassed this difficult hurdle using various strategies. First, microinjection delivers particles through a micropipette into live cells under a microscope4. While arguably the most controlled and least invasive method, this technique is relatively low-throughput because cells must be loaded one by one. Further, microinjection requires specialized equipment and is technically demanding.
Second, electroporation is a way to electro-inject proteins into cells via voltage-induced membrane disruption5,6,7. However, this method again requires specialized, expensive equipment, and the shock can cause cell stress and mortality. Further, cells must be trypsinized before electroporation and subsequently replated, limiting the timeframe at which cells can be investigated post-electroporation.
Third, cell membranes may be chemically modified for temporary, reversible permeabilization8,9. Streptolysin-O loading inserts an endotoxin into cell membranes, which forms temporary pores, allowing exogenous membrane-impermeable particles, including proteins and DNA plasmids, to enter cells10. After a 2 h recovery, about half the cells repair these pores and halt internalizing particles from the solution. However, this technique requires a long recovery time and is incompatible with cell types that cannot tolerate endotoxins.
Fourth, mechanical disruption loads particles into cells through physical perturbation of the cell membrane11. This can be done in multiple ways, including scratching, scraping, and rolling beads atop cells12,13. As early as 1987, beads have been used to load proteins into cells mechanically14. More recently, the bead loading technique has been optimized and adapted beyond proteins to include the loading of plasmids and RNA, as described here.
Bead loading is an easy, inexpensive, and fast method for loading protein and plasmids into adherent human cells. Glass beads are briefly rolled atop cells, temporarily disrupting their cellular membrane. This allows particles in solution to enter. As bead loading has low efficiency, it is best suited for single-molecule or single-cell microscopy experiments. Bead loading can introduce a wide variety of proteins, including fragmented antibodies (Fab),15,16 purified proteins like scFvs,17 intrabodies,18,19, or mRNA coat proteins, e.g., MS2 coat protein (MCP)20,21. Plasmid expression vectors can also be added to the protein solution and bead-loaded simultaneously22,23,24,25.
Beyond proteins and plasmids, molecules as large as 250 nm polystyrene beads have been introduced into cells via bead loading (personal&#160;communication). Bead loading is incredibly inexpensive, costing less than 0.01 USD per experiment in materials and requiring no additional expensive equipment. The cost is further reduced by minimizing the amount of probes used per experiment because only the cells in the central 14 mm-diameter microwell of an imaging chamber are loaded. It should be noted that the limited loading area means that bead loading is not ideal for bulk-cell loading.
This manuscript presents the bead loading process, including how to construct the bead loading apparatus and perform an experiment. It shows that proteins, RNA, and DNA can be loaded into various cell types and that two different, simultaneously bead-loaded proteins have highly correlated cellular concentrations and relatively low variance. Also discussed are variations in the protocol based on cell type and loading of protein, plasmid, or RNA. Although beads are thought to perforate and disrupt the cell membrane, when appropriately performed, the bead loading process dislodges only a small number of cells from the bottom of the imaging chamber. After a short recovery period, cells continue to grow and divide. This methodology is ideal for live-cell microscopy experiments, including single-molecule protein and RNA tracking, post-translational modification detection, observation of dynamic cellular mechanisms, or subcellular localization monitoring15,16,22,26,27.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10 cm cell culture dishes</td>
			<td>VWR</td>
			<td>82050-916</td>
			<td>Use to culture cells</td>
		</tr>
		<tr>
			<td>35 mm cell culture dishes</td>
			<td>Falcon</td>
			<td>353001</td>
			<td>Use to construct bead loader</td>
		</tr>
		<tr>
			<td>Attofluor Cell Chamber</td>
			<td>Thermo Fisher Scientific</td>
			<td>A7816</td>
			<td>Use to construct the custom bead loader</td>
		</tr>
		<tr>
			<td>DMEM, high glucose, no glutamine</td>
			<td>Thermo Fisher Scientific</td>
			<td>11960069</td>
			<td>Use in general cell culture</td>
		</tr>
		<tr>
			<td>Drierite Indicating Absorbents</td>
			<td>Thermo Fisher Scientific</td>
			<td>07-578-3B</td>
			<td>Store the bead loader in a desiccator with these absorbent pellets</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Atlas Biologics</td>
			<td>F-0050-A</td>
			<td>Use in general cell culture and as a supplement before bead loading</td>
		</tr>
		<tr>
			<td>Glass beads, acid washed, &#8804;106 &#181;m*</td>
			<td>Millipore Sigma</td>
			<td>G4649</td>
			<td>Sprinkle on cells to bead load plasmid DNA and proteins</td>
		</tr>
		<tr>
			<td>Glass bottom dishes, 35 mm, #1.5, 14 mm glass</td>
			<td>MatTek Corporation</td>
			<td>P35G-1.5-14-C</td>
			<td>Seed cells onto these chambers for imaging</td>
		</tr>
		<tr>
			<td>L-Glutamine-200 mM</td>
			<td>Thermo Fisher Scientific</td>
			<td>25030081</td>
			<td>Use to make DMEM + media</td>
		</tr>
		<tr>
			<td>Opti-MEM, Reduced Serum Medium</td>
			<td>Thermo Fisher Scientific</td>
			<td>31985070</td>
			<td>Optimal media for incubating cells before bead loading (optional step)</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>VWR</td>
			<td>52858-032</td>
			<td>waxy film used to construct bead loader</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Thermo Fisher Scientific</td>
			<td>15140-122</td>
			<td>Use to make DMEM + media</td>
		</tr>
		<tr>
			<td>Phenol-free DMEM</td>
			<td>Thermo Fisher Scientific</td>
			<td>31053036</td>
			<td>Use on cells before imaging</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS)</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM9625</td>
			<td>Working stock of sterile 1X PBS</td>
		</tr>
		<tr>
			<td>Sodium Hydroxide (NaOH)</td>
			<td>Thermo Fisher Scientific</td>
			<td>S318-500</td>
			<td>Use 2 M solution to wash glass beads</td>
		</tr>
		<tr>
			<td>Spectra Mesh Woven Filters, Polypropylene, 105 &#181;m opening</td>
			<td>Spectrum Labs</td>
			<td>148496</td>
			<td>Use to construct bead loader</td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Thermo Fisher Scientific</td>
			<td>25300062</td>
			<td>Use in general cell culture</td>
		</tr>
	</tbody>
</table>

bead loading proteins and nucleic acids into adherent human cells

Many live-cell imaging experiments use exogenous particles (e.g., peptides, antibodies, beads) to label or function within cells. However, introducing proteins into a cell across its membrane is difficult. The limited selection of current methods struggles with low efficiency, requires expensive and technically demanding equipment, or functions within narrow parameters. Here, we describe a relatively simple and cost-effective technique for loading DNA, RNA, and proteins into live human cells. Bead loading induces a temporary mechanical disruption to the cell membrane, allowing macromolecules to enter adherent, live mammalian cells. At less than 0.01 USD per experiment, bead loading is the least expensive cell loading method available. Moreover, bead loading does not substantially stress cells or impact their viability or proliferation. This manuscript describes the steps of the bead loading procedure, adaptations, variations, and technical limitations. This methodology is especially suited for live-cell imaging but provides a practical solution for other applications requiring the introduction of proteins, beads, RNA, or plasmids into living, adherent mammalian cells.

The most common application of bead loading is to introduce one or more types of protein into adherent human cells. To illustrate this, cells were bead-loaded with a solution of a Cy3- and Alexa488-conjugated Fab protein. Although not every cell in the microwell was bead-loaded, the cells that were loaded almost always had both Cy3- and Alexa488-labeled proteins together (Figure 2A). According to an earlier estimate, when 0.5 microgram&#160;of Fab diluted in 4 microliters is bead-loaded29, as in Figure 2A, each cell is loaded with roughly 106 Fab molecules.
Plasmid DNA encoding GFP (1 &#181;g of plasmid DNA, 1.8 &#181;L of a 557 ng/&#181;L solution) and 0.5 &#181;g of Cy3-labeled Fab&#160;was also introduced into cells via bead loading and subsequently expressed and visualized (Figure 2B). The GFP fluorescence indicated that the GFP-encoding plasmid was not only loaded into cells but also expressed. Thus, in the same cell, bead loading can introduce a protein probe (e.g., Cy3-labeled Fab) and reporter plasmid (e.g., GFP), as performed in this laboratory previously22,23,24. We determined that 40% of the cells were bead-loaded with Fab protein and 21% of the bead-loaded cells expressed the co-loaded plasmid, as shown in the representative fields-of-view in Figure 2B. Typically, each chamber is loaded with 1-2 &#181;g of plasmid, approximately the same amount as lipofection.
Bead-loaded cells express widely varying levels of plasmids (Figure 2C,D). To specifically measure this, we used the Fisher Ratio test to compare the distributions of protein and plasmid intensity data. The results showed that although proteins 1 and 2 had similar intensity distributions (p = ~1), each protein had a significantly smaller distribution than the plasmid (p = 3.2e-6 and 1.8e-5). Although this could be due to variability in how many plasmids are loaded per cell, the greater source of variability may arise from the many steps required for plasmid expression that are likely to vary greatly between cells, including being imported into the cell nucleus, transcription, and translation. In contrast, the levels of bead-loaded proteins had slight cell-to-cell variance, and the levels of two simultaneously loaded proteins were highly correlated with each other (Figure 2D,E).
Plasmid expression can be seen as early as 2-4 h post bead loading but may occur later depending on when optimal plasmid expression is obtained. We recommend performing a time course to determine the best window of expression for a specific plasmid spanning 2-24 h post bead loading. This can be done in one chamber with long timeframe imaging or by bead loading and staggering multiple chambers. Bead-loaded cells remain adherent and are healthy enough to grow and divide. Bead-loaded human U2OS cells were imaged directly before, directly after, and 24 h after bead loading. Proper bead loading had almost no noticeable effect on the number of cells or their morphology, as shown in Figure 3A&#160;(left, middle).
In contrast, poor bead loading with too many beads and excessive tapping force is depicted in Figure 3B. This caused much cell loss (large patches of the coverglass without cells and detached, floating, out-of-focus cells), poor cell morphology (cells appearing rounded up and poorly adhered), and clusters of beads remaining on the coverglass after bead loading. Though cells are thought to undergo mechanical damage during bead loading, cells grew and proliferated in the properly bead-loaded chamber, as evidenced by the increased number of cells 24 h after bead loading (Figure 3A, right). The effect on cell viability can be assessed through a variety of assays, such as a 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay, to compare bead-loaded to mock-loaded cells30. Further, this and previous work show that the bead-loaded cells undergo cell division (Figure 3C and Supplemental Video 1), and the timing of mitosis is not affected by bead loading31, which serves as further evidence for sustained cell health after bead loading.
Bead loading is a versatile technique, accommodating several adherent cell lines and various macromolecules. Here, this variety has been demonstrated by loading RPE1 and HeLa cell lines with Fab (Figure 4A,B). Table 1 provides further examples of bead loading in different cell lines, in this laboratory and beyond, and points out some of the nuanced differences between bead loading protocols from other labs. Of note, the diameter of glass beads used for loading varies greatly between laboratories, though the most efficient loading was found for small, 75 &#181;m diameter beads in several cell lines14. Further, this laboratory has begun bead loading RNA as well (data not shown). Figure 4C displays a representative U2OS cell bead-loaded with a Cy5-RNA 9mer and Cy3-DNA 28mer together.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62559/62559fig01.jpg" /> 
Figure 1: Bead loading apparatus, technique, and timeline <a href="https://www.jove.com/files/ftp_upload/62559/62559fig01large.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/62559/62559fig02.jpg" /> 
Figure 2. Bead loading introduces low variability in protein concentration but high variability in plasmid expression. (A) Cells were bead-loaded with 0.5 &#181;g of each of Alexa488-conjugated anti-H3K27 acetyl Fab (green) and Cy3-conjugated anti-RNAPII-Serine 5-phosphorylated Fab (red) in 4 &#181;L of bead loading solution. Cells were DAPI-stained (blue) and then live-imaged immediately. Scale bars = 20 &#181;m. (B) Cells were bead-loaded with 0.5 &#181;g of Fab protein (Cy3-conjugated anti-RNAPII-Serine 5-phosphorylated protein, red) and 1 &#181;g of plasmid encoding superfolder GFP-H2B (green) in 4 &#181;L of bead loading solution. After 24 h, cells were DAPI-stained (blue) and imaged live. Scale bars = 30 &#181;m. (C-E) Protein 1 (JF646-HaloLigand-labeled HaloTag-MCP), protein 2 (Cy3-conjugated anti-FLAG Fab), and a plasmid encoding EGFP (&#955;N-EGFP-LacZ) were bead-loaded together into cells. The total intensity in each fluorescent channel was measured in a 1.3 x 1.3 &#181;m patch in the cytoplasm of each cell. N = 25 cells. (C) Representative cells expressing the bead-loaded plasmid, &#955;N-EGFP-LacZ. The same imaging conditions and intensities were used for all cells. Spots are aggregates of the expressed protein. Scale bars = 10 &#181;m. (D) The chart shows each cell&#39;s total intensity of either protein 1, protein 2, or EGFP expressed from the plasmid. Each channel was normalized to the median. Bonferroni-corrected P values were calculated by the Fisher Ratio test to determine whether the distribution of protein or plasmid intensity data has the same variability. Each point represents a cell. (E) The total intensities for either both proteins, protein 1 and the plasmid, or protein 2 and the plasmid, are plotted against each other. Calculated R2 values are displayed. Each point represents a cell. Abbreviations: DAPI = 4&#8242;,6-diamidino-2-phenylindole; EGFP = enhanced green fluorescent protein; A.U. = arbitrary units; MCP = MS2 coat protein; RNAPII = RNA polymerase II. <a href="https://www.jove.com/files/ftp_upload/62559/62559fig02large.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/62559/62559fig03.jpg" /> 
Figure 3: Bead-loaded cells remain adherent and are healthy enough to grow and divide. (A) U2OS cells were bead-loaded with 0.5 &#181;g of Cy3-conjugated anti-FLAG Fab in 4 &#181;L of bead loading solution. The cells were imaged directly before, directly after bead loading, and 24 h after bead loading. Orange arrows identify areas where cells peeled off during bead loading. Scale bars = 2 mm. (B) Representative image of U2OS cells bead-loaded with components from (A) but with harsh tapping and too many beads. The red arrow identifies extra glass beads. Scale bar = 2 mm. (C) U2OS cells were loaded with 1.5 &#181;g of the 14.4 kbp plasmid smFLAG-KDM5B-15xBoxB-24xMS2, 0.5 &#181;g of Cy3-conjugated anti-FLAG Fab (green), 130 ng of HaloTag-MCP (magenta) in 8 &#181;L of bead loading solution. Directly before imaging, the HaloTag was stained with JF646-HaloLigand. The MS2 stem-loops of the mRNA transcribed from the reporter plasmid are labeled by MCP (magenta spots), and FLAG-tagged translated reporter protein is labeled by anti-FLAG Fab (green colocalization to mRNA). Mature Fab-labeled protein localizes to the nucleus. This cell was imaged 4-15 h after bead loading. Yellow arrows identify the cell nucleus before and nuclei after cell division. Scale bars = 5 &#181;m. Abbreviation: MCP = MS2 coat protein. <a href="https://www.jove.com/files/ftp_upload/62559/62559fig03large.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/62559/62559fig04.jpg" /> 
Figure 4: Variations in cell type loading material of the bead loading protocol. (A-B) RPE1 (A) and HeLa (B) cells were bead-loaded with 1.5 &#181;g of a nuclear Fab protein (anti-RNAPII-Serine 5-phosphorylation) in 4 &#181;L of loading solution. Each cell&#39;s nucleus (nuc) and cytoplasm (cyto) are marked. Cells were imaged 6 h after being bead-loaded. Scale bars = 5 &#181;m. (C) Human U2OS cells were bead-loaded with both Cy5-RNA 9mer (magenta) and Cy3-DNA 28mer (green) oligos, 10 picomoles of each, in 4 &#181;L of bead loading solution. Cells were imaged 4 h after being bead-loaded. All cell nuclei are highlighted by a dashed line. Scale bars = 5 &#181;m. Abbreviations: RNAPII = RNA polymerase II. <a href="https://www.jove.com/files/ftp_upload/62559/62559fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody style="font-size: 14px;">
		<tr>
			<td>Cell line</td>
			<td>Cell type</td>
			<td>Bead loading effectiveness</td>
			<td>Notes/reference</td>
		</tr>
		<tr>
			<td>Stem cells (human)</td>
			<td>Embryonic stem cells</td>
			<td>Difficult</td>
			<td>*Many cells peel off during bead loading if plated on gelatin-coated plates</td>
		</tr>
		<tr>
			<td>HEK 293</td>
			<td>Human embryonic kidney cells</td>
			<td>Difficult</td>
			<td>*Need to lay down a gel matrix to imaging chamber surface before bead loading. Tap gently when bead loading at first.</td>
		</tr>
		<tr>
			<td>Neurons (rat)</td>
			<td>Primary embryonic neurons (e-18), dissociated</td>
			<td>Very inefficient</td>
			<td>*Efficient bead loading of neurons was not observed using this standard bead loading protocol. This could be due to the non-adherent nature of neurons or from consequent damage to neural processes.</td>
		</tr>
		<tr>
			<td>MDCK (canine)</td>
			<td>Madin-Darby canine kidney cells</td>
			<td>See McNeil and Warder (1987)14</td>
			<td>*Low-efficiency bead loading14</td>
		</tr>
		<tr>
			<td>U2OS (human)</td>
			<td>Osteosarcoma</td>
			<td>Standard bead loading protocol</td>
			<td></td>
		</tr>
		<tr>
			<td>HeLa (human)</td>
			<td>Cervical cancer</td>
			<td>Standard bead loading protocol</td>
			<td></td>
		</tr>
		<tr>
			<td>RPE1 (human)</td>
			<td>Epithelial cells immortalized with hTERT</td>
			<td>Standard bead loading protocol</td>
			<td></td>
		</tr>
		<tr>
			<td>HFF (human)</td>
			<td>Primary foreskin fibroblasts</td>
			<td>See Besteiro et al. (2009)31&#160;</td>
			<td>*Modified protocol of tilting instead of tapping31</td>
		</tr>
		<tr>
			<td rowspan="3">BALB/c 3T3, NIH 3T3, and Swiss 3T3 (mouse)</td>
			<td rowspan="3">Embryonic fibroblasts</td>
			<td rowspan="3">See Gilmore and Romer (1996)32, Emerson et al. (2014)33 and McNeil and Warder (1987)14</td>
			<td>*425&#8211;600 &#956;m glass beads reported32</td>
		</tr>
		<tr>
			<td>*Used 200&#8211;300 &#956;m glass beads33</td>
		</tr>
		<tr>
			<td>*75 &#956;m glass beads gave better results than 400 &#956;m14</td>
		</tr>
		<tr>
			<td>DM (Indian muntjac)</td>
			<td>Skin fibroblasts</td>
			<td>See Manders, Kimura, and Cook (1999)34</td>
			<td></td>
		</tr>
		<tr>
			<td>CHO (hamster)</td>
			<td>Epithelial-like ovary cells</td>
			<td>See Memedula and Belmont (2003)35</td>
			<td>*Used 425&#8211;600 &#956;m glass beads35</td>
		</tr>
		<tr>
			<td>BAE (bovine)</td>
			<td>Bovine aortic endothelial cells (BAEC-11)</td>
			<td>See McNeil and Warder (1987)14</td>
			<td>*75 &#956;m glass beads gave better results than 400 &#956;m14</td>
		</tr>
		<tr>
			<td>PtK-2 (Potomus tridaclylis)</td>
			<td>Epithelial kidney cells</td>
			<td>See McNeil and Warder (1987)14</td>
			<td>*75 &#956;m glass beads gave better results than 400 &#956;m14</td>
		</tr>
		<tr>
			<td>HUVEC (human)</td>
			<td>Umbilical vein endothelial cells</td>
			<td>See Gilmore and Romer (1996)32</td>
			<td>*Used 425&#8211;600 &#956;m glass beads32</td>
		</tr>
		<tr>
			<td>J774 and J774.2 (mouse)</td>
			<td>monocyte macrophage cells</td>
			<td>See Becker et al. (2005)36 and McNeil and Warder (1987)14</td>
			<td>*Gentle agitation (instead of tapping) and 425&#8211;600 &#956;m glass beads36</td>
		</tr>
		<tr>
			<td>MS-5 (mouse)</td>
			<td>bone marrow stromal cells</td>
			<td>See Molenaar et al. (2003)37</td>
			<td></td>
		</tr>
		<tr>
			<td rowspan="2">WPE1-NB11 (human)</td>
			<td rowspan="2">prostate epithelial cells</td>
			<td>See Gilmore and Romer (1996)32 and</td>
			<td>*Used 425&#8211;600 &#956;m glass beads32</td>
		</tr>
		<tr>
			<td>Emerson et al. (2014)33</td>
			<td>*Used 200&#8211;300 &#956;m glass beads33</td>
		</tr>
	</tbody>
</table>
Table 1: Bead loading in different cell lines. For the cell lines that have not yet been bead-loaded in this laboratory, references and notes on variations in the protocol are provided.
Supplemental Video 1: Example of a bead-loaded cell undergoing cell division. U2OS cells were loaded with 1.5 &#181;g of the 14.4 kbp plasmid smFLAG-KDM5B-15xBoxB-24xMS2, 0.5 &#181;g of Cy3-conjugated anti-FLAG Fab (green), 130 ng of HaloTag-MCP (magenta) in 8 &#181;L of bead loading solution. Directly before imaging, the HaloTag was stained with JF646-HaloLigand. The MS2 stem-loops of the mRNA transcribed from the reporter plasmid are labeled by MCP (magenta spots), and FLAG-tagged translated reporter protein is labeled via anti-FLAG Fab (green colocalization to mRNA). Mature Fab-labeled protein localizes to the nucleus. This cell was imaged 4-15 h after bead loading. Scale bar = 10 &#181;m. Abbreviation: MCP = MS2 coat protein. <a href="https://www.jove.com/files/ftp_upload/62559/bead_load_cell_divides.avi" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about Bead Loading Proteins and Nucleic Acids into Adherent Human Cells at JoVE.com

Bead Loading Proteins and Nucleic Acids into Adherent Human Cells

Bead loading introduces proteins, plasmids, and particles into adherent mammalian cells. This cell loading technique is inexpensive, rapid, and does not substantially affect cell health. It is best suited for live-cell imaging.

Many live-cell imaging experiments use exogenous particles (e.g., peptides, antibodies, beads) to label or function within cells. However, introducing ...

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JoVE Journal

Biology

4.6K Views.  Colorado State University. Bead loading introduces proteins, plasmids, and particles into adherent mammalian cells. This cell loading technique is inexpensive, rapid, and does not substantially affect cell health. It is best suited for live-cell imaging. 

Video: Bead Loading Proteins and Nucleic Acids into Adherent Human Cells

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, many proteins are present as a freely diffusing population in free exchange with a sub-population that is tightly associated with a particular subcellular domain or structure. In situ subcellular fractionation allows the visualization of protein compartmentalization and can also reveal protein sub-populations that localize to specific structures. For example, removal of soluble cytoplasmic proteins and loosely held nuclear proteins can reveal the stable association of some transcription factors with chromatin. Subsequent digestion of DNA can in some cases reveal association with the network of proteins and RNAs that is collectively termed the nuclear scaffold or nuclear matrix. 
Here we describe the steps required during the in situ fractionation of adherent and non-adherent mammalian cells on microscope coverslips. Protein visualization can be achieved using specific antibodies or fluorescent fusion proteins and fluorescence microscopy. Antibodies and/or fluorescent dyes that act as markers for specific compartments or structures allow protein localization to be mapped in detail. In situ fractionation can also be combined with western blotting to compare the amounts of protein present in each fraction. This simple biochemical approach can reveal associations that would otherwise remain undetected.

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

In situ subcellular fractionation of mammalian cells on microscope coverslips allows the visualisation of protein localisation.

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, ...

Differentiation of Embryonic Stem Cells into Oligodendrocyte Precursors

Oligodendrocytes are the myelinating cells of the central nervous system. For regenerative cell therapy in demyelinating diseases, there is significant interest in deriving a pure population of lineage-committed oligodendrocyte precursor cells (OPCs) for transplantation. OPCs are characterized by the activity of the transcription factor Olig2 and surface expression of a proteoglycan NG2. Using the GFP-Olig2 (G-Olig2) mouse embryonic stem cell (mESC) reporter line, we optimized conditions for the differentiation of mESCs into GFP+Olig2+NG2+ OPCs. In our protocol, we first describe the generation of embryoid bodies (EBs) from mESCs. Second, we describe treatment of mESC-derived EBs with small molecules: (1) retinoic acid (RA) and (2) a sonic hedgehog (Shh) agonist purmorphamine (Pur) under defined culture conditions to direct EB differentiation into the oligodendroglial lineage. By this approach, OPCs can be obtained with high efficiency (&gt;80%) in a time period of 30 days. Cells derived from mESCs in this protocol are phenotypically similar to OPCs derived from primary tissue culture. The mESC-derived OPCs do not show the spiking property described for a subpopulation of brain OPCs in situ. To study this electrophysiological property, we describe the generation of spiking mESC-derived OPCs by ectopically expressing NaV1.2 subunit. The spiking and nonspiking cells obtained from this protocol will help advance functional studies on the two subpopulations of OPCs.

We describe a small molecule-based protocol for differentiation of mouse embryonic stem cells into oligodendrocyte precursor cells (OPCs). This protocol generates Olig2+NG2+ OPCs with high efficiency by 30 days of differentiation. We also describe a method to generate "spiking" OPCs that can fire action potentials.

Oligodendrocytes are the myelinating cells of the central nervous system. For regenerative cell therapy in demyelinating diseases, there is significant ...

Xenotransplantation of Human Stem Cells into the Chicken Embryo

The chicken embryo is a classical animal model for studying normal embryonic and fetal development and for xenotransplantation experiments to study the behavior of cells in a standardized in vivo environment. The main advantages of the chicken embryo include low cost, high accessibility, ease of surgical manipulation and lack of a fully developed immune system. Xenotransplantation into chicken embryos can provide valuable information about cell proliferation, differentiation and behavior, the responses of cells to signals in defined embryonic tissue niches, and tumorigenic potential. Transplanting cells into chicken embryos can also be a step towards transplantation experiments in other animal models. Recently the chicken embryo has been used to evaluate the neurogenic potential of human stem and progenitor cells following implantation into neural anlage1-6. In this video we document the entire procedure for transplanting human stem cells into the developing central nervous system of the chicken embryo. The procedure starts with incubation of fertilized eggs until embryos of the desired age have developed. The eggshell is then opened, and the embryo contrasted by injecting dye between the embryo and the yolk. Small lesions are made in the neural tube using microsurgery, creating a regenerative site for cell deposition that promotes subsequent integration into the host tissue. We demonstrate injections of human stem cells into such lesions made in the part of the neural tube that forms the hindbrain and the spinal cord, and into the lumen of the part of the neural tube that forms the brain. Systemic injections into extraembryonic veins and arteries are also demonstrated as an alternative way to deliver cells to vascularized tissues including the central nervous system. Finally we show how to remove the embryo from the egg after several days of further development and how to dissect the spinal cord free for subsequent physiological, histological or biochemical analyses.

In this paper we present a method for transplanting human stem cells into various regions of the central nervous system of the chicken embryo. This provides an in vivo model for assessing the proliferation and differentiation of various types of human stem cells in embryonic tissue environments.

The chicken embryo is a classical animal model for studying normal embryonic and fetal development and for xenotransplantation experiments to study the ...

NanoDrop Microvolume Quantitation of Nucleic Acids

Biomolecular assays are continually being developed that use progressively smaller amounts of material, often precluding the use of conventional cuvette-based instruments for nucleic acid quantitation for those that can perform microvolume quantitation.
The NanoDrop microvolume sample retention system (Thermo Scientific NanoDrop Products) functions by combining fiber optic technology and natural surface tension properties to capture and retain minute amounts of sample independent of traditional containment apparatus such as cuvettes or capillaries. Furthermore, the system employs shorter path lengths, which result in a broad range of nucleic acid concentration measurements, essentially eliminating the need to perform dilutions. Reducing the volume of sample required for spectroscopic analysis also facilitates the inclusion of additional quality control steps throughout many molecular workflows, increasing efficiency and ultimately leading to greater confidence in downstream results.
The need for high-sensitivity fluorescent analysis of limited mass has also emerged with recent experimental advances. Using the same microvolume sample retention technology, fluorescent measurements may be performed with 2 &mu;L of material, allowing fluorescent assays volume requirements to be significantly reduced. Such microreactions of 10 &mu;L or less are now possible using a dedicated microvolume fluorospectrometer.
Two microvolume nucleic acid quantitation protocols will be demonstrated that use integrated sample retention systems as practical alternatives to traditional cuvette-based protocols. First, a direct A260 absorbance method using a microvolume spectrophotometer is described. This is followed by a demonstration of a fluorescence-based method that enables reduced-volume fluorescence reactions with a microvolume fluorospectrometer. These novel techniques enable the assessment of nucleic acid concentrations ranging from 1 pg/ &mu;L to 15,000 ng/ &mu;L with minimal consumption of sample.

The use of NanoDrop microvolume systems as practical and efficient alternatives to traditional nucleic acid quantitation methodology is described through the demonstration of two microvolume nucleic acid quantitation protocols.

Biomolecular assays are continually being developed that use progressively smaller amounts of material, often precluding the use of conventional ...

Clonogenic Assay: Adherent Cells

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. Apart from documenting the method, the initial landmark study generated the first radiation-dose response curve for X-ray irradiated mammalian (HeLa) cells in culture1. Basically, the clonogenic assay enables an assessment of the differences in reproductive viability (capacity of cells to produce progeny; i.e. a single cell to form a colony of 50 or more cells) between control untreated cells and cells that have undergone various treatments such as exposure to ionising radiation, various chemical compounds (e.g. cytotoxic agents) or in other cases genetic manipulation. The assay has become the most widely accepted technique in radiation biology and has been widely used for evaluating the radiation sensitivity of different cell lines. Further, the clonogenic assay is commonly used for monitoring the efficacy of radiation modifying compounds and for determining the effects of cytotoxic agents and other anti-cancer therapeutics on colony forming ability, in different cell lines. A typical clonogenic survival experiment using adherent cells lines involves three distinct components, 1) treatment of the cell monolayer in tissue culture flasks, 2) preparation of single cell suspensions and plating an appropriate number of cells in petri dishes and 3) fixing and staining colonies following a relevant incubation period, which could range from 1-3 weeks, depending on the cell line. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cell lines with the use of an immortalized human keratinocyte cell line (FEP-1811)2. Also, our aims are to describe common features of clonogenic assays including calculation of the plating efficiency and survival fractions after exposure of cells to radiation, and to exemplify modification of radiation-response with the use of a natural antioxidant formulation.

The applicability of the clonogenic assay for evaluating reproductive viability has been established for more than 50 years. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cells.

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. ...

Concentration Determination of Nucleic Acids and Proteins Using the Micro-volume Bio-spec Nano Spectrophotometer

Nucleic Acid quantitation procedures have advanced significantly in the last three decades. More and more, molecular biologists require consistent small-volume analysis of nucleic acid samples for their experiments. The BioSpec-nano provides a potential solution to the problems of inaccurate, non-reproducible results, inherent in current DNA quantitation methods, via specialized optics and a sensitive PDA detector. The BioSpec-nano also has automated functionality such that mounting, measurement, and cleaning are done by the instrument, thereby eliminating tedious, repetitive, and inconsistent placement of the fiber optic element and manual cleaning.
In this study, data is presented on the quantification of DNA and protein, as well as on measurement reproducibility and accuracy. Automated sample contact and rapid scanning allows measurement in three seconds, resulting in excellent throughput. Data analysis is carried out using the built-in features of the software. The formula used for calculating DNA concentration is:
Sample Concentration = DF &middot; (OD260-OD320)&middot; NACF (1)
Where DF = sample dilution factor and NACF = nucleic acid concentration factor.
The Nucleic Acid concentration factor is set in accordance with the analyte selected1.
Protein concentration results can be expressed as &mu;g/ mL or as moles/L by entering e280 and molecular weight values respectively. When residue values for Tyr, Trp and Cysteine (S-S bond) are entered in the e280Calc tab, the extinction coefficient values are calculated as e280 = 5500 x (Trp residues) + 1490 x (Tyr residues) + 125 x (cysteine S-S bond). The e280 value is used by the software for concentration calculation.
In addition to concentration determination of nucleic acids and protein, the BioSpec-nano can be used as an ultra micro-volume spectrophotometer for many other analytes or as a standard spectrophotometer using 5 mm pathlength cells.

This communication presents data on the accuracy and reproducibility of the BioSpec-nano UV-VIS spectrophotometer for dsDNA and protein quantitation. Even with ultra-small volumes (1 to 2 L), reproducibility is excellent, while the automated wiping function improves throughput and results in minimal carryover for more precise results.

Nucleic Acid quantitation procedures have advanced significantly in the last three decades.  More and more, molecular biologists require consistent ...

Identifying Targets of Human microRNAs with the LightSwitch Luciferase Assay System using 3'UTR-reporter Constructs and a microRNA Mimic in Adherent Cells

MicroRNAs (miRNAs) are important regulators of gene expression and play a role in many biological processes. More than 700 human miRNAs have been identified so far with each having up to hundreds of unique target mRNAs. Computational tools, expression and proteomics assays, and chromatin-immunoprecipitation-based techniques provide important clues for identifying mRNAs that are direct targets of a particular miRNA. In addition, 3'UTR-reporter assays have become an important component of thorough miRNA target studies because they provide functional evidence for and quantitate the effects of specific miRNA-3'UTR interactions in a cell-based system. To enable more researchers to leverage 3'UTR-reporter assays and to support the scale-up of such assays to high-throughput levels, we have created a genome-wide collection of human 3'UTR luciferase reporters in the highly-optimized LightSwitch Luciferase Assay System. The system also includes synthetic miRNA target reporter constructs for use as positive controls, various endogenous 3'UTR reporter constructs, and a series of standardized experimental protocols.
Here we describe a method for co-transfection of individual 3'UTR-reporter constructs along with a miRNA mimic that is efficient, reproducible, and amenable to high-throughput analysis.

Identifying Targets of Human microRNAs with the LightSwitch Luciferase Assay System using 3&#039;UTR-reporter Constructs and a microRNA Mimic in Adherent Cells

MicroRNAs (miRNAs) are important regulators of gene expression and have been shown to play a role in numerous biological processes. To better understand miRNA-UTR interactions, we have created a genome-wide collection of 3 UTR luciferase reporters paired with a novel luciferase gene and assay reagent, the LightSwitch system.

MicroRNAs (miRNAs) are important regulators of gene expression and play a role in many biological processes.  More than 700 human miRNAs have been ...

Sequence-specific Labeling of Nucleic Acids and Proteins with Methyltransferases and Cofactor Analogues

S-Adenosyl-l-methionine (AdoMet or SAM)-dependent methyltransferases (MTase) catalyze the transfer of the activated methyl group from AdoMet to specific positions in DNA, RNA, proteins and small biomolecules. This natural methylation reaction can be expanded to a wide variety of alkylation reactions using synthetic cofactor analogues. Replacement of the reactive sulfonium center of AdoMet with an aziridine ring leads to cofactors which can be coupled with DNA by various DNA MTases. These aziridine cofactors can be equipped with reporter groups at different positions of the adenine moiety and used for Sequence-specific Methyltransferase-Induced Labeling of DNA (SMILing DNA). As a typical example we give a protocol for biotinylation of pBR322 plasmid DNA at the 5&#8217;-ATCGAT-3&#8217; sequence with the DNA MTase M.BseCI and the aziridine cofactor 6BAz in one step. Extension of the activated methyl group with unsaturated alkyl groups results in another class of AdoMet analogues which are used for methyltransferase-directed Transfer of Activated Groups (mTAG). Since the extended side chains are activated by the sulfonium center and the unsaturated bond, these cofactors are called double-activated AdoMet analogues. These analogues not only function as cofactors for DNA MTases, like the aziridine cofactors, but also for RNA, protein and small molecule MTases. They are typically used for enzymatic modification of MTase substrates with unique functional groups which are labeled with reporter groups in a second chemical step. This is exemplified in a protocol for fluorescence labeling of histone H3 protein. A small propargyl group is transferred from the cofactor analogue SeAdoYn to the protein by the histone H3 lysine 4 (H3K4) MTase Set7/9 followed by click labeling of the alkynylated histone H3 with TAMRA azide. MTase-mediated labeling with cofactor analogues is an enabling technology for many exciting applications including identification and functional study of MTase substrates as well as DNA genotyping and methylation detection.

DNA and proteins are sequence-specifically labeled with affinity or fluorescent reporter groups using DNA or protein methyltransferases and synthetic cofactor analogues. Depending on the cofactor specificity of the enzymes, aziridine or double activated cofactor analogues are employed for one- or two-step labeling.

S-Adenosyl-l-methionine (AdoMet or SAM)-dependent methyltransferases (MTase) catalyze the transfer of the activated methyl group from AdoMet to specific ...

Residue-specific Incorporation of Noncanonical Amino Acids into Model Proteins Using an Escherichia coli Cell-free Transcription-translation System

The canonical set of amino acids leads to an exceptionally wide range of protein functionality. Nevertheless, the set of residues still imposes limitations on potential protein applications. The incorporation of noncanonical amino acids can enlarge this scope. There are two complementary approaches for the incorporation of noncanonical amino acids. For site-specific incorporation, in addition to the endogenous canonical translational machineries, an orthogonal aminoacyl-tRNA-synthetase-tRNA pair must be provided that does not interact with the canonical ones. Consequently, a codon that is not assigned to a canonical amino acid, usually a stop codon, is also required. This genetic code expansion enables the incorporation of a noncanonical amino acid at a single, given site within the protein. The here presented work describes residue-specific incorporation where the genetic code is reassigned within the endogenous translational system. The translation machinery accepts the noncanonical amino acid as a surrogate to incorporate it at canonically prescribed locations, i.e., all occurrences of a canonical amino acid in the protein are replaced by the noncanonical one. The incorporation of noncanonical amino acids can change the protein structure, causing considerably modified physical and chemical properties. Noncanonical amino acid analogs often act as cell growth inhibitors for expression hosts since they modify endogenous proteins, limiting in vivo protein production. In vivo incorporation of toxic noncanonical amino acids into proteins remains particularly challenging. Here, a cell-free approach for a complete replacement of L-arginine by the noncanonical amino acid L-canavanine is presented. It circumvents the inherent difficulties of in vivo expression. Additionally, a protocol to prepare target proteins for mass spectral analysis is included. It is shown that L-lysine can be replaced by L-hydroxy-lysine, albeit with lower efficiency. In principle, any noncanonical amino acid analog can be incorporated using the presented method as long as the endogenous in vitro translation system recognizes it.

Residue-specific Incorporation of Noncanonical Amino Acids into Model Proteins Using an Escherichia coli Cell-free Transcription-translation System

An easy-to-use, cell-free expression protocol for the residue-specific incorporation of noncanonical amino acid analogs into proteins, including downstream analysis, is presented for medical, pharmaceutic, structural and functional studies.

The canonical set of amino acids leads to an exceptionally wide range of protein functionality. Nevertheless, the set of residues still imposes ...

Filtration Isolation of Nucleic Acids:  A Simple and Rapid DNA Extraction Method

FINA, filtration isolation of nucleic acids, is a novel extraction method which utilizes vertical filtration via a separation membrane and absorbent pad to extract cellular DNA from whole blood in less than 2 min. The blood specimen is treated with detergent, mixed briefly and applied by pipet to the separation membrane. The lysate wicks into the blotting pad due to capillary action, capturing the genomic DNA on the surface of the separation membrane. The extracted DNA is retained on the membrane during a simple wash step wherein PCR inhibitors are wicked into the absorbent blotting pad. The membrane containing the entrapped DNA is then added to the PCR reaction without further purification. This simple method does not require laboratory equipment and can be easily implemented with inexpensive laboratory supplies. Here we describe a protocol for highly sensitive detection and quantitation of HIV-1 proviral DNA from 100 &#181;l whole blood as a model for early infant diagnosis of HIV that could readily be adapted to other genetic targets.

We describe here a simple and rapid paper-based DNA extraction method of HIV proviral DNA from whole blood detected by quantitative PCR. This protocol can be extended for use in detecting other genetic markers or using alternative amplification methods.

FINA, filtration isolation of nucleic acids, is a novel extraction method which utilizes vertical filtration via a separation membrane and absorbent pad ...