A variety of optogenetic tools have been successfully developed based on specific LOV domain proteins possessing α-helices that change conformation in response to light (Weitzman and Hahn, 2014). We reasoned that fusing a LOV domain to a helical domain of Cre that is critical for its function could generate a single protein with light-dependent recombinase activity. We searched for candidate α-helices by inspecting the structure of the four Cre units complexed with two LoxP DNA targets (Ennifar et al., 2003; Guo et al., 1997; Figure 1a, b). Each subunit folds in two domains that bind to DNA as a clamp. It was initially reported that helices αA and αE of the amino-terminal domain, as well as helix αN of the C-terminal domain, participate to inter-units contacts (Guo et al., 1997); and this role of helix αN was later confirmed (Ennifar et al., 2003). Contacts between αA and αE associate all four amino-terminal domains (Figure 1a), and contacts involving αN lock the four carboxy-terminal domains in a cyclic manner (Figure 1b). These helices were therefore good candidates for manipulating Cre activity. We focused on αA and αN because their location at protein extremities was convenient to design chimeric fusions.

Figure 1

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We tested the functional importance of helices αA and αN by gradually eroding them. We evaluated the corresponding mutants by expressing them in yeast cells, where an active Cre can excise a repressive DNA element flanked by LoxP sites and thereby switch ON the expression of a green fluorescent protein (GFP) (Figure 1c). After inducing the expression of Cre mutants with galactose, we counted by flow cytometry the proportion of cells that expressed GFP and used this measure to compare recombinase activities of the different mutants (Figure 1d). As a control, we observed that the wild-type Cre protein activated GFP expression in all cells under these conditions. Mutants lacking the last two or the last three carboxy-terminal residues displayed full activity. In contrast, mutants lacking four or more of the C-ter residues were totally inactive. This was consistent with a previous observation that deletion of the last 12 residues completely suppressed activity (Rongrong et al., 2005). Our series of mutants showed that helix αN is needed for activity and that its residue E340 (located at position −4 from the protein end) is implicated. The role of this glutamic acid is most likely to stabilize the complex: the tetramer structure indicates that it interacts with residue R192 of the adjacent unit (Figure 1e). Interestingly, although D341 was not essential for activity, the salt bridges of this residue with R139 probably also reinforce attraction between the two protein units. Biomolecular simulations using a simplistic force-field model showed that the free-energy barrier for displacing the αN helix was much lower if E340 and D341 were replaced by alanines (Figure 1f). Consistent with this prediction, we observed that a double mutant E340A D341A lost ~10% of activity (Figure 1g). This mild (but reproducible) reduction of activity suggested that the double mutation E340A D341A led to a variant of Cre where multimerization was suboptimal.

We also tested the functional importance of α-helix A either in a normal context where the C-terminal part of Cre was intact or where it carried the destabilizing E340A D341A mutation (Figure 1g). Deletion of residues 2–37, which entirely ablated helix A, eliminated enzymatic activity (Figure 1g). Very interestingly, the effect of shorter deletions depended on the C-terminal context. When the C-terminus was wild-type, removing residues 2–21 (immediately upstream of helix A) had no effect and removing residues 2–28 (partial truncation of αA) decreased the activity by ~10%. When the C-terminus contained the E340A D341A mutation, deletions 2–21 and 2–28 were much more severe, reducing the activity by 12% and 80%, respectively. This revealed genetic interactions between the extremities of the protein, which is fully consistent with a cooperative role of helices αA and αN in stabilizing an active tetramer complex. From these observations, we considered that photo-control of Cre activity might be possible by fusing αA and αN helices to LOV domain photoreceptors.

We obtained successful fusions (Figure 2) after screening several types of candidate constructs. Our first strategy was to fuse the αN carboxy-terminal helix of Cre to the amino-terminal cap of the LOV domain of protein VVD, a well-characterized photosensor from N. crassa (Nihongaki et al., 2014; Zoltowski et al., 2009; Vaidya et al., 2011). The resulting chimeric protein, which contained the full-length Cre connected to VVD via four amino acids, did not display light-dependent recombinase activity (Figure 2—figure supplement 1). Introducing the (E340A D341A) double mutation in this chimeric protein reduced the overall activity – as expected from above – but did not cause light dependency. Changing the size of the linker or introducing different mutations at positions 340 and 341 also failed to generate light-dependency (Figure 2—figure supplement 1). We noted, however, that the activity of mutants (E340R D341A) and (E340R D341R) was markedly reduced, probably because attraction between E340 and R192 was changed into repulsion between two positively charged arginines.

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Our next strategy was based on a modified version of the AsLOV2 domain from Avena sativa, which had been optimized and used to build an optogenetic dimerizer by fusing its Jα C-ter helix to the bacterial SsrA peptide (Guntas et al., 2015). Instead, we fused Jα to the αA amino-terminal helix of Cre. Using the same GFP reporter system as described above for detecting in vivo recombination in yeast, we built a panel of constructs with various fusion positions and directly quantified their activity with and without blue-light illumination.

All fusions displayed reduced activity in both dark and light conditions compared to wild-type Cre. Although variability was high between independent transformants, we identified four constructs – corresponding to fusions of AsLOV2 to residues 17, 19, 27, and 32 of Cre, respectively – where the assay indicated a higher activity after light stimulation (Figure 2—figure supplement 2). To confirm this, we recovered the corresponding plasmids from individual yeast transformants, amplified them in bacteria to verify their sequence, and re-transformed them in yeast. This validated a differential activity between dark and light conditions for the constructs corresponding to fusion positions 19, 27, and 32 (Figure 2a). Fusion at position 32 (named LOV_Cre32) displayed the highest induction by light, with activity increasing from 15% in dark condition to 50% after 30 min of illumination. Although this induction was significant, a 15% activity of the non-induced form remained too high for most applications. We therefore sought to reduce this residual activity, which we did in two ways.

First, we randomized the residues located at the junction between AsLOV2 and Cre. We used degenerate primers and in vivo recombination (see Materials and methods) to mutagenize LOV_Cre32 at these positions and directly tested the activity of about 90 random clones. Six of them showed evidence of low residual activity in the dark, and we characterized them further by sequencing and re-transformation. For one clone, photo-induction was not confirmed after re-transformation (Figure 2b, clone ii). For the other five clones, residual activity was indeed reduced compared to LOV_Cre32, with the strongest reduction being achieved by an isoleucine insertion at the junction position (Figure 2b, clone v). However, this improvement was also accompanied by a weaker induced activity and a larger variability between independent assays.

As a complementary approach to reduce residual activity, we took advantage of the above-described genetic interaction between N-ter truncations and C-ter mutations targeting residues 340 and 341. We built another series of constructs where AsLOV2 fusions to αA helix were combined with the A340A341 double mutation. Results differed from those observed in the context of a wild-type αN (Figure 2—figure supplement 3), confirming genetic interaction between N-ter and C-ter mutations. In this assay, two constructs – corresponding to fusion positions 20 and 22 – showed higher recombination activity upon blue-light illumination. We extracted, sequence-verified, and re-tested them together with the one corresponding to fusion position 21. This revealed that fusion at position 20, but not 21 or 22, conferred light-dependent activity (Figure 2c). The construct corresponding to fusion of AsLOV2 at position 20 of the Cre A340A341 double mutant displayed a residual activity that was indistinguishable from the negative control and a highly reproducible induced activity of ~25%. Sequencing revealed that it also included a A->D mutation at the fusion junction (Supplementary file 1 – Supplementary Text S2). We called this construct LiCre and characterized it further.

We placed LiCre under the expression of the P_MET17 promoter and tested various lighting conditions. We first used the same illumination system as above, an LED box apparatus commercialized under the name ‘PAUL’, which was originally designed for viability-PCR assay. This device is suitable for continuous illumination at room temperature. We applied varying intensities and durations on cells that were cultured to stationary phase in the absence of methionine (full LiCre expression). Activity was very low without illumination and increased with both the intensity and duration of light stimulation. The minimal intensity required for stimulation comprised between 0.057 and 1.815 mW/cm². The highest activity (~65% of switched cells) was obtained with 90 min illumination at 36.3 mW/cm² (Figure 3a). At comparable duration and intensity (30 min and 36.3 mW/cm²), LiCre generated more recombination events in this experiment than in the previous one (Figure 3a vs. Figure 2c). This improvement likely results from the different expression systems and culture media that were used (induction of the P_MET17vs. P_GAL1 promoter). Extending illumination to 180 min did not further increase the fraction of switched cells. Remarkably, we observed that 2 min of illumination was enough to switch 5% of cells, and 5 min illumination generated 10% of switched cells (Figure 3b).

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We compared these performances with those of two previous systems that were both based on light-dependent complementation of a split Cre enzyme. We constructed plasmids coding for proteins CreN59-nMag and pMag-CreC60 described earlier (Kawano et al., 2016) and transformed them in our yeast reporter strain. Similarly, we constructed and tested plasmids coding for the previously described (Taslimi et al., 2016) proteins CRY2^L348F-CreN and CIB1-CreC. All four coding sequences were placed under the control of the yeast P_MET17 promoter. We analyzed the resulting strains as above after adapting light to match the intensity recommended by the authors (1.815 mW/cm² for nMag/pMag and 5.45 mW/cm² for CRY2^L348F/CIB1). As shown in Figure 3c, we validated the photoactivation of nMag/pMag split Cre in yeast, where activity increased about fourfold following 90 min of illumination, but we were not able to observe photoactivation of the CRY2^L348F/CIB1 split Cre system (Figure 3d). In addition, the photoactivation of nMag/pMag split Cre was not as fast as the one of LiCre since 30 min of illumination was needed to observe a significant increase of activity. This observation is consistent with the fact that dimerization of split Cre, which is not required for LiCre, probably limits the rate of formation of an active recombination synapse. Another difference was that, unlike LiCre, nMag/pMag split Cre displayed a mild but significant background activity in the absence of illumination (~6% of switched cells) (Figure 3c).

We then used another lighting device, a DMX-controlled LED spot, which provided more precise control on both the intensity and dynamics of illumination. We applied varying intensities on reporter cells expressing either LiCre, nMag/pMag split Cre, or CRY2^L348F/CIB1 split Cre (Figure 3e). Absolute efficiencies were weaker than those observed on the PAUL device. This difference is likely due to two factors: the presence of mirrors in the PAUL box (better distribution of light to the cells) and a difference of temperature, which was controlled at 30°C under the DMX spot but not in the PAUL box. Nonetheless, the relative efficiencies were very concordant between the two devices. The DMX spot stimulated an activity of nMag/pMag split Cre that was above background but much weaker than for LiCre, and it did not activate CRY2^L348F/CIB1 split Cre. This was true at all intensities tested. We observed the highest activation of nMag/pMag split Cre at 1.5 mW/cm² intensity, which matched the recommendation of its authors (Kawano et al., 2016). This calibration also revealed that 0.1 mW/cm² was enough – although clearly suboptimal – to stimulate LiCre above its residual activity. Intriguingly, we observed a weaker activity at 35.6 mW/cm² than at 17.9 mW/cm². However, this difference could be artifactual because it was within the range of variability observed in one of the conditions (3.7 mW/cm²). In parallel, we inspected if blue light at maximal intensity was toxic to yeast cells. We scored colony-forming units (CFUs) from cells exposed or not to 35.6 mW/cm² blue light (DMX spot) during 1 hr (Figure 3—figure supplement 1). We observed no significant differences in the number of CFUs generated by illuminated and non-illuminated samples, nor between LiCre-expressing and non-expressing cells. We conclude that this light exposure is safe regarding yeast viability.

We then interrogated the response of LiCre and nMag/pMag split Cre to periodic stimulations. We expected different response dynamics from these systems because AsLOV2 and VVD have different photocycle lifetimes, with a fast (~80 s) and slow (~18,000 s) decay rate of their adduct (activated) form, respectively (Pudasaini et al., 2015). At the optimal energy for nMag/pMag (1.5 mW/cm²), LiCre outperformed nMap/pMag split Cre when light episodes were prolonged but not when they were short (Figure 3f). Brief light pulses occurring 2 min apart were insufficient to activate LiCre but fully stimulated nMag/pMag split Cre (Figure 3f, 5% of lighting time). This is entirely consistent with a faster decay rate for the AsLOV2 domain of LiCre compared to the VVD domain of nMag/nMag. To specifically and more precisely characterize the response dynamics of LiCre, we applied periodic stimulations using a higher energy (35.6 mW/cm²) and a slow (2 min) or a fast (10 s) period (Figure 3g). As expected, the fast regime better stimulated recombination. Remarkably, light pulses of 0.5 s occurring 10 s apart were sufficient to fully stimulate LiCre. Altogether, these results show that, at least in the yeast cellular context, LiCre outperforms the two previous split systems in terms of efficiency, rapidity, and residual background activity.

To demonstrate the control of a biological activity by light, we built a reporter where Cre-mediated excision enabled the expression of the HIS3 gene necessary for growth in the absence of histidine. We cultured cells carrying this construct and expressing LiCre and spotted them at various densities on two HIS^- selective plates. One plate was illuminated during 90 min while the other one was kept in the dark, and both plates were then incubated for growth. After 3 days, colonies were abundant on the plate that had been illuminated and very rare on the control plate (Figure 4a). LiCre can therefore be used to trigger cell growth with light.

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We then sought to observe the switch in individual cells. To do so, we replaced GFP by mCherry in our reporter system so that the excitation wavelength of the reporter did not overlap with stimulation of LiCre. We expressed and stimulated LiCre (90 min at 3.63 mW/cm²) in cells carrying this reporter and subsequently imaged them over time. As expected, we observed the progressive apparition of mCherry signal in a fraction of cells (Figure 4b, c).

Although convenient for high-throughput quantifications, reporter systems based on the de novo production and maturation of fluorescent proteins require a delay between the time of DNA excision and the time of acquisition. We wished to bypass this limitation and directly quantify DNA recombination. For this, we designed oligonucleotides outside of the region flanked by LoxP sites. The hybridization sites of these primers are too distant for efficient amplification of the non-edited DNA template, but, after Cre-mediated excision of the internal region, these sites become proximal and PCR amplification is efficient (Figure 4d). We mixed known amounts of edited and non-edited genomic DNA and performed real-time qPCR to build a standard curve that could be used to infer the proportion of edited DNA from qPCR signals. After this calibration, we applied this qPCR assay on genomic DNA extracted from cells collected immediately after different durations of illumination at moderate intensity (3.63 mW/cm², PAUL device). Results were in full agreement with GFP-based quantifications (Figure 4e). Excision of the target DNA occurred in a significant fraction of cells after only 2 min of illumination, and we estimated that excision occurred in about 30% and 40% of cells after 20 and 40 min of illumination, respectively. To determine if DNA excision continued to occur after switching off the light, we re-incubated half of the cells for 90 min in the dark prior to harvest and genomic DNA extraction. The estimated frequency of DNA excision was strikingly similar to the one measured immediately after illumination (Figure 4f). This was fully consistent with the fast cycling properties of LiCre described above, with a rapid (tens of seconds) reversal of activated LiCre to its inactive state.

The qPCR assay also allowed us to compare the efficiency of light-induced recombination between cell populations in exponential growth or stationary phase. This revealed that LiCre photoactivation was about fourfold more efficient in non-dividing cells (Figure 4g). Although the reasons for this difference remain to be determined, this increase of LiCre photoactivation at stationary phase makes it particularly suitable for bioproduction applications, where metabolic switching is often desired after the growth phase.

LiCre offers a way to change the activities of cells without adding any chemical to their environment. This potentially makes it an interesting tool to address the limitations caused by metabolic burden, the natural trade-off between the fitness of host cells, and their efficiency at producing exogenous compounds (Wu et al., 2016). We therefore tested the possibility to use LiCre to control the production of a commercial compound with light.

Carotenoids are pigments that can be used as vitamin A precursors, antioxidants, or coloring agents, making them valuable for the food, agriculture, and cosmetics industries (Mata-Gómez et al., 2014). Commercial carotenoids are generally produced by chemical synthesis or extraction from vegetables, but alternative productions based on microbial fermentations offer remarkable advantages, including the use of low-cost substrates and therefore a high potential for financial gains. Bioproduction of carotenoids can be achieved by introducing recombinant biosynthesis pathways in host microorganisms, which offers the advantage of a well-known physiology of the host and optimizations by genetic engineering. For these reasons, strategies were previously developed to produce carotenoids in the yeast Saccharomyces cerevisiae. Expressing three enzymes (crtE, crtI, and crtYB) from Xanthophyllomyces dendrorhous enabled S. cerevisiae to efficiently convert farnesyl pyrophosphate (FPP) into β-carotene (Verwaal et al., 2007). FPP is naturally produced by S. cerevisiae from acetyl-CoA and serves as an intermediate metabolite, particularly for the production of ergosterol that is essential for cellular viability (Figure 5). This production is associated with a trade-off: redirecting FPP to β-carotene limits its availability for ergosterol biosynthesis and therefore impairs growth; and its consumption by the host cell can limit the flux toward the recombinant pathway. Consistently, metabolic burden associated with carotenoids production was shown to be substantial (Verwaal et al., 2010). A promising way to deal with this trade-off would be to favor the flux toward ergosterol during biomass expansion and, after enough producer cells are obtained, switch the demand in FPP toward β-carotene. We therefore explored if LiCre could offer this possibility.

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First, we tested if LiCre could allow us to switch ON the exogenous production of carotenoids with light. If so, one could use it to trigger production at the desired time of a bioprocess. We constructed a S. cerevisiae strain expressing only two of the three enzymes required for β-carotene production. Expression of the third enzyme, a bifunctional phytoene synthase and lycopene cyclase, was blocked by the presence of a floxed terminator upstream of the coding sequence of the crtYB gene (Figure 5b). Excision of this terminator should restore a fully functional biosynthetic pathway. As expected, this strain formed white colonies on agar plates, but it formed orange colonies after transformation with an expression plasmid coding for Cre, indicating that β-carotene production was triggered (Figure 5c). To test the possible triggering by light, we transformed this strain with a plasmid encoding LiCre and selected several transformants, which we cultured and exposed – or not – to blue light before spotting them on agar plates. The illuminated cultures became orange while the non-illuminated ones remained white. Plating a dilution of the illuminated cell suspension yielded a majority of orange colonies, indicating that LiCre triggered crtYB expression and β-carotene production in a high proportion of plated cells (Figure 5c). We quantified bioproduction by dosing total carotenoids in cultures that had been illuminated or not. This revealed that 72 hr after the light switch the intracellular concentration of carotenoids had jumped from background levels to nearly 200 μg/g (Figure 5d). Thus, LiCre allowed us to switch ON the production of carotenoids by yeast using blue light.

We then tested if LiCre could allow us to switch OFF with light the endogenous ergosterol pathway that competes with carotenoid production for FPP consumption. The first step of this pathway is catalyzed by the Erg9p squalene synthase. Given the importance of FPP availability for the production of various compounds, strategies have been reported to control the activity of this enzyme during bioprocesses, especially in order to reduce it after biomass expansion (Asadollahi et al., 2008; Xie et al., 2015; Tippmann et al., 2016). These strategies were not based on light but derived from transcriptional switches that naturally occur upon addition of inhibitors or when specific nutrients are exhausted from the culture medium. To test if LiCre could offer a way to switch ERG9 activity with light, we modified the ERG9 chromosomal locus and replaced the coding sequence by a synthetic construct comprising a floxed sequence coding for Erg9p and containing a transcriptional terminator, followed by a sequence coding for the catalytic domain of the 3-hydroxy3-methylglutaryl coenzyme A reductase (tHMG1) (Figure 5e). This design prepares ERG9 for a Cre-mediated switch: before recombination, Erg9p is normally expressed; after recombination, ERG9 is deleted and the tHMG1 sequence is expressed to foster the mevalonate pathway. Given that ERG9 is essential for yeast viability in the absence of ergosterol supplementation (Fegueur et al., 1991), occurrence of the switch can be evaluated by measuring the fraction of viable yeast cells prior to and after the induction of recombination. When doing so, we observed that expression of Cre completely abolished viability, regardless of illumination. In contrast, cultures expressing LiCre were highly susceptible to light: they were fully viable in the absence of illumination and lost ~23% of viable cells after light exposure (Figure 5f). Thus, LiCre offers the possibility to abolish the activity of the yeast squalene synthase by exposing cells to light.

Beyond yeast, LiCre may also have a large spectrum of applications on multicellular organisms. Therefore, we tested its efficiency in human cells. For this, we constructed a lentiviral vector derived from the simian immunodeficiency virus (SIV) and encoded a human-optimized version of LiCre with a nuclear localization signal fused to its N-terminus. To quantify the efficiency of this vector, we also constructed a stable reporter cell line where the expression of a membrane-located mCherry fluorescent protein could be switched ON by Cre/Lox recombination. We obtained this line by Flp-mediated insertion of a single copy of the reporter construct into the genome of Flp-In 293 cells (Figure 6a, see Materials and methods). Our assay consisted of producing LiCre-encoding lentiviral particle, depositing them on reporter cells for 24 hr, illuminating the infected cultures with blue light, and, 28 hr later, observing cells by fluorescence microscopy. As shown in Figure 6b, mCherry expression was not detected in non-infected reporter cells. In cultures that were infected but not illuminated, a few positive cells were observed. In contrast, infected cultures that had been exposed to blue light contained numerous positive cells. We sought to quantify this photoactivation and compare it to the one conferred by nMag/pMag split Cre. For this, we produced two additional sets of lentiviral particles: one encoding the originally described nMag/pMag split Cre (Kawano et al., 2016), and one encoding the 'Magnets-opti' coding sequence of ‘PA-Cre3.0’, where codons had been optimized to avoid intramolecular sequence homology (Morikawa et al., 2020). For consistency, all three constructs shared the same vector backbone and CMV promoter. We tested these systems in parallel as above using light energy that was either suited to LiCre (3.6 mW/cm²) or nMag/pMag split Cre (1.8 mW/cm²) and quantified recombination efficiencies by flow cytometry. As shown in Figure 6c, the residual activity of Magnets-opti in the dark was very high (~50% of switched cells), which was surprising. LiCre and the original nMag/pMag split Cre system displayed similar levels of induced activity at both low and high light energies; they also showed significant residual activity with 5–10% positive cells for LiCre in the dark and 10–15% for the original nMag/pMag. The highest fold induction was obtained with LiCre at 3.6 mW/cm², with 5% residual and 31% induced activities, respectively. LiCre can therefore be used to switch genetic activities in human cells with blue light.

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We built a structural model of LiCre to conceptualize its mode of activation (Figure 7a). We based this model on (i) the available structure of the Cre tetramer complexed with its target DNA (Guo et al., 1997), (ii) the available structure of AsLOV2 in its dark state (Guntas et al., 2015), and (iii) knowledge that the Jα helix of AsLOV2 domains unfolds after light activation (Swartz et al., 2002; Harper et al., 2003). From this model, we hypothesize that LiCre photoactivation may occur via two synergistic effects. First, the domain AsLOV2 likely prevents Cre tetramerization in the dark state simply because of its steric occupancy. The unfolding of the Jα helix in the light state may allow AsLOV2 to liberate the multimerizing interface. Second, because the Jα helix of AsLOV2 and the αA helix of Cre are immediately adjacent, they may disturb each other in their proper folding so that they cannot exist both simultaneously in their native conformation. The unfolding of Jα in the light state may therefore stimulate proper folding of αA, and thereby allow αA to bind to the adjacent Cre unit. This predicts two possible models of activation of recombinase activity, as depicted in Figure 7b. If LiCre is not bound to DNA in its dark state (Model 1), then photoactivation allows it to form a protein dimer bound to each LoxP sites. The two sites can then associate to form the synaptic complex. Alternatively, LiCre may already bind to DNA in the dark state (Model 2). In this case, photoactivation likely reinforces protein–protein interaction of the two bound units and it also allows the two dimers to assemble into a functional recombination synapse. Experiments interrogating LiCre:DNA interactions in dark and light conditions are now needed to test these models.

Figure 7

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Several tools already exist for inducing site-specific recombination with light. They fall in two groups: those that require the addition of a chemical and those that are fully genetically encoded. The first group includes the utilization of photocaged ligands instead of 4-hydroxy-tamoxifen to induce the activity of Cre-ER^T. This pioneering approach was successful in cultured human cells (Link et al., 2005) as well as fish (Sinha et al., 2010) and mouse (Lu et al., 2012). Later, a more complex strategy was developed that directly rendered the active site of Cre photoactivatable via the incorporation of photocaged amino acids (Luo et al., 2016). In this case, cells were provided with non-natural amino acids, such as the photocaged tyrosine ONBY, and were genetically modified in order to express three macromolecules: a specifically evolved pyrrolysyl tRNA synthetase, a pyrrolysine tRNA_CUA, and a mutant version of Cre where a critical amino acid such as Y324 was replaced by a TAG stop codon. The tRNA synthetase/tRNA_CUA pair allowed the incorporation of the synthetic amino acid in place of the nonsense mutation, and the resulting enzyme was inactive unless it was irradiated with violet or ultraviolet light. This strategy successfully controlled recombination in cultured human cells (Luo et al., 2016) and zebrafish embryos (Brown et al., 2018). We note that it presents several caveats: its combination of chemistry and transgenes is complex to implement, the presence of the tRNA synthetase/tRNA_CUA pair can generate off-target artificial C-terminal tails in other proteins by bypassing natural stop codons, and violet/ultraviolet light can be harmful to cells. More recently, a radically different chemical approach was proposed, which consisted of tethering an active TAT-Cre recombinase to hollow gold nanoshells (Morales et al., 2018). When delivered to cells in culture, these particles remained trapped in intracellular endosomes. Near-infrared photostimulation triggered activity by releasing the recombinase via nanobubble generation occurring on the particle surface. A fourth system is based on the chromophore phycocyanobilin, which binds to the PhyB receptor of A. thaliana and makes its interaction with PIF3 dependent on red light. Photostimulation of this interaction was used to assemble split Cre units into a functional complex in yeast (Hochrein et al., 2018). A major interest of these last two systems is to offer the possibility to use red light, which is less harmful to cells than blue or violet light and better penetrates tissues. However, all these strategies require to efficiently deliver chemicals to the target cells at the appropriate time before illumination, and their underlying chemistry can be expensive, especially for applications in the context of large volumes such as industrial bioprocesses.

Other systems, such as LiCre, are based on flavin chromophores that are readily available across a wide variety of organisms. This avoids the need of chemical additives. To our knowledge, there are currently three such systems. One is based on the sequestration of Cre between two large photo-cleavable domains (Zhang et al., 2017). The principle of light-induced protein cleavage is very interesting, but its application to Cre showed important limitations: a moderate efficiency (~30% of ON cells after the switch), the dependence on a cellular inhibitory chaperone, and the need of violet light. The two other systems are the CRY2/CIB1 and nMag/pMag split Cre (Taslimi et al., 2016; Kawano et al., 2016) that we evaluated here in comparison to LiCre. An important advantage of LiCre over these systems is that it is made of a single protein. The first benefit of this is simplicity. More efforts are needed to establish transgenic organisms expressing two open reading frames (ORFs) compared to a single one. This is particularly true for vertebrate systems, where inserting several constructs requires additional efforts for characterizing transgene insertion sites and conducting genetic crosses. For this reason, the two ORFs of the split Cre system were previously combined in a single construct, where they were separated either by an internal ribosomal entry site (IRES) or by a sequence coding a self-cleaving peptide (Taslimi et al., 2016; Kawano et al., 2016; Morikawa et al., 2020). Although helpful, these solutions have important limits: with an IRES, the two ORFs are not expressed at the same level; with a self-cleaving peptide, cleavage of the precursor protein can be incomplete, generating uncleaved products with unknown activity. This was the case for nMag/pMag split Cre in mammalian cells, where a non-cleaved form at ~72 kDa was reported and where targeted modifications of the cleavage sequence increased both the abundance of this non-cleaved form and the non-induced activity of the system (Morikawa et al., 2020). The second benefit of LiCre being a single protein is to avoid problems of suboptimal stoichiometry between the two protein units, which was reported as a possible issue for CRY2/CIB1 split Cre (Meador et al., 2019). A third benefit is to avoid possible intramolecular recombination between the homologous parts of the two coding sequences. Although not demonstrated, this undesired possibility was suspected for nMag/pMag split Cre because its two dimerizers derive from the same sequence (Morikawa et al., 2020). The other advantages of LiCre are its performances. In the present study, we used a yeast-based assay to compare LiCre with split Cre systems. Unexpectedly, although we used the improved version of the CRY2/CIB1 split Cre containing the CRY2-L348F mutation (Taslimi et al., 2016), it did not generate photo-inducible recombination in our assay. This is unlikely due to specificities of the budding yeast, such as improper protein expression or maturation, because the original authors reported activity in this organism (Taslimi et al., 2016). We do not explain this result, but it is consistent with previous studies reporting that photoactivation of the original version of the CRY2/CIB1 split Cre can be extremely low (Kawano et al., 2016), and that the induced activity of the CRY2-L348F/CIB1 system can also be low and highly variable (Morikawa et al., 2020). In contrast, we validated the efficiency of nMag/pMag split Cre and so did other independent laboratories (Morikawa et al., 2020; Takao et al., 2020; Allen et al., 2019; Weinberg et al., 2019). We observed a significant photoactivation of this system both in yeast and human cells. Recently, the nMag/pMag split Cre system was expressed in mice as a transgene – dubbed PA-Cre3.0 – which comprised the promoter sequence of the chicken beta actin gene (CAG) and the 'Magnets-opti' modified version of the original self-cleaving coding sequence (Morikawa et al., 2020). The authors reported that this strategy abolished residual activity and attributed this improvement to a reduction of the expression level of the transgene (Morikawa et al., 2020). In our human cell assay, where we used the CMV promoter, the 'Magnets-opti' system instead showed a much higher residual activity than the original nMag/pMag split Cre. This was surprising; it is probably very important to carefully calibrate the expression of Magnets-opti when using it. In contrast, LiCre displayed weaker residual activity than the original nMag/pMag split Cre in the dark. Importantly, LiCre also displayed higher induced activity – especially in yeast – and a faster response to light compared to nMag/pMag split Cre. This strong response probably results from its simplicity since the activation of a single protein involves fewer steps than the activation of two units that must then dimerize to become functional. Finally, we showed that the photocycle of active LiCre is faster than the one of active nMag/pMag split Cre. This offers higher temporal precision for the induction of recombination.

In conclusion, LiCre provides a cheap, simple, low-background, highly efficient and fast-responding way to induce site-specific recombination with light. Given that it works in both yeast and mammalian cells, it opens many perspectives from fundamental and biomedical research to industrial applications.

Plasmids, strains, and oligonucleotides used in this study are listed in Supplementary file 1 (Supplementary Tables S1, S2, and S3, respectively). LiCre plasmids are available from Addgene (http://www.addgene.org/) under accession numbers 166660, 166661 and 166663.

We ordered the synthesis of sequence LoxLEULoxHIS (Supplementary file 1 – Supplementary Text S1) from GeneCust, who cloned the corresponding BamHI fragment in plasmid pHO-poly-HO to produce plasmid pGY262. The P_TEF-loxP-KlLEU2-STOP-loxP-spHIS5 construct can be excised from pGY262 by NotI digestion for integration at the yeast HO locus. This way, we integrated it in a leu2 his3 strain, which could then switch from LEU+ his- to leu- HIS+ after Cre-mediated recombination (Figure 4e). To construct a GFP-based reporter, we ordered the synthesis of sequence LEULoxGreen (Supplementary file 1 – Supplementary Text S1) from GeneCust, who cloned the corresponding NheI-SacI fragment into pGY262 to obtain pGY407. We generated strain GY984 by crossing BY4726 with FYC2-6B. We transformed GY984 with the 4 kb NotI insert of pGY407 and obtained strain GY1752. To remove the ade2 marker, we crossed GY1752 with FYC2-6A and obtained strain GY1761. Plasmid pGY537 targeting integration at the LYS2 locus was obtained by cloning the BamHI-EcoRI fragment of pGY407 into the BamHI, EcoRI sites of pIS385. Plasmid pGY472 was produced by GeneCust, who synthesized sequence LEULoxmCherry (Supplementary file 1 – Supplementary Text S1) and cloned the corresponding AgeI-EcoRI insert into the AgeI, EcoRI sites of pGY407. We generated GY983 by crossing BY4725 with FYC2-6A. We obtained GY2033 by transformation of FYC2-6B with a 4 kb NotI fragment of pGY472. We obtained GY2207 by transformation of GY983 with the same 4 kb NotI fragment of pGY472. To generate GY2206, we linearized pGY537 with NruI digestion, transformed in strain GY855, and selected a LEU+ Lys- colony (pop-in), which we re-streaked on 5-FoA plates for vector excision by counter-selection of URA3 (pop-out) (Sadowski et al., 2007). Strain GY2214 was a diploid that we obtained by mating GY2206 with GY2207.

Mutations E340A D341A were introduced by GeneCust by site-directed mutagenesis of pSH63, yielding plasmid pGY372. We generated the N-terΔ21 mutant of Cre by PCR amplification of the P_GAL1 promoter of pSH63 using primer 1L80 (forward) and mutagenic primer 1L71 (reverse), digestion of pSH63 by AgeI and co-transformation of this truncated plasmid and amplicon in a trp1Δ63 yeast strain for homologous recombination and plasmid rescue. We combined the N-terΔ21 and the C-ter E340A D341A mutations similarly, but with pGY372 instead of pSH63. We generated N-terΔ28 and N-terΔ37 mutants, combined or not with C-ter E340A D341A mutations, by the same procedure where we changed 1L71 by mutagenic primers 1L72 and 1L73, respectively.

To generate a Cre-VVD fusion, we designed sequence CreCVII (Supplementary file 1 – Supplementary Text S1) where the Cre sequence from GENBANK AAG34515.1 was fused to the VVD-M135IM165I sequence from Zoltowski et al., 2009 via four additional residues (GGSG). We ordered its synthesis from GeneCust and co-transformed it in yeast with pSH63 (previously digested by NdeI and SalI) for homologous recombination and plasmid rescue. This generated pGY286. We then noticed an unfortunate error in AAG34515.1, which reads a threonine instead of an asparagine at position 327. We cured this mutation from pGY286 by site-directed mutagenesis using primers 1J47 and 1J48, which generated pGY339 that codes for Cre-4-VVD described in Figure 2—figure supplement 1. We constructed mutant C-terΔ14 of Cre by site-directed mutagenesis of pGY286 using primers 1J49 and 1J50, which simultaneously cured the N327T mutation and introduced an early stop codon. Mutants C-terΔ2, C-terΔ4, C-terΔ6, C-terΔ8, C-terΔ10, C-terΔ12 of Cre were constructed by GeneCust, who introduced early stop codons in pGY339 by site-directed mutagenesis. We generated Cre-VVD fusions carrying point mutations at amino acid positions 340 and 341, with or without truncation of the GGSG linker (Figure 2—figure supplement 1), by site-directed mutagenesis of pGY339.

To test LOV_Cre fusions, we first designed sequence EcoRI-LovCre_chimJa-BstBI (Supplementary file 1 – Supplementary Text S1) corresponding to the fusion of AsLOV2 with Cre via an artificial α-helix. This helix was partly identical to the Jα helix of AsLOV2 and partly identical to the αA helix of Cre. This sequence was synthesized and cloned in the EcoRI and BstBI sites of pSH63 by GeneCust, yielding pGY408. We then generated and directly tested a variety of LOV_Cre fusions. To do so, we digested pGY408 with BsiWI and MfeI and used this fragment as a recipient vector; we amplified the Cre sequence from pSH63 using primer 1G42 as the reverse primer, and one of primers 1M42 to 1M53 as the forward primer (each primer corresponding to a different fusion position); we co-transformed the resulting amplicon and the recipient vector in strain GY1761, isolated independent transformants, and assayed them with the protocol of photoactivation and flow cytometry described below. We generated and tested a variety of LOV_CreAA fusions by following the same procedure where plasmid pGY372 was used as the PCR template instead of pSH63. A transformant corresponding to LOV_Cre32 and showing light-dependent activity was chosen for plasmid rescue, yielding plasmid pGY415. A transformant corresponding to LOV_CreAA20 was chosen for plasmid rescue, yielding plasmid pGY416. Sanger sequencing revealed that the fusion sequence present in pGY416 was QID instead of QIA at the peptide junction (position 149 on LiCre sequence of Supplementary file 1 – Supplementary Text S2). All further experiments on LiCre were derived from the fusion protein coded by pGY416.

To introduce random residues at the peptide junction of LOV_Cre32 (Figure 2b), we first generated pGY417 using the same procedure as for the generation of pGY415 but with pSH47 instead of pSH63 as the PCR template so that pGY417 has a URA3 marker instead of TRP1. We then ordered primers 1N24, 1N25, and 1N26 containing degenerate sequences, used them with primer 1F14 to amplify the Cre sequence of pSH63, co-transformed in strain GY1761 the resulting amplicons together with a recipient vector made by digesting plasmid pGY417 with NcoI and BsiWI, and isolated and directly tested individual transformants with the protocol of photoactivation and flow cytometry described below. Plasmids from transformants showing evidence of reduced background were rescued from yeast and sequenced, yielding pGY459 to pGY464.

To replace the P_GAL1 promoter of pGY416 by the P_MET17 promoter, we digested it with SacI and SpeI, PCR-amplified the P_MET17 promoter of plasmid pGY8 with primers 1N95 and 1N96, and co-transformed the two products in yeast for homologous recombination, yielding plasmid pGY466. We changed the promoter of pGY415 using exactly the same procedure, yielding plasmid pGY465. We changed the promoter of pSH63 similarly using primer 1O83 instead of 1N96, yielding plasmid pGY502.

To express the nMag/pMag split Cre system in yeast, we designed sequence CreN-nMag-NLS-T2A-NLS-pMag-CreC partly (Supplementary file 1 – Supplementary Text S1) and ordered its synthesis from GeneCust. The corresponding BglII fragment was co-transformed in yeast for homologous recombination with pGY465 previously digested with BamHI (to remove AsLOV2 and part of Cre), yielding plasmid pGY488 that contained the full system. We then derived two plasmids from pGY488, each one containing one half of the split system under the control of the Met17 promoter. We obtained the first plasmid (pGY491, carrying the TRP1 selection marker) by digestion of pGY488 with SfoI and SacII and co-transformation of the resulting recipient vector with a PCR product amplified from pGY465 using primers 1O80 and 1O82. We obtained the second plasmid (pGY501, carrying the URA3 selection marker) in two steps. We first removed the pMag-CreC part of pGY488 by digestion with NdeI and SacII followed by Klenow fill-in and religation. We then changed the selection marker by digestion with PfoI and KpnI and co-transformation in yeast with a PCR product amplified from pSH47 with primers 1O77 and 1O89.

To express the CRY2^L348F/CIB1 split Cre system in yeast, we designed sequences CIB1CreCter and CRY2CreNter and ordered their synthesis from GeneCust, obtaining plasmids pGY526 and pGY527, respectively. To obtain pGY531, we extracted the synthetic insert of pGY527 by digestion with BglII and co-transformed it in yeast with the NdeI-BamHI fragment of pGY466 for homologous recombination. To obtain pGY532, we extracted the synthetic insert of pGY526 by digestion with BglII and co-transformed it in yeast with the SacI-BamHI fragment of pSH47 for homologous recombination.

To build a switchable strain for carotene production, we modified EUROSCARF strain Y41388 by integrating a LoxP-KlLEU2-T_ADH1-LoxP cassette immediately upstream the CrtYB coding sequence of the chromosomally integrated expression cassette described by Verwaal et al., 2007. This insertion was obtained by transforming Y41388 with a 6.6 kb BstBI fragment from plasmid pGY559 and selecting a Leu+ transformant, yielding strain GY2247. To obtain pGY559, we first deleted the crtE and crtI genes from YEplac195-YB_E_I (Verwaal et al., 2007) by MluI digestion and religation. We then linearized the resulting plasmid with SpeI and co-transformed it for recombination in a leu2Δ yeast strain with a PCR amplicon obtained with primers 1P74 and 1P75 and template pGY407. After Leu+ selection, the plasmid was recovered from yeast, amplified in bacteria, and verified by restriction digestion and sequencing.

We used CRISPR/Cas9 to build a switchable strain for squalene synthase. We cloned the synthetic sequence gERG9 (Supplementary file 1 – Supplementary Text S1) in the BamHI-NheI sites of the pML104 plasmid (Laughery et al., 2015) so that the resulting plasmid (pGY553) coded for a gRNA sequence targeting ERG9. This plasmid was transformed in GY2226 together with a repair-template corresponding to a 4.2 kb EcoRI fragment of pGY547 that contained LoxP-synERG9-T_ADH1-LoxP with homologous flanking sequences. The resulting strain was then crossed with Y41388 to obtain GY2236.

We used synthetic (S) media made of 6.7 g/l Difco Yeast Nitrogen Base without amino acids and 2 g/l of a powder that was previously prepared by mixing the following amino acids and nucleotides: 1 g of adenine, 2 g of uracil, 2 g of alanine, 2 g of arginine, 2 g of aspartate, 2 g of asparagine, 2 g of cysteine, 2 g of glutamate, 2 g of glutamine, 2 g of glycine, 2 g of histidine, 2 g of isoleucine, 4 g of leucine, 2 g of lysine, 2 g of methionine, 2 g of phenylalanine, 2 g of proline, 2 g of serine, 2 g of threonine, 2 g of tryptophane, 2 g of tyrosine, and 2 g of valine. For growth in glucose condition, the medium (SD) also contained 20 g/l of D-glucose. For growth in galactose condition (induction of P_GAL1 promoter), we added 2% final (20 g/l) raffinose and 2% final (20 g/l) galactose (SGalRaff medium). Media were adjusted to pH = 5.8 by addition of NaOH 1 N before autoclaving at 0.5 bar. For auxotrophic selections or P_MET17 induction, we used media where one or more of the amino acids or nucleotides were omitted when preparing S. For example, SD-W-M was made as SD but without any tryptophane or methionine in the mix powder.

Unless mentioned otherwise, quantitative tests were done by flow cytometry using yeast reporter strain GY1761. We used two devices for photoactivation. The first device was a PAUL apparatus from GenIUL: a box with mirrors on its walls and floor, and equipped with 460 nm blue LEDs on its ceiling. Using a NovaII photometer (Ophir Photonics), we measured that a 100% intensity on this apparatus corresponded to an energy of 36.3 mW/cm². We used Zomei ND filters when we needed to obtain intensities that were not tunable on this device. The second device was a computer-controlled DMX LED spot (450 nm) purchased from Neptune-LED (Grenoble, France), which we placed above a thermostated platform. The yeast reporter strain was transformed with the plasmid of interest, pre-cultured overnight in selective medium corresponding to conditions of transcriptional activation of the plasmid-borne Cre construct (SGalRaff-W for P_GAL1 plasmids, SD-W-M for P_MET17 plasmids, SD-W-U-M for split Cre systems) with no particular protection against ambient light. The saturated culture was transferred to two 96-well polystyrene flat-bottom Falcon sterile plates (100 μl per well) and one plate was illuminated at the indicated intensities while the other plate was kept in the dark. After the indicated duration of illumination, cells from the two plates were transferred to a fresh medium allowing expression of GFP but not cell division (SD-W-H or SD-W-U-H, strain GY1761 being auxotroph for histidine), and these cultures were incubated at 30°C for 90 min. Cells were then either analyzed immediately by flow cytometry or blocked in phosphate-buffered saline (PBS + 1 mM sodium azide) and analyzed the following day.

We acquired data for 10,000 events per sample using a FACSCalibur (BD Biosciences) or a MACSQuant VYB (Miltenyi Biotech) cytometer after adjusting the concentration of cells in PBS. We analyzed raw data files in the R statistical environment (http://www.r-project.org) using custom-made scripts based on the flowCore package (Hahne et al., 2009) from bioconductor (http://www.bioconductor.org). We gated cells automatically by computing a perimeter of (FSC-H, SSC-H) values that contained 40% of events (using 2D-kernel density distributions). We plotted the gates of all samples belonging to a single experiment (usually corresponding to the same day). This occasionally highlighted samples for which the gates were clearly shifted (outliers), indicating cell populations of unexpected size, and we therefore discarded these rare samples for further analysis. A threshold of fluorescent intensity (GFP or mCherry) was set to distinguish ON and OFF cells (i.e., expressing or not the reporter). To do this, we included in every experiment a negative control made of the reporter strain transformed with an empty vector and chose the 99.9th percentile of the corresponding 4000 fluorescent values (gated cells) as the threshold. No statistical power analysis was done prior to the experiment, no masking of samples IDs was applied during the experiments, and no data randomization was used.

For Figure 3g, we segmented individual cells on bright-field images using the ImageJ Lasso plugin. Then, we measured on the fluorescence images the mean gray value of pixels in each segmented area, providing single-cell measures of fluorescence. For each image, the background fluorescence level was quantified from eight random regions outside of cells and with areas similar to single cells. This background level was subtracted from the fluorescence level of each cell.

We strictly followed the procedure described in Verwaal et al., 2007, which consists of mechanical cell lysis using glass beads, addition of pyrogallol, KOH-based saponification, and extraction of carotenoids in hexane. Quantification was estimated by optical absorption at 449 nm using a Biowave spectrophotometer. No statistical power analysis was done prior to the experiment, no masking of samples IDs was applied during the experiments, and no data randomization was used.

We built a reporter construct for Cre-mediated recombination in human cells based on Addgene's plasmids 55779, containing a membrane-addressed mCherry sequence (Yost et al., 2007) (mCherry-Mem), and 51269, containing a zsGreen-based reporter of Cre recombination (Hermann et al., 2014). Re-sequencing revealed that 51269 did not contain three terminator sequences but only one between the LoxP sites. We applied a multi-steps procedure to (i) restore three terminators, (ii) replace zsGreen with mCherry-Mem, and (iii) have the final reporter in a vector suitable for targeted single-site insertion. First, we inserted a LoxP site between restriction sites NheI and HindIII of pCDNA5/FRT (Invitrogen) by annealing oligonucleotides 1O98 and 1O99, digesting and cloning this adaptor with NheI and HindIII, which yielded plasmid pGY519. Second, we replaced in two different ways the zsGreen sequence of 51269 by the mCherry-Mem sequence of 55779: either by cloning a SmaI-NotI insert from 55779 into EcoRV-NotI of 51269, yielding plasmid pGY520, or cloning a EcoRI-NotI from 55779 into EcoRI-NotI of 51269, yielding plasmid pGY521. Third, we inserted the HindIII-NotI cassette of pGY520 into the HindIII-NotI sites of pGY519, yielding plasmid pGY523. Fourth, we inserted the HindIII-NotI cassette of pGY521 into the HindIII-NotI sites of pGY519, yielding plasmid pGY524. Fifth, a HindIII-BamHI fragment of pGY523 containing one terminator and a BglII-EcoRI fragment of pGY524 containing another terminator were simultaneously cloned as consecutive inserts in the BglII-EcoRI sites of 51269. Finally, the resulting plasmid was digested with HindIII and BamHI to produce a fragment that was cloned into the HindIII-BglII sites of pGY524 to produce pGY525.

To establish stable cell lines, Flp-In T-REx 293 cells were purchased from Invitrogen (ThermoFisher) and transfected with both the Flp recombinase vector (pOG44, Invitrogen) and pGY525. Selection of clonal cells was first performed in medium containing 300 μg hygromycin (Sigma). After 2 weeks, we identified foci of cell clusters, which we individualized by transferring them to fresh wells. One of these clones was cultured for three additional weeks with high concentrations of hygromycin (up to 400 μg) to remove potentially contaminating negative cells. The resulting cell line was named T4-2PURE.

A synthetic sequence was ordered from GeneCust and cloned in the HindIII-NotI sites of pCDNA3.1 (Invitrogen V79020). This insert contained an unrelated additional sequence that we removed by digestion with BamHI and XbaI, followed by blunt-ending with Klenow fill-in. The resulting plasmid (pGY561) encoded LiCre optimized for mammalian codon usage, in-frame with a N-ter located SV40-NLS signal. This NLS-LiCre sequence was amplified from pGY561 using primers Sauci and Flard (Table S3), and the resulting amplicon was cloned in the AgeI-HindIII sites of the GAE0 Self-Inactivating Vector (Mangeot et al., 2004), yielding pGY577. To obtain a similar vector expressing nMag/pMag split Cre, we ordered its synthetic sequence from GeneCust, who cloned it into the AgeI-HindIII sites of pGY577, yielding pGY625. This plasmid was further modified by replacing the AgeI-EcoRV portion with a synthetic fragment corresponding to the ‘Magnets-opti’ coding sequence of ‘PA-Cre3.0" (Morikawa et al., 2020), yielding pGY626. Sequences of pGY577, pGY625, and pGY626 are provided in Supplementary file 1 – Supplementary Text S1. Lentiviral particles were produced in Gesicle Producer 293 T cells (TAKARA ref 632617) transiently transfected by an HIV-1 helper plasmid (45% of total DNA), a plasmid encoding the VSV-g envelope (15% of total DNA), and either pGY577, pGY625, or pGY626 (40% of total DNA) as previously described (Mangeot et al., 2004). Particle-containing supernatants were clarified, filtered through a 0.45 μm membrane, and concentrated by ultracentrifugation at 40,000 g before resuspension in 1× PBS (100-fold concentration). To quantify the particle load of these preparations, we designed a qPCR assay (primers 1Q74,1Q75) and calibrated it on a standard curve obtained by serial dilutions of plasmid DNA. These measures (as mean ± s.e.m. million particles per μl, n = 3 replicate measures) on the preparations used in Figure 6c were the following: 16 ± 0.7 for LiCre, 8.6 ± 0.4 for nMag/pMag split Cre, and 5.2 ± 0.04 for Magnets-opti.

About 3 × 10⁵ cells of cell line T4-2PURE were plated in two 6-well plates. After 24 hr, 100 μl of viral particles suspension were added to each well. After another 24 hr, one plate was illuminated with blue light (460 nm) using the PAUL apparatus installed in a 37°C incubator while the other plate was kept in the dark. For illumination, we applied a sequence of 20 min ON, 20 min OFF under CO₂ atmosphere, 20 min ON, where ON corresponded to 3.63 mW/cm² illumination (or 1.8 mW/cm² when indicated in Figure 6c). Plates were then returned to the incubator and, after 28 hr, were either imaged on an Axiovert135 inverted fluorescent microscope or treated with trypsin, resuspended in 1× PBS, and fixed with paraformaldehyde (PFA) prior to acquisitions on a MACSQuant flow cytometer. Flow-cytometry data was then processed as described above for yeast cells. Four samples were discarded from further analysis because the culture medium had become yellow and because flow cytometry revealed aberrant cell sizes (lower forward scatter [FSC] and very high side scatter [SSC]). Samples IDs were not masked during the experiments. We did not retype the T4-2PURE reporter cell line or the Gesicle Producer cell line, nor did we control them for mycoplasma contamination prior to the experiments because we used them in all the reporter assays that we applied to various constructs and it is extremely unlikely that a contamination would bias results for some constructs and not others.

We calculated the free-energy profile (reported in Figure 1f) for the unbinding of the C-terminal α-helix in the tetrameric Cre-recombinase complex (Ennifar et al., 2003) (PDB entry 1NZB) as follows. The software we used were the CHARMM-GUI server (Lee et al., 2016) to generate initial input files; CHARMM version c39b1 (Brooks et al., 2009) to setup the structural models and subsequent umbrella sampling by molecular dynamics; WHAM version 2.0.9 (http://membrane.urmc.rochester.edu/content/wham/) to extract the PMF; and VMD version 1.9.2 (Humphrey et al., 1996) to visualize structures. To achieve sufficient sampling by molecular dynamics, we worked with a structurally reduced model system. We focused thereby only on the unbinding of the C-terminal α-helix of subunit A (residues 334:340) from subunit F. Residues that did not have at least one atom within 25 Å from residues 333 to 343 of subunit A were removed including the DNA fragments. Residues with at least one atom within 10 Å were allowed to move freely in the following simulations; the remaining residues were fixed to their positions in the crystal structure. For the calculation of the double mutant A340A341, the corresponding residues were replaced by alanine residues. The systems were simulated with the CHARMM22 force field (GBSW and CMAP parameter file) and the implicit solvation model FACTS (Haberthür and Caflisch, 2008) with recommended settings for param22 (i.e., cutoff of 12 Å for non-bonded interactions). Langevin dynamics were carried out with an integration time step of 2 fs and a friction coefficient of 4 ps⁻¹ for non-hydrogen atoms. The temperature of the heat bath was set to 310 K. The hydrogen bonds were constrained to their parameter values with SHAKE (Ryckaert et al., 1977).

The PMF was calculated for the distance between the center of mass of the α-helix (residues 334:340 of subunit A) and the center of mass of its environment (all residues that have at least one atom within 5 Å of this helix). Umbrella sampling (Torrie and Valleau, 1977) was performed with 13 independent molecular dynamics simulations where the system was restrained to different values of the reaction coordinate (equally spaced from 4 to 10 Å) using a harmonic biasing potential with a spring constant of 20 kcal mol⁻¹ Å⁻¹ (GEO/MMFP module of CHARMM). Note that this module uses a pre-factor of ½ for the harmonic potential (as in the case of the program WHAM).

For each simulation, the value of the reaction coordinate was saved at every time step for 30 ns. After an equilibration phase of 5 ns, we calculated for blocks of 5 ns the PMF and the probability distribution function along the reaction coordinate using the weighted histogram analysis method (Kumar et al., 1992). A total of 13 bins were used with lower and upper boundaries at 3.75 and 10.25 Å, respectively, and a convergence tolerance of 0.01 kcal mol⁻¹. Finally, we determined for each bin its relative free energy ${\displaystyle F_i=-kT\mathrm{l}\mathrm{n}\left({\stackrel{-}{p}}_i\right)}$, where k is the Boltzmann constant, T the temperature (310 K), and ${\stackrel{-}{p}}_i$ the mean value of the probability of bin i when averaged over the five blocks. The error in the $F_i$ estimate was calculated with ${\sigma }_{F_i}=kT{\sigma }_{{\stackrel{-}{p}}_i}/{\stackrel{-}{p}}_i$, where ${\sigma }_{{\stackrel{-}{p}}_i}$ is twice the standard error of the mean of the probability. An offset was applied to the final PMF so that its lowest value was located at zero.

We grew 10 colonies of strain GY1761 carrying plasmid pGY466 overnight at 30°C in SD-L-W-M liquid cultures. The following day, we used these starter cultures to inoculate 12 ml of SD-W-M medium at OD₆₀₀ = 0.2. When monitoring growth by optical density measurements, we observed that it was fully exponential after 4 hr and until at least 8.5 hr. At 6.5 hr of growth, for each culture, we dispatched 0.1 ml in 96-well plate duplicates using one column (8 wells) per colony, stored aliquots by centrifuging 1 ml of the cell suspension at 3300 g and freezing the cell pellet at −20°C (‘exponential’ negative control), and re-incubated the remaining of the culture at 30°C for later analysis at stationary phase. We exposed one plate (Figure 4j ‘exponential’ cyan samples) to blue light (PAUL apparatus, 460 nm, 3.63 mW/cm² intensity) for 40 min while the replicate plate was kept in the dark (Figure 4j ‘exponential’ gray samples). We pooled cells of the same column and stored them by centrifugation and freezing as above. The following day, we collected 1 ml of each saturated, frozen, and stored cells as above (‘stationary’ negative control). We dispatched the remaining of the cultures in a series of 96-well plates (0.1 ml/well, two columns per colony) and exposed these plates to blue light (PAUL apparatus, 460 nm, 3.63 mW/cm² intensity) for the indicated time (0, 2, 5, 10, 20, or 40 min). For each plate, following illumination, we collected and froze cells from six columns (Figure 4h samples) and re-incubated the plate in the dark for 90 min before collecting and freezing the remaining six columns (Figure 4i, x-axis samples). For genomic DNA extraction, we pooled cells from six wells of the same colony (one column), centrifuged and resuspended them in 280 μl in 50 mM EDTA, and added 20 μl of a 2 mg/ml Zymolyase stock solution (SEIKAGAKU, 20 U/mg) to the cell suspension and incubated it for 1 hr at 37°C for cell wall digestion. We then processed the digested cells with the Wizard Genomic DNA Purification Kit from Promega. We quantified DNA on a Nanodrop spectrophotometer and used ~100,000 copies of genomic DNA as template for qPCR, with primers 1P57 and 1P58 to amplify the edited target and primers 1B12 and 1C22 to amplify a control HMLalpha region that we used for normalization. We ran these reactions on a Rotorgene thermocycler (Qiagen). This allowed us to quantify the rate of excision of the floxed region as N_Lox/N_Total, where N_Loxis the number of edited molecules and N_Total the total number of DNA template molecules. To estimate N_Lox, we prepared mixtures of edited and non-edited genomic DNAs at known ratios of 0%, 0.5%, 1%, 5%, 10%, 50%, 70%, 90%, and 100% and applied (1P57, 1P58) qPCR using these mixtures as templates. This provided us with a standard curve that we then used to convert Ct values of the samples of interest into N_Lox values. To estimate N_Total, we qPCR-amplified the HMLalpha control region from templates made of increasing concentrations of genomic DNA. We then used the corresponding standard curve to convert the Ct value of HMLalpha amplification obtained from the samples of interest into N_Total values. No statistical power analysis was done prior to the experiment, no masking of samples IDs was applied during the experiments, and no data randomization was used.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Fabien Duveau for critical reading of the manuscript and for signal quantifications from microscopy images; Grégory Batt for fruitful discussions; Maria Teresa Texeira for strains; Sandrine Mouradian, Véronique Barateau, and SFR Biosciences Gerland-Lyon Sud (UMS344/US8) for access to flow cytometers and technical assistance; Christian Marchet for advice on LED controllers; and developers of R, Bioconductor, VMD, QLC+, and Ubuntu for their software. This work was supported by the European Research Council under the European Union’s Seventh Framework Programme FP7/2007-2013 Grant Agreement no. 281359 and CNRS under the ‘MITI 80 Prime’ program grant READGEN.

© 2021, Duplus-Bottin et al.

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