As the best electron acceptor, oxygen plays a crucial role in biological systems. With this protocol, you can image the oxygen distribution in 2D using an optode. In addition to its high spatial and temporal resolution, this method does not require an additional reference dye, is extremely robust, and allows the acquisition of structural image.
This method can be applied to various areas of research in which oxygen plays a key role including ecology, medical research, bioprinting, and pressure-sensitive printing. The optode fabrication can be challenging as the amount of cocktail added to the support foil and the knife coating device drag speed may require optimization and practice. With this video contribution, we want to make this powerful technique accessible to researchers from other fields of study.
To prepare a planar oxygen optode, first use a film of water or 70%ethanol to fix a clean dust-free PET foil on a clean glass plate. Place a clean 120 micrometer knife coating device onto the foil and use a glass pipette to apply a line of sensor cocktail in front of the device. Next, drag the knife coating device slowly and uniformly over the PET foil to spread the cocktail evenly.
Then dry the finished planar oxygen sensitive optode in ambient air for one hour before drying overnight in a heated cabinet at 50 to 60 degrees Celsius. By the next morning, an approximately 12 micrometer thick layer will have been obtained. Store the optode protected from light until further use.
To prepare a rhizo-sandwich chamber, use light curable acrylic-based instant adhesive to glue microscope slides along the edges of a glass plate leaving one long edge open. Cut the planar optode into the required shape and size to fit into the space between the glued microscope slides and place the optode on the inside of the front glass plate with the coated side upwards. Tape one edge of the optode foil to the glass plate and add tap water between the glass plate and the optode foil.
Slowly lower the foil onto the water droplets allowing the optode to straighten itself out on the glass surface and use a soft tissue to carefully remove an air bubbles trapped between the optode and the plate. Then wipe the glass plate dry and tape the remaining edges of the foil to the plate. Next, use a flat glass plate to distribute 0.5 millimeters sieve sediment evenly across the plate to the same thickness as the microscope slide spacers.
Carefully clean the upper surface of the microscope slide to ensure that the second glass plate seals the chamber properly and apply silicone grease to the microscope slide surface. Then cover the sediment with a thin film of water while carefully avoiding the formation of air bubbles. Before placing a sample onto the sediment, carefully wash a single shoot of Littorella uniflora and position the shoot on a sediment with the leaves sticking out from the upper open side.
When the sample is in place, place the glass plate with the optode onto the sediment and apply gentle pressure to bring the optode in close contact with the plant roots and the surrounding sediment. Use clamps to fasten the plates together and dry the outer edges with tissue paper. Repeatedly add a few drops of water to the leaves to keep the plant hydrated throughout the assembly of the rhizo-sandwich and use vinyl electrical tape to tighten the rhizo-sandwich chamber.
Then seal the edges with modeling clay and additional electrical tape. To prepare for imaging, remove the covering foil from the optode area after incubation and position the rhizo-sandwich chamber with the glass wall with the optode upright against the aquarium wall. Use a spacer to press the rhizo-sandwich chamber against the wall.
Place the frequency domain-based luminescence lifetime camera equipped with an objective in front of the aquarium and the area of interest. Attach a suitable emission filter for imaging the indicator dye to the camera objective and fix the light guide in the LED excitation source. Then position the guide so that it will evenly illuminate the planar optode foil covering the area of interest.
In the imaging software, select the camera and select the LED in the LED control software. Set the LED intensity as needed and tick analog and sync to confirm that the LED is triggered by the external transistor-transistor logic. Manually focus the camera and adjust the objective aperture and set the internal modulation source, sine wave for the output waveform, additional phase sampling, eight-phase samples, phase order opposite, tap A B readout, and five kilohertz modulation frequency.
Adjust the exposure time until the region of interest statistics readout for the normalized luminescence intensity image is in the range of 0.68 to 0.72. Then click capture reference to start the acquisition of a reference measuring series. To calibrate the optode, use a gas mixing device to flush the water in a calibration aquarium with an ambient air nitrogen gas mixture with a known oxygen concentration monitoring the oxygen concentration with an external probe with an oxygen sensor.
Then acquire a series of images at different oxygen concentrations in the calibration chamber to obtain a proper curve fit to the acquired calibration data. When the system has been calibrated, switch off the lights applying irradiation to the plant and all of the other light sources and adjust the acquisition time based on the intensity of the image to ensure that the signal is neither oversaturated nor too weak for a good signal-to-noise ratio in the lifetime determination. When all of the oxygen lifetime images have been acquired, turn the lights back on to obtain a structural image and obtain an image with a ruler in the field of view to enable subsequent scaling of the acquired images.
Before imaging the sample, the optode must be calibrated. Following a quasi-exponential decay, the measured luminescence lifetime decreases as the oxygen concentration increases. This relationship can also be described using the simplified two-site model.
Once the optode has been calibrated, it is possible to determine the oxygen concentration by imaging the luminescence lifetime as observed in these images in which the distribution of the oxygen concentration in the rhizosphere of Littorella uniflora was imaged after light exposure to 500 micromole photons per meter squared per second for 12 hours and after 12 hours in darkness. In addition to lifetime images, structural images can also be acquired under external illumination while keeping the imaging geometry fixed to allow the oxygen imaging to be precisely correlated to the structural image. Oxygen concentration profiles across a single root for example can then be extracted from the images acquired in darkness and light.
It is critical to have good contact between the sample matrix and the optode to avoid unnecessary artifacts. If you are in doubt, remake the sandwich. The instantaneous acquisition of an image in a nondestructive manner enables monitoring of the oxygen environment.
The method might as well be combined with other optodes for pH or other analytes. Be sure to perform optode fabrication in a fume hood as the sensor cocktail contains chloroform. Planar optodes can be used to identify microbial communities in the sediment.
And after imaging, the sandwich can be opened up to sample these hot spots for microbial community analysis.
