OptoAssay—Light-controlled dynamic bioassay using optogenetic switches

Circumventing the limitations of current bioassays, we introduce a light-controlled assay, OptoAssay, toward wash- and pump-free point-of-care diagnostics. Extending the capabilities of standard bioassays with light-dependent and reversible interaction of optogenetic switches, OptoAssays enable a bidirectional movement of assay components, only by changing the wavelength of light. Demonstrating exceptional versatility, the OptoAssay showcases its efficacy on various substrates, delivering a dynamic bioassay format. The applicability of the OptoAssay is successfully demonstrated by the calibration of a competitive model assay, resulting in a superior limit of detection of 8 pg ml−1, which is beyond those of conventional ELISA tests. In the future, combined with smartphones, OptoAssays could obviate the need for external flow control systems such as pumps or valves and signal readout devices, enabling on-site analysis in resource-limited settings.


Main
Point-of care (POC) diagnostics is the key player for fast interventions that might have critical value for patient's outcome. For this purpose, POC testing that allows for rapid diagnostics in non-laboratory settings carried out by untrained personnel has become more commonplace over the last decade. Its importance has even more deepened during the ongoing Covid19 pandemic. One of the most used POC formats are paper-based devices like lateral flow assays (LFAs), where the sample is added on a cellulose-based test stripe and transported through capillary forces along the stripe. This, however, only allows for a uni-directional sample flow which fundamentally limits the flexibility in assay design and sample processing 1,2 . On the other hand, other POC devices offering bidirectional microfluidics suffer from expensive and bulky pumps or flow control systems, also requiring an additional energy source, that complicates their on-site use 3,4 .
To circumvent this restraint, we present for the first time the proof-of-concept of a light-controlled dynamic bioassay (OptoAssay). OptoAssay allows for bi-directional movement of biomolecules, enabling wash-free signal readout of bioassays. For this purpose, the biomolecules fused to a phytochrome interacting factor 6 (PIF6) can be released from and rebind to the plant photoreceptor phytochrome B (PhyB) by exposure to far-red or red light, respectively, thereby circumventing the need for external flow control devices. PhyB exhibits two distinct light absorbing conformations ( Figure 1A): a red-light absorbing Pr (red) state with λmax ~ 651 nm 5 nm and a far-red light absorbing, biologically active, Pfr (far red) state with λmax ~ 713 nm. In the active state, PhyB can bind proteins from a class called phytochrome interacting factors while in the inactive state this interaction is reversed 6,7 . Up to date, this switching behavior of the PhyB/PIF systems has been used in various applications to control, for example, gene expression 8 , cell signalling 9 or to create biohybrid materials 10 .
Here, we apply optogenetic switches for realizing the first light-controlled dynamic bioassay on a nitrocellulose substrate and successfully demonstrate its proof-of-principle, detecting a hexahistidinetag (referred to as his-tag) as analyte. Besides, we show the general versatility of the assay components on other substrates, namely agarose beads and poly(methyl methacrylate) (PMMA), employed for building POC devices.

Results
The concept of the OptoAssay consists of two distinct areas ( Figure 1B): a smaller receiver area where the signal readout is performed, is placed in the middle of the sender area into a notch so that the areas are not directly connected. These areas are later bridged by adding sample solution to both sides so that the assay components can diffuse from one area to the other. In this work, the OptoAssay is carried out as an immunoassay using antibodies. Herein, a competitive assay format was chosen, in which a competitor and the analyte in the sample compete for binding to the detection antibody. The sender area ( Figure 1C) is treated with neutravidin (nAv) so that the light sensitive components PhyB can be immobilized via a biotin-neutravidin interaction. Here, PIF6 can bind to PhyB or be released depending on the wavelength applied. On the other hand, antibodies directed against the molecule of interest are immobilized on the receiver area.
For the proof-of-principle assay, we use a his-tagged protein (see Methods) as analyte and his-tagged PIF6 as competitor, both of which are recognized by the anti his-tag antibody immobilized in the receiver area. The signal readout is performed by measuring the fluorescence intensity of the competitor complex, his-tagged PIF6, to which green fluorescent protein (GFP) is fused. For the initial OptoAssay configuration, the competitor complex is attached to PhyB on the sender area during red light exposure. The assay procedure itself comprises of two steps ( Figure 1B): first, far-red light illumination is used to release the competitor complex from PhyB. At the same time, the analyte is added. Now both the analyte and the competitor can compete for binding to the antibody immobilized on the receiver area. Second, red light is applied to re-associate unbound competitor complexes in the sender area and thus, to remove them from the receiver area. Due to the competitive assay format, the signal measured on the receiver area is inverse proportional to the concentration of the analyte.
In order to enable POC testing using the OptoAssay, a 3D printed PhotoBox that allows for illumination with red and far-red light has been built (Figure1 D,E) 11 . The illumination can be controlled with a smartphone that is linked via a Bluetooth module to the electronics of the PhotoBox. Images of the OptoAssay results can be then easily read out, evaluated, and further transmitted to, for example, medical facilities using a smartphone.
As substrate material for immobilization of assay components, we chose nitrocellulose since it is widely applied for LFA devices due to its non-specific affinity to proteins 12,13 . We first investigated an appropriate blocking method for nitrocellulose to cover unoccupied spaces after the protein of interest has been immobilized. Therefore, we examined the blocking performance of bovine serum albumin (BSA) and casein. The results ( Figure S1) suggest that albeit casein has a better blocking performance, it seems to mask the previous immobilized proteins, rendering them inaccessible. Conclusively, we employed BSA as blocking agent for subsequent experiments.
Since the PhyB/PIF interaction plays the key role in our experimental setup, we tested whether a competitor complex containing two PIF proteins shows better performance in terms of leakiness, i.e., fewer unspecific release during red-light illumination due to increased avidity. Therefore, the release of competitor complexes containing one or two PIF proteins was measured over a time range of 80 minutes. Additionally, to determine the unspecific release of the nitrocellulose membrane itself, the release of a biotinylated version of GFP-PIF6, immobilized directly on nAv, was determined.
According to our findings (Figure S2), the double PIF version has no advantages over the single version regarding the unspecific release (red light, 660 nm). In fact, the release of competitor molecules can be attributed to the unspecific release of the nitrocellulose substrate itself as the biotin-GFP-PIF6 version shows similar values than the two other non-released samples. However, far-red light illumination (740 nm) yields a higher release for the single PIF version, which is why we conducted all following experiments with this competitor version.
In order to find an assay architecture in terms of the assay area and their positioning, we tested three different variants regarding the diffusion time and fluid distribution on the distinct areas ( Figure S3).
For naked eye visualization, highly concentrated fluorescent protein mCherry (mCh) or the small molecule fluorescein was used. For both substances, the design variant where a small square shaped receiver area is enclosed by a larger square shaped sender area showed the best performance and was, therefore, employed for the following experiments.
For the proof-of-principle assay, we first demonstrated the light-induced competition of analyte and competitor. To simulate and verify the competitive mode of the OptoAssay, three conditions were tested: (i) red light illumination without analyte where no release of the competitor is expected, (ii) far-red light illumination without analyte where a release but no competition of analyte and competitor is possible, and (iii) far-red light illumination with analyte, where both a release and competition are expected. The fluorescence intensity measurements of the nitrocellulose membranes after illumination show a strong decrease in fluorescence signal of the sender membranes illuminated with far-red light compared to those illuminated with red light. Far-red light illuminated samples without analyte display a higher fluorescence in the receiver area than the ones where analyte was added (Figure 2A). This can also be confirmed by quantifying fluorescence intensities of the membranes ( Figure 2B). Also, the measurements of the supernatant collected (Figure 1confirm this observation as there is a lower fluorescence intensity for the far-red light illuminated samples without analyte than analyte-containing samples. When comparing the leakiness (i.e., unspecific release of competitor during red light illumination) to the red-light induced specific release by dividing the fluorescence intensity of the far-red by the red light illuminated sample, there is a 6.1-times increase in signals of the sample with analyte but only a 4.5-times signal increase for the sample without analyte. This could be due to the competitor binding to the receiver area and, therefore, leaving less competitor in the supernatant. While there is a high fluorescence intensity obtained on the receiver from the sample treated with far-red light and without analyte, only a low fluorescence is detectable for the far-red light illuminated samples with analyte. A quantification of the fluorescence intensities of receiver membranes after illumination shows a 115% signal increase when comparing far-red light illuminated samples with red light illuminated ones without analyte and only a 22% increase in signals of far-red light illuminated membranes with analyte. This results in a fluorescent signal increase of 163% when comparing far-red light illuminated samples with and without analyte.
The last step of the OptoAssay comprises of the rebinding of the unbound competitor molecules back to PhyB on the sender membrane which could enable a wash-free signal readout. To achieve this, one sample was illuminated first with far-red light to release the competitor and then with red light to rebind it. Control samples that were either illuminated with red or far-red light only were included as controls. The fluorescence images of the membranes ( Figure 2D) after illumination indicate that the control samples worked as anticipated. In the case of the sender membranes, the measured intensity of the far-red sample is lower than the red light one. Also, the binding of the competitor on the receiver membrane can be observed for the (far-red light illuminated) release samples ( Figure 2E). The rebinding samples (illuminated with far-red and then red light) display a slight increase in fluorescence intensity of sender membranes. The rebinding of the unbound competitor was also demonstrated by measuring the supernatant fluorescence ( Figure 2F) which corresponds the amount of free, unbound competitor. Here, the fluorescence intensity of the rebinding sample has only is only 61% compared to the release sample. This means that about 39% of the released competitor could be rebound to the sender membrane. When comparing the membrane intensities ( Figure 2E) before and after illumination, a reduction of fluorescence of the leakiness sample (red light) of 5% and 2% for sender and receiver, respectively, was observed. The release sample shows 36% reduction of intensity on the sender and a 23% increase on the receiver membrane. In the case of the rebinding sample, however, there is only a reduction of only 18% on the sender membrane which indicates a partial rebinding. The receiver membranes prove only an increase of 7%. For this experiment, lower amounts of competitor were applied compared to the previous experiment to ensure a higher rebinding efficiency, as depicted in Figure S4. The general lower fluorescence intensity of the rebinding samples, however, could be attributed to the longer incubation time of 80 minutes compared with 40 minutes used for the other samples. The longer incubation period might have led to higher dissociation of PhyB and PIF6 or even the unspecific release of nAv from the nitrocellulose membrane (see Figure S2).
Having successfully demonstrated the proof-of-concept of the OptoAssay on a nitrocellulose substrate, we aimed to the test the general functionality of the optogenetic components on other substrates. As an alternative planar substrate, we used PMMA which is low-priced and thus convenient for mass production of a potential POC device. A preliminary experiment ( Figure S5) showed that nAv directly absorbs to untreated PMMA which circumvents the need for a surface functionalization. Subsequently, we immobilized biotinylated PhyB on the nAv coated surface. With this setup, we could demonstrate a spatially resolved release of the competitor complex from a PMMA substrate ( Figure S6). However, we did not pursue with PMMA as a substrate for the OptoAssay because the total amount of immobilized photoreceptor was lower compared to nitrocellulose.
The next substrate tested were functionalized beads that are very often used as solid phase materials in bioassay applications in clinical and POC diagnostics 14,15 . In this setting, we verified the functionality of the optogenetic system on a bead-based platform where nAv functionalized agarose beads are employed to immobilize the biotinylated PhyB. We simulated the reversibility of releasing and rebinding the PIF6 molecule as the competitor from PhyB as described in 10 , immobilized on agarose beads ( Figure S7). We could again show a light-dependent release as well as partial rebinding of competitor molecules from and to the substrate suggesting the likely generic applicability of the OptoAssay in various assay formats.

Conclusion and Outlook
Here, we have successfully demonstrated the proof-of-principle of an OptoAssay that enables, in contrast to conventional POC tests like LFAs, a bi-directional sample flow without any pump or flow control systems for a wash-free signal readout. Through a photoreceptor that can light-dependently interact with a binding partner, a competitor molecule can be released and later removed from the detection area by applying far-red and red light. Although other photoswitchable assays already exist 16,17 , they only allow for detection of specific molecules, whereas the OptoAssay can be universally applied by fusing the molecule of interest to the phytochrome B interaction partner PIF6. For example, the integrating of a z domain 18 , an IgG-binding domain, antibodies can be attached to the PIF6 construct and act as detection antibodies to add even more versatility.
The system could also be further expanded by using or combining different optogenetic switches that respond to distinct wavelengths. For instance, the blue light receptor cryptochrome 2 (Cry2) that forms heterodimers with its interaction partner cryptochrome-interacting basic helix-loop-helix (CIB1), or another system, light-induced dimer (iLID), which comprises of the blue light receptor light oxygen voltage 2 domain of Avena sativa phototropin 1-SsrA (AsLOV2) and its binding partner stringent starvation protein B (SspB) could be employed 19 . However, both photoreceptors only enable an active associating with its interaction partner, the dissociation is usually a slower passive event induced by the absence of blue light. Therefore, the OptoAssay workflow might need to be adapted accordingly in order to use different photoreceptors.
Using a 3D printed PhotoBox along with a smartphone, the OptoAssay introduced can be employed for on-site applications. Herein, the PhotoBox could be extended by an excitation light and emission filter that allows for GFP detection using a smartphone. Various low-cost and easy sensing approaches in combination with a smartphone for a fluorescence readout have already been implemented 20 .
Finally, automated sample illumination and evaluation of the results via a smartphone app could be realized for user friendly operation.
Another issue that must be addressed is the overall operation time of the OptoAssay which is, at the moment, mainly limited by diffusion of the biomolecules employed. Herein, there are two important factors: the overall distance the molecules have to travel between the two areas (i.e., detection and immobilization zones) and the speed at which the molecules move. To decrease the distance, smaller assay areas could be designed and fabricated by micro/nanofabrication techniques. For increasing the speed, external energy, like ultrasonication, could be also applied to the system.
The OptoAssay provides the basics for a novel and dynamic bioassay format (independent of assay type and biomolecules employed, such as antibodies, proteins, or nucleic acids) through its light dependent two-way switch that can extend already existing POC devices and could pave the way for new classes of diagnostic devices in future, extending the capabilities of these state-of-the-art tools.  Fluorescence images of nitrocellulose membranes before and after illumination. Sender membranes were incubated with nAv and then blocked with BSA. Next, biotinylated PhyB was immobilized. The competitor complex (his-GFP-PIF6, 30 mg ml -1 ) was bound to PhyB under red light illumination. The receiver membranes were coated with anti his-tag antibody and subsequently blocked with BSA. For the measurement, sender and receiver membranes were assembled and covered with buffer or buffer containing the his-tagged analyte protein which was added at the same time as illumination started.
The samples were illuminated from above either with red or far-red light for 40 minutes. The membranes were washed under illumination with the corresponding light and subsequently imaged.

Plasmids and oligos
The plasmids generated and used in this study are described in Table 1 along with the templates and oligos for the polymerase chain reaction (PCR) to generate plasmids using Gibson cloning 21 . The sequences of the oligos used are listed in Table 2.  Protein production and purification

PIF6 constructs
For protein production, the corresponding plasmid (table xx)

Data analysis
The images were analyzed using the software ImageJ (University of Wisconsin-Madison, USA). The mean intensity values of the whole nitrocellulose membranes/PMMA pieces for the illuminated and not illuminated areas were analyzed with the area selection tool. In order to determine the final value of intensity, the mean background intensity was subtracted, and the obtained values were normalized.
For normalization, intensity values of plain buffer were subtracted from each value.     Figure 2D with the same contrast settings as Figure 2A. only. On the third sample, nAv was first adsorbed followed by blocking with 5% BSA. Then, GFP-PIF6biotin was added. The images were taken after washing the samples.