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The overall goal of the following experiment is to develop new fluorescent sensors based on Dun drummers with improved properties for pH imaging in vitro in living cells and in vivo. This is achieved by first conjugating pH sensitive dyes to a dendritic scaffold together with other moieties, such as targeting groups or pH in sensitive dyes. And in vitro measurements are taken to generate a pH calibration curve next to image pH in living cells.

The sensor is delivered by electroporation and pH maps are generated using confocal microscopy to image pH in the extracellular space of the brain. During physiological and pathological neural activity, the sensor is micro injected into the brain of anesthetized mice. Fluorescence data is then acquired using two photon microscopy.

Results obtained show that drimmer based sensors succeed to measure pH changes with high spatial and temporal resolution, both in living cells and in vivo. Although several flues dyes with pH sensitive behavior already been employed, the suffer of several drawbacks, such as poor mobility and cell leakage. Moreover, they do not allow the rash metric absolute measurement.

Here we demonstrated the conation of pH sensitive dyes to D genetic scaffold, and as the performances allow an accurate measurement with the special and temporal resolution, this method could be a useful tool to study physiological and pathological process regulated by pH in living cells. The disruption on the pH regulation in the neuro system is associated with several condition like stroke and epilepsy, and available pH sensor proved to be poorly effective leading to the design of the presented sensor. Though this met has been here applied pH h imaging, it is a general approach and can be applied to other biological relevant species.

By choosing the profit sensing dye, the easy and versatile synthetic approach makes these procedures straightforward. Demonstrating the procedure today Will be Giovanni re, Barbara ti, Marco Brody, and Sebastian Suli To conjugate pH indicators to Piam Demers. Begin by dissolving the rimer in anhydrous DMSO at a final concentration of 50 micromolar.

Prepare 10 micromolar stock solutions of fluorescein NHS and Tetraethyl RUMINE NHS, also known as TMR in anhydrous, DMSO in a micro centrifuge tube combined, one milliliter of G four peram dederer solution with eight equivalents, or 40 microliters of fluorescein and eight equivalents, or 40 microliters of TMR. The molar ratio in the mixture will reflect the amount of dyes loaded on the demer. Stir the solution at room temperature for 12 hours.

Next, dilute the mixture one to 10 with deionized water and load the reaction mixture in a dialysis bag with a molecular weight cutoff of 10 kilodaltons dialyze against deionized water and replace the water in the reservoir three times over 24 hours. The next day, transfer the solution to a glass vial and freeze it at minus 80 for a few hours. When the solution is completely frozen, transfer the vial to a lyophilizer and freeze dry for 24 hours.

A purple powder should be obtained, wavy obtained solid, and dissolve it in mqe water. At a final concentration of 10 micromolar. Dispense the solution in 50 microliter aliquots in 1.5 milliliter micro centrifuge tubes, and store them at minus 20 degrees Celsius.

For in vitro calibration, prepare a 500 nanomolar solution of D drummer in two millimolar PBS In a quartz Q vet. A very dilute PBS buffer is used to avoid abrupt changes of pH During the titration. Measure the emission spectra of fluorescein and TMR and optimize the optical settings of the fluorimeter to achieve a good signal to noise ratio.

Next, perform a pH titration by adding small volumes of 0.1 normal sodium hydroxide and 0.1 normal hydrochloric acid. After every addition, shake the Q vet to mix. Wait one minute for equilibration and then measure the pH by means of a pH.

Micro electrode emission spectra of fluorescein and TMR should be recorded for every step without any change. In the optical settings, plot the fluorescence intensity versus pH. For the titration, the rod domine signal should be unaffected by pH.

The fluorescein signal should be a sigmoidal curve and should be fitted with a single binding model with a PK of 6.4 for electroporation. Transfer six times 10 to the fifth hela cells in 72 microliters to a 1.5 milliliter micro centrifuge tube to the resuspended cells. Add dendri or aqueous solution at a concentration of two micromolar, add three milliliters, electroporation buffer to a micro puration tube.

Then using a 10 microliter electroporation tip aspirate the cell and rimer mixtures. Next, insert the micro pipette into the pipette station and set the micro puration pulse conditions Here. The pulse voltage is set to 1005 volts.

The pulse width is set to 35 milliseconds, and the pulse number is set to two after the pulse plate. 20 microliters of the electroporated cells onto glass bottom dishes with fresh medium without antibiotics. 12 hours after electroporation image the cells with a confocal microscope using a standard filter set for fluorescein and r domine.

Focus on the specimen and adjust laser power and detector gain to maximize the signal to noise ratio. If the electroporation was successful, cells should be brightly fluorescent in both channels. The localization depends on the size and charge of the D are used.Often.

Sun lysosomal localization, which is seen as small perinuclear vesicles is present due to endocytosis or com compartmentalization. If lysosomal localization is predominant in which most of the fluorescence is localized inside vesicles and pore signal is observed in the cytosol. This signifies toxicity and measurements should be discarded.

Acquire the two channels sequentially, acquire several images and average the images to improve image quality. To generate a pH calibration curve, use buffers with opfor at different phs to clump the intracellular and extracellular pH at the same value. Acquire at least 20 cell images per pH and measure at least five points from pH 5.5 to pH 7.5.

It reminds the pH B six are toxic to cells but can be tolerated for a short amount of time, so acquire images as short as possible. If you see sign of apoptosis, discard the cells and restart. Following acquisition.

Use image J or analogous software for data analysis. First, determine the green to red ratio. Then plot the ratio versus pH to produce a linear trend that will give a calibration curve that can be used to convert ratio to pH in future experiments.

For in vivo sample preparation, begin with C 57 black six J and female mice between postnatal day 28 and 70. After anesthetizing the mice with urethane, perform a toe pinch to ensure that the animal is fully anesthetized and apply eye ointment to reduce the cortical stress response and cerebral edema. During the surgery, inject two milligrams per kilogram body weight of dexamethasone, sodium phosphate intramuscularly.

Next, using surgical shears, shave the animal's head, then place a surgical drape and apply 2.5%lidocaine gel to the scalp. Use scissors to cut the flap of skin covering the skull of both hemispheres. Wash the exposed bone with saline and with forceps, gently remove the periosteum.

This will provide a better adhesion surface. For glue and dental cement, apply a custom made steel head post with a central imaging chamber and using cyanoacrylate, glue it into place in a plain approximately parallel with the skull over the cortical region of interest. Fix the post into place with white dental cement.

Fix the head of the mouse and perform a craniotomy of a two to three millimeter diameter by drilling over the region of interest. Frequently throughout the drilling procedure uses syringe to apply fresh sterile A CSF to the skull to minimize heating of the cortex during surgery. Following the craniotomy, transfer the animal to the imaging stage to keep the head still under the objective.

Screw the imaging chamber to the adjustable post. Apply a custom made plastic muzzle connected to oxygen. This will provide oxygen enriched air during the experiment to aid animal respiration.

To inject the sensor in the brain cortex, load a glass pipette with a tip diameter of four millimeters and a silver chloride electrode. With the one micromolar drimmer ER solution, the electrode will facilitate recording of extracellular field potentials with a microinjection set up. Insert the pipette into the cortex at a depth of approximately 150 microns.

Inject for one to two minutes at a pressure of 0.5 pounds per square inch to optimize the optical setup for imaging. Adjust the laser power to minimize photo bleaching and photo damage. Typically, the laser power employed is around 20 milliwatts, and the PMT gain is kept constant at 667 volts.

Set the laser wavelength to excite the sensor at 820 nanometers and direct the software to detect fluorescein and rod domine fluorescence simultaneously through standard FTSE and trit filters. Also set the acquisition frequency to a frame rate of two hertz for background correction. Acquire a dark frame with the laser shutter closed to measure the mean thermal noise arising in the PMTs and pedestal.

Finally, acquire pH dependent fluorescence images and store them for subsequent analysis. Analyze the data as before to assess the in vitro behavior of fluorescein and rumine. Loaded on the rimer, a pH titration was performed using a fluorimeter.

This figure shows a typical titration of a pH sensor. Fluorescein and rumine can be separately excited at 488 nanometers and 550 nanometers respectively, allowing minimum crosstalk between the two channels as can clearly be seen. Fluorescein shows a pH dependent behavior while R domine signal does not change significantly in the physiological pH range.

Therefore, the ratio of these two signals does not depend on sensor concentration, but only on pH. Next to deliver the indicator intracellularly hela cells were electroporated with the dederer sensors and confocal imaging was performed as described in this video. These confocal images show the cells in the fluorescein channel on the left, the R domine channel in the middle, and displayed as a pH ratio metric map on the right.

Strong fluorescein and r domine signals are observed, which co localize demonstrating the integrity of the sensor structure. A ratio metric map is reconstructed by dividing the two images pixel by pixel and is represented with a pseudo color scale. To calibrate the sensor response inside cells, pH clamping with ionophores was performed as shown here.

Indicators readily respond to pH with a change of the green to red ratio. The resulting data was used to generate a calibration curve for accurate pH measurements. The linear trend in the calibration demonstrates the ability of the sensor to respond to pH without any perturbation from the cellular environment.

To demonstrate how dendri based sensors can be employed for in vivo imaging, pH imaging was performed in the brain of a dendri or injected anesthetized mouse. Fluorescein and r domine signals were simultaneously obtained with 820 nanometer excitation, and the ratio map was built on a pixel by pixel basis similar to live cell measurements. Notably, the indicators localize in the extracellular space, the non fluorescent areas in the image identifies cellular bodies or small blood vessels to verify sensor response to pH.

We employed the use of hypercapnea. Indeed, carbon dioxide is known to alter the equilibrium of the carbonate buffers in blood and tissue resulting in an acidification of the tissue. As seen in this plot, the inhalation of 30%carbon dioxide is enough to induce a strong response of the sensor that is completely reversible upon rev ventilation of the mouse.

These results demonstrate the potential ofer based sensors to highlight physiological and pathological changes of pH in living cells and in vivo. A, our procedure is compatible with other measurement like electrophysiological recordings. This allowed to achieve simultaneously complementary information about the ongoing biological process.

This method, after its development, paved the way of researcher in cell biology and neurobiology to explore pH regulated processes in culture cells and in vivo.