Experimental Quantification of Interactions Between Drug Delivery Systems and Cells In Vitro: A Guide for Preclinical Nanomedicine Evaluation

This protocol provides quantitative data on how carrier systems interact with cells. Quantitative data is key for engineering and optimization. It allows us to move from yes or no biological questions, to how much questions.

This technique is useful to convert results from preclinical drug carrier performance, which were qualitative, into quantitative results. And this works even when you characterize your carrier on a different system than you characterize your cells. It can take a few iterations to find the optimal carrier dilution that fall within the linear or quantitative ranges of the nano side.

Take your time for finding the right conditions. To begin, mount the flow cell onto the laser module. Slowly flush the flow cell at the rate of 0.1 milliliter per second with one milliliter of distilled water.

If bubbles form within the flow cell, partially retract the suspension to merge the bubble with the air-liquid interface before proceeding. Then lock the entire laser module in place inside the instrument. Start the camera roughly halfway through flushing.

Be sure to confirm that the carrier debris is washed out. Select capture to open the capture settings tab and click start camera. Drive the system with one milliliter of air.

If any static carriers are visible on the screen, clean the flow cell according to the manufacturer’s instructions. Prepare the carriers for nanoparticle tracking analysis, by ensuring the carriers are well suspended via sonication or vortexing, depending on the carrier system involved. Dilute the carriers in water to prepare at least 0.6 to one milliliter of each sample with a carrier concentration between 1×10 to the seventh and 1×10 to the ninth carriers per milliliter.

Take out the laser module and place it upright. Drop the first carrier sample into a one milliliter syringe and attach the syringe to the tube inlet. Then carefully load the sample into the flow cell.

If bubbles form within the flow cell, retract the suspension partially to merge the bubble with the air-liquid interface before proceeding. Ensure the entire flow cell is filled with liquid, then pause the loading. Adjust the camera focus if needed to visualize individual carriers.

Make coarse focus adjustments with the rotating knob on the right hand side of the instrument. Make finer adjustments by selecting the hardware tab. Change the focus by adjusting the focus slider.

To ensure there is no oversaturation, select the optimal camera level by adjusting the slider within the capture tab. If the instrument is equipped with this accessory, load the syringe containing the carrier sample into the syringe pump. Under the SOP tab, select standard measurement to take five captures of 30 seconds each.

Enter base file name, and if desired, add additional sample information by clicking the advanced button which will open a modal dialogue with various choices. Press create and run script and wait for a pop-up to appear asking to please advance sample. If using the syringe pump, select the hardware tab and then syringe pump tab.

Set the desired infusion rate and press infuse. If not using the syringe pump, manually advance the sample. In the popup window, select OK to start capturing.

After each of the five captures, when the please advanced sample pop up reappears, check that the sample is still moving through the flow cell. Then select OK to proceed with the next capture. In the process tab, adjust the detection threshold slider between four and eight to correctly identify distinct carriers visible on the screen.

You can adjust the screen gain to aid visualization, it will not affect the downstream analysis. In the pop-up, press okay to initiate tracking analysis. Monitor the analysis progress by clicking on the analysis in single analysis tab.

Once the analysis is finished, look for an export settings prompt to appear. Confirm that the include PDF and include experiment summary is selected. Select any other export formats as desired.

In the results section of the PDF data export, to ensure the concentration measured is reliable, verify that the measured carrier concentration is between 1×10 to the seventh and 1×10 to the ninth carriers per milliliter and check for any error messages or messages of caution underneath the concentration measurement result. Repeat the procedure two or more times with different dilutions from the stock, and ensure that each sample’s concentration falls within the instrument’s linear range. Calculate the stock carrier concentration as described in the text manuscript.

Prepare the free or antibody conjugated fluorochrome solution as described in the text manuscript. Calculate the concentration of the stock solution from the concentration, the molecular weight, and Avogadro’s number as shown in the equation. Then perform a serial dilution of the dye in the carrier diluent to generate standard curve samples.

Then add the carrier sample to the measurement plate and measure fluorescence using a microplate reader. Generate the standard curve as described in the text manuscript. Calculate the absolute fluorescence intensity per carrier by dividing the bulk fluorescence by the carrier concentration.

Set up the flow cytometer for the final carrier cell experiment by determining optimal PMT voltage settings in the relevant channels. Run a negative control sample where the cells are not incubated with carriers to determine the background fluorescence. Prepare and re-suspend the flow cytometry quantitation beads.

Use the same buffer as used for the cell samples. If the bead populations are provided separately, pull them together. Run the flow cytometry quantitation bead and the carrier cell samples to determine the fluorescence intensity per cell.

Run the flow cytometer quantitation beads to generate a standard curve, converting the absolute fluorescent intensity into MFI. Use this to calculate the theoretical MFI of the carriers as described in the text manuscript. Run the cell carrier samples to determine the fluorescent intensity per cell of each sample.

Finally, calculate the number of carriers per cell for each sample by subtracting the background fluorescence from the measured fluorescence of the sample, and then dividing by the fluorescence per particle. For 633 nanometer Polymethacrylic acid core shell particles, the concentration and fluorescent intensity per carrier were directly quantifiable using the cytometer stream. In contrast, 100 nanometer super paramagnetic iron oxide nanoparticles were too small to detect individually and were analyzed using the bulk stream.

Labeled quantitation beads were used on multiple days to generate standard curves on a flow cytometer. The correlation between the measured MFI and the MESF value of the quantitation beads was linear and broadly similar between the measured dates. Time course experiments using fluorescent labeled HeLa cells with 235 nanometer polymethacrylic acid capsules from zero to 24 hours showed an increase in the MFI over time, indicating the capsules were associating with HeLa cells.

The difference in the apparent cellular response to carriers was stark, depending on relative or absolute quantification. The absolute quantification was independent of labeling intensity and thus more comparable. These techniques are the foundation for deeper analysis and comparison of particle performance.

We’re excited about using these techniques to parameterize mathematical models which will allow us to characterize particle performance independent of any particular experiment.