Light-dependent modulation of protein localization and function in living bacteria cells

doi:10.21203/rs.3.rs-1614433/v1

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Light-dependent modulation of protein localization and function in living bacteria cells

https://doi.org/10.21203/rs.3.rs-1614433/v1

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Most bacteria lack membrane-enclosed organelles to compartmentalize cellular processes. In lieu of physical compartments, bacterial proteins are often recruited to macromolecular scaffolds at specific subcellular locations to carry out their functions. Consequently, the ability to modulate a protein’s subcellular location with high precision and speed bears the potential to manipulate its corresponding cellular functions. Here we demonstrate that the CRY2/CIB1 system from Arabidopsis thaliana can be used to rapidly direct proteins to different subcellular locations inside live E. coli cells including the nucleoid, the cell pole, membrane, and the midcell division plane. We further show that such light-induced re-localization can be used to rapidly inhibit cytokinesis in actively dividing E. coli cells and that the CRY2/CIBN binding kinetics can be modulated by green light, adding a new dimension of control to the system. Finally, we tested this optogenetic system in three additional bacterial organisms, B. subtilis, C. crescentus and S. pneumoniae, providing important considerations for the applicability of this system in microbiology and bacterial cell biology.

Biological sciences/Biological techniques/Optogenetics

Biological sciences/Cell biology/Cell division/Cytokinesis

Compartmentalizing biological processes at specific subcellular locations for specialized functions is a general strategy employed by cells across all domains of life. Eukaryotic cells often achieve subcellular localization using membrane-enclosed organelles. Bacteria cells generally lack membranous organelles but can also achieve subcellular localization by directing molecules to different scaffolds made of protein, DNA, RNA, or the plasma membrane to form stable assemblies and/or phase-separated condensates¹. These subcellular scaffolds underlie a number of fundamental cellular processes including partitioning of cellular content, chromosome organization and segregation, cell elongation and division, as well as appendage growth and receptor clustering^1,2. Controlling the recruitment of proteins to subcellular scaffolds and/or modulating scaffolds’ assembly states has proven to be a useful tool to probe and manipulate their functions³.

Previously, chemically induced protein dimerization platforms such as rapamycin-based FKBP⁴, coumermycin-based GyrB⁵ and gibberellin-based⁶ systems have been employed to manipulate subcellular localizations of various proteins in both bacteria⁷ and eukaryotic⁸ cells. While relatively fast and robust, these methods lack the precise spatial targeting ability and are often irreversible on fast timescales⁴. In contrast, optogenetic-based systems use light-modulated binding of a photoreceptor either with itself or with its associated protein ligand and is reversible^9,10; the application of light activation can be precisely defined in space to target specific cell subpopulations and even within a single cell’s volume^11–16. Currently available light-induced protein-protein interaction systems have been extensively characterized in mammalian cell systems³ and have kinetic rates on timescales ranging from milliseconds to hours¹⁷. Despite the widespread use of optogenetic systems in eukaryotic cells, their use in bacteria is only beginning to emerge (reviewed in³). The major difficulty of their implementation resides in the bacteria cell’s 1000-fold smaller cellular volume compared to that of a mammalian cell¹⁸. Such a small cytoplasmic volume imposes strict requirements on the concentration and kinetic ranges of the optogenetic systems in order to achieve a high signal contrast and spatial resoltuion^18,19.

One particular system, the CRY2/CIB1 system from Arabidopsis thaliana, can be induced by a low dose of blue light (peak activation wavelength at 468 nm) and exhibits rapid heterodimerization kinetics with tight²⁰, but reversible¹⁹, binding. These properties make it an attractive candidate for use in model bacterial systems. The system is composed of the N-terminal domain (amino acids 1-170) of AtCIB1 (CIBN) and the photolyase homology region²¹ (amino acids 1-498) of AtCRY2 (CRY2)^19,22. Here we show that this optogenetic system can be used to rapidly target proteins to major subcellular compartments in living Escherichia coli cells (Fig. 1a, left). We further demonstrate that such an optogenetically controlled system can be used to inhibit cytokinesis in living E. coli cells. We observed that the association and dissociation kinetics of the system can be modulated by green light, adding a new dimension of control to suit different synthetic biological systems. Lastly, we tested the system in three additional bacterial model organisms, B. subtilis, C. crescentus and S. pneumoniae, providing important considerations for the applicability of this system in microbiology and in bacterial cell biology.

General considerations for using the CRY2/CIBN system in Escherichia coli cells

Previous studies have shown that CIBN can be tagged at either its N- or C-terminus without impeding its binding to CRY2, whereas CRY2 functions the best with its N-terminus free²³. Therefore, in the following experiments, we fused CIBN at its N- or C-terminus to an array of proteins that localize to a subcellular compartment of interest to serve as the bait for the recruitment platform. We then fused a fluorescent reporter, mCherry, to the C-terminus of CRY2 to serve as a universal prey (Fig. 1a, right). The use of the same CRY2-mCherry reporter to a variety of subcellular recruitment platforms allows us to compare recruitment kinetics between different systems.

Next, to attain the desired control of CRY2/CIBN expression levels inside small bacterial cells in a variety of experimental scenarios, we exogenously expressed the CRY2/CIBN fusion proteins either co-transcriptionally from a single plasmid where the two fusions were coupled under the control of a lac-inducible promotor, or independently from a two-plasmid system where the CRY2 and CIBN fusions were under the control of arabinose- and lac-inducible promoters, respectively (Fig. 1b). Whether the one- or two-plasmid expression system was employed depended on specific considerations for each experiment outlined below. We found that the flexibility in modulating the expression levels of the CRY2 and CIBN fusion proteins is important for minimizing light-independent background interaction¹⁹ while maintaining a fast light-dependent recruitment speed (discussed below).

Rapid, reversible, and light-dependent recruitment of cytoplasmic protein to chromosomal DNA

In bacterial cells, the nucleoid is the major subcellular location for all DNA-related processes. The ability to optically control chromosomal DNA topology (e.g., the formation of DNA loops or topological domains) or the binding of transcription factors on specific DNA sequences has proven useful to study transcription dynamics and gene expression^19,24,25 in mammalian and bacterial cells^24,26–29. Therefore, we chose the chromosomal DNA as the first platform to develop a light-dependent recruitment assay of cytoplasmic proteins.

To visualize the light-dependent chromosomal recruitment process, we used an E. coli strain harboring a 240X tetO array sequence inserted near oriC in its chromosome³⁰ (Fig. 2a). The presence of tandem tetO arrays enhances the mCherry signal strength and facilitates the characterization of the recruitment kinetics. We then fused CIBN to the C-terminus of TetR, the tetracycline repressor that binds tightly to its cognate tetO operator sequence³¹ and co-transcriptionally expressed the TetR-CIBN and CRY2-mCherry reporter fusions exogenously from a single plasmid (Fig. 1c, left). The use of the coupled expression system helps to maintain a low and near equimolar expression of the two fusions proteins to best accommodate the 1:1 binding ratio of TetR-CIBN and CRY2-mCherry, effectively minimizing the background signal that would arise from excess CRY2-mCherry reporter. In the absence of the blue (488 nm) activation light, CRY2-mCherry remained homogenously distributed in the cytoplasm as expected for freely diffusing CRY2-mCherry in its unbound state (Fig. 2b, left). Upon exposure to 488 nm activation light pulses (30 ms pulses at 84.6 W/cm² delivered every 5 seconds over a 200 second period), we observed rapid formation of CRY2-mCherry foci at locations representative of individual oriC sites in almost all cells in the view field (96 ± 1.3 %, ± s.e.m., mean ± standard error of the mean, N = 3 independent experiments with 1359 cells in total, Fig. 2b, right, Fig. 2c, top, Supplementary Video 1) across a large range of CRY2-mCherry expression levels in individual cells. Notably, the cytoplasmic distribution of CRY2-mCherry remained unchanged in control cells exposed to the 488 nm light but that did not contain the 240X tetO array (Fig. 2c, middle panel, Supplementary Video 2), or in cells that contained the tetO array but not exposed to the 488 nm activation light (Fig. 2c, bottom panel, Supplementary Video 3). Quantification of the CRY2-mCherry signal (percentage of fluorescence increase compared to that before activation) at foci positions over time showed that 90% recruitment was reached within 85 seconds (τ_0.9 = 85 ± 9 seconds, µ ± s.e.m., N = 3 independent experiments with 442 cells in total, τ_0.9 was used to show the extent of reaction completion as the curve was not well described by an exponential function, Fig. 2d). These results demonstrate that the formation of CRY2-mCherry foci is rapid and both light- and target-dependent.

In addition to its rapid association kinetics, the CRY2/CIBN system is also reversible in the absence of blue light¹⁹. When we allowed the CRY2/CIBN system to relax in the absence of blue light following the complete localization of CRY2-mCherry to oriC foci (Fig. 2e, first blue shaded region), we observed an almost complete reversion of CRY2-mCherry foci back to a uniform cytoplasmic distribution after ~ 40 minutes (Fig. 2e, Supplementary Fig. 1). The decay curve was well described by an exponential function with a relaxation time constant of ~ 10 minutes (𝜏_rev = 10 ± 2 min, µ ± s.e.m., N = 3 independent experiments with 142 cells in total, Fig. 2e, shaded grey region). This relaxation time scale is consistent with the reversion times measured previously in mammalian cells^17,19. Additionally, relaxed cells could be activated once again^17,19 to re-form the same oriC foci with the nearly identical recruitment rate and final plateau (Fig. 2e, second blue shaded region and Supplementary Video 4). In summary, these results demonstrate that the CRY2/CIBN system can be used to induce reversible recruitment of a cytoplasmic protein to chromosomal DNAs rapidly and efficiently in live E. coli cells.

Rapid, light-dependent recruitment of cytoplasmic protein to cell pole

In bacteria, asymmetric cell division is often achieved by partitioning proteins to each incipient daughter cell via cell pole localized protein scaffolds³². In symmetrically dividing bacteria such as E. coli, the two poles are also differentiated in age, and proteins can be targeted specifically to the old cell pole to produce a dimorphic cell population^33,34. Furthermore, the cell pole can be engineered into an optically controlled apical protein sink to present an inert space away from the cytoplasmic environment³⁵.

To develop a light-triggered cell pole recruitment platform, we took advantage of a protein called PopZ from Caulobacter crescentus. PopZ forms a stable, liquid droplet-like matrix enriched at the old cell pole when heterologously expressed in E. coli cells^36–38 (Fig. 2f). To visualize the cell pole localization of PopZ, we fused CIBN-GFP to the N terminus of PopZ and placed the fusion gene under a lac-inducible promoter on a plasmid (Fig. 1b, right). After optimizing the expression of the PopZ fusion, we observed large, stable CIBN-GFP-PopZ foci at the cell poles of nearly every cell (Supplementary Fig. 2). We then expressed the CRY2-mCherry fusion under the control of an arabinose-inducible promoter from a separate vector (Fig. 1b, right). Here we used the two-plasmid system for independent expression control of the two fusion proteins, because the liquid droplet nature of the PopZ foci can accommodate a very high expression level of CIBN-GFP-PopZ, which ensures a stable cell pole localization and to provide a sufficient number of binding sites for CRY2-mCherry.

As shown in Fig. 2g and Supplementary Video 5, following a single 100 ms blue light activation pulse (P₄₈₈ = 8.5 W/cm²), we observed rapid formation of CRY2-mCherry foci at the cell poles. Importantly, in cells exposed to blue light only expressing GFP-PopZ, but not CIBN-GFP-PopZ (Fig. 2h, middle panel and Supplementary Video 6), or in cells without blue light activation (Fig. 2h, bottom panel and Supplementary Video 7), we observed unchanged, homogenous distribution of CRY2-mCherry signal in the cytoplasm throughout the time course of the experiment. Quantification of the percent increase in CRY2-mCherry signal at the cell poles revealed that 90% recruitment was achieved within ~ 8 seconds (τ_0.9 = 8.3 ± 0.9 seconds, µ ± s.e.m., N = 3 independent experiments with 315 cells in total, Fig. 2i). Note that in this system, the recruitment of cytoplasmic CRY2-mCherry was sped up to ~ 10-fold compared to that in the DNA recruitment assay. The faster kinetics are most likely driven by the higher expression levels of both CIBN-GFP-PopZ and CRY2-mCherry in the system.

Formation of inner membrane-associated heteroclusters of CRY2/CIBN upon light activation

In many cell types, the inner membrane (IM) is the site of a diverse array of signaling pathways. In bacteria, the IM also serves as the assembly site for large macromolecular structures such as the divisome³⁹, the flagellum⁴⁰, or chemotaxis receptor clusters⁴¹. In mammalian cells, synthetic activation of signaling cascades can be achieved using cytoplasmic CRY2-tagged proteins that are recruited to membrane-anchored CIBN^{16,19,23,42,43}. To develop a light-dependent membrane recruitment assay in bacteria, we fused CIBN-GFP to an amphipathic helix from Bacillus subtilis (BsMTS)⁴⁴ and expressed it from a lac-inducible promotor independently from the arabinose-inducible CRY2-mCherry on a second plasmid. Here we used an exogenous membrane anchor to avoid potential interference with endogenous E. coli membrane proteins. Notably, the BsMTS-GFP-CIBN fusion showed uniform membrane localization as expected (Supplementary Fig. 3a, left). Upon blue light activation (50 ms pulses at 84.6 W/cm² delivered every 5 s), we observed the formation of dynamic CRY2-mCherry puncta that co-localized with CIBN-GFP-BsMTS puncta along the IM (Supplementary Fig. 3b, & Supplementary Video 8). The distributions of these puncta were distinct from the near- uniform distribution of CIBN-GFP-BsMTS along the IM surface prior to activation with blue light (Supplementary Fig. 3a, left), or that of directly membrane-targeted CRY2-GFP-BsMTS (Supplementary Fig. 3a, right). We reason that the puncta formation along the IM of BsMTS-CIBN-GFP and CRY2-mCherry upon light activation is likely indicative of the formation of individual membrane-tethered CIBN-CRY2 heteroclusters as previously predicted^12,23,45. We were unable to achieve uniform co-localization of CRY2-mCherry and CIBN on the IM of E. coli cells, despite the many strategies we employed (Supplementary Table 1, Reference 46). This ability is in contrast to what was observed previously in mammalian cells^12,19. Further experiments are required to investigate this phenomenon.

Rapid, light-dependent recruitment of cytoplasmic proteins to the midcell

In almost all bacteria studied to date, the essential process of cytokinesis is mediated by the formation of a large macromolecular complex, termed the divisome, at the future division plane of the cell⁴⁷. In E. coli, the divisome is comprised of more than 30 different proteins, which are recruited to the midcell by the essential tubulin homologue, FtsZ^39,48. FtsZ polymerizes at the midcell to assemble into a ring-like structure, and new data suggests that the dynamics of the Z-ring are essential to coordinate the action of the divisome for cell wall constriction and septum morphogenesis during cell division^49–51. Achieving a means to rapidly and reversibly deliver proteins to the divisome and/or modulate the assembly of the divisome at different cell division stages would provide a new way to probe the dependence of different divisome proteins and their functions on the assembly state of the divisome specifically at the division site⁵².

To achieve optically controlled delivery of proteins to the cell division plane, we first tested the ability to recruit CRY2-mCherry to the Z-ring by creating a C-terminal fusion of CIBN to ZapA (Fig. 3a). ZapA is a conserved Z-ring-associated protein, which has served as a faithful Z-ring marker in previous studies^53–56. Similar to the pole-recruitment platform, we expressed the ZapA-CIBN fusion independently from the CRY2-mCherry reporter using the two-plasmid expression system so that their expression levels could be optimized separately. Upon exposing cells to a single 100 ms pulse of 488 nm activation light (P₄₈₈ = 8.5 W/cm²), we observed rapid midcell localization of CRY2-mCherry fluorescence (τ_0.9 = 9.3 ± 0.15 seconds, µ ± s.e.m., N = 3 independent experiments with 443 cells in total, Fig. 3b-d, Supplementary Video 9). Similar to what was observed in the chromosomal DNA and cell pole recruitment assays, the midcell localization of CRY2-mCherry was dependent on the blue activation light (Fig. 3c). These results demonstrate that cytoplasmic proteins can be recruited to the Z-ring at the midcell rapidly and specifically using the CRY2/CIBN system.

Light-dependent destabilization of the Z-ring and inhibition of cell division

We next used the midcell recruitment platform to develop a Light-induced Inhibition of Cytokinesis (LInC) assay in live E. coli cells (Fig. 3e). In the LInC assay, we kept the same ZapA-CIBN fusion as the bait, but fused a Z-ring antagonist, MinC, to CRY2 to generate a CRY2-MinC fusion. MinC inhibits FtsZ polymerization in vitro and prevents Z-ring assembly at the cell poles by forming a high to low concentration gradient from the cell pole to the midcell with its membrane binding partner ATPase MinD⁵⁷. When highly overexpressed by itself, MinC antagonizes Z-ring assembly at the midcell as well^57–59. We verified that the CRY2-MinC fusion is fully functional as it complemented a minC deletion background (Supplementary Fig. 4). Therefore, by recruiting CRY2-MinC to FtsZ-associated ZapA-CIBN, we expect to increase the local concentration of MinC at the Z-ring, which may result in the destabilization of the Z-ring and subsequent inhibition of cell division (Fig. 3e). To visualize the progress of Z-ring destabilization in the LInC assay, we tagged the endogenous chromosomal copy of ZapA with mCherry as the Z-ring marker^53,54.

We first established a growth and expression conditions under which cells ectopically expressing the LInC system (ZapA-CIBN and CRY2-MinC) exhibited a near-WT midcell Z-ring morphology and underwent successful cell division cycles in the absence of blue light (Supplementary Note 1). Notably, these cells were moderately longer than uninduced cells due to the presence of higher than WT CRY2-MinC concentrations (L = 3.4 ± 0.4 µm, µ ± s.d., N = 3 independent experiments with 1465 cells in total compared to L = 2.3 ± 0.06 µm, µ ± s.d., N = 2 independent experiments with 379 cells in total for uninduced cells). We then imaged the Z-ring morphology of these cells before and after exposing them to 100 ms blue light pulses (P₄₈₈ = 84.6 W/cm²) every 10 seconds for a period of 5 minutes (Fig. 3f). Before the activation light exposure, we quantified that on average ~ 46% of all the cells harbored clear ZapA-mCherry localization at midcell, consistent with previous reports that the Z-ring only assembles after a significant portion of the chromosome is segregated⁶⁰. About 53% of these cells (53.3 ± 3.97%, µ ± s.e.m., N = 3 independent experiments with 1658 cells in total) exhibited considerably broader and more diffuse ZapA-mCherry midcell fluorescence upon exposure to blue light compared to that prior to light exposure (Fig. 3f & g), indicating the destabilization of the Z-ring. In uninduced cells or in cells without blue light activation, we observed significantly lower, ~ 40% (40.3 ± 15.0%, µ ± s.e.m., N = 2 independent experiments with 379 cells in total) or ~ 44%, (44.1 ± 1.68%, µ ± s.e.m., N = 3 independent experiments with 1433 cells in total), cells exhibiting diminished ZapA-mCherry midcell fluorescence after a 5-minute period, indicative of the natural disassembly process of the Z-ring during cell division. Interestingly, the extent of the Z-ring destabilization appeared to be dependent on the cell division stage (Fig. 3h, Supplementary Table 2). We observed that after Z-ring assembly, cells of relatively short lengths (4–8 µm in length, indicative of earlier cell division stages) showed significant destabilization of the Z-ring (Fig. 3h, blue) compared to cells of longer lengths, or cell of similar lengths but without the activation light (Fig. 3h, grey). Previous studies have shown that when the divisome matures and cell wall constriction initiates, the Z-ring becomes denser and denser until it finally and rapidly disassembles toward the end of the cell cycle^61,62. Therefore, it is possible that the LInC assay destabilizes the Z-ring effectively only before the Z-ring becomes too dense to be effectively depolymerized by MinC.

While we were able to achieve efficient and rapid Z-ring destabilization in cells judged by their widened Z-ring morphology and the reduction in the midcell localization percentage of FtsZ, it remained unclear whether these destabilized Z-rings were indeed defective in cell division. To address this, we monitored cell division for a long period of time in cells harboring the LInC system after induction of the system with blue light. As shown in Fig. 3i, when we subjected LInC cells to 100 ms pulses of 488-nm activation light (P₄₈₈ = 8.5 W/cm²) and 561-nm imaging light (P₅₆₁ = 8.7 W/cm²) every 5 minutes for a twelve-hour period, we observed that the majority of these cells ceased division and grew into long filaments instead. On average, ~ 76% (75.7 ± 12.3, µ ± s.e.m., N = 3 experiments with 33 cells) were unable to divide during the 12-hour period. Additionally, the average division time (𝜏_LInC = 355 ± 84.6 minutes, µ ± s.e.m., N = 3 experiments of 33 cells in total) of the remaining ~ 24% (24.3 ± 12.3, µ ± s.e.m., N = 3 experiments of 31 cells in total) cells that were able to divide eventually was nearly 1.5 times longer than the mean division time of the control group where the ZapA-CIBN bait fusion was absent (𝜏_control = 232 ± 31.2 minutes, µ ± s.e.m., N = 3 experiments of 31 cells in total). These observations indicated considerable deficiency of LInC cells in division. The inhibited cell division also correlated with significantly destabilized Z-rings—in all long, filamentous cells, the midcell fluorescence of ZapA-mCherry exhibited a wider spread, in contrast to the sharp midcell localization prior to light exposure (Fig. 3j). Note that the inhibition of cell division in these cells was not caused by photodamage due to the continuous illumination of the 488 nm activation light and the 561 nm imaging light, because cells under the same induction and illumination condition but ectopically expressing only CRY2-MinC grew and divided as wild type cells not exposed to light (Fig. 3i, top row). Furthermore, the inhibition was 488-nm light-dependent because cells expressing both CRY2-MinC and ZapA-CIBN in the absence of the blue light grew normally as WT cells (Fig. 3i, bottom row rightmost panel). Taken together, these results show that the CRY2/CIBN system can be used to destabilize the Z-ring rapidly and inhibit cell division in living E. coli cells in a light-dependent manner.

CRY2/CIBN complex dissociation and activated CRY2 concentration are sensitive to green light

Previous studies have shown that the light sensitivity of CRY2 is imparted by its flavin adenine dinucleotide (FAD) cofactor through a photocycle^52,53 (Fig. 4a). In this cycle, the redox state of CRY2’s FAD is thought to mediate a conformational change in CRY2 to form a signaling-active state that is competent to bind CIB⁶³. Briefly, the fully oxidated FAD^ox cofactor is in the resting inactive state. Following photoexcitation by blue light, FAD^ox accepts an electron from a nearby tryptophan followed by protonation from an aspartic acid to become the semi-reduced, stable semiquinone radical (FADH^o). This redox state is thought to trigger a conformational change in CRY2 (denoted as CRY2*) that allows for its binding to CIB^64,65. To complete the photocycle, semi-reduced FADH^o is reduced slowly to become FADH⁻, reversing the conformation change, after which it is oxidated rapidly back to its fully-oxidized resting state (FAD^ox). Previous studies have indicated that the semiquinone form FADH^o is the only green-absorbing species in FAD’s photocycle, and that green light can deplete FADH^o in vitro⁶⁵. In short-day entrained plants, green light leads to a delay in flowering, likely due to the increased reduction of FADH^o to FADH⁻ under green light exposure^54,55. However, previous characterization of CRY2/CIBN complex formation in vivo in heterologous expression systems showed no sensitivity to green light¹⁹.

We reason that if the association/dissociation kinetics of the CRY2/CIBN are indeed sensitive to green light, we could use green light (561 nm) in combination with blue activation light to fine tune the kinetic behavior of the CRY2/CIBN system. To investigate this possibility, we swapped the mCherry reporter with Halo⁶⁶, which can be covalently labeled with a bright, photostable far-red organic dye⁶⁷ (JF646), allowing us to avoid using green light to image the reporter. We verified that the extent of CRY2/CIBN association indicated by the percentage increase in cells expressing the Halo-labeled system exhibited a blue-light activation dose dependence (Fig. 4b) as expected¹⁹, and that the final association plateau could be further increased using subsequent multiple pulses of activation to recruit additional CRY2-Halo to the pole (Supplementary Fig. 5). These results indicate that CRY2-Halo can be activated in a similar manner as CRY2-mCherry by a 100 ms blue pulse, and that the final pole-enrichment of CRY2 is dependent on the available level of activated CRY2, which can be modulated by the intensity of the activation pulse.

Next, to investigate the influence of green light on CRY2/CIBN association, we monitored CRY2/CIBN complex formation at the cell poles using a single 100 ms activation pulse of 8.5 W/cm² blue light followed by a continuous illumination of green light of different intensities (561 nm, from 8.71 to 970 W/cm²) accompanying the imaging 647 nm light (50 ms exposure for 15 seconds). We found that the amount of CRY2 recruited to the cell pole decreased significantly with increasing green light intensity, accompanied by slight changes in the 90% recruitment value (Fig. 4c, left and right panels respectively, Supplementary Fig. 6, Supplementary Table 3). At the highest green light intensity (970 W/cm²), the pole-recruited CRY2 fraction was reduced to ~ 50% compared to that in the absence of the green light. The decreased level of recruitment but largely unaffected time constant are most consistent with decreased levels of activated CRY2* under the green light illumination condition; the green light potentially drives activated CRY2* back to the inactivated state even before it forms a complex with CIBN. Hence, the green light could modulate the level of activated CRY2* and subsequently the amount of CRY2*-CIBN complex formation in live E. coli cells.

To investigate whether green light might also modulate CRY2/CIBN’s dissociation kinetics, we light-induced CRY2-CIBN complex formation using blue light and then monitored the dissociation kinetics of the complex in the presence or absence of green light (561 nm, 473 W/cm², 100 ms pulses every 3 min, Fig. 4d). We found that while the fold of reduction of CRY2-CIBN complex formation indicated by the final dissociation plateau (Supplementary Fig. 7) did not appear to change dramatically, we observed a considerable decrease in the time constant of the dissociation reaction from 18.3 ± 8.2 minutes (N = 4 independent experiments with 91 cells in total) in the absence of green light to 12.7 ± 5.7 minutes (µ ± s.e.m., N = 4 independent experiments with 62 cells in total) in the presence of the green light, (Fig. 4d, left and right panel respectively, Supplementary Fig. 7, Supplementary Table 3). These observations suggest that the green light might also act upon activated CRY2* in complex with CIBN to drive it back to the inactivated from, effectively speeding up the dissociation of the CRY2*-CIBN complex. This process is possibly through the photoreduction of FADH^o within the CRY2*-CIBN complex. Further experiments are needed to test this possibility.

Adapting CRY2/CIBN to other bacterial species

The rod-shaped monoderm bacteria B. subtilis, the ovoid-shaped monoderm pathogen S. pneumoniae, and the crescent shaped diderm bacteria C. crescentus are three widely used model organisms for bacterial cell biology and genetic studies. Testing whether the CRY2/CIBN system could also be adapted to these organisms will provide first-of-its-kind information on the feasibility of developing new optogenetic tools in species other than E. coli. As we describe below, for each species, we tested a variety of expression, labeling and imaging conditions (Supplementary Information). Among these variables, we discovered that species-specific codon optimization and expression levels were most critical. For all the studies presented below, we used species-codon-optimized fusion proteins. We obtained variegated degrees of adaptation in these species, none of which were as robust as what we observed in E. coli. Nevertheless, our observations are important for further explorations of the CRY2/CIBN system in different bacterial species.

For B. subtills, we employed the Z-ring recruitment platform (Fig. 5a) as the Z-ring assembly pathway in B. subtilis is well established. We fused a B. subtilis codon-optimized CRY2 to the C-terminus of tdTomato and placed the fusion gene under the control of an IPTG inducible promoter at the chromosomal thrC locus (thrC::P_hyperspank-CRY2^Bs-tdTomato) in strain SC757 (see Supplementary Note 2 for full genotype and strain construction). We then expressed a zapA-CIBN fusion gene under control of the native zapA promoter (P_zapA-zapA-CIBN) at the amyE locus of the chromosome. Similar to E. coli, ZapA^Bs localizes to the midcell concurrently with FtsZ⁶⁸. When we grew B. subtilis cells expressing both the CRY2-tdTomato and ZapA-CIBN fusions in M9 + medium and exposed them to sequential pulses of blue and green light (50 ms pulses of 488 nm light at 1.28 W/cm² and 500 ms pulses of 561 nm light at 1.63 W/cm², delivered every 3 seconds). we observed blue light-dependent septal localization of CRY2-mCherry over the course of 8 minutes in 16 ± 6% cells (µ ± s.e.m.; N = 4 independent experiments with 810 cells in total) (Fig. 5b, c). The recruitment of CRY2-tdTomato to midcell localized ZapA-CIBN was slower than that in E. coli, with an initial lag period of ~ 150 sec followed by a largely linear increase phase to ~ 15% over 8 min (Fig. 5d). Further optimization of the expression and imaging conditions of the system using cells grown in casein hydrolysate (CH) media, a rich media used for conventional B. subtilis imaging experiments, produced variable degrees of midcell localization of CRY2-mCherry with similar kinetics (Supplementary Fig. 8a-c).

For C. crescentus, we used the PopZ localization platform, as cell pole in C. crescentus is an important location for asymmetric cell division and stalk growth^69–71. We expressed C. crescentus codon optimized mCherry-CRY2 fusion under the control of a xylose-inducible promoter (P_Xyl-mCherry-CRY2^Cc) and a CIBN-GFP-PopZ fusion under the control of an orthogonal vanillate-inducible (P_van-CIBN-GFP-popZ) promoter (see Supplementary Table 5). CIBN-GFP-PopZ localized to the cell pole in greater than 90% of cells even in the absence of vanillate induction due to the leaky expression of the vector (Supplementary Fig. 9a). However, despite our best effort in optimizing expression levels, growth conditions and imaging conditions (Supplementary Information), we only observed that maximally in ~ 30% of cells, diffusive, cytoplasmic mCherry-CRY2 relocated to cell poles. The pole localization is dependent on the expression of CIBN-GFP-PopZ but independent of vanillate induction or blue light activation (Supplementary Fig. 9a, b). The relatively low efficiency and light-independence in CRY2-mCherry’s pole localization are likely due to the less optimal folding of CRY2 and the small cell volume of C. crescentus cells respectively, as we observed heterogenous expression of CRY2-mCherry across cell populations (Supplementary Fig. 9a). This example highlights the challenge in optimizing the expression level of the CRY2-CIBN system in smaller bacterial cells such as C. crescentus.

For S. pneumoniae, we also employed the Z-ring recruitment platform. We constructed various bait (ZapA-CIBN or FtsZ-CIBN) and prey fusions (CRY2-mCherry, CRY2-Halo, or CRY2-tdTomato) and tested a variety of codon optimization strategies, genetic backgrounds, and growth and imaging conditions. However, despite our best efforts, we were unable to observe any midcell recruitment of CRY2 fusions (see Supplementary Note 3 & Supplementary Table 6). The underlying reason is unclear, but it may involve poor folding of expressed CRY2/CIBN domains in S. pneumoniae, blockage of CRY2/CIBN by interactions with pneumococcal proteins, absorbance of blue light by proteins in this bacterium, and/or limited availability of FAD cofactor for CRY2 binding.

The landscape of optogenetic systems to induce protein-protein interactions inside living cells is constantly growing^3,70. Optogenetic systems are especially useful for live-cell imaging studies due to their rapid induction, reversible binding and spatial precision^71,72. Here we demonstrate that the Arabidopsis thaliana CRY2/CIBN system (Fig. 1a), used extensively in mammalian cell systems, can be adapted to target a protein cargo to different subcellular structures including the chromosomal DNA, cell pole, inner membrane and the division plane at the midcell in living E. coli cells (Fig. 1b). During the optimization process we found that the expression levels of CRY2/CIBN system in bacteria cells can be varied to achieve recruitment halftimes ranging from the seconds to minutes timescales similar to those observed in mammalian cells^19,23. We then demonstrate that the CRY2/CIBN system can also be implemented to rapidly destabilize the cytokinetic ring and inhibit cell division in E. coli (Fig. 3), providing a rapid and spatially specific tool to manipulation bacterial cell division.

During our characterization of the CRY2/CIBN system in E. coli, we found that both the association and dissociation of CRY2 with CIBN can be modulated by green light (Fig. 4c). This effect not only enables another layer of system tuning akin to the blue light dose-dependent activation behavior (Fig. 4b) that we and others^19,25 have observed, but also provides direct evidence that green light inactivates CRY2 likely by acting on the semi-reduced FADH^o cofactor directly to reverse the active conformation of CRY2, hence lowering the amount of CRY2* available for complex formation and also speeding up the dissociation kinetics of previously formed CRY2*-CIBN complex. These results are consistent with previous experiments where green light exposure was found to rapidly convert CRY2’s cofactor from FADH^o to FAD_ox in vitro and inhibit CRY2/CIB1 activity in vivo⁶⁵.

In adapting the CRY2/CIBN system to other bacterial species, however, we observed that it was most successful in B. subtilis, less efficient in C. crescentus, and did not work in S. pneumoniae (Fig. 5, Supplementary Fig. 9, Supplementary Note 3). These observations highlight the feasibility and also challenges in implementing the optogenetic system in bacterial species other than E. coli, as specific codon optimization, cytoplasmic folding environment, and cell physiology need to be carefully considered on a species-by-species and need-specific basis.

In mammalian cells the CRY2/CIBN system has already been employed to optogenetically control cellular pathways including Cre recombination⁷¹, Raf/MEK/Erk²³, TGF-β⁷², NF-𝜅B¹⁵, Wnt/β-catenin¹¹, and RhoA¹¹. Most of these systems involve the binding of two proteins to initiate a signaling cascade. In bacteria, we envision that the CRY2/CIBN system can be similarly applied to a large number of processes to enable optogenetic control. For example, most of the two-component signaling (TCS) systems⁷³ in bacteria employ a histidine kinase that, upon stimulation, will recruit and phosphorylate its cognate response regulator resulting in the activation of downstream signaling⁷³. These protein pairs could be tagged and controlled in a similar manner as done in eukaryotes to allow for the pathway regulation in real-time. Furthermore, a number of pre-existing chemically inducible systems could be transformed into optogenetic systems by modifying their dimerization platforms. Potential candidate systems could include split ClpXP adaptors for optical control of protein degradation⁷, split T7 RNAP for optical control of gene expression²⁴ and split Cas9 for optical control of gene editing⁷³.

E. coli bacterial strains, plasmids, and growth conditions

All E. coli strains used for the cell pole recruitment, Z-ring recruitment, and cytokinesis inhibition experiments were derivatives of BW25113 (CGSC #7636) strain and were grown in M9 + Glucose minimal media (recipe is provided in Supplementary Table 4). The DNA recruitment background strain was previously constructed by the Sherratt Lab³⁰ (CGSC #12294) and was grown in EZ Rich Defined Media (EZRDM, Teknova Bio.) as described in Supplementary Table 4. Antibiotic requirements are specified in Supplementary Table 4 and the corresponding concentrations for each were chloramphenicol (150 µg/mL), streptomycin (50 µg/mL) and gentamycin (5 µg/mL). Inducer concentrations for respective strains and plasmid combinations are outlined in Supplementary Table 4 and described in more detail below. All bacterial cultures were grown in tubes covered in foil to minimize contact with ambient light. Plasmids expressing exogenous CRY2/CIBN fusion proteins with their associated sequences are described in Supplementary Table 4.

Recruitment of CRY2-mCherry to the DNA via TetR-CIBN in E. coli

Cultures were started from a single colony of the strain RM187 and grown in 6 mL EZ Rich Defined Medium (EZRDM, Teknova Bio.) containing chloramphenicol and gentamycin at 25 ºC with shaking until the cells entered early log-phase (OD600 between 0.1 and 0.2, ~ 19 hours). When the culture entered early log-phase the cells were induced with 60 µM IPTG and supplemented with 240 nM anhydrous-tetracycline (ATC) to avoid a replication block induced by binding of TetR-CIBN similarly as previously described. The cells were induced for 2 hours at 25 ºC with shaking. At the end of the induction period the cells were centrifuged for 10 minutes at 4110 rpm and the pellet was resuspended in 6 mL of EZRDM containing chloramphenicol and 240 nM ATC and grown for an additional 30 minutes at 25 ºC while shaking after which it was, again, centrifuged for 10 minutes at 4110 rpm and resuspended in 6 mL EZRDM containing chloramphenicol and gentamycin and allowed to grow an additional 2.5 hours at 25 ºC while shaking to allow the mCherry fluorophore to mature. Following the outgrowth period, the cells were prepared for imaging by centrifuging 1 mL of the culture at 13k rpm for 1 minute and then resuspending the pellet in 50 µL of fresh EZRDM. Following this, 0.5 µL of the resuspension was then placed on a gel pad and the pad and prepared for imaging as specified in the “Live cell imaging” section below.

Recruitment of CRY2-mCherry to the cell pole via CIBN-GFP-PopZ and the Z-ring via ZapA-CIBN in E. coli

Cultures were started from a single colony of the strain of interest and grown in M9 + Glucose minimal medium containing chloramphenicol and streptomycin at 37 ºC overnight until the cells reached stationary phase. The overnight culture was then diluted 1:100 in fresh of M9 + Glucose containing chloramphenicol and streptomycin and was allowed to grow at 37 ºC until it reached log-phase (OD600 between 0.1 and 0.2, ~ 3 hours) at which point the culture was induced with 0.4% arabinose and either 40 µM IPTG (ZapA fusion) or 100 µM IPTG (PopZ fusion). Following a 1-hour induction at 37 ºC the cells were prepared for imaging by centrifuging 1 mL of the culture at 13k rpm for 1 minute and then resuspending the pellet in 70 µL of fresh M9 + Glucose. Following this, 0.5 µL of the resuspension was then placed on a gel pad and the pad and prepared for imaging as specified in the “Live cell imaging” section below. The cells on the gel pad were then assembled into the imaging chamber and allowed to equilibrate on the microscope in the dark for 2 hours at ambient room temperature (RT).

Recruitment of CRY2-HaloTag to the cell pole via CIBN-GFP-PopZ using variable green (561nm) light in E. coli

Cultures were started from a single colony of the strain of interest and grown in M9 + Glucose minimal medium containing chloramphenicol and streptomycin at 37 ºC overnight until the cells reached stationary phase. The overnight culture was then diluted 1:100 in fresh of M9 + Glucose containing chloramphenicol and streptomycin and was allowed to grow at 37 ºC until it reached log-phase (OD600 between 0.1 and 0.2, ~ 3 hours) at which point the culture was induced. Following a 1-hour induction at 37 ºC the cells were prepared for imaging by centrifuging 1 mL of the culture at 13k rpm for 1 minute and then resuspending the pellet in 100 µL of fresh M9 + Glucose with 1 µM Janelia Fluor® 646 HaloTag® ligand (Promega Corp.) and allowed to incubate at RT for 2 hours covered in foil. Following this, 0.5 µL of the resuspension was then placed on a gel pad and the pad and prepared for imaging as specified in the “Live cell imaging” section below.

Rapid Z-ring decondensation experiments in E. coli

Cultures of strain RM077 and RM078 were started from a single colony and grown in LB at 37 ºC overnight until the cells reached stationary phase. The overnight LB culture was then diluted 1:200 in 3mL of M9 + Glucose and grown overnight at room temperature (RT, 25 ºC) until they reached log-phase (OD600 between 0.1 and 0.2) at which point the culture was induced with 10µM IPTG and 0.2% arabinose. Following a 2-hour induction at RT. Cells were prepared for imaging by centrifuging 1 mL of the culture at 13k rpm for 1 minute and then resuspending the pellet in 100 µL of fresh M9 + Glucose. Following resuspension, 0.5 µL of the resuspension was transferred to a 3% M9 + Glucose agarose gel pad containing 10µM IPTG and 0.2% arabinose for imaging.

Light induced inhibition of cytokinesis experiments in E. coli

Cultures were started from a single colony of the strain of interest (see Supplementary Table 4) and grown in LB containing chloramphenicol and/or streptomycin (depending on the strain background, see Supplementary Table 4) at 37 ºC overnight until the cells reached stationary phase. The overnight culture was then diluted 1:100 in fresh of M9 + Glucose containing chloramphenicol and streptomycin and was grown overnight at 25 ºC until it reached log-phase (OD600 between 0.1 and 0.2) at which point the culture was induced. Following a 2-hour induction at 25 ºC the cells were washed into fresh M9 + Glucose media lacking inducer and allowed to outgrow at 24 ºC for 2 hours. After the outgrowth, the cells were prepared for imaging by centrifuging 1 mL of the culture at 13k rpm for 1 minute and then resuspending the pellet in 40 µL of fresh M9 + Glucose. Following this, 0.5 µL of the resuspension was then placed on a gel pad and the pad and prepared for imaging as specified in the “Live cell imaging” section below. Control cells were placed in a chamber, wrapped in foil, left at RT in next to microscope for the duration of the experiment and imaged after the experiment ended.

B. subtilis culturing, sample preparation, and imaging

Cultures were started from a single colony of the strain SC757 grown on LB plates at 30 ºC for 16–20 hours. We note all the following culturing, sample preparation, and induction steps occurred in the dark to avoid pre-activation caused by ambient light.

For imaging in M9 + media (glucose minimal media with vitamins, see Supplementary Table 4), single colonies from fresh plates were then inoculated into M9 + with 100 µM IPTG to induce CRY2-tdTomato for 6–7 hours at 30 ºC shaking. When the culture entered early log phase between OD600 0.1 and 0.3, 500 µL of the cultures were harvested by centrifuging for 5 minutes at 5000 rpm and the pellet resuspended in 50 µL of M9 + by vortexing. 1 µL of the resuspended culture was spotted on 3% (w/v) M9 + agarose pad supplemented with 100 µM IPTG. The sample was allowed to equilibrate at the imaging temperature of 25 ⁰C for 1 hour prior to imaging.

For imaging in B. subtilis casein hydrolysate (CH) media⁷⁴ (B. subtilis rich media commonly used for imaging⁷⁵), single colonies from fresh plates were inoculated into CH media with 50 µM IPTG to induce CRY2-tdTomato for 16–20 hours at 25 ºC shaking in 10-fold dilution series. 500 µL of the cultures in early log phase between OD600 0.1 and 0.3, were harvested by centrifuging for 5 minutes at 5000 rpm and the pellet resuspended in 50 µL of CH media by vortexing. 1 µL of the resuspended culture was spotted on 3% (w/v) CH agarose pad supplemented with 50 µM IPTG. The sample was allowed to equilibrate at the imaging temperature of 25 ⁰C for 1 hour prior to imaging.

To image the cells, cells were placed into focus using the 561 nm imaging laser that was also used for image acquisition at 1.63 W/cm² measured 1 cm from the objective. The 488 nm laser power used to activate CRY2/CIBN system was 1.28 W/cm² measured 1 cm from the objective in an area 200 x 200 pixels (160 nm/pixel). Every 3 seconds, cells were pulsed with blue light for 500 ms to activate the system followed by 50 ms green light to acquire image for a total of 160 frames (8 min).

C. crescentus culturing, sample preparation, and imaging

Cultures were started from single colony of EG3979 and EG3988 grown on PYE supplemented with both 5 µg/mL gentamycin and 25 µg/mL kanamycin or only kanamycin, respectively, at 30 ºC for 48-72h. Colonies from fresh plates were then inoculated into PYE supplemented with appropriate antibiotics (1 µg/mL gentamycin and/or 5 µg/mL kanamycin) and grown overnight to an OD600 of ~ 0.3–0.6. Cultures were diluted to an OD600 of ~ 0.08 in 2 mL of appropriate media supplemented with 0.3% xylose and 0.5 mM vanillate for EG3979 or just xylose for EG3988 and induced for 2 hours. Control samples were left growing in appropriate media lacking inducers. All culture tubes were protected from ambient light.

500 µL of cells were concentrated 10X by spinning for 5 minutes at 5000 rpm, removing 450µL of media, and resuspending in left over 50 µL, 1 µL was spotted on PYE agarose pad containing gentamycin and kanamycin for EG3979 or just kanamycin for EG3988. Samples were allowed to equilibrate at the imaging temperature of 25 ºC for 30 minutes prior to image acquisition at 25 ºC.

Live cell imaging

Live cell imaging was performed on a custom-built optical setup routed to an Olympus IX71 inverted microscope with a 100X 1.49 NA oil-immersion objective (Olympus Inc.). The light was focused onto the chip of an EMCCD camera (iXon Ultra 897, Andor Technology) with a final pixel size of 160 x 160 nm. The imaging focal plane was controlled by a piezo-driven stage (ASI, Eugene, OR). The EMCCD camera, lasers and shutters were controlled by Metamorph™ software (Molecular Devices).

Excitation light was provided by solid state 488 nm (Coherent OBIS), 561 nm (Toptica Photonics) or 647 nm (Coherent OBIS) lasers. The fluorescence emission signal was collected using either a ZET 488/561 nm (Chroma Technology) or ZET 488/561/647 nm (Chroma Technology) dual-band dichroic depending on the imaging conditions used. For two-color experiments, the emission light from the RFP and GFP channels were split using a 525/50 (Chroma Technology) and 650/50 (Chroma Technology) filter set mounted inside of an Optosplit II beam-splitter system (Cairn Research) prior to focusing on the EMCCD camera chip. The 488 nm laser power used to activate the CRY2/CIBN system was the same as what was used to image GFP fusions at 8.08 W/cm² measured 1 cm from the objective for PopZ and ZapA experiments and 80.8 W/cm² measured 1 cm from the objective for the TetR experiments measured using a Thor Labs Inc. Power Meter). The 561 nm laser power used to image the mCherry fusions was 8.71 W/cm² measured 1 cm from the objective for the PopZ and ZapA experiments and 93.3 W/cm² measured 1 cm from the objective for the TetR experiments unless otherwise noted in the text.

For two-color experiments using the 647 nm laser, the emission light from the RFP and GFP channels were split using a 556 nm long-pass filter (T556lpxr, Chroma Technology) in the Optosplitter followed by 700/75 (Chroma Technology) filter for the RFP emission light prior to focusing on the EMCCD camera chip.

To avoid activation of the CRY2/CIBN system by the LED bright-field lamp (LDB100F System, Prior Scientific Inc.), we placed a 715nm long-pass filter (RG715, Thor Labs Inc.) or a 665nm long-pass filter (ET665lp, Chroma Technology) filter after the condenser and the BF lamp intensity was adjusted such that the measured 488 nm light after the filter was measured to be ~ 200 nW at 1 cm from the sample plane.

Gel pads used to support the living bacteria cells were prepared by melting 3% w/v solution of agarose powder (SeaPlaque™ GTG™, Lonza Scientific) and either EZRDM or M9 + Glucose for 0.5-1 hour at 70 ºC. The melted gel was then transferred to a gasketed, cover-glass (FCS2, Bioptechs Inc.) and allowed to polymerize while sandwiched between the cover-glass and a cleaned coverslip (40 CIR-1, VWR Inc.) for 4 hours at 25 ºC (room temperature). 0.5 µL of the cells to be imaged were placed on the gel pad and allowed to dry (~ 3 minutes) before a fresh coverslip was placed on top and the imaging chamber (FCS2, Bioptechs) was assembled.

Generally, the blue light intensities and pulse sequences used to activate CRY2/CIBN in various imaging configurations were chosen to minimize phototoxicity of cells to blue light while maximizing the amount of activated CRY2. Whenever possible, we tried to minimize both the intensity and the number of blue pulses we delivered over the course of an acquisition. However, if the observed level of complex formation was not sufficient under a particular blue light induction scheme, we increased the number of pulses and or the intensity of blue light used to activate the system accordingly.

For the DNA recruitment experiments, the cells were irradiated with 30 ms pulses of 488 nm and 561 nm light delivered consecutively every 5 seconds using a 30 ms exposure time for each.

For the cell pole and Z-ring recruitment experiments the cells were first imaged for 2.5 seconds with 561 nm light after which they were irradiated with 488nm light for 100ms and then imaged with 56 nm light by streaming for 15 seconds using a 50ms exposure time.

For the cell pole recruitment experiments where the green (561 nm) light was varied: the cells were first imaged for 2.5 seconds with both 647 nm and 561 nm light after which they were irradiated with 488 nm light for 100 ms and then imaged with 647 nm and 561 nm light by streaming for 15 seconds using a 50 ms exposure time.

For the fast Z-ring decondensation experiments, cells were initially imaged with 561 nm light after which they were irradiated with 50 ms pulses of 488 nm light delivered consecutively every 10 seconds for a 5-minute period. The cells were then imaged again with 561 nm light at the end of the 5-minute period.

For the Light-induced Inhibition of Cytokinesis (LInC) experiments, cells were irradiated with alternating 100 ms pulses of 488 nm and 561 nm every five minutes for a period of 12 hours.

Quantifying the percentage increase in the DNA recruitment experiments in E. coli

To quantitatively characterize the accumulation of CRY2-mCherry at chromosomal sites, we calculated the increase in the Weber contrast⁷⁷ of single cells. This calculation was used because chromosomal DNA spots were diffusive in both the lateral and azimuthal directions (relative to the z-axis of our objective) over the course of the experiments. This type of movement made it difficult to track individual spots and accurately estimate their intensities using the method we applied to our cell pole and midcell platforms (see above). Since the accumulation of CRY2-mCherry from a uniform distribution to a single concentrated location inside the cell body is equivalent to an increase in the Weber Contrast of a single cell image, we estimated the accumulation using the following method. First, we segmented the cells with observable spots in the last frame of the time lapse movie. There are normally 1 or 2 spots in a segmented cell region. Then we subtracted the background intensity of each image using ImageJ⁷⁸. Finally, the normalized contrast for a single cell, j, in frame i is defined as:

$$\:{C}_{norm}\left(i,j\right)=\:\frac{{I}_{\text{max}i,j}-{I}_{\text{min}i,j}}{⟨{I}_{i,j}⟩-{I}_{\text{min}i,j}}$$

Where $\:{C}_{norm}(i,j)$ reflects the extended accumulation of CRY2-mCherry in the DNA spot and $\:{I}_{\text{max}i,j}$ and $\:{I}_{\text{min}i,j}$ are the intensity of the brightest and darkest pixels in the cells, respectively. The contrast is normalized by the mean intensity, 〈I_i,j ⟩, of all the pixels in this region considering cell-to-cell variations in expression levels. Figure 2d and 2e were calculated using this method with the y-axis in Fig. 2e converted to percent increase ($\:\%\:Inc)$ by:

$$\:\%\:Inc\:\left(t\right)=\:\left(\frac{{C}_{norm}\left(t\right)}{{C}_{norm}(t=0)}-1\right)\times\:100$$

for ease of comparison with other experiments.

Quantifying the RFP and/or GFP signal in cell pole and Z-ring recruitment experiments

A variation on a previously published custom MATLAB script⁷⁶ used to quantify FRAP data was created to measure the GFP and RFP signals in the live-cell recruitment experiments. Briefly, cells were manually selected and cropped using a maximum intensity projection image of the entire recruitment stream (created in ImageJ) to avoid any selection bias. The max intensity image was also used to manually crop both the total cell area as well as the recruitment site area (either Z-ring or cell pole). For the Z-ring experiments the selected area was cropped manually using a rectangular selection. For the cell pole experiments the PopZ region was cropped using a fixed circle with a 2-pixel radius. To calculate the percent increase at target loci, we first calculated the fraction $\:Frac\left(t\right)\:$of the fluorescence intensity $\:{I}_{target}\left(t\right)\:$at the target site at a given time t$\:\:$versus that of the whole cell $\:{I}_{cell}\left(t\right)$:

$$\:Frac\left(t\right)=\frac{{I}_{target}\left(t\right)}{{I}_{cell}\left(t\right)}\:$$

We then converted this fraction to percent increase ($\:\%\:Inc)$ by:

$$\:\%\:Inc\:\left(t\right)=\:\left(\frac{Frac\left(t\right)}{Frac(t=0)}-1\right)\times\:100$$

The percentage increase curves of all cells were then averaged to obtain the mean and the associated standard error.

We noticed that our experimental setup introduced a slight variation in the time between activation with blue light and acquisition with either green or far-red light. This variation was because in some experiments (Fig. 2i and 3d), the CRY2/CIBN association kinetics were too fast that the microscope hardware speed of switching blue-light activation to green-light was not fast enough to capture the true time 0 after switching, and hence the first datapoint did not start at true time 0. To account for this delay, we created a custom MATLAB script to determine the true time delay between activation and acquisition frames using the imaging metadata obtained by Metamorph. We then used the actual time of switching when aggregating data and/or making measurements. Therefore, some of the time traces did not start at precisely time 0.

Quantifying the Halo signal in cell pole dissociation experiments in E. coli

Individual well-isolated cells were cropped from whole field of view time-course image stacks starting 5 minutes after CRY2/CIBN association was activated using blue light. The individual cell image stacks were then registered in x-y space using the ImageJ StackReg⁷⁹ plug-in to minimize the shifting of the cells over long timescales due to growth. The depletion of cell pole foci was then measured using the Weber contrast-based methodology employed for the TetR recruitment platform described above. The resulting traces for individual cells were averaged together and the exponential decay curve was fit to

$$\:f\left(t\right)=A{e}^{-\left(\frac{1}{{\tau\:}_{off}}\right)(t-5)}+C$$

to extract the dissociation time constant (𝜏_off).

Quantification of Z-ring decondensation in E. coli

Cells were segmented using the MiSiC deep learning-based cell segmentation algorithm⁸⁰. After segmentation, the resulting cell masks were fed into MicrobeJ⁸¹ to quantify the mean ZapA-mCherry intensity at across the long axis of the cell and cell lengths. All cells were visually inspected to remove any cells that lacked a Z-ring at the start of the experiment. The midcell localization fraction of the Z-ring was measured by dividing the integrated ZapA-mCherry fluorescence intensity within a 3-pixel window (480 nm width) about the midcell maximum (approximate Z-ring position) by the total integrated ZapA-mCherry fluorescence intensity of the whole cell both before and after a 5-minute blue light induction. The change in the midcell localization fraction before and after blue light induction was calculated, normalized by the midcell fraction before induction and multiplied by a factor of 100 to obtain the percent change in Z-ring intensity at midcell. The percentage of cells that both harbored a Z-ring prior to induction with blue light and showed a reduction in the midcell fraction of ZapA-mCherry intensity at midcell greater than 4% were deemed as “decondensed”. This decondensation threshold was applied to every cell in each experimental population to determine the fraction of cell that underwent significant decondensation in each experimental condition. Bootstrapping was employed to estimate the standard error of the mean. Briefly, a subpopulation of cells (500 for the induced cells and 300 for the uninduced cells) were randomly sampled and used to calculate the percentage of cells that underwent decondensation. This calculation was repeated 100 times to obtain the standard error of the mean.

Quantification of inhibition of cytokinesis in E. coli

Cells were manually tracked throughout the duration of the experiment to determine if and when a successful division occurred. Cells were cropped and the average midcell intensity across the long-axis of the cells (long-axis projection) was calculated using a custom MATLAB script.

Quantification of maximal enrichment and reduction of CRY2-mCherry and CRY2-HaloTag

In order to reduce the effects of noise on the maximal fold enrichment and maximal fold reduction of CRY2-HaloTag signal at the cell poles (i.e., estimation of the plateau value), the final plateau value was calculated by taking the mean of the last ten datapoints of individual cell trajectories.

Statistics

All experimental replicates (N) were noted in the main text along with the number of data points used in the analysis where applicable. All error bars and significance values were calculated using MATLAB by a statistical test specified in the figure legend. p-values < 0.05 were accepted as statistically significant. The different significance levels indicated as stars in figures correspond to *p value < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001 or n.s. where no significant difference was observed.

MATERIALS AVAILIBILITY

All plasmids and strains used in this work are readily available from the corresponding author.

DATA AVAILIBILITY

The datasets generated during and/or analyzed during the current study are available from the corresponding author.

CODE AVAILIBILITY

The code used to analyze all data in the current study is available from the corresponding author.

COMPETING FINANCIAL INTERESTS

The authors declare no competing financial interests.

AUTHOR CONTRIBUTIONS

R.M. constructed and characterized all the E. coli strains and fusions and proposed and carried out all E. coli experiments as well as quantified and interpreted all E. coli data unless otherwise noted. C.H.B. assisted in developing methodologies for fold enrichment calculations with R.M. and J.X and assisted in analysis of CRY2/CIBN green light effects. X.Y. developed the Normalized Contrast method used to analyze the DNA localization data and performed the data analysis using this method. J.W.M., together with R.M., designed strains for the light induced inhibition of cytokinesis assay. K.E.B. and A.J.P. performed experiments in S. pneumoniae while H.C.T.T. and Y.M.H. constructed strains with oversight by M.E.W. on experimental design. E.L.S. constructed strains in C. crescentus while E.L.S. and A.J.P. performed experiments with oversight by E.G. on experimental design. S.C. constructed B. subtilis strains while A.J.P. performed experiments with oversight by K.S.R. on experimental design. A.J.P. and R.M. performed data analysis on data acquired from B. subtilis or C. crescentus experiments. J.X. planned and directed the project and, together with R.M. and A.J.P., designed experiments, interpreted the data and wrote the manuscript.

ACKNOWLEDGMENTS

We would like to thank T. Bernhardt and J. Buss for the BW25113 (zapA-mCherry) strain. Additionally, we would like to thank N. Yehya for helpful insight and discussions. This work was supported by NIH R01 GM086447, NIH R35 GM136436 (to J.X.), NIH T32 GM008403 (to R.M.), and NIH F32 GM150262 (to A.J.P.), R35 GM131767 to (M.E.W.), R35 GM136221 (to E.G.), T32 GM144272 (to E.L.S.). K.S.R. was funded by the Intramural Research Program of the NIH, the National Cancer Institute, the Center for Cancer Research.

Surovtsev IV, Jacobs-Wagner C (2018) Subcellular Organization: A Critical Feature of Bacterial Cell Replication. Cell 172:1271–1293
Thanbichler M, Shapiro L (2008) Getting organized — how bacterial cells move proteins and DNA. Nat Rev Microbiol 6:28–40
Kolar K, Knobloch C, Stork H, Žnidarič M, Weber W (2018) OptoBase: A Web Platform for Molecular Optogenetics. Acs Synth Biol 7:1825–1828
Inobe T, Nukina N (2016) Rapamycin-induced oligomer formation system of FRB–FKBP fusion proteins. J Biosci Bioeng 122:40–46
Farrar MA, Olson SH, Perlmutter RM (2000) [31] Coumermycin-induced dimerization of GyrB-containing fusion proteins. Methods Enzymol 327:421–IN5
Miyamoto T et al (2012) Rapid and orthogonal logic gating with a gibberellin-induced dimerization system. Nat Chem Biol 8:465–470
Davis JH, Baker TA, Sauer RT (2011) Small-molecule control of protein degradation using split adaptors. Acs Chem Biol 6:1205–1213
Voß S, Klewer L, Wu Y-W (2015) Chemically induced dimerization: reversible and spatiotemporal control of protein function in cells. Curr Opin Chem Biol 28:194–201
Losi A, Gardner KH, Möglich A (2018) Blue-Light Receptors for Optogenetics. Chem Rev 118:10659–10709
Goglia AG, Toettcher JE (2018) A bright future: optogenetics to dissect the spatiotemporal control of cell behavior. Curr Opin Chem Biol 48:106–113
Bugaj LJ, Choksi AT, Mesuda CK, Kane RS, Schaffer DV (2013) Optogenetic protein clustering and signaling activation in mammalian cells. Nat Methods 10:249–252
Taslimi A et al (2014) An optimized optogenetic clustering tool for probing protein interaction and function. Nat Commun 5:4925
Kawano F, Suzuki H, Furuya A, Sato M (2015) Engineered pairs of distinct photoswitches for optogenetic control of cellular proteins. Nat Commun 6:6256
Guntas G et al (2015) Engineering an improved light-induced dimer (iLID) for controlling the localization and activity of signaling proteins. Proc Natl Acad Sci 112:112–117
DeFelice MM et al (2019) NF-κB signaling dynamics is controlled by a dose-sensing autoregulatory loop. Sci Signal 12:eaau3568
Benedetti L et al (2018) Light-activated protein interaction with high spatial subcellular confinement. Proc Natl Acad Sci 115:201713845
Hallett RA, Zimmerman SP, Yumerefendi H, Bear JE, Kuhlman B (2016) Correlating in Vitro and in Vivo Activities of Light-Inducible Dimers: A Cellular Optogenetics Guide. Acs Synth Biol 5:53–64
Xiao J, Dufrêne YF (2016) Optical and force nanoscopy in microbiology. Nat Microbiol 1:16186
Kennedy MJ et al (2010) Rapid blue-light-mediated induction of protein interactions in living cells. Nat Methods 7:973–975
Shao K et al (2020) The oligomeric structures of plant cryptochromes. Nat Struct Mol Biol 27:480–488
Sancar A (2003) Structure and Function of DNA Photolyase and Cryptochrome Blue-Light Photoreceptors. Chem Rev 103:2203–2238
Liu H et al (2008) Photoexcited CRY2 Interacts with CIB1 to Regulate Transcription and Floral Initiation in Arabidopsis. Science 322:1535–1539
Duan L et al (2017) Understanding CRY2 interactions for optical control of intracellular signaling. Nat Commun 8:547
Pu J, Zinkus-Boltz J, Dickinson BC (2017) Evolution of a split RNA polymerase as a versatile biosensor platform. Nat Chem Biol 13:432–438
Yamada M, Suzuki Y, Nagasaki SC, Okuno H, Imayoshi I (2018) Light Control of the Tet Gene Expression System in Mammalian Cells. Cell Rep 25:487–500e6
Castillo-Hair SM, Baerman EA, Fujita M, Igoshin OA, Tabor JJ (2019) Optogenetic control of Bacillus subtilis gene expression. Nat Commun 10:3099
Jayaraman P et al (2016) Blue light-mediated transcriptional activation and repression of gene expression in bacteria. Nucleic Acids Res 44:6994–7005
Olson EJ, Hartsough LA, Landry BP, Shroff R, Tabor JJ (2014) Characterizing bacterial gene circuit dynamics with optically programmed gene expression signals. Nat Methods 11:449–455
Hensel Z, Weng X, Lagda AC, Xiao J (2013) Transcription-Factor-Mediated DNA Looping Probed by High-Resolution, Single-Molecule Imaging in Live E. coli Cells. Plos Biol 11:e1001591
Lau IF et al (2003) Spatial and temporal organization of replicating Escherichia coli chromosomes. Mol Microbiol 49:731–743
Ramos JL et al (2005) The TetR Family of Transcriptional Repressors. Microbiol Mol Biol R 69:326–356
Lasker K et al (2020) Selective sequestration of signalling proteins in a membraneless organelle reinforces the spatial regulation of asymmetry in Caulobacter crescentus. Nat Microbiol 1–12. 10.1038/s41564-019-0647-7
Molinari S et al (2019) A synthetic system for asymmetric cell division in Escherichia coli. Nat Chem Biol 15:917–924
Mushnikov NV, Fomicheva A, Gomelsky M, Bowman GR (2019) Inducible asymmetric cell division and cell differentiation in a bacterium. Nat Chem Biol 15:925–931
Lim HC, Bernhardt TG (2019) A PopZ-linked apical recruitment assay for studying protein–protein interactions in the bacterial cell envelope. Mol Microbiol 112:1757–1768
Laloux G, Jacobs-Wagner C (2013) Spatiotemporal control of PopZ localization through cell cycle–coupled multimerization. J Cell Biology 201:827–841
Bowman GR et al (2008) A Polymeric Protein Anchors the Chromosomal Origin/ParB Complex at a Bacterial Cell Pole. Cell 134:945–955
Ebersbach G, Briegel A, Jensen GJ, Jacobs-Wagner C (2008) A Self-Associating Protein Critical for Chromosome Attachment, Division, and Polar Organization in Caulobacter. Cell 134:956–968
McQuillen R, Xiao J (2020) Insights into the Structure, Function, and Dynamics of the Bacterial Cytokinetic FtsZ-Ring. Annu Rev Biophys 49:1–33
Macnab RM (2003) How Bacteria Assemble Flagella. Annu Rev Microbiol 57:77–100
Sourjik V (2004) Receptor clustering and signal processing in E. coli chemotaxis. Trends Microbiol 12:569–576
Katsura Y et al (2015) An optogenetic system for interrogating the temporal dynamics of Akt. Sci Rep-uk 5:14589
Mühlhäuser WWD, Hörner M, Weber W, Radziwill G (2017) Light-Regulated Protein Kinases Based on the CRY2-CIB1 System. Methods Mol Biology Clifton N J 1596:257–270
Szeto TH, Rowland SL, Habrukowich CL, King GF (2003) The MinD membrane targeting sequence is a transplantable lipid-binding helix. J Biol Chem 278:40050–40056
Che DL, Duan L, Zhang K, Cui B (2015) The Dual Characteristics of Light-Induced Cryptochrome 2, Homo-oligomerization and Heterodimerization, for Optogenetic Manipulation in Mammalian Cells. Acs Synth Biol 4:1124–1135
McQuillen RJ (2022) Optogenetic Control of Protein Organization in Living Escherichia coli Cells. Johns Hopkins University
Goehring NW, Beckwith J (2005) Diverse paths to midcell: assembly of the bacterial cell division machinery. Curr Biol 15:R514–R526
Lutkenhaus J, Du SE (2013) coli Cell Cycle Machinery. Prokaryotic Cytoskeletons, vol 84. Springer, Cham, pp 27–65
Yang X et al (2019) FtsW exhibits distinct processive movements driven by either septal cell wall synthesis or FtsZ treadmilling in E. coli. Biorxiv 850073. 10.1101/850073
McCausland JW et al (2019) Treadmilling FtsZ polymers drive the directional movement of sPG-synthesis enzymes via a Brownian ratchet mechanism. Biorxiv 857813. 10.1101/857813
McQuillen R, Xiao J (2020) Insights into the Structure, Function, and Dynamics of the Bacterial Cytokinetic FtsZ-Ring. Annu Rev Biophys 49:309–341
Goehring NW, Gueiros-Filho F, Beckwith J (2005) Premature targeting of a cell division protein to midcell allows dissection of divisome assembly in Escherichia coli. Gene Dev 19:127–137
Galli E, Gerdes K (2012) FtsZ-ZapA-ZapB Interactome of Escherichia coli. J Bacteriol 194:292–302
Buss J et al (2013) In vivo organization of the FtsZ-ring by ZapA and ZapB revealed by quantitative super‐resolution microscopy. Mol Microbiol 89:1099–1120
Fu G et al (2010) In Vivo Structure of the E. coli FtsZ-ring Revealed by Photoactivated Localization Microscopy (PALM). PLoS ONE 5:e12680
Buss J et al (2015) A Multi-layered Protein Network Stabilizes the Escherichia coli FtsZ-ring and Modulates Constriction Dynamics. Plos Genet 11:e1005128
Rowlett VW, Margolin W (2013) The bacterial Min system. Curr Biol 23:R553–R556
Hu Z, Lutkenhaus J (2000) Analysis of MinC Reveals Two Independent Domains Involved in Interaction with MinD and FtsZ. J Bacteriol 182:3965–3971
Justice SS, García-Lara J, Rothfield LI (2000) Cell division inhibitors SulA and MinC/MinD block septum formation at different steps in the assembly of the Escherichia coli division machinery. Mol Microbiol 37:410–423
Romberg L, Levin PA (2003) Assembly Dynamics of the Bacterial Cell Division Protein FtsZ: Poised at the Edge of Stability. Annu Rev Microbiol 57:125–154
Coltharp C, Buss J, Plumer TM, Xiao J (2016) Defining the rate-limiting processes of bacterial cytokinesis. Proc Natl Acad Sci 113:E1044–E1053
Lan G, Daniels BR, Dobrowsky TM, Wirtz D, Sun S (2009) X. Condensation of FtsZ filaments can drive bacterial cell division. Proc Natl Acad Sci 106:121–126
Liu B, Liu H, Zhong D, Lin C (2010) Searching for a photocycle of the cryptochrome photoreceptors. Curr Opin Plant Biol 13:578–586
Bouly J-P et al (2007) Cryptochrome Blue Light Photoreceptors Are Activated through Interconversion of Flavin Redox States. J Biol Chem 282:9383–9391
Banerjee R et al (2007) The Signaling State of Arabidopsis Cryptochrome 2 Contains Flavin Semiquinone. J Biol Chem 282:14916–14922
Los GV et al (2008) HaloTag: A Novel Protein Labeling Technology for Cell Imaging and Protein Analysis. Acs Chem Biol 3:373–382
Grimm JB, Brown TA, English BP, Lionnet T, Lavis LD (2017) Synthesis of Janelia Fluor HaloTag and SNAP-Tag Ligands and Their Use in Cellular Imaging Experiments. 1663:179–188
Gueiros-Filho FJ, Losick R (2002) A widely conserved bacterial cell division protein that promotes assembly of the tubulin-like protein FtsZ. Gene Dev 16:2544–2556
Perez AJ et al (2021) FtsZ-Ring Regulation and Cell Division Are Mediated by Essential EzrA and Accessory Proteins ZapA and ZapJ in Streptococcus pneumoniae. Front Microbiol 12:780864
Lindner F, Diepold A (2021) Optogenetics in bacteria – applications and opportunities. Fems Microbiol Rev 46:fuab055
Repina NA, Rosenbloom A, Mukherjee A, Schaffer DV, Kane RS (2017) At Light Speed: Advances in Optogenetic Systems for Regulating Cell Signaling and Behavior. Annu Rev Chem Biomol 8:13–39
Tischer D, Weiner OD (2014) Illuminating cell signalling with optogenetic tools. Nat Rev Mol Cell Bio 15:551–558
Boekel J et al (2015) A split-Cas9 architecture for inducible genome editing and transcription modulation. Nat Biotechnol 33:139–142
Sterlini JM, Mandelstam J (1969) Commitment to sporulation in Bacillus subtilis and its relationship to development of actinomycin resistance. Biochem J 113:29–37
Kim EY, Tyndall ER, Huang KC, Tian F, Ramamurthi KS (2017) Dash-and-Recruit Mechanism Drives Membrane Curvature Recognition by the Small Bacterial Protein SpoVM. Cell Syst 5:518–526e3
Yang X et al (2017) GTPase activity–coupled treadmilling of the bacterial tubulin FtsZ organizes septal cell wall synthesis. Science 355:744–747
Pelli DG, Bex P (2013) Measuring contrast sensitivity. Vis Res 90:10–14
Schindelin J et al (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9:676–682
Thevenaz P, Ruttimann UE, Unser M (2021) A pyramid approach to subpixel registration based on intensity. Ieee T Image Process 7:27–41
Panigrahi S et al (2020) MiSiC, a general deep learning-based method for the high-throughput cell segmentation of complex bacterial communities. Biorxiv 10.07.328666 (2020) 10.1101/2020.10.07.328666
Ducret A, Quardokus EM, Brun YV (2016) MicrobeJ, a tool for high throughput bacterial cell detection and quantitative analysis. Nat Microbiol 1:16077

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Light-dependent modulation of protein localization and function in living bacteria cells

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Abstract

Figures

INTRODUCTION

RESULTS

Rapid, reversible, and light-dependent recruitment of cytoplasmic protein to chromosomal DNA

Rapid, light-dependent recruitment of cytoplasmic protein to cell pole

Formation of inner membrane-associated heteroclusters of CRY2/CIBN upon light activation

Rapid, light-dependent recruitment of cytoplasmic proteins to the midcell

Light-dependent destabilization of the Z-ring and inhibition of cell division

CRY2/CIBN complex dissociation and activated CRY2 concentration are sensitive to green light

Adapting CRY2/CIBN to other bacterial species

DISCUSSION

METHODS

Live cell imaging

Quantifying the RFP and/or GFP signal in cell pole and Z-ring recruitment experiments

Quantification of maximal enrichment and reduction of CRY2-mCherry and CRY2-HaloTag

Statistics

Declarations

MATERIALS AVAILIBILITY

DATA AVAILIBILITY

CODE AVAILIBILITY

COMPETING FINANCIAL INTERESTS

AUTHOR CONTRIBUTIONS

ACKNOWLEDGMENTS

References

Additional Declarations

Supplementary Files

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