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Small RNA-mediated Cry toxin silencing allows Bacillus thuringiensis to evade Caenorhabditis elegans avoidance behavioral defenses

Abstract Pathogen avoidance behavior protects animal hosts against microbial pathogens. Pathogens have evolved specific strategies during coevolution in response to such host defenses. However, these strategies for combatting host avoidance behavioral defenses remain poorly understood. Here, we used Caenorhabditis elegans and its bacterial pathogen Bacillus thuringiensis as a model and determined that small RNA (sRNA)-mediated Cry toxin silencing allowed pathogens to evade host avoidance behavioral defenses. The B. thuringiensis strain YBT-1518, which encodes three nematicidal cry genes, is highly toxic to C. elegans. However, the expression of the most potent toxin, Cry5Ba, was silenced in this strain when YBT-1518 was outside the host. Cry5Ba silencing was due to the sRNA BtsR1, which bound to the RBS site of the cry5Ba transcript via direct base pairing and inhibited Cry5Ba expression. Upon ingestion by C. elegans, Cry5Ba was expressed in vivo by strain YBT-1518. Cry5Ba silencing may allow B. thuringiensis to avoid nematode behavioral defenses and then express toxins once ingested to kill the host and gain a survival advantage. Our work describes a novel model of sRNA-mediated regulation to aid pathogens in combating host avoidance behavioral defenses.

INTRODUCTION
Pathogen avoidance behavior is a highly important phenomenon across all animal taxa and protects host organisms against microbial pathogens and danger (1)(2)(3). A host generally faces the following two choices when challenged by pathogens: combat the threatening organism by erecting a physical barrier or activating physiological or cellular defenses (4)(5)(6); or avoid the organism by moving away from pathogen-rich regions. This pathogen avoidance behavior minimizes the risk of infection and economizes host resources (1)(2)(3). Therefore, avoidance behavior is critical for the survival of all animals, even human beings, in their natural ecological niches and has affected the evolution of foraging strategies, habitat choice, social interactions and mate choice (3,7,8).
The molecular genetics underlying host avoidance behavior have been partially explored in the nematode Caenorhabditis elegans, which is a powerful model for dissecting genetic and biochemical mechanisms in animal hosts (1,3). When C. elegans is infected, a complex innate immune response is triggered, involving interconnected signaling cascades such as the MAPK (9) and TGF-ß pathways (10), among others. Furthermore, C. elegans has evolved behavioral defenses in response to harmful microbes. For example, C. elegans engages in host−protective avoidance behavior in response to pathogen derived molecules such as surfactants (11) and quorum−sensing signaling molecules (12)(13)(14)(15). Several distinct mechanisms are involved in pathogen avoidance in C. elegans, including G protein-like (16), insulin-like (17), and Toll-like receptor signaling (2), as well as neuronal serotonin- (15) and HECT domain-containing E3 ubiquitin ligase-based pathways (14).
Pathogens have evolved specific strategies in response to such host defenses during coevolution (1,18). However, the mechanisms by which pathogens combat host avoidance behavioral defenses remain poorly understood. Several studies have shown that pathogens deceive C. elegans via surface molecule modification to prevent detection (1) or produce certain molecules that attract worms (19).
Bacillus thuringiensis is a Gram-positive, spore-forming soil bacterium that produces parasporal crystal inclusions during the sporulation phase of its growth cycle (20). This bacterium has been used as a specific, safe, and effective bio−insecticide to control various pests and nematodes for >70 years (20). B. thuringiensis strain YBT−1518, which encodes three nematicidal cry genes, namely, cry55Aa, cry6Aa, and cry5Ba, is highly toxic to C. elegans. We pre-

DNA manipulation and transformation
DNA manipulation was performed according to the methods described by Sambrook and Russell (23). DNA from gels or PCR products was purified using DNA purification kit (QIAGEN, Shanghai). Primers (Supplementary  Table S2) were synthesized, and all constructs were confirmed by Sanger sequencing (AuGCT, Beijing). Escherichia coli transformation was performed utilizing a standard chemical-competent method (23), and B. thuringiensis was transformed by electroporation as described previously (24).

Microscopy observation
For general observation, after B. thuringiensis was grown in LB medium to complete sporulation, the crystals and spores were harvested for fuchsin staining and examined with an optical microscope (Olympus, USA) using an oil immersion lens with 10 × 100-fold magnification. For GFP reporter observation, B. thuringiensis was grown in LB medium and the cells were harvested at the vegetative stage and observed under a fluorescence microscope (Primostar-FL2) using an oil immersion lens with 10 × 100-fold magnification.

RNA extraction and transcription analyses
Total RNA was extracted from B. thuringiensis strains using TRIzol reagent (Invitrogen, California, USA). RNA samples were treated with DNase I (Roche, Basel, Switzerland), then precipitated with ethanol and re−dissolved in RNase-free water. The purified total RNA was quantified by NanoDrop ND-1000 spectrophotometer (Labtech, Wilmington, MA, USA). The qRT-PCR analysis of cry5Ba and BtsR1 was carried out using two-step methods with the primers listed in Supplementary Table S2. 2 g total RNA from each sample was used for cDNA synthesis with Superscript II reverse transcriptase kit (Invitrogen, Carlsbad, CA, USA). qRT-PCR was then conducted with Power SYBR Green PCR Master Mix in an Applied Biosystems 7300 Real Time PCR System (Applied Biosystems, Foster City, CA, USA). 16S rRNA was used as an internal control. At least three independent biological samples were tested and fold-changes between samples were plotted. The Northern blot analysis of cry5Ba and BtsR1 was performed as described by Sambrook and Russell (23) with the probes listed in Supplementary Table S2. The 5 -end FAM-labeled BtsR1 and cry5Ba RNA probes were synthesized (AuGCT, Beijing, China). Total RNA per lane (5 g) was separated on a 1.2% agarose gel, and then transferred to a positively charged nylon membrane and UV-crosslinked. The membrane was blocked and washed, and the labeled RNA was detected with a Typhoon Scanner system (GE, Fairfield, CT, USA). The 16S rRNA was used as a reference. At least three independent biological samples were tested. Data from Northern blot analysis were quantified with ImageJ software (available via http://rsbweb.nih.gov/ij/).

Crystal protein purification and detection
Spore and crystal mixtures were collected and washed as described previously (25). The spore crystal pellets were then resuspended in solubilization buffer (50 mM Na 2 CO 3 , 25 mM dithiothreitol, pH 10.5) and incubated at 37 • C for 2 h. The insoluble material that remained was removed by centrifugation, and the solubilized proteins from the supernatant were analyzed by 10% SDS-PAGE. Cry5Ba expression was detected by Western blot using a Cry5Ba antibody as previously described (26).

Prediction and structural analysis of small RNA BtsR1
We constructed a database that contains all the YBT-1518 intergenic region sequences (IGRs) based on YBT-1518 genome annotation. Among these sequences, the bacterial promoter regions were predicted by BPROM (http://linux1. softberry.com/) and Neural Network Promoter Prediction program (http://www.fruitfly.org/seq tools/promoter.html); while, the Rho-independent transcription terminators were identified using the TransTermHP program (27). The IGRs that contained potential promoter or terminator were retained as potential sRNAs. We then scanned the retained IGRs database using the coding region and promoter sequences of cry5Ba by a local BLASTN program. Finally, a putative sRNA, BtsR1, was predicted that could bind with strong complementarity to the cry5Ba mRNA. The transcription start site of BtsR1 were ascertained by 5 RACE technique using SMARTer® RACE 5 /3 kit (Clontech, CA, USA) with primer in Supplementary Table S2.

Site-directed mutagenesis of BtsR1
Bases were individually mutated according to the secondary structure of BtsR1, which was constructed online (http: //rna.tbi.univie.ac.at/). Each mutated base was encoded on a primer (as shown in Supplementary Table S2), and we applied reverse PCR to obtain a fragment containing the mutated base, then linked the mutated base fragments in parallel with Cry5Ba.

Detection of beta-galactosidase activity
Different B. thuringiensis cultures (2 ml) in LB medium were collected at the indicated time points. The ␤-galactosidasespecific activities were measured according to the previously described method (28). The recombinant strains were cultivated in LB medium at 37 • C, and the cultured cells were collected and suspended in 665 l of Z-buffer and betamercaptoethanol, followed by the addition of 55 l of chloroform and 55 l of SDS. The mixture was incubated in a 30 • C water bath for 5 min and activated with 160 l of ONPG at 30 • C for 90-120 min. The reaction was stopped by adding 400 l of a 1 M Na 2 CO 3 solution; and the action times were recorded. We detected the A 420 absorbance values of these reagents by ELISA. Finally, A 420 was converted into Miller units. The assays were conducted in triplicate, and the results were shown as the mean ± SE.

The RNA:RNA EMSA assay
In vitro-transcribed RNAs of pcry5Ba (200 nt, −100 to +100 regions of cry5Ba mRNA including the ribosome binding site, RBS) and pcry5BaM (187 nt, −100 to +100 regions of cry5Ba mRNA without the predicted 13 base pairs that interact with BtsR1) were created. Full-length BtsR1 RNA was synthesized with a FAM label at the 5 end (AuGCT, Beijing). RNAs were assessed with an RNA6000 NANO chip on the Agilent Bioanalyzer 2100 system. FAM-BtsR1 (10 nM) was incubated in the presence or absence of pcry5Ba RNA (0, 100, 500, 1000, 1500, 2000, 2500 and 5000 nM) or pcry5BaM RNA (0, 100, 500, 1000, 1500, 2000, 2500 and 5000 nM) for the EMSA reactions, which were performed as previously described (29). Samples were electrophoresed through a 5% Tris/Borate/EDTA (TBE) gel at 200 V for 25 min, and then transferred to a positively charged nylon membrane and UV-crosslinked. The membrane was blocked and washed, and the labeled RNA was detected with a Typhoon Scanner system (GE, Healthcare).

Competitive RNA:RNA EMSA
The RNAs of pcry5Ba and pcry5BaM as well as seven selected BtsR1 mutants were transcribed in vitro. FAM-BtsR1 (10 nM) was incubated in the presence or absence of pcry5Ba RNA (0 to 1000 nM) to conduct the competitive EMSA reactions. Then, the seven selected, unlabeled BtsR1 mutants were added into each reaction system at a concentration of 5000 nM to compete with the FAM−BtsR1 interaction with pcry5Ba RNA. Reactions were performed as previously described (Liu et al., 2012). Samples were electrophoresed through a 5% TBE mini-gel at 200 V for 25 min, then transferred to a positively-charged nylon membrane and UV-crosslinked. The membrane was blocked and washed, and the labeled RNA was detected using a Typhoon Scanner system (GE Healthcare).

Construction of the cry5Ba deletion mutant strain
DNA fragments corresponding to the regions upstream and downstream of the cry5Ba gene in YBT-1518 chromosomal were generated by PCR using the primers listed in Supplementary Table S2. The two amplified fragments were digested with BamHI−XbaI and BamHI−KpnI, respectively, and cloned into the temperature-sensitive plasmid pHT304-Ts. Then, the spectinomycin resistance gene from plasmid pBMB2062 was digested with BamHI and inserted between the above two fragments. The resulting plasmid was transformed into strain YBT-1518 and cultivated in LB medium (containing 25 g/ml erythromycin) for 8 h. Then, the transformants were cultivated at 42 • C for 4 days to eliminate the un-integrated plasmid. Erythromycin-resistant (25 g/ml) but spectinomycin sensitive (20 g/ml) colonies were harvested. The correct mutant strain was confirmed by PCR and sequencing (AuGCT, Beijing) and named as YBT-1518( cry5Ba).

Construction of the BtsR1 depletion strain
Endogenous BtsR1 was depleted by overexpressing the 5 -UTR of cry5Ba mRNA including BtsR1 target region. DNA fragments corresponding to the kanamycin promoter in plasmid pHT315-papha3 gfp, the 5 -UTR of cry5Ba gene (−100 to +100), and the terminator of the cry5Ba mRNA were generated by PCR and then fused together by SOE-PCR using the primers listed in Supplementary Table S2. The fusion gene was confirmed by sequencing (AuGCT, Beijing), then digested with KpnI−BamHI and cloned into pHT304 vector. The resulting plasmid was transformed into strain YBT-1518 to generate the BtsR1 depletion strain YBT-1518(BtsR1-dep).

Nematode bioassays
B. thuringiensis strains against C. elegans were tested using bioassays as previously described (30).

Nematode pumping rate assays
Pumping rate assays were conducted as previously described (31) to quantitatively assess the feeding rates of individual animals. L4 animals (20)(21)(22)(23)(24)(25)(26)(27)(28)(29)(30) were selected and transferred to mutant or wild-type B. thuringiensis plate. The times of pharynx movement (pumps) in 30 s of each animal was observed and recorded using a dissecting microscope after the indicated times. This assay was incubated at room temperature with three independent repeats.

Nematode feeding assays
L4 animals (20)(21)(22)(23)(24)(25)(26)(27)(28)(29)(30) were transferred to a plate containing mutant or wild-type B. thuringiensis cultures to determine fractions of animals in a population that are feeding. Each animal was observed by using a dissecting microscope for a maximum of 10 s, and scored as 'feeding' if rhythmical backward movements of the grinder were observed. Otherwise, the animals were recorded as not feeding. This assay was incubated at room temperature and observed after the indicated times. Three independent repeats were performed for each assay.

Nematode chemotaxis assays
Chemotaxis assays were conducted based on standard chemotaxis assay protocols (13,32) with some modification. Briefly, 25 l of control B. thuringiensis suspensions (OD 600 = 1.0) were placed 1 cm away from the edge of a 9-cm Petri dish filled with NG medium (22). The same amount of a test B. thuringiensis suspension was placed on the opposing side and acted as the counter attractant. Approximately 50−200 J4 stage animals were placed at the center of the plate. All nematodes were previously fed on E. coli OP50. Plates were then sealed with parafilm and stored at room temperature in the dark. Animals could move freely for 1-2 h before they were immobilized with 1 ml of 10 mM sodium azide applied to the spots of bacteria. In most cases, animals quickly entered one lawn and remained there for the duration of the assay. The number of nematodes at each bacterial spot was recorded. A chemotaxis index was used to score the nematode responses and was calculated as follows: (number of nematodes on the test bacteria -number of nematodes on control bacteria)/total number of nematodes counted. This calculation provided a chemotaxis score ranging from −1.0 (complete repulsion from test bacteria) to 1.0 (complete attraction to test bacteria). A score of ∼0 indicates equal numbers of nematodes on all bacterial spots. Five plates were assessed per replicate, and the procedure was repeated at least thrice.

Growth kinetics of B. thuringiensis in C. elegans
Doses of approximately 10 3 vegetative cells of different B. thuringiensis were fed to L4 worms in 48-well plates. Each treatment of 200 l/well consisted of 160 l of S-Medium, 20 l of L4 N2 worms (∼30−50), and 20 l of the B. thuringiensis cultures. Then, the plates were cultured at 20 • C for 24−48 h. Worms were collected, washed (with sterile water three times and M9 buffer three times), and centrifuged at 100 × g. Finally, 20 worms from each group were homogenized for 5−10 s to break up the tissues, then the tissues were diluted and spread on plates to count the numbers of B. thuringiensis.

Data analysis
Data analyses were performed using SPSS (Statistical Package for the Social Sciences) version 13.0 (SPSS, Chicago, IL, USA), and the mean ± SE are shown. Student's t-test were performed for statistical comparisons between two values, and differences were significant at the following thresholds: *P < 0.05; **P < 0.01; ***P< 0.001. Panels a, b, c contains crystals produced by strains YBT-1518/pHT304, BMB171/pHT304-cry5Ba, and YBT-1518/pHT304-cry5Ba, respectively. The red arrows indicate bipyramidal-shaped crystals, black arrows indicate rice-shaped crystals, and blue arrows indicate spores. All crystals are observed under an optical microscope and shown at the same magnification (10 × 100-fold).

Nematicidal Cry toxin Cry5Ba was silenced in B. thuringiensis strain YBT-1518
In our previous study, we observed Cry5Ba toxin silencing in the strain YBT-1518 (21). We confirmed this observation by evaluating the expression of Cry5Ba in YBT-1518 using SDS-PAGE, Western blot analysis, and microscopy. The expression of Cry5Ba in strain BMB171, which is a crystalliferous mutant of B. thuringiensis subsp. Kurstaki that does not encode cry genes and is nontoxic to nematodes, was used as a control (21). According to the SDS-PAGE results, after growth at 36 h, YBT-1518 produced two major proteins corresponding to the Cry55Aa and Cry6Aa proteins. However, the 140-kDa band for Cry5Ba protein was not detected ( Figure 1A). Cry5Ba was also undetectable by Western blot using a Cry5Ba specific antibody ( Figure 1B). The expression of Cry5Ba proteins in strain BMB171 results in the formation of bipyramidal-shaped crystals ( Figure 1C, panel b). However, microscopic observation only indicated the presence of rice-shaped crystals in YBT-1518 ( Figure  1C, panel a), which were formed by Cry6Aa and Cry55Aa (21). These results confirm that the Cry5Ba is silenced in strain YBT-1518.
Cryptic or silent cry genes often lack functional promoters (33) Figure S2) YBT-1518. We further demonstrated that Cry5Ba silencing is not caused by competition from Cry55Aa and Cry6Aa (Supplementary Figure S3), and does not relate to regulation by flanking sequences (Supplementary Figure S4). Detailed information is described in the Supplementary Information. In this case, the cryptic behavior of Cry5Ba in YBT-1518 appears to be attributable to a mechanism that differs from those mentioned above. When an additional copy of the cry5Ba gene was introduced into YBT-1518 via a low copy vector pHT304 under the control of its own promoter, Cry5Ba protein band was detected by SDS-PAGE ( Figure 1A) and western blot ( Figure 1B). Bipyramidal-shaped and rice-shaped crystals were also observed in the recombinant strain ( Figure 1C, panel c). Thus, the cryptic nature of Cry5Ba expression in YBT-1518 was dependent on the copy number of the cry5Ba gene.

Cry5Ba silencing in YBT-1518 occurred at the posttranscriptional level
To further understand the mechanisms underlying this silencing, we asked if the regulation occurred at the transcriptional level or a later step. Transcription profiles of cry5Ba mRNA in strains YBT-1518/pHT304, YBT-1518/pHT304-cry5Ba and BMB171/pHT304-cry5Ba at different growth phases were assessed by RT-PCR (Supplementary Figure S5) and Northern blot ( Figure  2A). The results showed that cry5Ba was highly transcribed from stationary phase to late stationary phase (12−36 h) in strains YBT-1518/pHT304-cry5Ba and BMB171/pHT304-cry5Ba. However, cry5Ba transcription in YBT-1518 reached the highest levels at 12 h, and then decreased sharply during stationary phase through late stationary phase (16−24 h). These results indicate that cry5Ba transcripts may be destabilized during later growth stages in YBT-1518. To confirm this observation, we perform an mRNA stability assay using rifampicin with the above three strains. The strains were cultured for 16 h, then 100 g/ml rifampicin was added to stop new transcription. The mRNA of cry5Ba was detected by northern blots of total RNA isolated at different times after addition of rifampicin ( Figure 2B). The results showed that the half-lives of the cry5Ba mRNA in strains YBT-1518/pHT304-cry5Ba and BMB171/pHT304-cry5Ba (with half-life of 5.3 and 5.8 min, respectively) was similar, but was significantly shorter (half-life of 1.0 min) in the strain YBT-1518/pHT304 ( Figure 2C).
The expression of Cry5Ba protein in the above strains was also evaluated using Western blot ( Figure 2D). Cry5Ba was undetectable at all growth phases in the wild type strain YBT-1518. By contrast, in YBT-1518/pHT304-cry5Ba and BMB171/pHT304-cry5Ba, Cry5Ba was detectable from 12−36 h in very large amounts, which is consistent with the cry5Ba transcription profiles in these strains. Thus, the silencing of Cry5Ba expression occurs at the posttranscriptional level in YBT-1518. The 16S rRNA was used as a reference gene. The relative quantification of the northern blots is presented in (C), with the mRNA level at 0 min was set to 100% for each strain. (D) Expression profile analysis of Cry5Ba proteins by Western blot. The total protein samples were prepared at different growth phases from cultured B. thuringiensis strains. Cry5Ba expression was detected with Cry5Ba antibody.

Cry5Ba was negatively regulated by a small RNA BtsR1
sRNAs negatively regulate the expression of target genes at the post-transcriptional level (35)(36)(37). Therefore, we hypothesized that the expression of Cry5Ba may be regulated by specific sRNAs in YBT-1518. Using the YBT-1518 genome sequence (38), we identified a putative sRNA (designated as BtsR1) that could bind with strong complementarity to the cry5Ba mRNA. BtsR1 is located on the antisense strand between the lysR (encoding a LysR family transcriptional regulator) and merR (encoding a MerR family transcriptional regulator) genes. The size of the BtsR1 was predicted to be 51 nt from the transcriptome data of YBT-1518. The transcription start site (+1) of BtsR1 was determined by 5 -RACE ( Figure 3A). The targetRNA program (39) gave a high score to a pairing between the BtsR1 intergenic region and the 5 -UTR of cry5Ba mRNA. A 13- bp sequence in BtsR1 is complementary to the 5 -UTR of the cry5Ba mRNA, including the 6-bp RBS ( Figure 3B). BtsR1 folds into a stable structure ( Figure 3C) with a Gibbs free energy of −14.72 kcal/mol as predicted by the Lo-cARNA tool (40). The putative region of complementarity to the RBS of cry5Ba mRNA was in a single-stranded region, which is consistent with the interaction site features reported in the sRNA (41,42). The conservation of the BtsR1 sequence was analyzed using BLASTn against the Bacillus genomes deposited in the non-redundant (NR) database. A total of 88 genomes (28 B. thuringiensis, 16 B. cereus, and 44 B. anthracis) contained a BtsR1-like sequence with 100% coverage and 98−100% identity. Within these genomes, the BtsR1 core sequences are highly conserved, except for positions +4, +19, +22 and +47 (Supplementary Figure S6). Interestingly, the predicted BtsR1−cry5Ba interaction is only observed in B. thuringiensis strain YBT-1518, because among the above 88 genomes, only YBT-1518 encodes cry5Ba.
We performed a northern blot to determine if the BtsR1 sRNA is produced in vivo. Using a BtsR1 specific probe, we detected a transcript consistent with the predicted length (51 nt) of BtsR1 and it showed the same size with that of the synthesized FAM labeled 51 nt BtsR1 ( Figure 3D). This result confirmed that BtsR1 is a true sRNA. To check the transcription profile of BtsR1, RT-PCR (Supplementary Figure S7) and northern blot ( Figure 3E) were carried out using total RNA isolated from different growth phases of YBT-1518. The results showed that BtsR1 was transcribed from exponential phase to late stationary phase (9-36 h) of growth. However, BtsR1 is not stably transcribed at 9 and 36 h.
To confirm whether BtsR1 inhibited the expression of Cry5Ba, we attempted but failed to construct a BtsR1 deletion mutant strain. Therefore, we constructed a BtsR1 depletion strain by overexpressing the 5 -UTR of cry5Ba mRNA including its putative BtsR1 binding site but without the Cry5Ba ORF. BtsR1 levels in the BtsR1 depletion strain are 85−94% lower than wild type ( Figure 4A), demonstrating the 5 -UTR of cry5Ba can be used to deplete BtsR1. The cry5Ba mRNA is more stable when the endogenous BtsR1 is depleted (half-life of 4.8 min), which is significantly longer than that of strain YBT-1518/pHT304 (1.3 min) ( Figure 4B). The results of SDS-PAGE and western blot showed that Cry5Ba expressed well when the endogenous BtsR1 is depleted ( Figure 4C). We also overexpressed the BtsR1 in cis or in trans with Cry5Ba ( Figure  4D) to confirm whether BtsR1 regulates the expression of Cry5Ba in the BtsR1 absent strain BMB171 and the wildtype strain YBT-1518. The results showed that either when BtsR1 was expressed in cis or in trans, Cry5Ba translation in both BMB171 and YBT-1518 was suppressed, as determined by SDS-PAGE. Although a small amount of Cry5Ba expression was still detectable in BMB171, the expression was significantly suppressed compared to expression levels in the absence of BtsR1 ( Figure 4E). This finding was further validated by Western blot analysis ( Figure 4F). Thus, our data suggests that the sRNA BtsR1 is essential for the negative regulation of Cry5Ba expression in YBT-1518.

BtsR1 directly interacted with cry5Ba mRNA by base-pairing with the RBS
The majority of bacterial sRNAs exert their regulatory functions by base-pairing to short stretches within the 5 -UTR of target mRNAs (41,42) to prevent 30S ribosome entry and inhibit translation initiation (37). The predicted interaction region of BtsR1 in the cry5Ba mRNA spans a 13-nt 5 -UTR region including the RBS (Figure 3B), indi-cating that BtsR1 may inhibit the translation initiation of Cry5Ba by base-pairing with its RBS.
To confirm this hypothesis, we used GFP as reporter to test the BtsR1-mediated regulation of the cry5Ba mRNA. The cry5Ba 5 -UTR was cloned as a translational fusion with gfp into the vector pHT315, resulting in pHT315-pcry5Ba'gfp. BtsR1 with its own promoter was inserted into the vector pBMB2062. Then, both plasmids were transferred into strains YBT-1518 and BMB171, as shown in Figure 5A. B. thuringiensis carrying the plasmid pHT315-papha3 gfp (43), containing the gfp gene under the control of the constitutive kanamycin promoter, was used as a positive control. We verified GFP protein expression for these strains utilizing fluorescence microscopy. The positive control strains and recombinant strains YBT-1518/pHT315-pcry5Ba'gfp and BMB171/pHT315-pcry5Ba'gfp showed substantial fluorescence signals after 36 h, compared with the negative control strains which harbored plasmid pHT315. However, no significant fluorescence signals were observed during the same growth phase when both pHT315-pcry5Ba'gfp and pBMB2062-BtsR1 were present ( Figure 5B).
The effect of BtsR1 on cry5Ba expression was also tested using a lacZ fusion. The strains harboring pHT304-pcry5Ba'lacZ alone exhibited high beta-galactosidase activity from 12 h to 36 h of growth. The strains harboring pHT304-pcry5Ba'lacZ together with pBMB2062-BtsR1 demonstrated significantly lower beta-galactosidase activity at all tested growth phases ( Figure 5C) compared to the strains harboring pHT304-pcry5Ba'lacZ alone (P = 0.0021 in BMB171 background; P = 0.0024 in YBT-1518 background, t-test). This result indicated that BtsR1 suppressed the expression of lacZ under the control of cry5Ba pro-moter. Thus, we concluded that BtsR1 acts on the 5 -UTR of the cry5Ba mRNA.
To further confirm that the region in the cry5Ba mRNA targeted by BtsR1 is the RBS, we constructed a mutant cry5Ba 5 -UTR lacking the RBS (pcry5BaM) and detected direct interactions in an EMSA assay. The wildtype (pcry5Ba) and mutant cry5Ba 5 -UTR (pcry5BaM) were synthesized via in vitro transcription ( Figure 5D). The 5 -end FAM-labeled BtsR1 (BtsR1-FAM) and increasing concentrations of unlabeled pcry5Ba and pcry5BaM RNAs were hybridized and resolved by native PAGE. FAMlabeled full-length BtsR1 formed higher-molecular-weight complexes with the wild type cry5Ba 5 -UTR (pcry5Ba), whereas no shift was observed for the mutant cry5Ba 5 -UTR (pcry5BaM) ( Figure 5E). This result indicated that BtsR1 pairs within the cry5Ba 5 -UTR region at its RBS. Together, we concluded that BtsR1 interacts directly with the 5 -UTR of cry5Ba mRNA by base-pairing at its RBS to inhibit Cry5Ba translation initiation.

Mapping key base-pairing sites in BtsR1 to the RBS of cry5Ba mRNA
To confirm the key base-pairing sites of the BtsR1 interaction with the RBS of cry5Ba mRNA, we applied base scanning mutagenesis and substituted each of the 13 bases in BtsR1 that are predicted to have complementarity to the cry5Ba mRNA ( Figure 6A). Each of the BtsR1 variants was cloned into pBMB2062 and transferred into strain YBT-1518/pHT304-cry5Ba. SDS-PAGE was conducted to determine whether mutagenesis influenced the expression of Cry5Ba. Ten of the 13 mutants retained the ability to inhibit Cry5Ba expression; while the other three BtsR1 mutants (BtsR1M5, BtsR1M11 and BtsR1M13, corresponding to the sites of +5, +11 and +13, respectively) lost the ability to inhibit Cry5Ba expression ( Figure 6B).
Translational lacZ reporter fusions were constructed to confirm these results. Plasmids containing each of the 13 BtsR1 mutants were transferred into strain YBT-1518/pHT304-pcry5Ba'lacZ. The ␤-galactosidase activities of these recombinant strains were tested in the presence of each BtsR1 mutant. No BtsR1-dependent translational repression was evident in the strains encoding BtsR1M5, BtsR1M11, and BtsR1M13 mutants. However, the strains encoding of the other 10 BtsR1 mutants demonstrated significant inhibition of ␤-galactosidase activity ( Figure 6C). These results identify the bases in BtsR1 relevant for inhibition of Cry5Ba expression.
Competitive EMSA assays were performed to confirm the interactions between the seven selected BtsR1 mutants and cry5Ba mRNA. When present at a high concentration (5000 nM), four mutants retained the ability to bind to the cry5Ba mRNA, while the other three mutants lost the ability to bind to the cry5Ba mRNA. We assessed the ability of the mutants to outcompete the interactions between Cry5Ba mRNA and FAM-labeled BtsR1. The four BtsR1 mutants that retained the ability to inhibit Cry5Ba expression could compete and interrupt the BtsR1-Cry5Ba mRNA interaction, while the three mutants that lost the ability to inhibit Cry5Ba expression exhibited no competition ( Figure 6D). These results are consistent with our cry5Ba expression studies and the secondary structure prediction results. Thus, the single-stranded-exposed bases of BtsR1 are important for interaction with cry5Ba mRNA, and the +5, +11 and +13 sites are the key base-pairing sites for this interaction.
We predicted the secondary structures of all 13 BtsR1 mutants using the LocARNA tool (40). The ten BtsR1 mutants that retained the ability to inhibit Cry5Ba expression folded into stable structures similar to that of wild type BtsR1 (Supplementary Figure S8). In contrast, the secondary structures of the three BtsR1 mutants that lost their ability to inhibit Cry5Ba expression differed compared to that of wild type BtsR1 ( Figure 6E). In particular, the predicted single-stranded exposed regions of BtsR1 that are complementary to the RBS of cry5Ba mRNA formed a double-stranded stem structure. This change may result in the inability of these three BtsR1 mutants to bind to the cry5Ba mRNA. Our results are consistent with previous reports indicating that the single-stranded-exposed bases of sRNAs are more likely to be responsible for regulation than bases in stems (41,42).

Cry5Ba was expressed when YBT-1518 was ingested in vivo by nematodes
Strain YBT-1518 has high toxicity to C. elegans and possesses three nematicidal cry genes, namely, cry55Aa, cry6Aa, and cry5Ba (21). When expressed in strain BMB171, Cry5Ba exhibits the highest activity among these three Cry toxins against nematodes (44). However, the Cry5Ba toxin expression in YBT-1518 is silenced when outside the host (21), and we showed that the sRNA BtsR1 directly binds to cry5Ba mRNA and represses the expression of Cry5Ba. This raises the possibility that the Cry5Ba toxin may not contribute to the toxicity of YBT-1518 to the nematode due to the negative regulation by BtsR1. To evaluate whether the BtsR1 mediated Cry5Ba regulation affects the toxicity of YBT-1518, nematode bioassays were conducted with YBT-1518 and BtsR1 mutant strains. To our surprise, YBT-1518 exhibited high toxicity against C. elegans N2 similar to those of strains YBT-1518/pHT304-cry5Ba and the BtsR1 depletion strain YBT-1518(BtsR1-dep), in which both express Cry5Ba (t-test, P > 0.05). Furthermore, the cry5Ba knock-out mutant strain YBT-1518( cry5Ba) lost most of the toxicity against C. elegans N2 compared with that of wild type strain YBT-1518 (t-test, P < 0.001) (Figure 7A). These results indicated that Cry5Ba is the leading factor for the toxicity of YBT-1518 to nematodes.
The most likely explanation for such situation is that the Cry5Ba may be expressed after YBT-1518 have been ingested in vivo by the nematode host, resulting in high toxicity against C. elegans. To confirm this hypothesis, we determined the production of Cry5Ba in vivo by Western blot. L4 C. elegans N2 animals were infected with YBT-1518, YBT-1518/pHT304-cry5Ba, YBT-1518(BtsR1-dep) and YBT-1518( cry5Ba), respectively. Then, the bacteria were isolated from 400 to 500 infected C. elegans (at 36 hpi) individuals by grinding the worms. The same amounts of bacterial cells cultured in LB medium for 36 h were used as controls. As expected, Cry5Ba was indeed expressed when YBT-1518 in the host, as detected by the presence of the Cry5Ba signal band ( Figure 7B).
Then, we compared the BtsR1 transcription profiles in YBT-1518 at different phases during bacterial growth in C. elegans and in LB medium by qRT-PCR. For in vivo infection, the bacteria were isolated from 400 to 500 YBT-1518-infected C. elegans individuals by grinding the worms. The transcription level of BtsR1 under LB medium conditions at the same time points were set to 1. Compared with that in the LB medium condition, BtsR1 showed significant low-level transcription when YBT-1518 was isolated from the host, with downregulation by 141-fold, 187-fold and 351-fold from 12 to 36 hpi, respectively ( Figure 7C). Thus, BtsR1 transcription was repressed in vivo, which may and Cry55Aa protein bands are indicated on the right. (C) Translational reporter fusions (lacZ) were performed to confirm the inhibitory functions of BtsR1 mutants. ␤-galactosidase activity was tested in the presence of each BtsR1 mutant and shown as the average of three independent assays. The ␤galactosidase activity of strain YBT-1518/pHT304-pcry5Ba'lacZ was used as a control (CK). The error bar indicates the SD. Cells were cultured in LB medium at 24 h. (D) Competitive EMSA assay was performed to confirm the interactions between selected BtsR1 mutants and cry5Ba mRNA. The wild type cry5Ba 5 -UTR pcry5Ba and selected BtsR1 mutants were synthesized via in vitro transcription systems. The 5 -end FAM-labelled wild type BtsR1 (FAM-BtsR1, 10 nM) and 1000 nM of pcry5Ba mRNA was hybridized and resolved by native PAGE. Excessive of unlabeled BtsR1 mutants (5000 nM) were used as competitors in these reaction systems. (E) Secondary structures of the three BtsR1 mutants that lost the ability to bind cry5Ba 5 -UTR. responsible for the induced expression of Cry5Ba when in the host.

YBT-1518 benefited from BtsR1-mediated Cry5Ba silencing, which facilitated better ingestion and colonization in nematodes
The ability of YBT-1518 to silence Cry5Ba under medium conditions and express this toxin under host conditions is an interesting phenomenon. It has been reported that the Cry5Ba could rapidly induce nematode feeding cessation as an avoidance behavior to this toxin (3,31). Therefore, YBT-1518 potentially silences Cry5Ba toxin expression to confuse C. elegans, resulting in host ingestion.
To test this hypothesis, we analyzed the feeding cessation induced by B. thuringiensis strains in the pres-ence or absence of Cry5Ba. The strains YBT-1518, YBT-1518( cry5Ba), YBT-1518/pHT304-cry5Ba, and YBT-1518(BtsR1-dep) were fed to C. elegans N2 L4 animals. Strain BMB171 was used as a control. Pumping rate assays indicate ordinary feeding behavior of C. elegans N2 when exposed to BMB171 for 5 min to 2 h and the lack of any significant avoidance. C. elegans N2 feeding on strains YBT-1518 and YBT-1518( cry5Ba) exhibited only a slight decrease in pumping rate compared to BMB171 at the 5-30 min timepoints (t-test, P<0.05). However, the feeding behavior of C. elegans N2 on YBT-1518/pHT304-cry5Ba and YBT-1518(BtsR1-dep) demonstrated very strong avoidance at all tested time points compared with feeding on BMB171 (t-test, P < 0.001). The animals ingested the bacteria up to five min, and then all animals ceased feeding between 10 min and 8 h ( Figure 8A). The kinetics of feed- ing cessation induced by these strains when they were presented as the only food option confirmed the above results. Animal feeding ceased in a significant fraction of the population 5 min after transfer onto YBT-1518/pHT304-cry5Ba and YBT-1518(BtsR1-dep) in the entire population from 30 min to 8 h. No significant feeding cessation at any tested time point was observed in animals exposed to BMB171. When exposed to strains YBT-1518 or YBT-1518( cry5Ba), feedings also ceased in a fraction of the population, but this fraction was significantly lower than observed for YBT-1518/pHT304-cry5Ba and YBT-1518(BtsR1-dep) at all tested time points (t-test, P < 0.001) ( Figure 8B). Thus, the BtsR1 mediated Cry5Ba silencing decreases the feeding cessation defense behavior in the C. elegans host.
Given that the expression of Cry5Ba by B. thuringiensis strains induced more significant host feeding cessation than that of strains without Cry5Ba, we asked which food is preferred by the nematode host when offered jointly. We tested the chemotaxis behavior of C. elegans N2 following exposure to B. thuringiensis strains with or without the presence of Cry5Ba by a two-choice preference assay (Figure 8C). Two treatments were conducted where YBT-1518 with YBT-1518/pHT304-cry5Ba or YBT-1518 with YBT-1518(BtsR1-dep) were the only food choices. The preference of C. elegans for different bacteria was measured in a binary choice assay in which animals migrated towards one of two bacterial lawns on opposite sides of a plate. A choice index of 1.0 represents a complete preference for B. thuringiensis strains without Cry5Ba, while an index of −1.0 represents a complete preference for B. thuringiensis strains producing Cry5Ba. An index of 0 represents an equal distribution for the two types of bacteria. In both groups, the animals strongly preferred the B. thuringiensis strains that did not The average pumping rates of animals when exposed to B. thuringiensis strains with or without Cry5Ba. (B) The fractions of animals that were feeding when exposed to B. thuringiensis strains with or without Cry5Ba. (C) Schematic representation of the two-choice preference assays. The black dot at the center of the plate represents the starting point of the N2 worms. (D, E) The worm choice indexes in response to B. thuringiensis strains with or without Cry5Ba tested in the two-choice preference assay. (F) Cry5B-expressing B. thuringiensis results in poor colonization in vivo. All data are presented as the mean ± SE (n = 3), and significant differences are as follows: *P < 0.05; **P< 0.01 and ***P< 0.001. The lack of any symbol indicates no significant difference. express Cry5Ba ( Figure 8D, and E). These results demonstrate that BtsR1-mediated Cry5Ba expression silencing results in a feeding preference for YBT-1518 in C. elegans.

DISCUSSION
The most intriguing feature of the B. thuringiensis strain YBT-1518 is its ability to silence expression of the toxin Cry5Ba during free living growth in medium (Figure 1, (21)) and then to express this toxin when inside a host ( Figure  7B). This specialized behavior was mediated by a specific sRNA, BtsR1, which inhibited the expression of Cry5Ba by directly base-pairing with the RBS sites of the cry5Ba 5 -UTR (Figures 4-6). Small RNAs are involved in the regulation of gene expression and have been reported in many bacteria to regulate many processes, including DNA replication, nutrient metabolism, oxidative stress, iron homeostasis, toxicity, and persistence (37,41). However, the identification of sRNA and its actions on gene expression has not been reported in B. thuringiensis. Our study presents the identification of the first sRNA in B. thuringiensis, BtsR1 ( Figure 3). In addition, the mechanisms that regulate Cry protein expression in B. thuringiensis have been widely studied, and factors such as promoters (45), stable mRNA factors (including STAB-SD sequences (46)), accessory proteins (such as P19 or P20 (47)), and chaperones (including ORF2 (48)) have been identified. Thus far, there has been no report of an sRNA affecting Cry protein expression. Here we showed that an sRNA, BtsR1, bound to the RBS site of the cry5Ba mRNA via direct base pairing and inhibited Cry5Ba expression (Figure 4-6). Our present work describes a novel mode of Cry expression regulation mediated by sRNA in B. thuringiensis.
The ability of YBT-1518 to silence Cry5Ba under medium conditions and express this toxin under host conditions is a very interesting ecological phenomenon. C. elegans is an abundant bacteria-feeding metazoan in soil (49). B. thuringiensis has been isolated from diverse environments, primarily from the soil (50). The soil represents a survival space for the coevolution of B. thuringiensis and C. elegans, and C. elegans may provide adequate nutrition for B. thuringiensis strains in soil (51). C. elegans has evolved certain strategies as defenses against pathogens. One of these strategies involve the evolution of physical barriers, such as the pharyngeal grinder to destroy pathogens (52). Another strategy is the activation of cellular defenses to express AMPs or lysozyme via innate immunity signing pathways such as p38 MAPK (53), JunK-like MAPK (54), and DAF-2 insulin-like growth factor (55). Additionally, C. elegans engages in pathogen avoidance behavioral defenses (1,2). As a pathogen, B. thuringiensis has also evolved certain strategies to respond to nematodes, such as the formation of spores to escape the pharyngeal grinder or the expression of virulence factors to destroy host physical barriers (44,56). In this study, the ingestion of Cry5B-expressing B. thuringiensis resulted in rapid and severe feeding cessation by nema-todes ( Figure 8A and B), subsequent low levels of feeding preference ( Figure 8D and E), and poor colonization ability in vivo ( Figure 8F), compared to Cry5B-silencing B. thuringiensis. YBT-1518 benefited from sRNA-mediated Cry5Ba silencing, which facilitates ingestion by the host (Figure 8D and E) and colonization in vivo ( Figure 8F). Therefore, YBT-1518 potentially silenced Cry5Ba toxin expression regulated by BtsR1 to subvert C. elegans and evade pathogen avoidance behavioral defenses. Indeed, we also demonstrated that YBT-1518 expressed Cry5B in vivo (Figure 7B) following ingestion due to a decrease in BtsR1 transcription ( Figure 7C). This may allow B. thuringiensis to avoid nematode defenses and then express toxins in vivo to kill the host and gain a survival advantage. Our work describes a novel mechanism of sRNA-mediated regulation that allows pathogens to combat avoidance behavioral defenses in the host.
Additionally, rich soils harbor large populations of nematodes and microbes, thereby exerting a powerful selective force on both the microbes and nematodes. Given that nematodes demonstrate sensitive avoidance responses, pathogens that gain fitness advantage by infecting the host are selected. These pathogens have evolved certain strategies to conceal their presence, including surface molecule modifications that cannot be detected by host (1) or the production of molecules that attract worms (19). Conversely, selective pressure exists on nematodes to detect this microbial deceit, potentially resulting in a co-evolutionary arms race consisting of repeated cycles of microbial adaptation followed by host counter-adaptation. These dynamics likely shaped both the evolution of the microbe and the host. B. thuringiensis produced Cry toxin at a huge cost, both in terms of large pure mass (>25% of the total dry weight of cell) and complex metabolism (57). The production of multiple crystal proteins is energy-consuming but presumably benefits the population by facilitating pathogenic infections (58). There are three crystal protein genes exist in YBT-1518. In this study, the silencing of a specific Cry toxin increases the likelihood of YBT-1518 feeding by nematodes, at least in the context of a recombinant strain producing the Cry5Ba toxin ( Figure 8). Cry toxin production is very costly, and hosts represent a fixed pool of resources. Therefore, expression of the Cry toxin solely during the host infection process conserves resources for B. thuringiensis survival. Cry5Ba toxin expression in the host and environment mediated by sRNA BtsR1 may be related to reduced fitness costs and derive from host selection, revealing the selective pressures associated with host avoidance behavior that shape pathogen evolution.
In summary, we describe an unusual strain of B. thuringiensis, YBT-1518, which possesses the following unique feature: it silenced the expression of the toxin Cry5Ba under medium-growth conditions and expressed this toxin under host conditions. This intriguing regulation represents a new model of pathogen response to host avoidance behavioral defense associated with a novel form of sRNA-mediated regulation of cry gene expression. Cry toxin synthesis involved enormous energy consumption by B. thuringiensis. Conversion to toxin production in YBT-1518 appears to increase its competitive ability by reducing toxin synthesis in nutritionally deficient environments.


extfieldsofstudy:
Physics
Materials Science

text:
Painting with bacteria: Smart templated self assembly using motile bacteria

External control of the swimming speed of `active particles' can be used to self assemble designer structures in situ on the micrometer to millimeter scale. We demonstrate such reconfigurable templated active self assembly in a fluid environment using light powered strains of Escherichia coli. The physics and biology controlling the sharpness and formation speed of patterns is investigated using a bespoke fast-responding strain.

pairs. These release their energy stepwise along an electrochemical potential ladder of respiratory enzymes located in the inner cell membrane, generating a PMF of ≈ −150 mV. The electron pair ultimately passes to and reduces O 2 to water. Thus, with no O 2 , PMF = 0 and swimming ceases [17]. If cells under anaerobic conditions can express proteorhodopsin (PR) [3], a green-photon-driven proton pump [20], then they will swim only when suitably illuminated: these are living analogues of synthetic light-activated colloidal swimmers. [9,10,11,12] We show below that the speed with which such cells respond to changes in illumination is crucial for successful bacterial STASA. For this work, we constructed a PR-bearing mutant (AD10) that stops much faster when illumination ceases than previously-reported [3,21], by deleting the unc gene cluster [22] encoding the F 1 F o -ATPase membrane protein complex (see SI §1.1), so that these enzymes cannot act in reverse in darkness to continue to export protons and sustain a PMF [23].
We suspended cells in phosphate motility buffer at optical density OD 8 (cell-body volume fraction ≈ 1.1%) [17] and sealed 2 µL into 20 µm high flat capillaries, where cells swim in two dimensions but have enough room to 'overtake' each other in all three spatial dimensions. Differential dynamic microscopy (DDM) [24] returned an averaged speedv ≈ 30 µm s −1 and β ≈ 20% of non-motile organisms at OD=1 under fully-oxygenated conditions. (Note that 'non-motile' = cells that can never swim; 'stationary' = non-swimming cells capable of motility when illuminated.) Motile cells were allowed to swim until O 2 was depleted andv dropped abruptly to zero after a few minutes [17] (see SI §1.2 and Fig. S1(a)). After these cells were left in the dark for ≈ 10 min, green illumination was turned on (510 -560 nm, intensity I ≈ 5 mW cm −2 at the sample). The stationary cells accelerated uniformly before saturating, Fig. 1. Fitting the data tov(t) =v sat t/(t+τ on ) givesv sat = 9.5 µm s −1 , τ on = 30 s. When illumination ceased,v dropped within τ off 1 s, but never quite to zero -it is unclear why a few cells (< 1%) continued to swim.v sat increased with I, Fig. 1 (inset), up to 27 µm s −1 . {v, β, v sat , τ on } changed over hours as cells aged.
The discharging of the PMF through the membrane (capacitance C 10 −14 F) and rotary motors (total resistance R 10 14 Ω) upon cessation of illumination should take RC ∼ 1 s (see SI §1.3 for details), which explains the observed τ off . Consistent with this interpretation, τ off is approximately independent of the starting speed of decelerating cells (see SI §1.2 and Fig. S2(a)). The observed τ on ≈ 30 s is likely controlled by the rate constant [25] for stator units to come on and off motors, k stator ≈ 0.  Our STASA protocol uses a spatial light modulator (SLM) to project patterns of illumination onto a uniform field of cells rendered stationary by O 2 exhaustion. We first projected a positive mask with bright features on a dark background, Fig. 2a (inset). The width of the features, 90 µm, is comparable to the persistence length (i.e. the length scale over which the cell's motion can be approximated as a straight line) of smooth-swimming E. coli [26]. Over time, the mask pattern was replicated in the cell population with increasing clarity: cells swim out of illuminated areas, stop within τ off ∼ 1 s, and accumulate at the edges just outside the boundary of these areas, Fig. 2b (also see SI movie M1). Next we abruptly switched to a negative mask (dark pattern against bright background), Fig. 2c (inset). Edge-accumulated cells start swimming, about half of which arrive inside the dark features immediately and stop within τ off , rapidly outlining the inside edge of the masked areas, Fig. 2c.
Thereafter, the dark areas progressively filled up: bacteria continually arrive from the outside, which acts as an essentially infinite reservoir of swimming cells, Fig. 2d-e (also see SI movies M2 and M3). Other patterns can be 'painted' using this method, Fig. 2g. In contrast to static 'bacterial painting' recently demonstrated [27], our method is dynamic and reconfigurable in real time within our liquid medium.
To understand how fast such reconfigurations of spatial patterns can take place, we experimented with square masks, Fig. 2f, using two initial conditions: a (dark) field of stationary cells (v(0) = 0, i.e., diffusive), and a field of (illuminated) cells that have reached speed saturation (v(0) =v sat ). We studied the evolution of cell density (see Methods and SI §1.5) and speed inside bright squares against a dark background. Starting with diffusive cells,v(t) =v sat t/(t + τ on ) ≈ at with a =v sat /τ on for t τ on . The time to empty an L × L square should scale as The measured cell density evolution, ρ tot (t), in illuminated squares of different sizes is shown in Fig. 3a, from which we extracted t D , the time for ρ tot to drop to 1/e of its decay. As expected, t D (L) ∼ L 0.5 , Fig. 3c. Fitting Eqn.
(1) gives a = 0.05 µm s −2 , which is a reasonable average acceleration over the first ∼ 100 s for the data in On the other hand, starting with cells all swimming (on average) atv sat , a square should empty in From the measured ρ tot (t) in this case, Fig. 3b, we deduced a 'box emptying time' and find, Fig. 3c between bright and dark areas acts as an absorber of swimmers arriving with an effective diffusive process. Due to conservation of mass the diffusive propagation of the peak is thus expected, although quantitative aspects remain to be elucidated.
Finally, we seek to understand what controls the sharpness of the self-assembled features using our method.
A stark demonstration of the answer comes from projecting a step-function pattern of light-dark illumination on initially uniformly swimming cells (see SI §1.7 for details). Figure 4 shows the resulting density of cells (from the intensity variance, see SI §1.6) for two strains, AD10, with τ stop 1 s, and AD57, a strain lacking the unc cluster deletion and having τ off > 1 min (see Fig. S7 for images). In the case of AD10, the illumination pattern is clearly replicated in cell density within 1 min, and the steady state shows a sharp feature with half width 10 µm. For AD57, it takes 1 min to establish a corresponding density pattern, which, in the steady state, is 50 µm wide.
Thus, a short enough τ off is necessary for the fast establishment of sharp features using our STASA protocol.  In sum, we have demonstrated how to perform smart templated self assembly using a PR-bearing strain of E. coli with unc cluster deletion dispersed in anaerobic motility buffer. Spatial patterns of light are reproduced sharply in cell density, with the time scale of pattern formation controlled by the 'on time' and the sharpness of the features controlled by the 'off time'. This opens up many opportunities. Thus, e.g., it should now be possible to test a fundamental prediction that the product of density and speed is constant in a system with spatiallydependent activity [28]. Practically, we can go beyond a recent demonstration [13] of the control of lithographed micro-devices using light-activated bacteria: STASA allows the assembly and real-time reconfiguration of the devices themselves using the same bacteria [16]. On the other hand, if cell density is used to trigger the secretion of adhesive biopolymers, bacterial STASA offers a route for manufacturing bespoke 10-100 µm microstructures, which remains a challenging length scale for current 3D printing.

Methods Cells
The Escherichia coli used in this work (AD10 & AD57) are smooth-swimming mutants of AB1157 transformed by a plasmid expressing SAR86 γ-proteorhodopsin (a gift from Jan Liphardt, UC Berkley), similar to the original study [3]. To achieve better light-switching response, we also deleted the unc gene complex, thus removing the coupling between PMF and ATP [22].
Overnight cultures were grown aerobically in 10 mL Luria-Bertani Broth (LB) using an orbital shaker at 30 • C and 200 rpm. A fresh culture was inoculated as 1:100 dilution of overnight-grown cells in 35 mL tryptone broth (TB) and grown for 4 h (when an optical density of OD 600 ≈ 0.2 was reached). The production of proteorhodopsin (PR) was induced by adding arabinose to a concentration of 1 mmol as well as the necessary cofactor all-transretinal to 10 µmol in the growth medium. Cells were incubated under the same conditions for a further hour to allow protein expression to take place and then transferred to motility buffer (MB, pH = 7.0, 6.2 mM K 2 HPO 4 , 3.8 mM KH 2 PO 4 , 67 mM NaCl and 0.1 mM EDTA). Cells were re-suspended in MB after a single filtration (0.45 µm HATF filter, Millipore), yielding high-concentration stock solutions (OD ≈ 8 to 10), which were then diluted with MB as required.
Suspensions were loaded into 2 µL sample chambers (SC-20-01-08-B, Leja, NL, 6 mm×10 mm×20 µm), giving a quasi-2D environment for E. coli swimming. The chamber was sealed using vaseline to stop flow and oxygen supply, and placed onto a microscope for video recording. E. coli initially swims under endogenous metabolism, which depletes the oxygen within the sample cells [17]. Once oxygen is depleted, the cells stop swimming unless illuminated by green light.

Optical setup
The samples were observed using a Nikon TE2000 inverted microscope with a phase contrast objective (PF 10×/0.3).
Time series of ≈ 40 s-movies (100 frames per second) were recorded using a fast CMOS camera (MC 1362, Mikrotron). A long-pass filter (RG630, Schott Glass) was introduced into the bright-field light path to ensure that the light required for imaging did not activate PR. The light controlling the swimming behavior of the bacteria was provided by an LED light source (Sola SE II, Lumencor) filtered to a green wavelength range (510 -560 nm) to overlap with the absorption peak of PR [3]. This light was reflected off a digital mirror device (DLP6500, Texas Instruments), which was imaged onto the microscope sample plane using a demagnifying relay telescope, leading to a resolution of 2.7 µm per pixel. The light was coupled into the transmission path by introducing a dichroic mirror in between the long working distance phase contrast condenser and the sample.
This 'trans' geometry allowed us to illuminate uniformly a circular area of ≈ 2.9 mm in diameter, larger than the objective's field of view. Computer control of the digital mirror device gave us precise spatial and temporal control of the illumination pattern.
Motility and density measurements.
The motility of the samples was quantified using differential dynamic microscopy (DDM) [29], which yields the average swimming speedv and fraction of non-motile bacteria β from low magnification movies [24,30,17]. By comparing the amplitude of the DDM signal to that of a reference sample, it can also give the relative bacteria density ρ/ρ 0 (see SI §1.5 for details). However, the spatial resolution of this method of density measurement is very limited. In experiments where we need to measure density profiles with sharp spatial variations, we used instead the local variance of the image intensity to provide estimates of local cell density (see SI §1.6 for details).  AD10 was constructed using our laboratory stock of E.coli AB1157 [31]. All of the genes present within the operon encoding the ATP synthase complex (atpI, atpB, atpE, atpF, atpH, atpA, atpG, atpD, atpC) were deleted in a single step by allelic replacement using a recombinant pTOF24 vector [32] containing 400bp homology arms flanking the deletion on each side. This resulted in the ∆unc strain AD10 with a deletion of 7504bp (corresponding to position 3915552-3923056 of the E.coli K12 MG1655 chromosome). P1 transduction was used to delete the cheY gene in both this strain and the parental strain (to generate AD57) using JW1871 (BW25113 cheY) as donor [33]. Diagnostic PCR reactions were performed to confirm genotypes. Both strains were transformed with plasmid pBAD-His C (a kind gift from Jan Liphardt, UC Berkley) encoding the SAR86 γ-proteobacterial photorhodopsin variant [21].

Saturation, startup and stopping behaviour
Here we give further information and detailed data supporting the claims made in the main text about the dependence of average swimming speed,v, on light intensity (I), startup time (τ on ), and stopping time (τ off ).  Fig. S1(a). Figure S1: (a) Swimming speed of AD10 (sample OD = 1) during oxygen depletion and for different light intensities: Within ≈ 7 min of sealing the sample oxygen was depleted and swimming ceased. Green light was switched on at ≈ 20 min. Illumination intensity was changed at various points throughout this experiment to provide data for the speed vs. intensity plot in the main manuscript (inset, Fig. 1). (b) A light-sensitive strain with unc-cluster deletion (AD10) shows a rapid decrease in swimming speed as soon as the illumination is stopped (black symbols). However, for strains which do not have the unc gene cluster deleted (AD57) a significant fraction of the population (see inset showing the motile fraction α = 1 − β, lines are guides to the eye) continues to swim at reduced speed for several minutes, giving a much slower decay inv (red symbols; solid red line: exponential fit, giving τ ≈ 60 s).
We claimed in the main text that deleting the unc gene cluster responsible for encoding the ATP synthase complex significantly reduced τ off . Figure S1(b) shows the behaviour of two strains of proteorhodopsin-powered E. coli: AD57 and AD10, without and with unc gene cluster deletion respectively. When illumination was switched off after the cells have reached steady state in constant illumination, the average swimming speed of both strains showed a sharp drop. In the case of AD10, this sharp drop was from ∼ 17 to 4 µm s −1 over 1 s.
The drop was much smaller for AD57, from ∼ 14 to 10 µm s −1 , again over 1 s. Thereafter, a significant fraction of AD57 cells kept on swimming with their average speed decreasing exponentially with a time constant of ∼ 60 s.
A simple resistor-capacitor equivalent circuit for the cell membrane and flagella motor was used in the main text to explain the observed τ off ≈ 1 s. We claimed that this was supported by the observation that τ off was independent of the starting speed from which cell decelerated. Figure S2(a) shows evidence for this claim.
In the main text, we suggested that the long τ on of cells that have been kept in prolonged darkness was controlled by the time needed to reassemble the motor from individual stator units. We supported this claim by saying that τ on was much shorter for cells that had not been in low illumination for very long. Figure S2(b) shows evidence for this. When illumination alternated between high and low with a period of 20 s, we find τ 1 s, comparable to τ off . Figure S2: (a) Average swimming speed measured with DDM as green light is switched off (at t = 0). There is a very rapid speed drop τ off ≈ 1 s as the light is switched off, which is independent of the initial swimming speed (at the resolution limit of our technique). (b) AD10 bacteria adapt their swimming speed within 1s as the green light is alternated between low and high intensity. In this case τ on ≈ τ off . Figure S3: Equivalent circuit of the bacterial PMF generation and dissipation: R mot = resistance of flagella motors, C mem = membrane capacitance, V res = PMF generated by respiratory enzymes, R res = variable resistor modelling the effect of oxygen concentration on respiratory enzymes, V PR = proteorhodopsin-generated PMF, R PR = variable resistor modelling the effect of light on PR. (Redrawn after [3].)

Circuit model
The effect of proteorhodopsin (PR) on the proton motive force (PMF) can be modelled using a simple equivalent circuit, Fig. S3 [3]. We use this model for our experiments by interpreting R res as the effect of oxygen concentration on the respiratory circuit: R res → ∞ when oxygen is exhausted. The effect of light on PR is modelled as a variable resistor, R PR , which Walters et al. have argued to be related to the incident light intensity I by R PR (I) = R ∞ PR (I + K)/I (with K a constant). Using the assumption that the average swimming speed is proportional to PMF, this leads to a similar intensity dependence of the speedv sat (I) =v max I/(I + I 1/2 ) under complete oxygen starvation. Fitting this to our experimental data gives a maximum swimming speed v max = 28.4(7) µm s −1 and an intensity to reach half this speed of I 1/2 = 10.7(4) mW cm −2 , suggesting that our PR is significantly more efficient than the one used by Walter et al. (for which I 1/2 = 60 mW cm −2 ) [3].
We suggest that the stopping time is controlled by the discharge of the PMF of the equivalent capacitance of the membrane through the equivalent resistance of the flagella motor, i.e. τ off = R mot C mem . The specific capacitance of bacterial membranes is ≈ 0.6 µF cm −2 [34]. A 2 µm×1 µm spherocylindrical E. coli cell has 2π µm 2 surface area, giving C mem ≈ 4 × 10 −14 F. We have previously estimated [17] that the proton current through a motor is ≈ 2 × 10 4 H + s −1 , which is driven by a PMF of ≈ −150 mV, giving R mot ≈ 0.5 × 10 14 Ω per motor. On average, each cell has, say, ≈ 5 motors. These constitute resistors in parallel, so that R mot ≈ 0.1 × 10 14 Ω. Thus, τ off ∼ R mot C mem 1 s, as claimed in the main text.

Additional data for cells leaving illuminated squares
In the main text we reported measurements of how long it took to empty an illuminated square (side L) of cells, starting from swimming or stationary cells. The emptying time scales as L and L 1/2 respectively. This is the scaling expected for random ballistic swimmers leaving a box of size L. This is appropriate for our experiments, as the persistence length of our smooth swimming bacteria is comparable to the box size. Here we report the average speed of cells remaining in the illuminated square,v r , as a function of time in the two cases. Starting from swimming cells, our simplistic model presented in the main manuscript assumed a constant swimmming speedv r =v sat , whereas the actually measured speedv r decreased with time, Fig. S4

Differential dynamic microscopy
Differential dynamic microscopy (DDM) is a fast, high-throughput technique to measure the dynamics of particle suspensions such as colloids [29] or swimming microorganisms [24,30,17]. For swimming bacteria, DDM measures motility parameters averaged over ∼ 10 4 cells from ≈ 30 s long movies via the differential image correlation function g( q, τ ), i.e. the power spectrum of the difference between pairs of images delayed by time τ . Under appropriate imaging conditions and for isotropic motion, g( q, τ ) is related to the intermediate scattering function f (q, τ ), which is the q th Fourier component of the density temporal autocorrelation function, via Here, B(q) relates to the background noise and A(q) is the signal amplitude. Fitting f (q, τ ) to a suitable swimming model of E. coli, we obtain the average,v, and width, σ, of the speed distribution P (v), the fraction of non-motile bacteria β and their diffusion coefficient D [30]. Provided the system is dilute enough such that its structure factor S(q) ≈ 1, A(q) is proportional to the particle density [35,36]. Thus, in two otherwise identical samples with different cell densities ρ 1,2 , We verified this using bacteria suspensions of different cell concentrations (measured by optical density), Fig. S5.

Estimating cell density using intensity variance
Although DDM in the form of Eq. 4 is useful for extracting average cell densities over large uniform areas, its strictly limited spatial resolutions makes it unsuitable for assessing cell densities that vary rapidly in space, such as from images shown in Fig. 2 of the main text to illustrate STASA, or the band of high density bacteria shown Figure S5: Cell density deduced from DDM ( §1.5) and intensity variance (section §1.6) measurements for a series of 7 samples starting at a measured optical density of OD = 5.6 and diluted successively by factors of 2. Both analysis methods were applied to the same movies and yield consistent results in quantitative agreement with the prepared sample densities throughout the measured range. Both data sets have been calibrated by assuming the data point for OD 1.4 was measured correctly.
in Fig. S6(a). In the latter case, neither the intensity along a line nor the average of such line profiles, Fig. S6(b) and (c) respectively, bears much resemblance to the cell density variation visible to direct inspection. This is related to the nonlinearities at high phase shifts and artifacts inherent to phase contrast imaging [37].
Interestingly, the single line profile, Fig. S6(b), shows that the intensity fluctuates more in the high-density region. Indeed, the averaged horizontal variance profile, Fig. S6(d), shows qualitative agreement with the visuallyobserved density profile of the band, suggesting that the variance (with background subtracted) could be used to quantify local density. Figure S5 shows that, indeed, densities of uniform samples deduced from variance measurements agree quantitatively with densities deduced from DDM using Eq. 4 and from optical density measurements. At least within the low density limit, this scaling of intensity variance with cell density is not surprising: a (dark) cell on a uniform (light) background will lower the mean intensity slightly and introduce a characteristic amount of variance relative to this background. As more cells are added they initially increase the variance by the same characteristic amount (as the mean background hardly changes), leading to the observed proportionality. For phase contrast imaging, phase-dark object are often surrounded by a lighter halo, so that the net effect of cells on mean intensity is rather small while at the same time leading to a significant characteristic variance per object.

STASA for AD10 vs AD57
We demonstrate the effect of τ stop on STASA by projecting a bright-dark intensity step onto cells that had been uniformly illuminated for ≈ 10 min (so that the population had reached saturation speed,v AD10 = 17.5 µm s −1 andv AD57 = 11.8 µm s −1 ). Therefore there is initially very little effect on the cell density, and only after a much longer time a far more gradual pattern emerges, as demonstrated by the profiles of the intensity variance (reproduced from the main text in Fig. S7). Figure S7: Spatial control of a) AD10 strain compared to b) AD57 strain. Phase contrast images (Nikon PF 20x/0.5 objective) of the samples just before (0 min) as well as 1 min and 16 min after blocking the green light on the right half of the image (boundary position indicated by green lines). The corresponding variance profiles (normalised by the mean variance under uniform illumination) are shown underneath the images. The numerically smoothing (bold lines; using a binomial smoothing filter with n = 201) of the raw data (faint lines) highlights the underlying change in cell density. For the AD57 strain the density difference develops much more slowly and shows a much more gradual transition, making it less suitable for STASA.

Movies
Movie M1: 40 s movie showing the initial formation of the 'UoE' pattern. This corresponds to the run-up to fig. 2a in the main manuscript. The frame-rate has been reduced from the 100 fps of the original movie to 2 fps and is played back at 5× the real speed. One pixel corresponds to 1.4 µm.
Movie M2: Movie showing initial 40 s just after the inversion of the 'UoE' pattern. This corresponds to the transition between fig. 2c and d in the main manuscript. Frame-rate adjusted as for movie M1.
Movie M3: Timelapse image series of the UoE pattern being formed and inverted. This illumination pattern is switched on at time '0:00' when the sample is still uniform (corresponding to movie M1) and becomes clearer over time. After 9 min the pattern is inverted (corresponding to movie M2) and again becomes more pronounced over time. The outline of the letter 'o' is marked to highlight the switching of the boundary as the pattern is inverted.