We thank Stephan Uphoff for discussions. 
R.C. was supported by Linacre College, Oxford University. A.P. was supported by the German Academic Exchange Service (DAAD), the German National Academic Foundation and EPSRC. M.S. was supported by the Wellcome Trust. A.N.K. was supported by a UK BBSRC grant (BB/H01795X/1), and a European Research Council Starter grant (261227).

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The authors have nothing to disclose.

Many parameters can be varied during cell electroporation and data acquisition depending on the biological system of interest and the precise nature of the experiment (cell-level or single-molecule analysis). For example, when electroporating DNA into bacteria, 0.25 to 5 pmol of labeled dsDNA fragments leads to a low internalization efficiency, allowing direct single-molecule detection (i.e., without the need for photobleaching beforehand). Above 5 pmol dsDNA, cells tend to be heavily loaded, a regime better suited for single-cell analysis. All labeled DNAs should also be previously gel-purified in order to remove any trace of free dye (non-reacted fluorophore) from the DNA stock solution. Besides, potential issues with DNA degradation, particularly for smFRET experiments, can be addressed by using DNAs with unnatural nucleic acids, or motifs that protect exonuclease-accessible termini such as hairpin loops.
Another adjustable parameter in electroporation is the field strength applied during electroporation. Low field strength (~1 kV/cm) will lead to a low loading efficiency appropriate for single-molecule studies. Higher field strengths (up to 1.8 kV/cm) will increase the loading efficiency; however, there is an inverse correlation between field strength and cell viability after electroporation (see Figure 4). For reference, a normal field strength used for bacteria and yeast electroporation is ~1.5 kV/cm. The time constant, representing the length of this decay, is a convenient parameter to follow, since the time constant drops as soon as any arcing phenomenon occurs in the cuvette. Under normal settings, the time constant should be greater than 4 ms; lower values will lead to low loading efficiency or even non-loaded damaged cells. Most electroporators offer other degrees of freedom (such as &#8220;pulse truncation&#8221; or &#8220;pulse shape&#8221;) which can be modified to tune both cell loading and viability. We applied this method to both bacteria and yeast, however similar procedures should also allow the internalization of labeled biomolecules into mammalian cells using appropriate electroporator settings since their membrane is actually less complex (single lipid bilayer) and since electroporation has already been used with such cells 21.
When internalizing labeled proteins, all free dye needs to be removed from the labeled protein stock solution prior electroporation. Free dye molecules, due to their smaller size, can be internalized preferentially over the proteins of interest, and are difficult to discriminate during data analysis (despite their expected faster diffusion). As a guide, for a sample of organically labeled protein to be suitable for electroporation, the amount of remaining free dye should be below 2% (detected using fluorescent scanning of a SDS-PAGE)22. This process is particularly important, as some molecules might stick to the outer membranes of electroporated bacteria or yeast. In this respect, the negative control sample should display fluorescence intensity per cell clearly lower than electroporated cells, ideally as low as the autofluorescence level of empty cells (cells which have not been incubated with any fluorescently labeled biomolecules nor electroporated, Figure 2).
As with dsDNA, the internalization efficiency of labeled proteins is linked to the amount of biomolecules added to the cells prior to electroporation. However, other parameters, such as size and charge, play a role in internalization. Small proteins exhibit high internalization efficiencies, whereas larger proteins (up to 98 kDa) can be successfully internalized but with lower efficiency (Figure 5)26. The isolelectric point of the protein, potential interactions with the cell membrane and other physicochemical parameters also influence cell loading during electroporation. As a result, users need to optimize experiments for their own system, knowing that a high initial concentration of labeled protein (&#62;50 &#956;M) will give the best chance for successful loading. Electroporation also offers a new tool to perturb and analyze cellular function by introducing proteins and other biomolecules into cells (either labeled or unlabeled). The T7 RNA polymerase experiments (Figure 5C) present such an example of an experiment where we can introduce a biomolecule that can change gene expression in vivo using electroporation.
When performing single-molecule fluorescence experiments, TIR illumination is usually favored over other illumination modes as it offers the best signal-to-noise ratio by exciting only fluorophores within a thin section above the coverslip surface (~100 nm). However imaging labeled biomolecules diffusing inside living microorganisms might require deeper illumination (up to 0.8 &#181;m for E. coli). Deeper illumination is achieved in HILO mode, while preserving a high signal-to-noise ratio. On the other hand, wide field imaging is particularly important for step-wise photobleaching analysis, where the user is estimating the number of internalized molecules by photobleaching an entire loaded cell with high laser power and dividing the initial cell fluorescence intensity by the unitary intensity produced by a single molecule (single photobleaching step, Figure 3). Widefield imaging is also required for long-term molecule tracking in order to localize the diffusing molecules of interest even if their trajectories cover the entire cell volume.
In this protocol, we present how electroporation, a standard technique for biologists and biochemists for delivering nucleic acids in cells, constitutes a simple method for delivering fluorescent biomolecules in various cell types. This novel, high-throughput technique offers a unique tool to observe labeled molecules in their native environment. In addition biomolecules labeled with fluorophores covering a wide range of wavelengths, electroporation can deliver molecules modified with many chemical groups, such as unnatural nucleotides and amino acids, metal chelators, crosslinkers, and caging groups. If the biological system of interest is not essential to the cell development, the gene encoding for the target protein can also be deleted (or knocked-down), ensuring that the proteins observed after internalization represent all (or most) of the intracellular protein pool. In essence, electroporation can &#8220;transplant&#8221; the flexibility of in vitro bioconjugates into living cells and therefore benefit efforts in synthetic biology, systems biology, and in vivo single-molecule detection.

Most fluorescence studies inside living cells depend on protein fusions with fluorescent proteins (FPs), such as GFP1. These fluorescent tags allow studies of the copy number, diffusion pattern or localization of proteins involved in processes such as gene expression or membrane transport2-7. FPs offer high labeling specificity, easy implementation, and are available in a large inventory of variants with various photophysical and chemical properties1. However, organic fluorophores remain the prime choice for in vitro experiments due to their greater photostability (up to 100-fold more stable than FPs)8,9, small size (up to 100-fold smaller volume than FPs) and ease of intramolecular labeling (mainly through the use of cysteine residues). All these factors are particularly important for single-molecule fluorescence and FRET studies10.
Several internalization methods combining the advantages of organic labeling and in vivo detection have been introduced over the past decade; however, such methods either employ relatively large polypeptides tags (e.g., TMP, HALO, or 20 kDa SNAP tags)11-14, require the use of unnatural amino acids15, or are limited to large, single-membrane eukaryotic cells (e.g., scrape loading, syringe loading, microinjection)16-19.
This protocol describes a novel, straightforward and high-throughput internalization method that couples the advantages of organic fluorophores with in vivo observation. To develop this technique, we adapted the electroporation procedure commonly used to transform cells with plasmid DNA20,21 in order to load microorganisms, such as E. coli or S. cerevisi&#230; with organically labeled biomolecules. The protocol consists of 4 simple steps: incubation of cells with labeled biomolecules, electroporation, cell recovery, and cell washing to remove non-internalized biomolecules. Here, we present this electroporation protocol, as well as the cell imaging and data analysis processes to study cell-based and single-molecule fluorescence and FRET signals.
Electroporation relies on discharging a high-voltage electric field across a low ionic strength cell suspension to form transient membrane pores through which biomolecules can enter cells (Figure 1)20,21. Just as with transformation of bacteria or yeast with plasmid DNA, cells have to be prepared prior to electroporation to ensure their electrocompetency. This procedure, consisting of several washing steps with water, increases the membrane permeability and lowers the ionic strength of the cell solution to avoid arcing in the electroporation cuvette. In this protocol, cells can be prepared as described below (See PROTOCOL: 1.1) or bought from commercial providers.
<img alt="Figure 1" src="/files/ftp_upload/52208/52208fig1.jpg" /> 
Figure 1: Schematic representation of the internalization protocol. From left to right: add a few microliters of labeled biomolecules to the aliquot of electrocompetent cells (doubly-labeled DNA fragments and bacteria in this example); incubate 1 to 10 min&#160;on ice and transfer to a pre-chilled electroporation cuvette; electroporate and then add 0.5-1 ml rich medium to the cells immediately after; incubate at 37 &#176;C (or the temperature required by the organism, e.g., 29 &#176;C for yeast) to let the cells recover; perform 5 washing steps to remove any excess non-internalized labeled molecules; resuspend the final pellet in 100-200 &#956;l of PBS buffer and pipette 10 &#956;l on an agarose pad; cover the pad with a cleaned coverslip and image on a fluorescence microscope (in wide field mode or HILO mode).
Electrocompetent cells are incubated with the labeled biomolecules just before electroporation, which can be performed using standard electroporators found in most biochemistry laboratories. Immediately after electroporation, cells are incubated in a rich medium allowing their recovery before washing (Figure 1). The excess of non-internalized labeled biomolecules is first removed by washing in a buffer containing a fairly high concentration of salt and some detergent (See PROTOCOL: 3.3). The presence of salt disrupts non-specific electrostatic interactions formed by non-internalized labeled biomolecules which otherwise may stick on the outer membrane. Similarly, the presence of detergent in the washing buffer disrupts non-specific hydrophobic interactions.
While DNA internalization is straightforward (Figure 2), precautions need to be taken when internalizing labeled proteins using electroporation. First, the stock sample of organically labeled protein might still contain a small percentage of free dye. Free dye molecules are much smaller than proteins and might therefore be internalized preferentially. To ensure that the vast majority of the observed internalized fluorescent molecules correspond to the protein of interest, the initial protein sample should contain less than ~2% free dye (Figure 5)22. The excess of non-internalized labeled proteins can also stick to the outer cell membrane after electroporation; this phenomenon is protein-specific and needs to be checked for each new protein. We propose several options that allow the removal of non-internalized proteins from the loaded cell sample (See PROTOCOL: 3.3.3).
Finally, cells are resuspended in a small volume of phosphate buffer and pipetted onto an agarose pad, allowing their imaging on a fluorescence microscope. Immobilization on agarose pads is a simple and efficient way of imaging cells on a coverslip without harming their integrity. The pad should contain a low-fluorescence culture medium.
Cell imaging can be performed either in widefield, total internal reflection fluorescence (TIRF) or using HILO (Highly Inclined and Laminated Optical Sheet) microscopy. In the HILO configuration, the laser beam penetrates deeper into the specimen than in TIRF, yet does not illuminate the entire sample as for widefield, allowing a greater signal-to-noise ratio23. Depending on the laser power and time resolution used, internalized biomolecules can be counted (using stepwise-photobleaching analysis, Figure 3), localized, or tracked24-28. Internalization of doubly labeled constructs with a FRET pair of fluorophores allows the quantification of FRET both at single-cell or single-molecule levels (Figure 6).
Different parameters can be varied depending on the desired output and the biological system studied. First, the amount of internalized material per cell can be tuned by altering the concentration of labeled biomolecules added to the cells prior electroporation (Figure 2). Electroporation field strength will also influence both the loading efficiency and cell viability; as expected, while the loading efficiency increases with increasing field strength, the viability of electroporated cells decreases (Figure 4A). Both parameters can be quantified by recording the percentage of loaded and dividing cells after electroporation. This viability assay coupled with fluorescence imaging also verifies the observation of internalized biomolecules in living cells and allow continuous observation over several generations (Figure 4B).
In summary, this protocol allows the internalization of fluorescently labeled DNA and protein molecules into E.coli or S. cerevisi&#230;26. Individual molecules labeled with organic fluorophores can be tracked with high spatiotemporal resolution for timescales an order of magnitude longer than FPs. Finally, this method is compatible with widefield, TIRF and confocal detection, as well as pulsed excitation schemes, such as ALEX (alternating laser excitation28,29).

<table border="0" cellpadding="0" cellspacing="0" width="834">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Name of Material/ Equipment</td>
			<td style="width:227px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:321px;">ElectroMax DH5-alpha Comptent cells</td>
			<td style="width:227px;">Invitrogen</td>
			<td style="width:119px;">11319-019</td>
			<td>or any other commercial or lab-mage electrocompetant bacteria or yeast.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">EZ Rich Defined Madia</td>
			<td style="width:227px;">Teknova</td>
			<td style="width:119px;">M2105</td>
			<td>low fluorescence rich media</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MicroPulser Electroporation Apparatus&#160;</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">165-2100</td>
			<td>or any classical electroporator for microorganism transformation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Certified Molecular Biology agarose</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">161-3100</td>
			<td>low fluorescence agarose for agarose pad</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:321px;">Microscope coverslips No 1.5 thickness</td>
			<td style="width:227px;">Menzel</td>
			<td style="width:119px;">BB024060SC</td>
			<td>remove background particles by heating slides in furnace at 500 &#176;C for 1h</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Single-molecule fluorescence microscope</td>
			<td style="width:227px;">Home-built</td>
			<td style="width:119px;"></td>
			<td>described in REFs</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Localization software</td>
			<td style="width:227px;">Custom-written, available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB and C++ software package that can be adapted for localization analysis.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Tracking software</td>
			<td style="width:227px;">Available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB implementation by Blair and Dufresne.&#160;</td>
		</tr>
	</tbody>
</table>

internalization and observation of fluorescent biomolecules in living microorganisms via electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Sample preparation
The different steps of the protocol are presented as schematics in Figure 1. As an example, we represented the loading of bacteria with doubly labeled (donor and acceptor dyes) DNA fragments. Representative results for DNA internalization are shown in Figure 2. For each electroporated sample, data for empty cells and non-electroporated cells were also recorded (Figure 2). &#8220;Empty cells&#8221; correspond to electrocompetent cells neither incubated with fluorescent biomolecules nor electroporated; their intensity in the fluorescent channel reflects the autofluorescence level under identical experimental conditions (laser power, time resolution, temperature, etc.). &#8220;Non-electroporated cells&#8221; (also called &#8211;EP, i.e., minus EP) correspond to a negative control in which electrocompetent cells have been incubated with the fluorescent biomolecules but not electroporated. These non-electroporated cells should exhibit a fluorescence level similar to the autofluorescence of the empty cells and significantly lower than the fluorescence intensity displayed by loaded, electroporated cells. This confirms the removal of any non-internalized labeled biomolecules that could have adhered to the outer cell membrane.
<img alt="Figure 2" src="/files/ftp_upload/52208/52208fig2.jpg" /> 
Figure 2: Representative results for the internalization of dsDNA labeled with different fluorophores at different concentrations in bacteria (A-E) and yeast (F). Left to right: white-light, fluorescence and overlay images. -/+ EP denotes incubation without/with electroporation. Scale bars: 3 &#181;m. A. Cy3B dsDNA, 10 pmol, E. coli. B. ATTO647N dsDNA, 10 pmol, E. coli. C. Alexa647 dsDNA, 5 pmol, E. coli. D. Cy3B dsDNA, 100 pmol, E. coli. E. ATTO647N dsDNA, 100 pmol, E. coli. F. ATTO647N dsDNA, 30 pmol, Yeast. This figure has been modified from reference 26. <a href="https://www.jove.com/files/ftp_upload/52208/52208fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Counting the number of internalized biomolecules per cell
The procedure to estimate the number of internalized labeled biomolecules per cell using photobleaching analysis is presented in Figure 3 and Supplementary Movie 2, together with representative results obtained with different concentrations of labeled DNA. Cell loading efficiency increases with the initial amount of incubated labeled DNA, allowing the user to tune the number of labeled molecules per cell from a &#8220;single-molecule&#8221; level (&#60;10, Supplementary Movie 2B) to an &#8220;ensemble&#8221; level (&#62;10, Supplementary Movie 2A). A robust way of estimating the percentage of loaded cells is to count the number of electroporated cells displaying a cell intensity above the average cell intensity of non-electroporated cells plus 3 times their standard deviation (i.e., Av(I&#8211;EP)+3*StdDev(I&#8211;EP) where Av = average, I = intensity per pixel, Std. Dev. = standard deviation and &#8211;EP = non-electroporated) as shown in Figure 3.
<img alt="Figure 3" src="/files/ftp_upload/52208/52208fig3.jpg" /> 
Figure 3: Counting the number of internalized molecules using photobleaching analysis. (A) Single-cell photobleaching analysis. Examples of fluorescence intensity timetraces (blue: raw data;&#160;red: fit; insets:&#160;WL&#160;and&#160;fluorescence images&#160;of&#160;E.&#160;coli&#160;loaded with ATTO647N-labeled dsDNA before and after bleaching). Top: single-step bleaching event. Middle: cell containing &#177;3 molecules showing bleaching and blinking. Bottom: cell containing &#62;10 steps corresponding to at least 10 molecules. (B) Histogram of single-step height intensities from an automated step-fitting algorithm from 57 cells containing less than 6 distinguishable steps. Single-Gaussian fit is centered at 11 &#177; 3 a.u., corresponding to a unitary fluorophore intensity of 8100 photons per second. The asterisk marks the bin gathering all the step heights above or equal 50 a.u. (C) Histogram of internalized molecules per cell electroporated with different amounts of ATTO647N dsDNA, calculated after dividing the initial fluorescence intensity by the unitary fluorophore intensity. Top to bottom: empty cells (i.e., not incubated with fluorescent molecules and not electroporated), non-electroporated (but incubated with fluorescent molecules, named &#8211;EP), and electroporated cells incubated with 10 and 100 pmol dsDNA (named +EP). Empty and non-electroporated cells correspond to autofluorescence, whereas electroporated cells show a broad distribution of internalized molecules, with a higher proportion of highly loaded cells at 100 pmol (&#8805; 4 molecules, see asterisk-marked bin). Internalization efficiency (fraction of cells with Int. &#62; mean + 3x Std. Dev. of non &#8211;EP sample) for the 10 and 100 pmol samples was 94% and 90%, respectively. Mean number of internalized molecules per cell: 121 &#177; 106 molecules for 10 pmol dsDNA, and 176 &#177; 187 molecules for 100 pmol dsDNA. Settings: 100 ms exposure, widefield illumination. Scale bars: 1 &#956;m. This figure has been modified from reference 26. <a href="https://www.jove.com/files/ftp_upload/52208/52208fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Cell loading and viability
In addition to changing the amount of labeled biomolecules added to the cells prior to electroporation, the user can tune the amount of internalized molecules by choosing different field strengths during electroporation (Figure 4, Supplementary Movie 1). Higher field strengths lead to greater internalization efficiencies, but lead to a slight decrease of cell viability. For protein internalization, the use of a filtration step may help remove non-internalized labeled proteins (see 3.3.3.1.1). In such cases, cell filtration ensures that observed fluorescent proteins are indeed internalized inside the bacterial cytoplasm; we note, however, that filtration also has a negative impact on cell viability (for more details, see REF 22).
<img alt="Figure 4" src="/files/ftp_upload/52208/52208fig4.jpg" /> 
Figure 4: Influence of electroporation voltage upon cell loading and viability. (A) Bar chart representing the effect of electroporation field strength on loading efficiency (red bars) and viability of the cells (green bars). 84 &#177; 8 % of the non-electroporated cell (0 kV/cm) divide after 1 h on an agarose pad at 37 &#176;C. Under the same conditions, 78 &#177; 3 % and 49 &#177; 3 % of the cells electroporated at 0.9 kV/cm and 1.8 kV/cm respectively divide after 1 hr. For loading efficiency, 73 &#177; 8 % of the cells are loaded at 0.9 kV/cm, while 92 &#177; 6 % of the cells are loaded at 1.8 kV/cm. The error bars represent the standard deviation calculated from three independent measurements; more than 200 cells were analyzed for each sample and each repeat. Overall loading efficiency increases with electroporation voltage to the slight detriment of cell viability. (B) Cell-based fluorescence measurements over several generations show that the overall fluorescence intensity is shared equally between both daughter cells. Cell 1 and 2 refers to the cell number in the white light image (left) and fluorescence image at t = 0. Scale bar: 1 &#181;m). This figure has been modified from reference 26.
Protein internalization
Representative results for protein internalization are in Figures 5A &#38; B. It is particularly important to remove as much of the remaining free (unreacted) dye from the protein sample as possible prior to electroporation. In the examples in Figures 5A &#38; B, the Cy3b-labeled Klenow Fragment sample (Cy3b-KF, where KF is the Klenow fragment of the E. coli DNA Polymerase I, 66 kDa) only contains 1% free dye; such dye contribution to the overall cell loading is negligible. Comparisons of the electroporated sample of interest with both the non-electroporated cells (incubated with the same amount of labeled protein) as well as cells electroporated with the equivalent amount of free dye constitute two required controls to ensure that the observed fluorescent molecules are indeed internalized labeled proteins.
<img alt="Figure 5" src="/files/ftp_upload/52208/52208fig5.jpg" /> 
Figure 5: Protein internalization in live bacteria. (A) Representative fluorescence overlay field of views. Cells electroporated at 1.4 kV voltage with 50 pmol RNAP &#969; subunit from a protein stock solution that contained only 1% free Cy3b dye. Non-electroporated (non &#8211;EP) and empty cells are defined as previously. Free dye was internalized at the same concentration as in the RNAP &#969; electroporated sample. Imaging in wide field mode, 532-nm excitation at 1 mW, 50 ms exposure. (B) Distribution of uncorrected cell-averaged intensities for samples in (A), given in proportion of the total cell count. More than 400 cells per sample were segmented. This figure has been modified from reference 22. (C) Internalization of unlabeled T7 RNA polymerase (T7 RNAP, 98 kDa) into electrocompetent DH5&#945; carrying the pRSET-EmGFP plasmid encoding emerald GFP (EmGFP) under control of a T7 promoter. Left: Schematic of assay. Middle: fluorescence overlay. Right: histograms of cell-based fluorescence intensities for the non-electroporated sample (top) and cells incubated and electroporated with T7 RNAP (bottom); approximately 11% of the electroporated cells show high fluorescence intensity (fraction of cells with Int. &#62; mean + 3x Std. Dev. of non &#8211;EP sample) indicating expression of EmGFP. The asterisks indicate bins gathering all the intensities above or equal 1,100 a.u. Scale bar: 3 &#956;m. This figure has been modified from reference 26.
Figure 5C presents another application of protein electroporation. Here, the electroporated protein is unlabeled, but its internalization triggers an observable fluorescent response. This experiment verifies the presence and functionality of electroporated proteins in the cell cytoplasm. Unlabeled T7 RNA polymerase (98 kDa) was internalized into E. coli strain DH5&#945; containing a plasmid encoding for fluorescent protein EmGFP under the control of a T7 promoter26. As the gene for T7 RNA polymerase is absent in DH5&#945;, EmGFP expression in our experiments requires that functional T7 RNA polymerase is introduced into the cells via electroporation (Figure 5C). Following electroporation with 1 pmol T7 RNAP, &#62;11% of the cells (blue bar, Figure 5C) exhibited fluorescence higher than the negative control (incubated with the same amount of T7 RNAP, but not electroporated). This result establishes that a proportion of the T7 RNAP molecules internalized by electroporation retain their integrity in vivo and can perform their intended functions in the cell cytoplasm.
In vivo FRET at the single-molecule and single-cell levels
Finally, the internalization and analysis of doubly labeled species in living bacteria is presented in Figure 6 and Supplementary Movie 3. As fluorescent protein fusions are not ideal for in vivo smFRET studies, the ability to deliver doubly labeled biomolecules into living cells using electroporation is one of the great assets of this method. Figure 6A presents single-cell FRET analysis of bacteria loaded with different FRET DNA standards (using Cy3B and Atto647N fluorophores as donor-acceptor FRET pair). Cells are electroporated with 20 pmol of three short doubly-labeled dsDNA FRET standards with apparent FRET efficiencies (E*) of 0.17, 0.48, and 0.86 in vitro (previously determined26). All DNAs enter cells efficiently (Figure 6A, left) and the main peak of each single-cell E* distribution agrees well with the in vitro results (Figure 6A, right). In the intermediate- and high-FRET samples, cell populations with lower E* than expected are observed, presumably due to a combination of acceptor bleaching and photophysical inactivity, variable cell loading (thus, variable signal-to-noise ratio) and DNA degradation.
<img alt="Figure 6" src="/files/ftp_upload/52208/52208fig6.jpg" /> 
Figure 6: Representative results for single-cell and single-molecule FRET observation in living bacteria. Ensemble and smFRET studies in single bacteria. (A) Analysis of cells loaded with 20 pmol each of three DNA FRET standards exhibiting low (~0.17), intermediate (~0.48), and high (~0.86) FRET (as measured using in vitro single-molecule measurements; see REF 26). Left: white light and green/red (FRET) fluorescence overlay images (Scale bar: 3 &#956;m). Examples of FRET values from different cells are indicated (white). Right (top to bottom): uncorrected cell-based FRET (E*) histograms for donor only (dark green), low (light green), intermediate (yellow), and high (red) FRET DNA standards. (B&#8211;D) In vivo smFRET. Cells are loaded with 0.25 pmol intermediate-FRET DNA (panel B), 0.25 pmol high-FRET DNA (panel C), and 5 pmol doubly-labeled KF (panel D). Left column: green/red fluorescence overlay of single frame before and after acceptor photobleaching. Middle column: time traces corresponding to the molecule in the yellow circle. FRET efficiencies, donor emission intensities, and acceptor emission intensities are displayed in blue, green, and red, respectively. Right column: FRET histograms of donor only molecules (green) and donor-acceptor molecules (yellow, red, and gray) from 20 time traces for each sample. Scale bars: 3 &#956;m for A, 1 &#956;m for B&#8211;D. This figure has been modified from reference 26. <a href="https://www.jove.com/files/ftp_upload/52208/52208fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To observe smFRET in vivo for DNA or protein samples, low amounts (0.25 pmol) of intermediate- and high-FRET DNA standards (Figure 6B, C) or 5 pmol of double-labeled KF (Alexa647/Cy3B fluorophores as FRET pair, Figure 6D) are electroporated into E. coli. Such concentrations led to many cells loaded with few (n = 1&#8211;10) labeled molecules, allowing direct localization, tracking, and FRET monitoring for single molecules. Some molecules diffuse freely, whereas others appear immobile or diffuse slowly (Supplementary Movie 3). Timetraces of immobilized doubly-labeled biomolecules (Figure 6, middle) last for 1 to 30 s and show the hallmarks of smFRET: anticorrelated changes in the donor and acceptor fluorescence upon acceptor bleaching (for example, t~16 sec; Figure 6B, middle), followed by single-step donor bleaching (for example, t~;19 sec; Figure 6B). FRET distributions generated from such timetraces (Figure 6, right) result in a mean value that is in excellent agreement with published in vitro studies26,31,32. These results establish the capability for quantitative smFRET studies on internalized DNAs and proteins, and suggest that proteins maintain their integrity and structure upon electroporation and internalization (as supported by the T7 RNAP internalization experiments).
Supplementary Movie 1: Cell viability. Left: White light images. Right: fluorescence images. Animated GIF animated showing the division after electroporation (1.8 kV/cm) of bacteria loaded with 10 pmol Atto647 labeled DNA. The overall apparent decrease of fluorescence is due to the dilution of labeled DNA upon cell division and also partially to the photobleaching which occurs during each measurement.
Supplementary Movie 2: Cell-based photobleaching studies. A. Representative example of a heavily loaded cell (containing &#62;100 Atto647N-labeled DNA molecules). Top left, white light image of the cell of interest (red rectangle). Top right, movie of the loaded cells showing their fluorescence decay over several minutes. Bottom, time trace of the overall fluorescence intensity decay of the cell of interest. Organic fluorophores can exhibit photobleaching lifetimes 2 orders of magnitude higher than FPs (here, ~41 sec for Atto647N). B. Representative example of a cell loaded with less than 10 labeled molecules (3 in this case). Top, same as panel A. Bottom, time trace of the overall fluorescence intensity of the cell of interest showing single step bleaching and/or blinking corresponding to single organic fluorophores. The average height of these steps corresponds to the single-molecule unitary intensity (here ~12 a.u.) used to estimate the initial number of internalized molecules per cell. Movies under continuous red laser excitation at 300 &#181;W power and 100 msec per frame.
Supplementary Movie 3: In vivo single-molecule FRET. Top: Cells loaded with 0.25 pmol high-FRET DNA (as in Figure 6C) continuously monitored at 50 ms per frame under nTIRF illumination using 1 mW green (532 nm) laser. Each frame is a green/red (FRET) fluorescence overlay of each channel. Diffusing and immobile red (intact) and green (single active label) DNA molecules can be observed. Bottom: Time trace corresponding to the molecule in the yellow circle. FRET efficiencies, donor emission intensities, and acceptor emission intensities are displayed in blue, green, and red, respectively. Anti-correlated acceptor bleaching event (red-to-green transitions) corresponds to the signature of single-molecule FRET.

Watch this Scientific Journal Video about Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation at JoVE.com

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

internalization-observation-fluorescent-biomolecules-living

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16.7K Views.  University of Oxford. Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

Video: Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and...

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Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

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This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

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Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron (dgn)-destabilized Green Fluorescent Protein (GFP)-based Reporter Protein

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. Maintenance of proteasome activity was implicated in many key cellular processes, like cell's stress response 3, cell cycle regulation and cellular differentiation 4 or in immune system response 5. The dysfunction of the ubiquitin-proteasome system has been related to the development of tumors and neurodegenerative diseases 4, 6. Additionally, a decrease in proteasome activity was found as a feature of cellular senescence and organismal aging 7, 8, 9, 10. Here, we present a method to measure ubiquitin-proteasome activity in living cells using a GFP-dgn fusion protein. To be able to monitor ubiquitin-proteasome activity in living primary cells, complementary DNA constructs coding for a green fluorescent protein (GFP)&#x2013;dgn fusion protein (GFP&#x2013;dgn, unstable) and a variant carrying a frameshift mutation (GFP&#x2013;dgnFS, stable 11) are inserted in lentiviral expression vectors. We prefer this technique over traditional transfection techniques because it guarantees a very high transfection efficiency independent of the cell type or the age of the donor. The difference between fluorescence displayed by the GFP&#x2013;dgnFS (stable) protein and the destabilized protein (GFP-dgn) in the absence or presence of proteasome inhibitor can be used to estimate ubiquitin-proteasome activity in each particular cell strain. These differences can be monitored by epifluorescence microscopy or can be measured by flow cytometry.

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron dgn-destabilized Green Fluorescent Protein GFP-based Reporter Protein

A method to monitor ubiquitin-proteasome activity in living cells is described. A degron-destabilized GFP- (GFP-dgn) and a stable GFP-dgnFS fusion protein are generated and transduced into the cell using a lentiviral expression vector. This technique allows to generate a stable GFP-dgn/GFP-dgnFS expressing cell line in which ubiquitin-proteasome activity can be easily assessed using epifluorescence or flow cytometry.

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. ...

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific properties, such as binding affinity, agonist or antagonist behavior, or stability, relative to the native sequence. Protein design lies at the center of current advances drug design and discovery. Not only does protein design provide predictions for potentially useful drug targets, but it also enhances our understanding of the protein folding process and protein-protein interactions. Experimental methods such as directed evolution have shown success in protein design. However, such methods are restricted by the limited sequence space that can be searched tractably. In contrast, computational design strategies allow for the screening of a much larger set of sequences covering a wide variety of properties and functionality. We have developed a range of computational de novo protein design methods capable of tackling several important areas of protein design. These include the design of monomeric proteins for increased stability and complexes for increased binding affinity.
To disseminate these methods for broader use we present Protein WISDOM (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>), a tool that provides automated methods for a variety of protein design problems. Structural templates are submitted to initialize the design process. The first stage of design is an optimization sequence selection stage that aims at improving stability through minimization of potential energy in the sequence space. Selected sequences are then run through a fold specificity stage and a binding affinity stage. A rank-ordered list of the sequences for each step of the process, along with relevant designed structures, provides the user with a comprehensive quantitative assessment of the design. Here we provide the details of each design method, as well as several notable experimental successes attained through the use of the methods.

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

We developed computational de novo protein design methods capable of tackling several important areas of protein design. To disseminate these methods we present Protein WISDOM, an online tool for protein design (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>). Starting from a structural template, design of monomeric proteins for increased stability and complexes for increased binding affinity can be performed.

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific ...

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

Tracking Drug-induced Changes in Receptor Post-internalization Trafficking by Colocalizational Analysis

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and overall cell responsiveness to ligands and is, itself, influenced by intra- and extracellular conditions, including ligand-induced signaling. Optimized for use with monolayer-plated cultured cells, but extendable to free-floating tissue slices, this protocol uses immunolabelling and colocalizational analysis to track changes in intracellular receptor trafficking following both chronic/prolonged and acute interventions, including exogenous drug treatment. After drug treatment, cells are double-immunolabelled for the receptor and for markers for the intracellular compartments of interest. Sequential confocal microscopy is then used to capture two-channel photomicrographs of individual cells, which are subjected to computerized colocalizational analysis to yield quantitative colocalization scores. These scores are normalized to permit pooling of independent replicates prior to statistical analysis. Representative photomicrographs may also be processed to generate illustrative figures. Here, we describe a powerful and flexible technique for quantitatively assessing induced receptor trafficking.

Receptor trafficking modulates signaling and cell responsiveness to ligands and is, itself, responsive to cell conditions, including ligand-induced signaling. Here, we describe a powerful and flexible technique for quantitatively assessing drug-induced receptor trafficking using immunolabeling and colocalizational analysis.

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. To improve the mating efficiency of B. Xylophilus and to increase the chances of mating observation, virgin adults were cultured and used for the investigation. Eggs were obtained by keeping the nematodes in water and allowing the females to lay eggs for 10 min. The second-stage juveniles (J2) were synchronized by incubating the eggs for 24 h at 25 &#176;C in the dark, and the early J4 were obtained by culturing the J2 with grey mold, Botrytis cinerea, for another 52 h. At this time point, most J4 nematodes could be clearly distinguished as being male or female using their genital morphology. The male and female J4 were collected and cultured separately in two different Petri dishes for 24 h to get virgin adult nematodes. A virgin male and a virgin female were paired in a drop of water in the well of a concave slide. The mating behavior was filmed with a video recorder under a stereomicroscope. The whole period of the mating process was 82.8 &#177;3.91 min (mean &#177;SE) and could be divided into 4 different phases: searching, contacting, copulating, and lingering. The mean minutes of duration were 21.8 &#177; 2.0, 28.0 &#177; 1.9, 25.8 &#177; 0.7 and 7.2 &#177; 0.5, respectively. Eleven sub-behaviors were described: cruising, approaching, encountering, touching, hooping, locating, attaching, ejaculating, separating, quiescence, and roaming. Interestingly, obvious intra-sexual competition was observed when one female was grouped with 3 males or one male with 3 females. This protocol is useful and valuable, not only in investigating the mating behavior of B. xylophilus, but also in acting as a reference for ethological studies of other nematodes.

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A protocol for investigating the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, is presented. Behavioral features of B. xylophilus are described in the mating process.

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. ...