Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The goal of this procedure is to internalize and visualize biomolecules such as DNA or proteins labeled with organic Fluor inside living microorganisms using a method based on electroporation. This is accomplished by first incubating electro competent cells with fluorescently labeled biomolecules before transferring them into an electroporation vete. By applying a high voltage across the vete, transient pore formations on both the cell wall and cell membrane will enable biomolecular diffusion into the cells.

Next, the cells are incubated in a rich medium to allow them to recover before the excess non internalized biomolecules are removed. Finally, the cells are transferred onto a low fluorescence minimal nutrient arose pad, and then a glass cover slip is overlaid on top to form a glass agro sandwich. Ultimately, thees essence microscopy is used to analyze the location, diffusion pattern, and dynamical properties of the labeled biomolecules inside the living cells.

We first had the idea for this method when we realized the limitations in live cell imaging. Due to the fast glitching of photo fluorescent proteins such as GFP, we reasoned that by loading living cells with biomolecules labeled with organic fluoro force, we would transplant in living microorganisms. The main advantages of this force that is brightness, photo stability, and small size.

We then hope to follow the localization and diffusion of DNA or proteins in their native environments on much longer timescales, which would match many important biological processes. The main advantage of these techniques over existing methods, such as microinjection, is that it can also be applied to microorganisms that are smaller than the diameter of typical microinjection needle. An additional advantage of electroporation is that a large number of cell can be loaded at the same time with fluorescent biomolecules and visualize in parallel those making electroporation a particularly eye throughput technique.

Although this method can provide insight into the diffusion and dynamics behavior of fluorescently labeled biomolecules in vivo, it can also be applied in other contexts such as to electrode biomolecules that are not labeled, but it can, for example, trigger a physiological response when internalized by the cell To construct aros pads for bacteria or yeast fluorescent microscopy begin by preparing some two x low fluorescence media, then melt a solution of 2%aros in distilled water with a lab microwave promptly. Add one matching volume of the hot HA rose to the media and gently pipette the mixture up and down. Avoiding bubble formation immediately pipette approximately 200 microliters of the arose mixture onto a standard size microscope cover slip.

Leave the cover slip on the bench for a minute at room temperature until the gel solidifies or temporary storage place a second cleaned cover slip over the fresh pad and gently apply pressure from the top to flass in the egg rose, starting with a stock of fluorescent biomolecules prepared in a low ionic strength buffer. Transfer up to five microliters of the biomolecules into either a 20 microliter aliquot of competent bacteria, or 50 microliters of competent yeast cells. The amount of fluorescent biomolecules incubated with the cells directly correlates with the amount of biomolecules internalized per cell.

After electroporation, incubate the mixture on ice for up to 10 minutes. At the same time, select an electroporation vete of appropriate electrode spacing and chill on ice. Transfer the cell mixture into the pre chilled vete.

Gently tap the vete a few times to remove any bubbles from the solution. Remove any moisture from the vete with a paper towel and place the vete into the electro ator. Apply a high voltage pulse to the cells just after this process.

Verify that the time constant displayed on the electro ator is between four and six milliseconds. Lower time constants are often due to the presence of excess salt or bubbles within the qve, and it may decrease or completely inhibit uptake of biomolecules into the cells. Higher voltage pulses improve the cell loading, but can also affect the cell viability immediately after electroporation.

Add 500 microliters of rich media and transfer the cells into a fresh 1.5 milliliter tube. Let the cell membrane recover by incubating the bacteria of 37 degrees Celsius or yeast a 29 degrees Celsius for two to 10 minutes. After biomolecule incorporation and growth recovery, spin down the cells for one minute in a four degrees Celsius pre chilled centrifuge and discard the supine agent Reese.

Suspend the pellet in 500 microliters of buffer and repeat the spin down sate and removal and buffer resus suspension processes. Three more times. These wash steps will remove all traces of non internalized biomolecules from the cell suspension.

In the case of labeled protein internalization and additional purification step is performed by pipetting the cell suspension into a 1.5 milliliter tube fitted with a 0.22 micron membrane filter on top. Separate the cells from the buffer by centrifugation while ensuring there's enough liquid remains above the filter to avoid the cells drying out and discard the flow through fraction. Below the filter, add 500 microliters of fresh PBS over the cells and repeat the combination of a spin down followed by fresh PBS edition.

Two more times. Perform a final spin down at 3, 300 G for one minute. To remove the previous PBS wash, re suspend the cells with 150 microliters of PBS.

Finally, remove the upper cover slip from the previously made aros pad. Load the cells onto the pad by pipetting 10 microliters of the cell suspension dropwise. After placing a new cleaned cover slip on top of the arose pad, the sandwich cells are now ready for fluorescent microscopy.

Negative controls such as non-rated controls, which have been incubated with labeled biomolecules or empty cells, which haven't been electro rated nor incubated, should also be prepared in parallel with any loaded samples, loaded cells can be imaged on any appropriate fluorescence microscope. While wide field imaging mode allows cell level imaging and analysis, turf or high low illumination or optimal for single molecule observation and tracking. To prevent excess photobleaching of the biomolecules, please make sure all on stage laser illuminations are rather turned off or blocked with an opaque filter prior to sample loading.

Furthermore, switch off the E-M-C-C-G camera gain to prevent over exposure damage to the camera. For an inverted microscope setup, place the Aris pad sandwich onto the microscope stage with the cell covered side facing downwards towards the bottom objective. Then turn on the top side halogen light and bring the cells into focus under transmitted white light microscopy mode prior to turning on the laser, take a photograph the cells under white light transmission mode.

This photo will be used as a positional reference map that will correlate between regions of biomolecular fluorescence to actual locations of cells within the same field of view. For viability measurement, use an objective heater set at the optimum temperature and record sequential images of the same field of view under white light illumination every 30 minutes without moving the stage, turn off the top topside illumination and protect your sample from all other ambient light sources in the lab. Then switch on the CCD camera and begin data acquisition.

Start recording the fluorescent ion just before switching on the laser illumination. After performing fluorescent microscopy, begin the data analysis by opening all photographs with an image processing software such as Image J to start normalize all fundamental image parameters such as brightness or contrast across all images by pressing the set button and select the propagate to all other images. Option one can qualitatively ascertain if the cell loading was successful as loaded cells should be significantly brighter than the non electroporated cells used as a negative control for a given image.

Use the free hand selection tool and encircle different regions of fluorescence with hand-drawn polygons. Compare the fluorescence intensity among all pertinent polygons by choosing the measurement option from the analyze menu. Repeat this process manually or using an automated script to extract the intensity per pixel of all the cells in each sample and negative controls.

When an electro rated cell is loaded with one or several fluorescent biomolecules, its average emission intensity will be larger than the average intensity of a negative control sample. To evaluate the cell viability manually count the percentage of dividing intact by non dividing and damaged cells. In the same field of view for which data were recorded every 30 minutes over several divisions, it isn't always possible to count the number of biomolecules internalized per cell directly, particularly when the biomolecules diffuse fast or when cells are loaded with more than five labeled biomolecules.

Cell-based photobleaching analysis allows the evaluation of the number of labeled biomolecules per cell in such conditions. Using the freehand selection button, trace the outline of a given fluorescent cell and extract and plot the raw cell fluorescence as a function of time due to photobleaching effects. The intensity versus time graph will be an exponentially decreasing curve, the heavily loaded cells, but for cells containing less than 10 biomolecules, clear steps should be distinguishable.

Calculate the average autofluorescence after photobleaching by averaging the raw fluorescent intensity values near the flat tail end portion of the experiment, which could be, for example, from the last 50 to 100 image frames in the time-lapse series. After determining the baseline also intensity, calculate the time dependent photobleaching intensity by subtracting the baseline autofluorescence from the raw fluorescence data, then block the resulting photobleaching intensity data and quantize the height of each step manually or using an algorithm. This process should be repeated for several cells exhibiting more than 100 steps overall in order to get an accurate estimation of the average step height, this value corresponds to the unitary fluoro four intensity due to bleaching of a single fluoro four.

Finally, the number of internalized molecules per cell can be calculated by dividing the baseline subtracted cell intensity at time equals zero by the unitary fluoro. Four intensity for this electroporation protocol. The amount of fluorescently labeled DNA or protein that can be internalized into bacteria or yeast can be controlled by adjusting the amount of DNA probes present within the electroporation buffer.

When the biomolecular internalization rate is high, the photobleaching lifetime of the labeled biomolecule, which can be found by first plotting the fluorescent intensity as a function of time and then extracting its exponential time constant gives information about the photo stability of the dyes in vivo. In the case of low incorporation rates, quantization of the fluorescent intensity will provide a means for approximating the total number of biomolecules within a given cell. Electroporation can also be used to incorporate proteins into bacteria or yeast.

In the case of fluorescently labeled protein. Since free fluorophores themselves can be internalized very efficiently, it is important to remove all excess free and non covalently attached fluorophores from the protein stock. Finally, doubly labeled by our molecules can also be internalized using our method to allow single molecule fret observation in living cells over several seconds.

While attempting this procedure, it is important to remember to also prepare and analyze the control samples such as non electro cells, which are incubated with a labeled biomolecule or empty cells, which are neither electroporated nor incubated with a labeled molecule. These controls allow us to evaluate the level of cell autofluorescence and deficiency of washing steps, and as such are necessary to assess whether the electroporation experiment has been successful After its developments. This technique opened new avenues for researchers in the field of biophysics and third biology to explore the diffusion and dynamical behavior of label biomolecule in living microorganisms.

And this on timescale one to two orders of magnitude longer than with autofluorescent proteins such as GFPs. It transferred the flexibility of organic d labeling in vivo making single molecule threat experiments much more feasible for the study of intra and intermolecular dynamics in living cells.