We thank Sofie Lindegaard Jakobsen (University of Copenhagen) and Lars Borregaard Pedersen (Aarhus University) for technical assistance. Funding for this study was obtained from a Sapere-Aude Advanced grant from the Independent Research Fund Denmark (DFF-1323-00065B; MK), project grants from the Independent Research Fund Denmark | Natural Sciences (DFF-8021-00308B; MK) &#38; Technical and Production Sciences (DFF-8022-00301B and DFF-4184-00515B; MK), the Danish National Research Foundation (DNRF136), and the Poul Due Jensen Foundation (KK).

1. Fabrication of planar O2 optode
<ol>
	<li>Dissolve 1.5 mg of the luminescent O2 indicator dye platinum(II)-5,10,15,20-tetrakis-(2,3,4,5,6-pentafluorphenyl)-porphyrin (PtTFPP) and 100 mg of polystyrene (PS) in 1 g of chloroform to get the so-called &#39;sensor cocktail&#39;. 
	NOTE: The cocktail can be kept in a closed, gas-tight glass vial for a few hours in the fridge and darkness until further use.</li>
	<li>Fix a clean, dust free polyethylene terephthalate (PET) foil (size dependent on the application) on a cleaned glass plate with the help of a water or ethanol (70%) film (Figure 1A).</li>
	<li>Place the cleaned knife coating device (120 &#181;m) on the foil and apply a line of the sensor cocktail in front of the device using a glass pipette (Figure 1B). Then, drag the knife-coating device slowly and uniformly over the PET foil to spread the cocktail evenly. 
	NOTE: All materials and tools have to be cleaned thoroughly and the fabrication should be done in a dust-free environment, such as a fume hood, flow bench or underneath a point suction device. To avoid heterogeneities in the final sensor foil, the steps following the application of the sensor cocktail onto the foil should be done swiftly, as the chloroform evaporates fast.</li>
	<li>Dry the finished planar O2-sensitive optode in ambient air for 1 h and then over-night in a heating cabinet at 50-60 &#176;C, resulting at a final layer thickness after solvent evaporation of ~12 &#181;m. Store the produced optodes in darkness (e.g., in a paper envelope) until further use (Figure 1C). 
	NOTE: Planar O2 optodes can be stored dry and in darkness for several months to years prior to use. A final layer thickness ranging from 1-20 &#181;m has proven to deliver good results, with sufficient luminescence signal and adequate response times.</li>
</ol>
2. Rhizo-sandwich chamber
<ol>
	<li>Clean two glass plates (24.5 x 14 cm2, thickness: 4 mm) with 96% ethanol.</li>
	<li>Use light-curable, acrylic-based instant adhesive (see Table of Materials) to glue microscope slides (76 x 26 mm2, thickness: 1 mm) along the edges of the first glass plate (i.e., the rear chamber side), while leaving one long edge open. Use a glasscutter to shorten microscope slides as needed. 
	CAUTION: Cutting glass can cause sharp edges and should be handled with care. 
	NOTE: The microscope slides function as spacers between the front and back, and depending on the thickness of the roots and plant size, multiple layers of microscope slides can be glued on top of each other.</li>
	<li>Cut the planar optode into the required shape and size to fit into the space between the glued microscope slides. Place it on the inside of the front glass plate with the coated side upwards, to allow contact with the sample of interest when pressed against it.</li>
	<li>Tape one edge of the optode foil to the glass plate and add a few drops of tap water in between the glass plate and the optode foil (Figure 2A). Slowly lower the foil on these water droplets allowing it to straighten itself out on the glass surface.</li>
	<li>Carefully remove air bubbles trapped in-between the planar optode and the glass plate using a soft tissue, while avoiding scratching of the sensor coating. Wipe the glass plate dry and tape the remaining edges of the optode foil to the glass plate (Figure 2B). 
	NOTE: A tape with suitable adhesion under water should be chosen.</li>
	<li>Sieve the sediment using a mesh size of 0.5 mm. Place a spoon of wet sediment on the first glass plate (Figure 2C). 
	NOTE: The mesh size should not be larger than half the spacer thickness.</li>
	<li>Distribute the sediment evenly and adjust it to the same thickness as the microscope slide spacers using a flat glass plate. Carefully clean the upper surface of the microscope slides to ensure that the second glass plate seals the chamber properly.</li>
	<li>Apply silicon grease to the microscope slide surface. Cover the sediment with a thin water film, whilst carefully avoiding the formation of air bubbles.</li>
	<li>Carefully wash a single shoot of Littorella uniflora and place it on the sediment, with the plant leaves sticking out from the upper open side (Figure 2D).</li>
	<li>Place the second glass plate, with the optode attached to it, on the sediment and apply gentle pressure to bring the optode in close contact with the plant roots and the surrounding sediment. 
	NOTE: Air bubbles trapped in the sediment can be removed by tilting the glass plates while bringing them together.</li>
	<li>Fasten the glass plates together using clamps (Figure 2E). Dry the outer edges with tissue paper. Keep the leaves moisturized throughout the entire assembly of the rhizo-sandwich (e.g., by frequent addition of a few drops of water).</li>
	<li>Tighten the rhizo-sandwich chamber using vinyl electrical tape. Seal the edges with modeling clay and additionally tape them with vinyl electrical tape (Figure 2F). 
	NOTE: If there are many air bubbles in the sediment, or sediment grains in between the spacer microscope slides and the second glass plate, the chamber should be reassembled as pore water can leak out (repeat steps 2.4 - 2.8).</li>
	<li>Use an opaque plastic to cover the rhizo-sandwich, but leave a slit in the foil for the plant leaves to stick out. Cut a window in the plastic foil, so it can be opened for the experiments by unfolding. Close the window during acclimatization times using rubber bands (Figure 2G) to protect the optode from photo bleaching while the plant is incubated. 
	NOTE: As algal growth could potentially interfere with O2 concentrations measured, we recommend trying to minimize it, by using filtered water, pre-cleaned experimental equipment and removing algae upon formation.</li>
</ol>
3. Rhizo-sandwich chamber incubation
<ol>
	<li>Place the rhizo-sandwich chamber in a water tank (32 x 7 x 28 cm3) in a slightly tilted position to encourage root growth against the planar optode.</li>
	<li>Fill the water tank with enough water to fully submerge the plant leaves.</li>
	<li>Establish a 14 h light, 10 h dark cycle for acclimatization of the plant using a time-controlled lamp. Place an air-stone or a water pump in the tank to ensure aeration and mixing of the water (Figure 2H).</li>
</ol>
4. Imaging
<ol>
	<li>Imaging setup
	<ol>
		<li>Remove the plastic foil covering the planar optode in the rhizo-sandwich chamber. Position the chamber with the glass wall with the optode upright against the aquarium wall. Use a spacer to press the rhizo-sandwich chamber against the aquarium wall. 
		NOTE: The overall thickness of the aquarium wall plus the rhizo-sandwich chamber wall should not get too thick, however, glass thicknesses for aquaria walls for luminescence imaging are recommended with &#62; 1 cm, in order to reduce spatial cross-talk by increasing the attenuation of the scattered light. It is important, to use the same material for both glass walls (same refractory index), in order to minimize light scattering at the material interface; as this would lead to a blurred image as well12.</li>
		<li>Place the frequency-domain-based luminescence lifetime camera equipped with an objective (see Table of Materials) in front of the aquarium and the area of interest (roots of the aquatic plant Littorella uniflora, which are in direct contact with the planar optode) (Figure 3). 
		NOTE: The camera might be placed on a lab stand to allow easy height adjustment of the camera. The position of the lab stand should be marked and kept fixed. Additionally, the camera can be taped to the lab stand to avoid accidental movement of the camera during the experiment.</li>
		<li>Screw a suitable emission filter for imaging PtTFPP as indicator dye (see Table of Materials) on the camera objective, to remove inferences from the excitation source. 
		NOTE: Screw-on filters are ideal, but square filters may also be used with either an appropriate adaptor, or by careful taping them to the objective.</li>
		<li>Connect a LED excitation source (see Table of Materials) to the modulation and dark gate output of the camera. 
		NOTE: The former delivers the modulation signal for the light source, while the latter switches off the light during image readout of the image sensor. Connect the LED excitation source and camera to a computer. Background light should be minimized during image readout, by either darkening the entire room or putting a dense, black cloth over the entire set-up. In the latter case, it is important to ensure sufficient ventilation to avoid heating of the camera.</li>
		<li>Fix the light guide in the LED excitation source and position it to evenly illuminate the planar optode foil covering the area of interest. 
		NOTE: In the used LED excitation source it is possible to switch between 3 different LEDs (460 nm, 528 nm, 625 nm), the intensity of which can be adjusted via the control software.</li>
	</ol>
	</li>
	<li>Settings and camera operation 
	NOTE: For the described experiments, we used a frequency-domain-based lifetime camera in combination with a dedicated module for lifetime imaging in a commercially available software package (see Table of Materials).
	<ol>
		<li>Select the camera in the chosen software prior to use. 
		NOTE: The software and camera drivers need to be installed prior to imaging following the manufacturers guidelines.</li>
		<li>Open the LED control software (again installed prior to starting the experiment) and choose the suitable LED (here: 528 nm) by ticking off standby. Set the LED intensity as needed (here to 30%). Ensure that the LED is triggered by the external TTL; this is done by ticking analog and sync for the LED. 
		NOTE: The LED intensity needs to be adjusted individually, as too high laser power can lead to accelerated photo bleaching of the indicator or reference dye.</li>
		<li>Focus the camera and adjust the aperture of the objective manually (in the present study use f = 2.8). 
		NOTE: It is important to focus the camera on the planar optode and not on the aquarium glass; this can be ensured by taking an image with a ruler for scale, and focusing on the shadow of the ruler on the optode, rather than on the actual ruler.</li>
		<li>Set the following parameters within the software&#39;s camera control panel: internal modulation source; sine wave for the output waveform; additional phase sampling (Yes); 8 phase samples, phase order opposite, Tap A + B readout; 5 kHz modulation frequency. 
		NOTE: These parameters affect image quality and can be changed if needed. The manufacturer of the camera provides guidelines about the individual parameters (The camera manufacturer is releasing guidelines and updates whenever the software gets updated).</li>
		<li>Take a reference image prior to experiments. 
		NOTE: This can be done either by imaging a calibration standard (a luminescent dye with a known lifetime (ns or &#181;s)), or by using the reflected light of the LED. In the latter case, the emission long pass filter needs to be removed from the objective and the known lifetime can be set to 1 ns.</li>
		<li>Adjust the exposure time in the calibration section of the dedicated imaging software until the ROI Statistics readout (in the bottom of this panel) for the normalized luminescence intensity image is in the range of 0.68 - 0.72. 
		NOTE: Now the reference lifetime (e.g., 1 ns) is given as input to the software.</li>
		<li>Press Capture reference to start the acquisition of a reference measuring series. 
		NOTE: When finished, the reference data are stored and either single or time lapse measurements can be done on samples.</li>
	</ol>
	</li>
	<li>Calibration of the O2 optode
	<ol>
		<li>Position a piece of a planar O2-sensitive optode in a (small) glass aquarium. Fix the planar optode on the glass wall of the calibration chamber as described previously (see section 2.3). Place the calibration aquarium in front of the camera. Ensure even illumination by the LED, as well as that the optode fills the entire field of view. 
		NOTE: The planar optode should be from the same piece of foil or made from the same sensor cocktail as the foil used in the actual experiment.</li>
		<li>Fill the aquarium with the same liquid medium as used in the experiments. 
		NOTE: Using different media for calibrations and experiments can influence the measurement, (e.g., by changing the sensor response and/or the O2 solubility). Thus, calibration should be done in the same medium, and at the same temperature as the actual experiment. Fluctuations in temperature will affect the luminescence signal and should be avoided. However, if the temperature cannot be kept stable, temperature compensation needs to be done by calibrating the O2-sensitive optode (multiple points) at different (relevant) temperatures and subsequent recalculation of the values.</li>
		<li>Adjust the O2 concentration within the calibration aquarium by flushing the water with an air/N2 gas-mixture of known O2 concentration, using a gas-mixing device. Ensure that the water is well equilibrated with the used gas-mixture by aerating for a sufficient time (depends on the flow rate and size of the aquarium). 
		NOTE: We recommend monitoring the O2 level in the calibration aquarium with an external, calibrated O2 sensor with temperature compensation (e.g., using a fiber optic or electrochemical O2 sensor).</li>
		<li>Take a series of images at different O2 concentrations in the calibration chamber. 
		NOTE: At least five different O2 concentrations should be measured in order to enable a proper curve fit to the acquired calibration data. It is important to measure at 0 hPa (anoxic conditions), and then distribute the other values over the dynamic range of your specific indicator dye. Here we used PtTFPP as the O2-sensitive indicator dye immobilized in a polystyrene matrix. Images were taken at 0, 48, 102, 156, and 207 hPa; 207 hPa corresponds to 100% air saturation at the given salinity and pressure.</li>
	</ol>
	</li>
	<li>Imaging the sample
	<ol>
		<li>Place the sample in front of the camera and ensure even illumination.</li>
		<li>Switch off the light supplying irradiation to the plant (and all other light sources) just prior to acquiring the luminescence lifetime image of the plant. Adjust the acquisition time based on the intensity image, ensuring that the signal is neither oversaturated nor too weak for a good signal to noise (S/N) ratio in the lifetime determination.</li>
		<li>Expose the plant to varying light conditions (e.g., light/ dark) and acquire a set of images.</li>
		<li>Switch on the light in the room to acquire a Structural image. 
		NOTE: When the background light is switched on, the camera will not measure a realistic lifetime image. However, the intensity image now shows the entire field of view as seen through the semitransparent optode.</li>
		<li>Take an image with a ruler or alike in the field of view to enable later scaling of the acquired images.</li>
	</ol>
	</li>
</ol>
5. Data analysis
<ol>
	<li>Export the phase lifetime and intensity images directly from the dedicated imaging software, using the macro provided by the camera manufacturer.</li>
	<li>Perform further image analysis using a freely available image analysis software (see Table of Materials).</li>
	<li>Open the phase lifetime images of the calibration in the image analysis software and determine the mean of the entire image using the measure function. Plot the measured lifetimes against the known O2 concentrations to determine the calibration function (Figure 4A).</li>
	<li>Calculate &#964;0/&#964; from all the data (&#964;0 is the measured phase lifetime in the absence of O2). Plot these values versus the known O2 concentrations (Figure 4B).</li>
	<li>Determine the parameters Ksv and f from the calibration plot, using the simplified two-site model for dynamic collisional quenching (equation 3)38,39 where [Q] is the O2 concentration. Define the fit-function in the data analysis software, which then determines Ksv and f.</li>
</ol>
(3) <img align="center" src="/files/ftp_upload/60191/60191eq3.jpg" />
<ol start="6">
	<li>Open the acquired sample images in the image analysis software to convert the imaged lifetimes into O2 concentrations, using the determined parameters Ksv, f and &#964;0. 
	NOTE: As an alternative approach also the acquired calibration phase lifetime values (Figure 4A) can be used directly. In this case, an exponential fit using the curve fit function is used for calibration.</li>
	<li>Open the image with the ruler next in the image analysis software and measure a known distance using the measuring tool. Set this measurement as global scale under Set scale.</li>
</ol>

The author Gerhard Holst is an employee of PCO AG which manufactures the camera system used in this article. PCO AG financially contributed to the publication and open access costs of this article.

In this protocol, the entire work-flow from optode preparation to O2 image analysis is covered. By following this protocol, chemical images can be obtained using the novel frequency-domain-based luminescence lifetime camera. Depending on the application, the planar optodes can be fabricated in various sizes and layer thickness of the sensor layer ranging from robust 50-100 &#181;m thick planar optodes of several tenths of square centimeter to microscope cover slips with &#60;1 &#181;m thick sensor layers6,40. The potential of this method was demonstrated with a particular application, but is not only limited to O2 imaging in plant rhizospheres12,28.
This method has several benefits when compared to pure luminescence intensity-based chemical imaging methods. Luminescence lifetime imaging is not, or at least far less, affected by uneven illumination, uneven optode thickness, and photo bleaching25. Also, this method avoids the use of an additional reference dye common in ratiometric imaging17,37. In comparison to other lifetime based camera systems, such as commonly used gated time-domain cameras8,26, the novel camera system and protocol presented here can deliver comparable results. In a recent publication, the analytical characteristics of those two systems were compared and it was found that the frequency-domain-based luminescence lifetime camera system is at least comparable to the discontinued time-domain-based predecessor8.
We have presented the simplest O2 optode consisting only of an indicator in a polymer matrix. Besides multiple other possible O2 indicators20 that might be used additives can be included, i.e., scattering agents such as TiO2 or diamond powder2 to increase sensor signal while reducing transparency of the optode. Also additional dyes might be used to enhance signal intensity via energy transfer41.
For planar optode fabrication, we recommend using a gap in the knife-coating device of 75 - 120 &#181;m to yield a final sensor layer thickness of around 7.5 to 12 &#181;m after solvent evaporation (around 10% of the used gap), when using the described sensor cocktail composition. This is a good compromise between signal intensity, which can be modified by higher dye loading, or by choosing indicator and reference dyes of higher brightness, and response time. An increase in layer thickness results in an increase in response time, as the time span required for the analyte to reach a thermodynamic equilibrium in the sensing layer with the surrounding media increases12.
Optodes, as described here, react to changes in O2 concentration within a few seconds17 while still having a sufficiently strong luminescence signal. Ultrathin sensor coatings with sub-second response times can be realized with spin-coating6. If the support or the knife-coating device are not well-cleaned, it might result in inhomogeneous sensor layers. Also, when the cocktail is not completely homogenous or applied too rapidly after spreading in front of the coating device such an unwanted result can be observed. Therefore, it might need some practice to prepare optimal optodes.
The method can be used to image samples which can be put in close contact to the optode, such as certain marine animals42, biofilms6 and soils31 just to name a few. We present a standalone setup using an objective, however, the camera can easily be coupled to a microscope for higher resolution chemical imaging43.
While time-domain based luminescence lifetime imaging enabled suppression of background fluorescence26, this is an issue when using the new frequency-domain-based camera system8. Due to the continuous image acquisition, this camera will record any background fluorescence of the sample that can be excited by the selected LED and emits in the selected spectral window as defined by the emission filter on the camera objective. This will result in an apparently lower lifetime and consequently in false readings. In case you work with samples with a significant intrinsic fluorescence overlapping with the O2 sensor excitation and emission, it is essential to apply an extra optical isolation on top of the optode, by coating an additional layer containing carbon black2,17. Thus, only luminescence emitted from the planar optode will reach the camera. In order to check for background luminescence an image without the optode can be taken, which then would exclusively show intrinsic luminescence of the sample. It is also possible to add scattering agents such as TiO2 or diamond powder2,44, to the sensor cocktail, to increase luminescence intensity of the indicator dye. However, this can also lead to faster photo bleaching and TiO2 is a known photo-catalyst, which can impair photostability of a dye41. A further aspect to consider is background light. When imaging luminescence lifetimes, background light needs to be avoided as efficiently as possible. Therefore, this imaging method requires the setup to be placed in a dark environment and any external light source needs to be temporarily switched off during image acquisition.
In summary, luminescence lifetime imaging is a robust chemical imaging method that can be adapted to many different applications. This protocol (see section 1 - 5) covers all the essential steps to generate an O2 image and uses the currently most flexible frequency-domain luminescence lifetime imaging system, which can replace the discontinued gated time-domain camera for 2D O2 imaging with planar optodes.

The distribution and dynamics of dissolved gases and ions in sediments and soils provide key information on biogeochemical processes such as microbial respiration1,2, or radial oxygen loss from plant roots3,4,5, and the chemical microenvironment of microbes6,7, plant rhizospheres5,8,9 and animal burrows10,11,12. Biological and chemical activity in such diffusion-limited environments can create steep gradients of chemical substrates or products of biogeochemical processes. In particular, O2 availability has a huge impact on biogeochemical processes and thus the biology and ecology of a system13. Therefore, analyzing O2 concentrations at high spatial and temporal resolution is of prime importance in aquatic and terrestrial sciences. First, electrochemical and optical microsensors14,15 were developed to measure this important analyte. Later, 2 dimensional (2D) imaging of O2 with planar optodes was introduced12,16,17,18,19, which enabled the visualization and quantification of the heterogeneous O2 distribution in soils and sediments.
Planar O2 optodes consist of an O2-sensitive indicator dye20, which is dissolved in a suitable polymer21. The indicator dye is excited at specific optical wavelengths and emits red-shifted light upon relaxation in the form of luminescence. In the presence of O2, the excited indicator dye can transfer its energy to the O2 molecule upon collision, which is referred to as collision-based luminescence quenching22. Therefore, the luminescence intensity as well as the luminescence lifetime are reduced with increasing O2 concentration23. In an ideal case the change in intensity and lifetime follows the Stern-Volmer equation (equation 1) using either the luminescence intensity or lifetime in the absence (I0; &#964;0) or presence (I, &#964;) of O2 at a given concentration [Q]. The Stern-Volmer constant (Ksv) is a measure for the sensitivity of the optode towards O2; KSV is dependent on environmental variables such as temperature and pressure.
(1) <img align="center" src="/files/ftp_upload/60191/60191eq1.jpg" />
Recording such changes in luminescence over a planar sensor foil with a camera system can be used to visualize the corresponding changes in O2 distribution. Initially, simple luminescence intensity-based O2 imaging was used18. However, such methodology is very sensitive to external interferences, which compromise the reliability of the results due to heterogeneous illumination, fluctuations in the excitation source or camera, as well as uneven distribution of the indicator dye within the planar optode.
Some of these limitations can be alleviated by using planar optodes for ratiometric imaging17,24, where the O2-sensitive indicator dye is co-immobilized in the polymer layer of the planar optode with an insensitive reference dye emitting in a different spectral range than the O2-indicator. Based on emission images acquired in two spectral windows, the O2-sensitive emission signal is divided by the reference signal, generating a ratio image that is less prone to the above mentioned interferences5,17. The method requires the use of a second dye, which ideally can be excited by the same excitation source, but emits at a different wavelength (without significant spectral overlap), in another spectral window of the camera (e.g., in another color channel of an RGB camera).
Alternatively, O2 imaging can be based on quantifying the O2-dependent change in luminescence lifetime of the indicator dye, which is not affected by uneven illumination or heterogeneities in the indicator concentration25. First luminescence lifetime-based O2 imaging systems were based on time-domain measurements with a gate-able charged coupled device (CCD) camera system26, where a pulsed excitation source is used and luminescence images are taken over defined time intervals within the excitation or emission of the indicator8,23,27. From such images, the luminescence lifetime can be determined and correlated to the corresponding O2 concentration in a calibration. Subsequently, luminescence lifetime images for a given sample pressed against the planar optode can be converted to images of the corresponding 2D distribution of O2 concentration. This system has been used in many applications both in the laboratory and in situ16,28, but the essential gate-able CCD camera is no longer commercially available.
Recently, a different luminescence lifetime camera system was released, which acquires images in the frequency-domain8. The system relies on a continuously modulated light source for excitation. This can either be a sinusoidal or square wave instead of a pulsed excitation, which is used for image acquisition in the time-domain. This modulation results in a modulated luminescence emission of the O2 indicator dye, which is phase-shifted by an angle, &#966;, which is depended on the luminescence lifetime of the indicator dye (&#964;) (see equation 2).
(2) <img align="center" src="/files/ftp_upload/60191/60191eq2.jpg" />
The change between excitation and emission amplitude (i.e., the so-called modulation index or depth (amplitude divided by the constant luminescence part)) is also dependent on the luminescence lifetime. So, by setting a known modulation frequency the special CMOS image sensor within the camera is able to measure the luminescence lifetime in the ns to &#181;s range as described in detail elsewhere 8,29,30. A general guide on the operation principle can be found (using the following link <a href="https://www.youtube.com/watch?v=xPAB_eVWOr8" target="_blank">https://www.youtube.com/watch?v=xPAB_eVWOr8</a>).
In the following protocol, we demonstrate the use of the novel camera system for imaging the distribution of O2 concentration around the roots of the aquatic freshwater plant Littorella uniflora in 2D9,31. We would like to emphasize that this method is by no means limited to this application. Oxygen-sensitive optodes or sensor particles27 in combination with various imaging methods have been used in medical research32, in bioprinting33, for pressure sensitive paints34,35, or to study photosynthetic systems2,36,37, just to name a few other fields of application.

<table><tbody><tr><td>Air pump with air stone and water pump</td><td>Local aquarium store</td><td></td><td></td></tr><tr><td>Chloroform</td><td>Sigma Aldrich</td><td>67-66-3</td><td></td></tr><tr><td>DC4 silicone compound</td><td>Dow Corning GmbH</td><td>2793695</td><td></td></tr><tr><td>Gas mixer</td><td>V&amp;ouml;gtlin Instruments GmbH</td><td>red-y compact meter GCM</td><td>This is just one possible instrument. Several companies offer gas mixing devices</td></tr><tr><td>Glass plates and aquaria</td><td>Local aquarium or hardware store</td><td></td><td></td></tr><tr><td>ImageJ Software</td><td>ImageJ</td><td></td><td>Freely available imaging software (imagej.nih.gov/ij/index.html)</td></tr><tr><td>Knife-coating device</td><td>&lt;br/&gt; &lt;br/&gt; BYK-GARDNER GMBH byk.com</td><td>2021</td><td>This is a four sided film applicator enabling easy variation of the film thickness. Other versions are also available. We recommend a thickness of the applied film between 75-120 &amp;micro;m, which yields a final sensor layer thickness of ~10% of the applied thickness before solvent evaporation.</td></tr><tr><td>LED lamp, Reflector PAR38</td><td>Megaman</td><td>MM17572</td><td></td></tr><tr><td>LED LEDHUB</td><td>Omicon Laserage, Germany</td><td></td><td>Can be configured with a variety of LEDs. For the presented example, the green LED (528 nm) is essential</td></tr><tr><td>LOCTITE AA 3494</td><td>Henkel AG &amp;amp; Co. KGaA</td><td>NA</td><td>Acrylic-based instant adhesive</td></tr><tr><td>NIS Elements AR Software</td><td>Nikon Inc</td><td></td><td>Software package used for image acquisition</td></tr><tr><td>pco.flim</td><td>PCO AG, Germany</td><td></td><td>Frequency domain based luminescence lifetime camera</td></tr><tr><td>platinum(II)-5,10,15,20-tetrakis-(2,3,4,5,6-pentafluorphenyl)-porphyrin (PtTFPP)</td><td>Frontier Scientific</td><td>PtT975</td><td>O&lt;sub&gt;2&lt;/sub&gt; indicator</td></tr><tr><td>polyethylene terephthalate (PET) foil</td><td>Goodfellow</td><td>320-992-72</td><td>Such foils might also be found from other providers and serve as solid support</td></tr><tr><td>Polystyrene (PS)</td><td>Sigma Aldrich</td><td>9003-53-6</td><td>Polymer matrix</td></tr><tr><td>Schott RG610 filter</td><td>www.uviroptics.com</td><td></td><td>Here 52mm screw on Filters can obtained. Other sources offer square glass filters from Schott glass that can be fixed in front of the objective</td></tr><tr><td>Vinyl electrical tape</td><td>Scotch, Super 33+</td><td>NA</td><td></td></tr><tr><td>Zeiss Makro Planar 2/100 with Hama C for Nikon adaptor</td><td>delivered with the camera</td><td></td><td>Here any other objective might also be used in combination with an adaptor if the objective does not have a C-mount</td></tr></tbody></table>

luminescence lifetime imaging of o2 with a frequency-domain-based camera system

We describe a method to image dissolved oxygen (O2), in 2D at high spatial (&#60; 50-100 &#181;m) and temporal (&#60; 10 s) resolution. The method employs O2 sensitive luminescent sensor foils (planar optodes) in combination with a specialized camera system for imaging luminescence lifetime in the frequency-domain. Planar optodes are prepared by dissolving the O2-sensitive indicator dye in a polymer and spreading the mixture on a solid support in a defined thickness via knife coating. After evaporation of the solvent, the planar optode is placed in close contact with the sample of interest - here demonstrated with the roots of the aquatic plant Littorella uniflora. The O2 concentration-dependent change in the luminescence lifetime of the indicator dye within the planar optode is imaged via the backside of the transparent carrier foil and aquarium wall using a special camera. This camera measures the luminescence lifetime (&#181;s) via a shift in phase angle between a modulated excitation signal and emission signal. This method is superior to luminescence intensity imaging methods, as the signal is independent of the dye concentration or intensity of the excitation source, and solely relies on the luminescence decay time, which is an intrinsically referenced parameter. Consequently, an additional reference dye or other means of referencing are not needed. We demonstrate the use of the system for macroscopic O2 imaging of plant rhizospheres, but the camera system can also easily be coupled to a microscope.

As an application example for the new imaging system, we show 2D O2 imaging of a complex biological sample (i.e., the rhizosphere of the aquatic plant Littorella uniflora).
First, the method describes the fabrication of a planar sensor film, a so-called planar optode. As seen in Figure 1, such an optode is made of a thin layer of an optical indicator in a polymer matrix that is spread on a transparent support. By following the described protocol, a homogeneous sensor film with a uniform thickness, as defined by the gap of the knife coating device, is obtained. If the produced optode has a patchy sensor material distribution (e.g., holes in the coating, shows stripes, or dye aggregates (this can be evaluated visually, and visually with the help of an UV lamp)), the protocol needs to be repeated and all materials need to be thoroughly cleaned using acetone.
Once the planar optode is prepared, the sample can be brought in close contact with the sensing layer of the planar optode, as shown here with the planar optode integrated in a rhizo-sandwich chamber, where the roots of a plant within a surrounding sediment matrix can be positioned in close contact to the planar optode (Figure 2). If prepared correctly, the rhizo-sandwich chamber should be easily moveable from one aquarium (incubation) to the other (measurement). If not constructed correctly, the rhizo-sandwich chamber may be instable, lose sediment or contain air bubbles. Visual examination of the rhizo-sandwich chamber directly after assembly is thus recommended.
The given protocol enables frequency-domain-based luminescence lifetime imaging of the sample in contact with the planar optode using the frequency-domain-based luminescence lifetime camera. More details on this camera system such as the mode of image acquisition and scientific complementary metal-oxide-semiconductor (SCMOS) camera characteristics are given in recent publications8,29.
The setup itself is rather simple and only includes the camera that controls a light source (in this case, a LED excitation source) and the sample with the optode (Figure 3). Ensure that all parts are correctly connected and that the sample is illuminated homogeneously. Background light needs to be avoided while preforming measurements.
Prior to imaging the sample, the optode needs to be calibrated. As seen in Figure 4A, the measured luminescence lifetime decreases with increasing O2 concentration following a quasi-exponential decay. This relationship can also be described using the simplified two-site model (Figure 4B and equation 3). In the given example, the parameters needed to subsequently calculate the O2 concentration were as followed; &#964;0 = 56.26 &#181;s, Ksv = 0.032 hPa-1 and f = 0.86.
Performing a calibration is also an ideal way to test that the system is working correctly. If all components are installed as described here (or within the manufacturers guidelines), the measured lifetime should show the same O2 dependence as seen in Figure 4. In addition, for the same combination of O2 sensing materials (polymer and dye), the measured &#964;0 should be in the same range (&#177; a few &#181;s) as measured here (mainly influenced by the experimental temperature). If unable to obtain a similar calibration curve, ensure that all steps were followed correctly. Sometimes the optode is accidentally fixed with the sensitive side facing the glass wall rather than the sample, or the acquired images are over- or underexposed.
With the calibration parameters, it is possible to determine the O2 concentration by imaging the luminescence lifetime (&#964;). This is demonstrated in Figure 5A,B, where the distribution of O2 concentration in the rhizosphere of Littorella uniflora was imaged in darkness and after light exposure to 500 &#181;mol photons m-2 s-1 for 12 h, respectively. Due to the photosynthetic activity of the plant, the O2 concentration in the rhizosphere increased after light exposure. Besides lifetime images, also &#34;structural&#34; images can be acquired under external illumination, while keeping the imaging geometry fixed. In this way, O2 images can be precisely correlated to the structural image (Figure 5C), cross sections or regions of interest. As an example, O2 concentration profiles across a single root were extracted from the image acquired in darkness and light, respectively (Figure 5D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60191/60191fig1a.jpg" /> 
Figure 1: Fabrication of a planar O2 optode. (A) A PET foil is fixed on a glass plate and the knife-coating device is placed on the foil. (B) The prepared sensor cocktail is spread on the PET foil as a thin line in front of the knife-coating device. (C) The knife-coating device is moved downwards to spread the sensor cocktail as a thin film on the PET foil, which after solvent evaporation results in a ready to use planar optode. <a href="https://www.jove.com/files/ftp_upload/60191/60191fig1alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60191/60191fig2a.jpg" /> 
Figure 2: Rhizo-sandwich chamber assembly with integration of a planar O2 optode. (A) The optode is fixed on one of the glass plates using a water film. (B) The optode is glued to the plate with electric tape. (C) Sediment is filled into the opposing plate with the attached spacers (i.e., microscope slides). (D) The plant roots are placed on the evenly spread out sediment. (E) The rhizo-sandwich chamber is closed and temporarily fixed with clamps. (F) Fully closed and assembled rhizo-sandwich chamber. (G) To protect the optode from light exposure by the incubation lamp and to avoid algal growth a plastic cover is placed over the assembled rhizo-sandwich chamber. (H) The rhizo-sandwich chamber incubated in an aquarium. <a href="https://www.jove.com/files/ftp_upload/60191/60191fig2alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60191/60191fig3a.jpg" /> 
Figure 3: Imaging setup containing the frequency-domain-based luminescence lifetime camera, with the objective focused at the sample with the optode from behind via the transparent aquarium and rhizo-sandwich chamber walls. The light guide of the LED excitation source is positioned to illuminate the sample evenly. <a href="https://www.jove.com/files/ftp_upload/60191/60191fig3alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60191/60191fig4a.jpg" /> 
Figure 4: Calibration curves for planar O2 optode. (A) Different phosphorescence lifetimes measured at the respective O2 concentrations in the water-filled calibration chamber. (B) Stern-Volmer plot of the calibration data fitted using the simplified two-site model for dynamic collisional quenching (equation 3). <a href="https://www.jove.com/files/ftp_upload/60191/60191fig4alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60191/60191fig5a.jpg" /> 
Figure 5: Lifetime imaging of the O2 distribution in the rhizosphere of the aquatic plant Littorella uniflora. (A) O2 distribution after keeping the plant under light for 12 h at approximately 500 &#181;mol photons m-2 s-1. (B) O2 distribution after keeping the plant in darkness for 1 h. (C) Structural image of the plant roots as seen through the planar optode. (D) Cross-sectional O2 concentration profile (the location is indicated by the yellow line in panel A and B) after 12 h in light (red) and 1 h in darkness (black). Adapted with permission from (Koren, K., Mo&#223;hammer, M., Scholz, V. V., Borisov, S.M., Holst, G., K&#252;hl, M. Luminescence Lifetime Imaging of Chemical Sensors - A Comparison between Time-Domain and Frequency-Domain Based Camera Systems. Analytical Chemistry. 91 (5), 3233-3238, doi: 10.1021/acs.analchem.8b05869 (2019)). Copyright (2019) American Chemical Society. <a href="https://www.jove.com/files/ftp_upload/60191/60191fig5alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Luminescence Lifetime Imaging of O2 with a Frequency-Domain-Based Camera System at JoVE.com

Luminescence Lifetime Imaging of O2 with a Frequency-Domain-Based Camera System

We describe the use of a novel, frequency-domain luminescence lifetime camera for mapping 2D O2 distributions with optical sensor foils. The camera system and image analysis procedures are described along with the preparation, calibration and application of sensor foils for visualizing the O2 microenvironment in the rhizosphere of aquatic plants.

Luminescence Lifetime Imaging of O2 with a Frequency-Domain-Based Camera System

We describe a method to image dissolved oxygen (O2), in 2D at high spatial (&#60; 50-100 &#181;m) and temporal (&#60; 10 s) resolution. The method employs ...

luminescence-lifetime-imaging-o2-with-frequency-domain-based-camera

Research

JoVE Journal

Chemistry

9.1K Views.  University of Copenhagen. Als de beste elektronen acceptor, zuurstof speelt een cruciale rol in biologische systemen. Met dit protocol u de zuurstofverdeling in 2D beeld geven met behulp van een optode. Naast de hoge ruimtelijke en temporele resolutie vereist deze methode geen extra referentiekleurstof, is uiterst robuust en maakt het mogelijk om structureel beeld te verwerven.Deze methode kan worden toegepast op verschillende onderzoeksgebieden waarin zuurstof een sleutelrol speelt, waaronder ecologie, medisch...

Video: Luminescence Lifetime Imaging of O2 with a Frequency-Domain-Based Camera System

Cerebral Blood Oxygenation Measurement Based on Oxygen-dependent Quenching of Phosphorescence

Monitoring of the spatiotemporal characteristics of cerebral blood and tissue oxygenation is crucial for better understanding of the 
neuro-metabolic-vascular relationship. Development of new pO2 measurement modalities with simultaneous monitoring of pO2 in larger fields of view with higher 
spatial and/or temporal resolution will enable greater insight into the functioning of the normal brain and will also have significant impact on diagnosis 
and treatment of neurovascular diseases such as stroke, Alzheimer's disease, and head injury.
Optical imaging modalities have shown a great potential to provide high spatiotemporal resolution and quantitative imaging of pO2 based on hemoglobin absorption in visible and near infrared range of optical spectrum. However, multispectral measurement of cerebral blood oxygenation relies on photon migration through the highly scattering brain tissue. Estimation and modeling of tissue optical parameters, which may undergo dynamic changes during the experiment, is typically required for accurate estimation of blood oxygenation. On the other hand, estimation of the partial pressure of oxygen (pO2) based on oxygen-dependent quenching of phosphorescence should not be significantly affected by the changes in the optical parameters of the tissue and provides an absolute measure of pO2. Experimental systems that utilize oxygen-sensitive dyes have been demonstrated in in vivo studies of the perfused tissue as well as for monitoring the oxygen content in tissue cultures, showing that phosphorescence quenching is a potent technology capable of accurate oxygen imaging in the physiological pO2 range. 
Here we demonstrate with two different imaging modalities how to perform measurement of pO2 in cortical vasculature based on phosphorescence lifetime imaging. In first demonstration we present wide field of view imaging of pO2 at the cortical surface of a rat. This imaging modality has relatively simple experimental setup based on a CCD camera and a pulsed green laser. An example of monitoring the cortical spreading depression based on phosphorescence lifetime of Oxyphor R3 dye was presented. In second demonstration we present a high resolution two-photon pO2 imaging in cortical micro vasculature of a mouse. The experimental setup includes a custom built 2-photon microscope with femtosecond laser, electro-optic modulator, and photon-counting photo multiplier tube. We present an example of imaging the pO2 heterogeneity in the cortical microvasculature including capillaries, using a novel PtP-C343 dye with enhanced 2-photon excitation cross section.
Click here to view the related article <a href="http://www.jove.com/Details.php?ID=1731">Synthesis and Calibration of Phosphorescent Nanoprobes for Oxygen Imaging in Biological Systems.</a>

We present an experimental procedure for measuring the partial pressure of oxygen (pO2) in cerebral vasculature based on oxygen-dependent quenching of phosphorescence. Animal preparation and imaging procedures were outlined for both large field of view CCD-based imaging of pO2 in rats and 2-photon excitation based imaging of pO2 in mice.

Monitoring of the spatiotemporal characteristics of cerebral blood and tissue oxygenation is crucial for better understanding of the 
...

Neuroscience

Fluorescence Lifetime Imaging of Molecular Rotors in Living Cells

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular events. Viscosity is one of the key parameters affecting the diffusion of molecules and proteins, and changes in viscosity have been linked to disease and malfunction at the cellular level.1-3 While methods to measure the bulk viscosity are well developed, imaging microviscosity remains a challenge. Viscosity maps of microscopic objects, such as single cells, have until recently been hard to obtain. Mapping viscosity with fluorescence techniques is advantageous because, similar to other optical techniques, it is minimally invasive, non-destructive and can be applied to living cells and tissues.
Fluorescent molecular rotors exhibit fluorescence lifetimes and quantum yields which are a function of the viscosity of their microenvironment.4,5 Intramolecular twisting or rotation leads to non-radiative decay from the excited state back to the ground state. A viscous environment slows this rotation or twisting, restricting access to this non-radiative decay pathway. This leads to an increase in the fluorescence quantum yield and the fluorescence lifetime. Fluorescence Lifetime Imaging (FLIM) of modified hydrophobic BODIPY dyes that act as fluorescent molecular rotors show that the fluorescence lifetime of these probes is a function of the microviscosity of their environment.6-8 A logarithmic plot of the fluorescence lifetime versus the solvent viscosity yields a straight line that obeys the F&ouml;rster Hoffman equation.9 This plot also serves as a calibration graph to convert fluorescence lifetime into viscosity.
Following incubation of living cells with the modified BODIPY fluorescent molecular rotor, a punctate dye distribution is observed in the fluorescence images. The viscosity value obtained in the puncta in live cells is around 100 times higher than that of water and of cellular cytoplasm.6,7 Time-resolved fluorescence anisotropy measurements yield rotational correlation times in agreement with these large microviscosity values. Mapping the fluorescence lifetime is independent of the fluorescence intensity, and thus allows the separation of probe concentration and viscosity effects. 
In summary, we have developed a practical and versatile approach to map the microviscosity in cells based on FLIM of fluorescent molecular rotors.

Fluorescence Lifetime Imaging (FLIM) has emerged as a key technique to image the environment and interaction of specific proteins and dyes in living cells. FLIM of fluorescent molecular rotors allows mapping of viscosity in living cells.

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular ...

Bioengineering

Water in Oil Emulsions: A New System for Assembling Water-soluble Chlorophyll-binding Proteins with Hydrophobic Pigments

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. Their functionality critically depends on their specific organization within large and elaborate multisubunit transmembrane protein complexes. In order to understand at the molecular level how these complexes facilitate solar energy conversion, it is essential to understand protein-pigment, and pigment-pigment interactions, and their effect on excited dynamics. One way of gaining such understanding is by constructing and studying complexes of Chls with simple water-soluble recombinant proteins. However, incorporating the lipophilic Chls and BChls into water-soluble proteins is difficult. Moreover, there is no general method, which could be used for assembly of water-soluble proteins with hydrophobic pigments. Here, we demonstrate a simple and high throughput system based on water-in-oil emulsions, which enables assembly of water-soluble proteins with hydrophobic Chls. The new method was validated by assembling recombinant versions of the water-soluble chlorophyll binding protein of Brassicaceae plants (WSCP) with Chl a. We demonstrate the successful assembly of Chl a using crude lysates of WSCP expressing E. coli cell, which may be used for developing a genetic screen system for novel water-soluble Chl-binding proteins, and for studies of Chl-protein interactions and assembly processes.

This manuscript describes a simple and high-throughput method for assembling water-soluble proteins with hydrophobic pigments that is based on water-in-oil emulsions. We demonstrate the effectiveness of the method in the assembly of native chlorophylls with four variants of recombinant water-soluble-chlorophyll binding proteins (WSCPs) of Brassica plants expressed in E. coli.

Chlorophylls (Chls) and bacteriochlorophylls (BChls) are the primary cofactors that carry out photosynthetic light harvesting and electron transport. ...

Imaging Amyloid Tissues Stained with Luminescent Conjugated Oligothiophenes by Hyperspectral Confocal Microscopy and Fluorescence Lifetime Imaging

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease afflicting primarily the central nervous system, and systemic amyloidosis where serum amyloid A, transthyretin and IgG light chains deposit as amyloid in liver, carpal tunnel, spleen, kidney, heart, and other peripheral tissues. Amyloid has been known and studied for more than a century, often using amyloid specific dyes such as Congo red and Thioflavin T (ThT) or Thioflavin (ThS). In this paper, we present heptamer-formyl thiophene acetic acid (hFTAA) as an example of recently developed complements to these dyes called luminescent conjugated oligothiophenes (LCOs). hFTAA is easy to use and is compatible with co-staining in immunofluorescence or with other cellular markers. Extensive research has proven that hFTAA detects a wider range of disease associated protein aggregates than conventional amyloid dyes. In addition, hFTAA can also be applied for optical assignment of distinct aggregated morphotypes to allow studies of amyloid fibril polymorphism. While the imaging methodology applied is optional, we here demonstrate hyperspectral imaging (HIS), laser scanning confocal microscopy and fluorescence lifetime imaging (FLIM). These examples show some of the imaging techniques where LCOs can be used as tools to gain more detailed knowledge of the formation and structural properties of amyloids. An important limitation to the technique is, as for all conventional optical microscopy techniques, the requirement for microscopic size of aggregates to allow detection. Furthermore, the aggregate should comprise a repetitive &#946;-sheet structure to allow for hFTAA binding. Excessive fixation and/or epitope exposure that modify the aggregate structure or conformation can render poor hFTAA binding and hence pose limitations to accurate imaging.

Amyloid deposition is a hallmark of different diseases and afflicts many different organs. This paper describes the application of luminescent conjugated oligothiophene fluorescence staining in combination with fluorescence microscopy techniques. This staining method represents a powerful tool for detection and exploration of protein aggregates in both clinical and scientific setups.

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find ...

Biochemistry

Visualization of Ambient Mass Spectrometry with the Use of Schlieren Photography

This manuscript outlines how to visualize mass spectrometry ambient ionization sources using schlieren photography. In order to properly optimize the mass spectrometer, it is necessary to characterize and understand the physical principles of the source. Most commercial ambient ionization sources utilize jets of nitrogen, helium, or atmospheric air to facilitate the ionization of the analyte. As a consequence, schlieren photography can be used to visualize the gas streams by exploiting the differences in refractive index between the streams and ambient air for visualization in real time. The basic setup requires a camera, mirror, flashlight, and razor blade. When properly configured, a real time image of the source is observed by watching its reflection. This allows for insight into the mechanism of action in the source, and pathways to its optimization can be elucidated. Light is shed on an otherwise invisible situation.

This paper presents a protocol for the visualization of gaseous streams of an ambient ionization source using schlieren photography and mass spectrometry.

This manuscript outlines how to visualize mass spectrometry ambient ionization sources using schlieren photography. In order to properly optimize the mass ...

Time-resolved Photophysical Characterization of Triplet-harvesting Organic Compounds at an Oxygen-free Environment Using an iCCD Camera

Here, we present a sensible method of the acquisition and analysis of time-resolved photoluminescence using an ultrafast iCCD camera. This system enables the acquisition of photoluminescence spectra covering the time regime from nanoseconds up to 0.1 s. This enables us to follow the changes in the intensity (decay) and emission of the spectra over time. Using this method, it is possible to study diverse photophysical phenomena, such as the emission of phosphorescence, and the contributions of prompt and delayed fluorescence in molecules showing thermally activated delayed fluorescence (TADF). Remarkably, all spectra and decays are obtained in a single experiment. This can be done for solids (thin film, powder, crystal) and liquid samples, where the only limitations are the spectral sensitivity of the camera and the excitation wavelength (532 nm, 355 nm, 337 nm, and 266 nm). This technique is, thus, very important when investigating the excited state dynamics in organic emitters for their application in organic light-emitting diodes and other areas where triplet harvesting is of paramount importance. Since triplet states are strongly quenched by oxygen, emitters with efficient TADF luminescence, or those showing room temperature phosphorescence (RTP), must be correctly prepared in order to remove any dissolved oxygen from solutions and films. Otherwise, no long-lived emission will be observed. The method of degassing solid samples as presented in this work is basic and simple, but the degassing of liquid samples creates additional difficulties and is particularly interesting. A method of minimizing solvent loss and changing the sample concentration, while still enabling to remove oxygen in a very efficient and a repeatable manner, is presented in this work.

Here, we present a method of the spectroscopic characterization of organic molecules by means of time-resolved photoluminescence spectroscopy on the nanosecond-to-millisecond timescale in oxygen-free conditions. Methods to efficiently remove oxygen from the samples and, thus, limit luminescence quenching are also described.

Here, we present a sensible method of the acquisition and analysis of time-resolved photoluminescence using an ultrafast iCCD camera. This system enables ...

Synthesis and Calibration of Phosphorescent Nanoprobes for Oxygen Imaging in Biological Systems

Oxygen measurement by phosphorescence quenching [1, 2] consists of the following steps: 1) the probe is delivered into the medium of interest (e.g. blood or interstitial fluid); 2) the object is illuminated with light of appropriate wavelength in order to excite the probe into its triplet state; 3) the emitted phosphorescence is collected, and its time course is analyzed to yield the phosphorescence lifetime, which is converted into the oxygen concentration (or partial pressure, pO2). The probe must not interact with the biological environment and in some cases to be 4) excreted from the medium upon the measurement completion. Each of these steps imposes requirements on the molecular design of the phosphorescent probes, which constitute the only invasive component of the measurement protocol. Here we review the design of dendritic phosphorescent nanosensors for oxygen measurements in biological systems. The probes consist of Pt or Pd porphyrin-based polyarylglycine (AG) dendrimers, modified peripherally with polyethylene glycol (PEG's) residues. For effective two-photon excitation, termini of the dendrimers may be modified with two-photon antenna chromophores, which capture the excitation energy and channel it to the triplet cores of the probes via intramolecular FRET (F&ouml;rster Resonance Energy Transfer). We describe the key photophysical properties of the probes and present detailed calibration protocols.

We present principles of oxygen measurements by phosphorescence quenching and review design of porphyrin-based dendritic nanosensors for oxygen imaging in biological systems.

Oxygen measurement by phosphorescence quenching [1, 2] consists of the following steps: 1) the probe is delivered into the medium of interest (e.g. blood ...

Biology

Electron Spin Resonance Micro-imaging of Live Species for Oxygen Mapping

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment of live cells with micron-scale resolution1. Oxygen is one of the most important molecules in the cycle of life. It serves as the terminal electron acceptor of oxidative phosphorylation in the mitochondria and is used in the production of reactive oxygen species. Measurements of oxygen are important for the study of mitochondrial and metabolic functions, signaling pathways, effects of various stimuli, membrane permeability, and disease differentiation. Oxygen consumption is therefore an informative marker of cellular metabolism, which is broadly applicable to various biological systems from mitochondria to cells to whole organisms. Due to its importance, many methods have been developed for the measurements of oxygen in live systems. Current attempts to provide high-resolution oxygen imaging are based mainly on optical fluorescence and phosphorescence methods that fail to provide satisfactory results as they employ probes with high photo-toxicity and low oxygen sensitivity. ESR, which measures the signal from exogenous paramagnetic probes in the sample, is known to provide very accurate measurements of oxygen concentration. In a typical case, ESR measurements map the probe's lineshape broadening and/or relaxation-time shortening that are linked directly to the local oxygen concentration. (Oxygen is paramagnetic; therefore, when colliding with the exogenous paramagnetic probe, it shortness its relaxation times.) Traditionally, these types of experiments are carried out with low resolution, millimeter-scale ESR for small animals imaging. Here we show how ESR imaging can also be carried out in the micron-scale for the examination of small live samples. ESR micro-imaging is a relatively new methodology that enables the acquisition of spatially-resolved ESR signals with a resolution approaching 1 micron at room temperature2. The main aim of this protocol-paper is to show how this new method, along with newly developed oxygen-sensitive probes, can be applied to the mapping of oxygen levels in small live samples. A spatial resolution of ~30 x 30 x 100 &mu;m is demonstrated, with near-micromolar oxygen concentration sensitivity and sub-femtomole absolute oxygen sensitivity per voxel. The use of ESR micro-imaging for oxygen mapping near cells complements the currently available techniques based on micro-electrodes or fluorescence/phosphorescence. Furthermore, with the proper paramagnetic probe, it will also be readily applicable for intracellular oxygen micro-imaging, a capability which other methods find very difficult to achieve.

This protocol describes a method for micron-scale three-dimensional imaging of oxygen concentration in the immediate environment of live cells by electron spin resonance microscopy.

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment ...

Flexible Measurement of Bioluminescent Reporters Using an Automated Longitudinal Luciferase Imaging Gas- and Temperature-optimized Recorder (ALLIGATOR)

Luciferase-based reporters of cellular gene expression are in widespread use for both longitudinal and end-point assays of biological activity. In circadian rhythms research, for example, clock gene fusions with firefly luciferase give rise to robust rhythms in cellular bioluminescence that persist over many days. Technical limitations associated with photomultiplier tubes (PMT) or conventional microscopy-based methods for bioluminescence quantification have typically demanded that cells and tissues be maintained under quite non-physiological conditions during recording, with a trade-off between sensitivity and throughput. Here, we report a refinement of prior methods that allows long-term bioluminescence imaging with high sensitivity and throughput which supports a broad range of culture conditions, including variable gas and humidity control, and that accepts many different tissue culture plates and dishes. This automated longitudinal luciferase imaging gas- and temperature-optimized recorder (ALLIGATOR) also allows the observation of spatial variations in luciferase expression across a cell monolayer or tissue, which cannot readily be observed by traditional methods. We highlight how the ALLIGATOR provides vastly increased flexibility for the detection of luciferase activity when compared with existing methods.

Flexible Measurement of Bioluminescent Reporters Using an Automated Longitudinal Luciferase Imaging Gas- and Temperature-optimized Recorder ALLIGATOR

Genetically encoded luciferase is a popular non-invasive reporter of gene expression. Use of an automated longitudinal luciferase imaging gas- and temperature-optimized recorder (ALLIGATOR) enables longitudinal recording from bioluminescent cells under a wide range of conditions. Here we show how ALLIGATOR may be used in the context of circadian rhythms research.

Luciferase-based reporters of cellular gene expression are in widespread use for both longitudinal and end-point assays of biological activity. In ...

Fluorescence Lifetime Macro Imager for Biomedical Applications

This paper presents a new photoluminescence lifetime imager designed to map the molecular oxygen (O2) concentration in different phosphorescent samples ranging from solid-state, O2-sensitive coatings to live animal tissue samples stained with soluble O2-sensitive probes. In particular, the nanoparticle-based near-infrared probe NanO2-IR, which is excitable with a 625 nm light-emitting diode (LED) and emits at 760 nm, was used. The imaging system is based on the Timepix3 camera (Tpx3Cam) and the opto-mechanical adaptor, which also houses an image intensifier. O2 phosphorescence lifetime imaging microscopy (PLIM) is commonly required for various studies, but current platforms have limitations in their accuracy, general flexibility, and usability.
The system presented here is a fast and highly sensitive imager, which is built on an integrated optical sensor and readout chip module, Tpx3Cam. It is shown to produce high-intensity phosphorescence signals and stable lifetime values from surface-stained intestinal tissue samples or intraluminally stained fragments of the large intestine and allows the detailed mapping of tissue O2 levels in about 20 s or less. Initial experiments on the imaging of hypoxia in grafted tumors in unconscious animals are also presented. We also describe how the imager can be re-configured for use with O2-sensitive materials based on Pt-porphyrin dyes using a 390 nm LED for the excitation and a bandpass 650 nm filter for emission. Overall, the PLIM imager was found to produce accurate quantitative measurements of lifetime values for the probes used and respective two-dimensional maps of the O2 concentration. It is also useful for the metabolic imaging of ex vivo tissue models and live animals.

This paper describes the use of a new, fast optical imager for the macroscopic photoluminescence lifetime imaging of long decay emitting samples. The integration, image acquisition, and analysis procedures are described, along with the preparation and characterization of the sensor materials for the imaging and the application of the imager in studying biological samples.

This paper presents a new photoluminescence lifetime imager designed to map the molecular oxygen (O2) concentration in different phosphorescent samples ...

Luminescence Lifetime Imaging of O₂ with a Frequency-Domain-Based Camera System

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