MogR is a ubiquitous transcriptional repressor affecting motility, biofilm formation and virulence in the Bacillus cereus group

Flagellar motility is considered an important virulence factor in a range of different pathogenic bacteria. In Listeria monocytogenes the transcriptional repressor MogR regulates motility in a temperature-dependent manner by directly repressing flagellar- and chemotaxis genes. The only other bacteria known to carry a mogR homolog are members of the Bacillus cereus group, which includes both motile and non-motile species. Furthermore, the motility locus in B. cereus group bacteria appears to be more closely related to L. monocytogenes than to Bacillus subtilis. In this study we show that in Bacillus thuringiensis, overexpression of MogR results in non-motile host cells devoid of flagella. Global gene expression profiling showed that 110 genes were differentially regulated by MogR overexpression, including flagellar motility genes, but also genes associated with virulence, stress response and a biofilm lifestyle. Accordingly, phenotypic assays showed that MogR also affects cytotoxicity and biofilm formation in B. thuringiensis. Overexpression of a MogR variant mutated in two key amino acids within the putative DNA binding domain restored phenotypes to those of a wild type empty vector control strain. In line with this, the introduction of these mutations resulted in complete loss in the ability of MogR to bind to its putative target site within the flagellar motility locus in vitro. In contrast to L. monocytogenes, MogR appears to be regulated in a growth-phase dependent and temperature-independent manner in B. thuringiensis 407. Interestingly, mogR was found to be conserved also in non-motile B. cereus group species with highly reduced motility loci, such as Bacillus mycoides and Bacillus pseudomycoides, and is expressed in non-motile Bacillus anthracis, providing further indications of an expanded and unexpected set of functions for MogR in B. cereus group species, in addition to motility regulation. In conclusion, MogR constitutes a novel B. cereus group pleiotropic transcriptional regulator, acting as a repressor of motility genes, and affecting the expression of a variety of additional genes involved in biofilm formation and virulence.


Introduction
In many bacterial species, flagella have been demonstrated to be important to virulence functions, including reaching the optimal host site, colonization or invasion, maintenance at the infection site, post-infection dispersal, protein secretion and more (Chaban et al., 2015). Temperature-dependent regulation of motility in the human pathogen Listeria monocytogenes has been described in a series of studies published by Higgins and co-workers, in which motility was shown to be regulated by the transcriptional repressor MogR and its antirepressor GmaR (Gründling et al., 2004;Kamp and Higgins, 2009;Shen et al., 2009;Kamp and Higgins, 2011), the genes for which are widely distributed among different Listeria spp. (Smith and Hoover, 2009). L. monocytogenes is a foodborne facultative intracellular pathogen, which uses flagellum-based motility when present in its extracellular environmental niche. During mammalian infection however, motility genes are downregulated by MogR-dependent repression upon sensing of mammalian physiological temperature (37 °C in the human). In this system, the GmaR anti-repressor functions as the temperature sensor, by antagonizing MogR repression activity at temperatures below 37 °C . The activity of GmaR is dependent on the transcriptional activation by DegU at low temperatures, and a temperature-dependent, posttranscriptional mechanism limits GmaR production to temperatures below 37 °C (Kamp and Higgins, 2009). This system allows L. monocytogenes to switch from an environmental and extracellular motile bacterium to an intracellular pathogen. Inside host cells, flagella are not required as L. monocytogenes cells instead move by actin-based motility, and downregulation of flagella during infection is thought to aid bacterial evasion of the host innate immune system (Hayashi et al., 2001;Li et al., 2017).
The only other known bacteria carrying a homolog to Listeria spp. mogR are species from the Bacillus cereus group (B. cereus sensu lato) (Gründling et al., 2004;Fagerlund et al., 2010), which is a group of closely related Gram-positive spore-forming bacteria of considerable medical and economic importance. The group comprises at least seven species, including B. cereus (sensu stricto), Bacillus anthracis, Bacillus thuringiensis, Bacillus weihenstephanensis, Bacillus mycoides, Bacillus pseudomycoides, and Bacillus cytotoxicus, which, like L. monocytogenes, can be isolated from the environment, e.g. soil, air and water. In contrast to L. monocytogenes however, B. cereus is an extracellular opportunistic pathogen. The majority of strains of B. cereus sensu stricto, B. thuringiensis, B. weihenstephanensis and B. cytotoxicus are motile by peritrichous flagella, while B. anthracis, B. mycoides, and B. pseudomycoides are described as non-motile (Twine et al., 2009;Guinebretière et al., 2013). B. cereus group bacteria can cause foodborne disease (emetic or diarrhoeal syndrome) and serious opportunistic infections in man, but also encompasses strains that are used as probiotics (Stenfors Arnesen et al., 2008;Bottone, 2010;Cutting, 2011), some of which have been suggested to have separate species status (e.g. Bacillus toyonensis; Jimenez et al., 2013). The obligate mammalian pathogen B. anthracis is the cause of anthrax disease, being endemic in several parts of the world, and has been used as a biological terror agent. B. thuringiensis is an entomopathogenic bacterium, frequently used as a biopesticide to protect crops against insect pests, however this species also carries virulence factors shared with B. cereus sensu stricto (Damgaard, 1995;Gaviria Rivera et al., 2000;Swiecicka et al., 2006;Celandroni et al., 2014;Kim et al., 2015) and has caused human infections similar to those caused by B. cereus (Samples and Buettner, 1983;Jackson et al., 1995;Damgaard et al., 1997;Hernandez et al., 1998;Ghelardi et al., 2007b). Although strains in the B. cereus group to a certain degree follow ecological diversification patterns through evolutionary time (Guinebretière et al., 2008), B. cereus and B. thuringiensis isolates are intermingled in global MLST-or k-mer-based phylogenetic analyses of a high number of non-biased isolates (Kolstø et al., 2009;Bazinet, 2017), while the thermotolerant species B. cytotoxicus forms a separate and phylogenetically remote clade within the B. cereus group population.
The bacterial flagellum is a complex molecular structure made up of about 25 different proteins. In most bacteria studied to date, expression of flagellar genes is subject to hierarchical regulation to ensure the sequential expression required for proper flagellum assembly (Smith and Hoover, 2009;Erhardt et al., 2010). In contrast, L. monocytogenes and B. cereus group strains appear to lack this transcriptional cascade control of flagellar biosynthesis (Smith and Hoover, 2009;Chiara et al., 2015). B. cereus group bacteria also lack  D , a common key regulator of motility genes in bacteria, indicating a different mode of transcriptional regulation for the motility genes. In this study, we aimed to investigate functional roles of the MogR homolog identified in B. cereus group bacteria, and whether motility regulation in the B. cereus group more closely resembles that of L. monocytogenes rather than other Bacillus species.

Sequence analysis
Motility proteins that were orthologs between species were identified by amino acid sequence searches (BLASTP) performed using BLAST v.2.6.0+ (Altschul et al., 1990;Altschul et al., 1997) between all pairwise combinations of the motility loci from B. thuringiensis 407, L. monocytogenes EGD-e, and B. subtilis 168 (RefSeq accession numbers NC_018877.1, NC_003210.1, and NC_000964.3, respectively; Glaser et al., 2001;Barbe et al., 2009;Sheppard et al., 2013). Results for reciprocal best BLASTP hits between each pair were included in Supplementary Table S1 if the BLASTP alignments had a percentage of identical matches above 20%, a bit score greater than 30 and covered at least 25% of each amino acid sequence. Comparisons between the motility locus in different B. cereus group strains were obtained using the Integrated Microbial Genomes (IMG) browser (Markowitz et al., 2010; http://img.jgi.doe.gov), by searching for orthologous genome neighborhoods to genes from the B. thuringiensis 407 motility operon. Annotations for the B. cereus ATCC 10987 motility cluster and the B. mycoides DSM 2048 mogR gene were corrected using EasyGene (Larsen and Krogh, 2003). To perform an exhaustive comparative analysis of the genetic structure of the motility locus, genome sequences of 106 subgroup I strains of the B. cereus group that had been sequenced to a minimum of scaffold level were downloaded from NCBI (November 25, 2019), and a local BLAST database was indexed from the corresponding subgroup I proteomes.

Strains and growth conditions
The strains used in this study are presented in Table 1. B. thuringiensis 407 Cryˉ (also sometimes referred to as Bt407) is an acrystalliferous strain cured of its cry plasmid (Lereclus et al., 1989). It is genetically close to the B. cereus type strain ATCC 14579 (Tourasse et al., 2006).
Unless otherwise stated, B. thuringiensis 407 cultures were inoculated with 1% of an overnight culture and grown at 30 °C and 200 rpm in Luria Bertani (LB) broth or in bactopeptone medium (1% w/v bactopeptone, 0.5% w/v yeast extract, 1% w/v NaCl). For cloning and expression in Escherichia coli, ampicillin at 50 or 100 µg ml -1 , kanamycin at 50 µg ml -1 and/or erythromycin at 400 µg ml -1 was used. Erythromycin at 10 µg ml -1 was used to maintain the pHT304-Pxyl plasmid constructs in B. thuringiensis 407. For induction of gene expression from the xylA promoter on pHT304-Pxyl, xylose was added to the growth medium at 1 mM or as otherwise stated. For induction of MBP-MogR in E. coli BL21(DE3), 0.3 mM isopropyl-β-D-thiogalactopyranoside (IPTG) was used. Growth curves were prepared in bactopeptone medium, pH 7.0 with cultures grown with shaking at 220 rpm, using 50 ml culture volumes in 250 ml baffled flasks.

Reverse transcription quantitative PCR (RT-qPCR)
For analysis of mogR, flaA and flaB expression throughout the bacterial growth phase, RT-qPCR was performed essentially as described by Fagerlund et al. (2016). Briefly, cells grown in bactopeptone medium at 30 °C were incubated in an equal volume of ice-cold methanol for 5 minutes before harvesting by centrifugation. Cells were lysed using a Precellys 24 Tissue Homogenizer (Bertin) and RNA was isolated using the RNeasy Mini or Midi Kits (Qiagen). After treatment with DNase and further purification, cDNA synthesis was performed in duplicate for each sample using SuperScript III Reverse Transcriptase (Invitrogen). For all samples, a negative control reaction without reverse transcriptase was included. RT-qPCR was carried out with a LightCycler 480 Real-Time PCR System (Roche) using primers listed in Supplementary Table S2. The three genes gatB, rpsU, and udp, shown to be stably expressed throughout the B. cereus life cycle (Reiter et al., 2011), were used as reference genes, and were included for each sample and on each plate. The second derivative maximum method in the LightCycler 480 software (Roche) was utilized to obtain a quantification cycle (Cq) value for each reaction. The expression of each target gene in each biological replicate was converted into E Cq values (Pfaffl, 2001) and then normalized to the geometric mean of the E Cq values obtained for the three reference genes. Finally, averages and standard deviations were calculated from the normalized expression values from the biological replicates.
For analysis of flaA and flaB expression in the B. thuringiensis 407 strains overexpressing MogR and MogR QN→AA (see below), the same procedure was used except that gatB and rpsU were used as reference genes.

Cloning of mogR for expression in B. thuringiensis 407
The low-copy number E. coli/Bacillus shuttle vector pHT304-Pxyl, in which xylR and the xylA promoter from B. subtilis was inserted into the pHT304 cloning site (Arantes and Lereclus, 1991) allowing xylose-inducible expression of downstream cloned genes, was used for overexpression studies. The sequence encoding MogR from B. thuringiensis 407 (BTB_RS08390) was PCR-amplified using primers 5'-gtcggatccgaattgtgaaaggatgagg-3' and 5'-taaggtacctcttccttcttcggaacg-3' and genomic DNA as the template. The PCR product was inserted into pHT304-Pxyl using the primer-incorporated BamHI and KpnI restriction sites (underlined), creating pHT304-Pxyl-mogR and placing mogR under transcriptional control of the xylA promoter.
The plasmids were verified by sequencing. The constructed pHT304-Pxyl plasmids and the empty vector were introduced by electroporation into B. thuringiensis 407 (Masson et al., 1989).

Motility assay
Swimming ability was determined on 0.3% LB soft agar plates with 1 mM xylose and 10 µg ml -1 erythromycin added. A 5 µl drop of culture grown in LB broth overnight at 30 °C was spotted on each agar plate. The plates were wrapped in plastic and incubated for 7 hours at 30 °C. Each independent assay was performed with two or three technical replicates.

Atomic force microscopy (AFM)
AFM imaging and analysis was performed using a Nanowizard AFM microscope (JPK Instruments). Bacterial cell culture was grown in LB broth with erythromycin (10 µg ml -1 ) and xylose (10 mM) at 37 °C to an OD600 of 3. One ml of culture was washed three times and finally resuspended in 0.9% NaCl. Cells were then diluted (15:50) in a 10 mM magnesium/Tris buffer, pH 7.5. Ten µl was applied onto freshly cleaved mica surfaces mounted on a glass slide, and allowed to adhere for 10 minutes followed by washing (10×100 µl) using sterile dH2O. Excess water was carefully removed, and the slide gently dried using a nitrogen gas jet stream. Images were recorded in intermittent-contact mode at room temperature in air using a MicroMasch NSC35/AIBS cantilever. AFM images were analyzed using The NanoWizard IP Image Processing Software (JPK Instruments).

SDS-PAGE and Western immunoblotting
For detection of flagellin, for each strain analyzed, two parallel bacterial culture samples (10 ml LB broth) were harvested by centrifugation (4100 × g, 4 °C) after 3.5 hours growth at 30 C (OD600 ~1.2)one for extraction of surface proteins, and one for whole cell protein extraction. The cell pellets were resuspended in 1 ml PBS (pH 7.1) and kept on ice. For extraction of surface proteins, the washed cells were centrifuged for 5 minutes at 16,000 × g and 4 °C and resuspended in an equal volume of 2×SDS-PAGE sample buffer before incubation at 95 °C for 5 minutes. The supernatant was collected by centrifugation as before.
The whole cell fraction was prepared by washing the cell pellet in PBS followed by centrifugation for 5 minutes at 16,000 × g and 4 °C. The pellet was then resuspended in 500 µl PBS and lysed using a Precellys 24 Tissue Homogenizer (Bertin). The supernatant was collected by centrifugation for 8 minutes as before. Whole cell supernatant (21 µl) with 7 µl 4×SDS-PAGE buffer and 5 µl of the surface protein fraction were separated on 12% SDS-PAGE gels as described below.
For detection of Hbl, Nhe and CytK, cultures were harvested after 4.5 hours growth. Supernatant samples were collected by centrifugation, concentrated 40-fold by precipitation with four volumes of ice-cold acid acetone:methanol (1:1 v/v), stored at -20 °C overnight, and harvested by centrifugation for 30 minutes at 16,000 × g and 4 °C. Then, pellets were left to evaporate at 4 °C overnight and resuspended in 2×SDS-PAGE sample buffer. Samples were diluted 20-fold with MQ H2O. Concentrated supernatant samples (10 µl) were separated on 10% SDS-PAGE gels.

Microarray analysis
Cultures of B. thuringiensis 407 harboring either pHT304-Pxyl or pHT304-Pxyl-mogR were grown in LB broth containing 10 mM xylose at 37 °C for 3 hours, and then incubated in an equal volume of ice-cold methanol for 5 minutes before pellets were harvested by centrifugation at 2800 × g for 20 minutes. For extraction of RNA, cells were lysed using a Precellys 24 Tissue Homogenizer (Bertin) and RNA was isolated using the RNeasy Mini Kit (Qiagen) and subjected to on-column DNase-treatment using the RNase-Free DNase Set (Qiagen). cDNA synthesis, labelling and purification was performed as described (Gohar et al., 2008). Microarray slides were printed at The Microarray core facility of the Norwegian University of Science and Technology (NTNU). Design, printing, prehybridization, hybridization and scanning of the slides and analysis of the data was performed as described (Gohar et al., 2008). Each microarray experiment was based on four slides, all biological replicates. P-values were computed using a false discovery rate (FDR) of 0.05.
The microarray slides contain 70-mer oligonucleotide probes designed to detect open reading frames (ORFs) in the following strains: B. anthracis Ames, B. anthracis A2012, and B. cereus ATCC 14579, in addition to selected genes from B. cereus ATCC 10987 (Kristoffersen et al., 2007). All probe sequences on the microarray were analyzed by BLAST for hits to the annotated genes of a B. thuringiensis 407 draft genome sequence (as of April 30, 2009; the gene lists are based on the GenBank annotations as of this date; accession no. ACMZ00000000.1). Only probes with 93% identity or greater to a transcript/feature sequence of B. thuringiensis 407 were included in the analysis. Of the predicted B. thuringiensis 407 genes, 1719 genes, most of which were hypothetical genes, did not have corresponding probes on the array (these include the 761 genes on contigs 00213 and 00060, which appear to be plasmid-borne). COG categories were obtained for the analysed genes as reported in the IMG database (http://img.jgi.doe.gov).

Cytotoxicity assay
The Vero cell cytotoxicity assay was performed as described by Lindbäck and Granum (2006), and measures the percentage inhibition of C 14 -leucine incorporation in cells due to the cells being subjected to toxins, calculated relative to a negative control where cells were not subjected to toxin sample. Samples of early stationary phase cultures grown to an OD600 of 2.4 were collected and 100 µl or 150 µl samples (for deletion mutants and overexpression strains, respectively) were applied to the cytotoxicity assay. The assays were performed for three independent biological replicates with two technical replicates in each assay.

In vivo toxicity in insect larvae
The virulence-related properties of MogR were assessed by comparing the killing effect of the B. thuringiensis 407 MogR overexpression and empty vector control (pHT304-Pxyl plasmid) strains, in both wild type and ∆flaAB backgrounds, by infection (force feeding) in 5 th instar Galleria mellonella larvae. G. mellonella eggs were hatched at 28 °C and the larvae reared on beeswax and pollen. For infection experiments, groups of 20 to 25 G. mellonella larvae, weighing about 200 mg each, were used. As MogR overexpression is activated from the pHT304-Pxyl promoter by the addition of xylose, we tried for the first time to evaluate if activation could occur in the insect intestine as well. Therefore, xylose (20 mM) was both added to the LB growth medium of the four strains, as well as to the bacterial inoculums and the toxin alone control (Cry1C) at time zero (time point of force feeding). Larvae were force fed a second time with 10 µl 20 mM xylose 5 hours later (in order to again activate MogR expression from the plasmid). Infections were otherwise performed as previously described (Fedhila et al., 2006), by force feeding with 10 µl of a mixture containing 4-5×10 6 of vegetative bacteria (exponential growth OD600 ~1 in LB) with 20 mM xylose and 3 µg of activated Cry1C toxin to overcome the B. thurinngiensis 407 strain being cry negative. The larvae in the control group were fed PBS buffer or Cry1C toxin and xylose in corresponding amounts to the samples containing bacterial inocula. The chosen dose was expected to result in about 70% (± 5%) mortality at 37 °C after 48 hours.

Biofilm assays
The ability to form biofilms was determined using a glass tube screening assay (Houry et al., 2010). Briefly, exponential phase cultures were diluted into HCT medium (Lecadet et al., 1980;Lereclus et al., 1982) to an OD600 of 0.01, and 2 ml was inoculated into sterile 6 ml glass tubes. The tubes were incubated for 72 hours at 37 °C. The biofilm was subsequently collected by removing the culture medium with a Pasteur pipette and thoroughly vortexing in 2 ml PBS before measuring the OD600 of the suspension of biofilm cells. Each strain was tested in five biological replicates, each with 3 or 4 technical replicates.
The ability to form biofilms in polyvinylchloride (PVC) microtiter plates was determined using a crystal violet biofilm screening assay (Auger et al., 2006). Briefly, fresh bactopeptone medium was inoculated with 0.5% exponential phase culture, transferred to 96well plates (Falcon #353911) and incubated for 24, 48 and 72 hours at 30 °C. The biofilm was subsequently washed using PBS, stained using 0.3% crystal violet, solubilized with acetone:methanol (1:3 v/v), and transferred to flat-bottomed microtiter plates (Falcon #353915) for determination of the absorbance of the solubilized dye at 570 nm. Each strain was tested in triplicate. Statistical analyses were performed separately for each time point, as described under "Statistical analyses" below.

Expression of MBP-tagged MogR in E. coli
The sequence encoding MogR from B. thuringiensis 407 (BTB_RS08390) was PCR amplified using primers 5'-atgtatcaccacacagcaattaatgtattag-3' and 5'gcgcggatccttattactgtgttacggtcataacttgtcc-3' and genomic DNA as the template. The PCR product was cloned into the pMAL-p5x vector (New England Biolabs) using the XmnI and BamHI restriction sites (underlined) according to the manufacturer's instructions, allowing expression of MBP-MogR. A construct expressing the MBP-MogR QN→AA protein was created by site-directed mutagenesis of pMAL-p5x-mogR using the QuikChange II Site-Directed Mutagenesis Kit (Stratagene) with the same primers as for pHT304-Pxyl-MogR QN→AA . Constructs were transformed into E. coli BL21(DE3) cells. The plasmids were verified by sequencing. MBP-MogR and MBP-MogR QN→AA proteins were expressed in E. coli BL21(DE3) and purified using the manufacturer's manual "Method I -Total cell extract". In short, 100 ml LB broth was inoculated with a 1 ml overnight culture containing cells harbouring fusion plasmid. Cells were grown at 37 °C with shaking to an OD600 of ~0.5. IPTG was added to a final concentration of 0.3 mM and cells were induced for 2 hours. The cells were harvested by centrifugation at 4500 × g for 10 minutes and frozen at -20 °C overnight, sonicated in an ice-water bath, and then centrifuged at 20,000 × g for 20 minutes at 4 °C. The supernatants were diluted 1:6 in Column Buffer (20 mM Tris-HCl, 200 mM NaCl, 1 mM EDTA). MBP-MogR proteins were purified by amylose affinity chromatography (Poly-Prep Chromatography column, 0.8 × 4 cm, Bio-Rad). The identity and purity of proteins was confirmed by SDS-PAGE (Coomassie stain) and the quantity determined by Bradford assay.

Electrophoretic mobility shift assay (EMSA)
DNA probes were generated by PCR using chromosomal DNA from B. thuringiensis 407 and the following primer combinations: 5'-cgggtgtcaactaaaaattcg-3' / 5'caactatcataatatcaccttttcgg-3' (PflaA probe), 5'-ttacagaaatgaaatttacggataac-3' / 5'ccttatcctttctgtctggtc-3' (PhblC probe), 5'-tccgtatgtaattccgtttcaaga-3' / 5'-aatttcctgcttgacccctt-3' (Pctrl1 probe). Corresponding 5' biotinylated probes were generated for flaA and hblC with the same primers with biotin labelling. ctrl1 was chosen as a control for binding to B. thuringiensis DNA and is a PCR product from inside a randomly chosen gene from B. thuringiensis 407 (BTB_RS08990 encoding an elongation factor G-binding protein). A commercial EMSA kit (LightShift Chemiluminescent, Thermo Scientific) was used to detect the binding of MogR protein to DNA following the manufacturer's instructions. Unlabelled Epstein Barr nuclear antigen DNA supplied with the kit was used as negative non-Bacillus DNA control. DNA binding reactions were conducted in 20 ml of 10×Binding Buffer (100 mM TrisHCl, 500 mM KCl, 10 mM DTT, pH 7.5) with 50 ng µl -1 polydeoxyinosinicdeoxycytidylic acid (poly dI-dC). 1 µg of MBP-MogR or MBP-MogR QN→AA were added to approximately 2 fmol biotinylated DNA probe and incubated for 20 minutes at room temperature. A total of 20 ml of each binding reaction was loaded on a 5% SDS-PAGE gel (0.5×TBE) and resolved for 1 hour at 100 V.

Quantification of cyclic di-GMP by LC-MS/MS
LC-MS/MS analysis was based on a method described by Spangler et al. (2010), with some modifications, as previously described (Fagerlund et al., 2016), using a Thermo Scientific LTQ XL Linear Ion Trap Mass Spectrometer (Thermo Scientific) and separation on a 5 mm × 1 mm I.D. Nucleodur C18 Pyramid precolumn and a 50 mm × 1 mm I.D. Nucleodur C18 pyramid analytical column (both from Marchery-Nagel), with an electrospray ionization (ESI) source operated in the positive ionization mode to interface the HPLC and the MS.

Statistical analyses
Minitab v.17 software was used for statistical analysis. Unless stated otherwise, all data were analyzed using analysis of variance (ANOVA) followed by Tukey's post hoc test for pairwise comparisons. P-values <0.05 were regarded as significant.

The motility gene loci in B. thuringiensis and L. monocytogenes are closely related
A cluster of approximately 45-50 genes with homology to flagellar-based motility-and chemotaxis genes is present in most B. cereus group strains. Although B. cereus group bacteria are phylogenetically more closely related to B. subtilis than to L. monocytogenes, some aspects of organization of the B. thuringiensis 407 motility gene cluster were found to more closely resemble that of L. monocytogenes. Bidirectional BLAST analysis showed that a greater number of motility protein orthologs is shared between B. thuringiensis 407 and L. monocytogenes EGD-e (41 orthologous proteins, average alignment percentage identity 46%) than between B. thuringiensis 407 and B. subtilis 168 (31 orthologous proteins, 37% average sequence identity) (Supplementary Table S1). Furthermore, only other Listeria spp. and species of the B. cereus group are known to have orthologs to the L. monocytogenes motility gene repressor MogR (Gründling et al., 2004;Smith and Hoover, 2009), potentially indicating that elements of motility regulation may be shared between these species (Figure 1 and Supplementary Table S1).

Non-motile B. anthracis and B. pseudomycoides have retained mogR
As has previously been noted (Klee et al., 2010), comparative alignments between the motility loci of different B. cereus group strains indicate that the flagellin gene locus is an evolutionary hotspot, as the number of flagellin genes in each strain varies between one and five copies, and several strains also contain additional sets of genes in this locus. Isolates belonging to the B. cereus group may be divided into seven phylogenetic subgroups (I to VII) and for which, with the exception of B. pseudomycoides (group I) and B. cytotoxicus (group VII), species designations do not strictly follow the phylogenetic group designations. Instead, isolates belonging to the different phylogenetic subgroups have distinct ecotypes and preferred growth temperature ranges (Guinebretière et al., 2008). An analysis of B. cereus group genomes revealed striking differences between the motility gene clusters in strains belonging to the various phylogenetic groups (Figure 2). The motility gene clusters of strains in phylogenetic subgroups III, IV and V were overall highly similar to that of B. thuringiensis 407, with a few exceptions mainly related to insertion of transposable elements and variation in the number of flagellin genes. Of note, strains of B. anthracis, which constitutes a highly clonal lineage within subgroup III, all carry an 11.5 kb insertion in place of the second flagellin gene ( Phylogenetic subgroup I corresponds to the species B. pseudomycoides, but also includes strains for which an original identification as B. cereus or B. mycoides has not yet been revised (Guinebretière et al., 2008). BLASTP searches were performed against available subgroup I proteomes using the corresponding protein sequences for the genes from the motility locus of B. thuringiensis 407 as query (Figure 2). Among six fully sequenced strains (single scaffold per replicon or closed genome) belonging to subgroup I (219298, BTZ, Rock 3-44, DSM 12442, Rock 1-4 and Rock 3-17), all contained a severely reduced motility gene cluster, and strikingly, the only conserved motility-related gene in the motility locus of these strains is mogR (Figure 2). The absence of all motility genes except mogR in all six fully sequenced group I strains analyzed, as well as in all except for one (AFS092012) of the 100 additional group I strains annotated as B. pseudomycoides at the NCBI and for which a For each strain, the first and last gene (shown in red) are conserved genes outside of the motility gene locus itself (orthologs to BTB_RS08235 and BTB_RS08480, respectively). These are included to indicate that the flanking regions and location of the motility locus are generally conserved between strains. The last gene of the intact locus with a predicted function mapping within motility, flgG (light blue), is indicated by a filled square above the gene. For genes shown in a color other than red, identical color indicates that genes are from the same orthologous group (top COG hit), except light yellow which indicates no COG assignment. The gmaR gene is indicated with a red box. In the B. anthracis strain, the filled triangles above genes indicate the location of nonsense or frameshift mutations. The brackets in the representation of the Bc Rock 3-44 sequence indicates that this locus is split between two contigs. Roman numerals next to the strain names indicate the phylogenetic subgroup within the B. cereus group to which the strain belongs (Guinebretière et al., 2008 genome sequence at scaffold level was available, indicates that absence of motility is likely a general characteristic of this genetic group, in line with the description of B. pseudomycoides as a nonmotile species (Nakamura, 1998). The predicted MogR proteins of the B. pseudomycoides group show ~56% and ~30% pairwise identity to the corresponding orthologs in B. thuringiensis 407 and L. monocytogenes EGD-e, respectively. A multiple sequence alignment demonstrated that the L. monocytogenes MogR residues predicted to make base-specific DNA contacts in L. monocytogenes (Shen et al., 2009) have been conserved in the MogR proteins from all examined B. cereus group strains, including those belonging to non-motile species (Supplementary Figure S1). Interestingly, an ortholog of gmaR, encoding a temperature-controlled antirepressor of MogR in L. monocytogenes (Kamp and Higgins, 2009), is found in B. cytotoxicus strains (cluster VII), but is absent in most B. cereus group strains belonging to other phylogenetic clusters.

Flagellin gene expression drops following a sharp increase in mogR expression
B. thuringiensis 407 produces two flagellins, structural proteins building the flagellum main filament, from chromosomal genes flaA and flaB . Expression of mogR (BTB_RS08390), flaA (BTB_RS08400) and flaB (BTB_RS08395) was followed at the transcriptional level throughout the bacterial growth phase using RT-qPCR. In contrast to previous proteomics studies which indicated that in B. thuringiensis 407 only the first flagellin gene (flaA) was expressed in early stationary phase , results showed that both flaA and flaB are indeed expressed at the RNA level, although expression of flaA was approximately four-fold higher than that of flaB ( Figure 3). The two fla genes reached maximum expression at 3.5 hours, at the transition between exponential and stationary growth phase, in agreement with corresponding data obtained using a transcriptional fusion between lacZ and the flaA promoter (Houry et al., 2010). These data also corresponded well with microscopic observations of cultures sampled throughout the growth phase. The expression of mogR increased 64-fold between the 2.5 hour and 3.5 hour time-point (Figure 3), peaking at 4 hours, at which time flaA and flaB expression was rapidly decreasing. This is in parallel with the expression pattern found for mogR in B. anthracis (Bergman et al., 2006). While downregulation of motility-related genes upon entry to the stationary phase is common in both Gram-negative and Gram-positive bacteria (Ramirez Santos et al., 2005;Han et al., 2017), it was nevertheless interesting to note that expression of the flaA and flaB genes dropped after 3.5 hours, subsequent to the sharp increase in mogR transcription ( Figure 3).

Overexpression of MogR renders B. thuringiensis 407 non-flagellated and nonmotile
A MogR overexpression strain was constructed using the pHT304-Pxyl expression vector carrying a xylose-inducible promoter, as well as an isogenic strain overexpressing a version of MogR where two mutations were introduced: Q119A and N120A (plasmids and strains used are listed in Table 1). In the mutated protein (referred to as MogR QN→AA ), alanine substitutions were made for two of the six amino acids corresponding to residues in L. monocytogenes MogR that were shown to make base-specific contacts with DNA, in cognate DNA recognition sites positioned upstream of L. monocytogenes genes that were subject to direct MogR transcriptional repression (Shen et al., 2009;Supplementary Figure S1). Cellular growth rates were not affected by MogR or MogR QN→AA overexpression (relative to an empty vector control) at 25 °C, 30 °C or 37 °C (Supplementary Figure S2).
Motility of the MogR overexpression strain was investigated by swimming assays employing 0.3% LB agar plates, showing that cells overexpressing MogR were non-motile, and that the repression of motility was dependent on the two amino acid residues (Q119 and N120) mutated in MogR QN→AA , at 30 °C ( Figures 4A,B), as well as at 25 °C and 37 °C. Swimming  Table 1). Expression was induced with 1 mM xylose and cells were grown at 30 °C. motility was also monitored by light microscopy and found to be completely lost upon MogR overexpression at all time points and temperatures examined (every half hour between 1 and 7 hours of cultivation, at 25 °C, 30 °C and 37 °C). To determine whether the reduced motility was due to a loss of flagellar structure or flagellar rotation, strains were analyzed by Atomic Force Microscopy (AFM). AFM analyses showed that the MogR overexpression strain was completely devoid of flagellar structures, while overexpression of MogR QN→AA restored the empty vector control phenotype ( Figures 4D,E,F). In accordance, a Western blot of the MogR overexpression strain using anti-flagellin antibodies showed that flagellin proteins were below the level of detection in whole cell extracts and severely attenuated in outer cell surface extracts compared to in the vector control strain, while being expressed at comparable levels in the B. thuringiensis 407 wild type strain and in B. thuringiensis 407 overexpressing MogR QN→AA ( Figure 4G). In a ΔflaAB negative control strain in which the genes encoding the FlaA and FlaB flagellins had been deleted, no flagellin proteins could be detected neither in the whole cell extract nor on the cell surface. In L. monocytogenes, flagellin genes were previously found to be directly repressed by MogR, and in order to investigate if MogR function was conserved in B. thuringiensis 407, the expression of flagellin genes was investigated in the MogR overexpression strain relative to the empty vector and MogR QN→AA control strains, by quantitative RT-PCR. A 30-fold (flaA) and 50-fold (flaB) reduction in transcription, respectively, was seen in the MogR overexpression strain relative to the empty vector control, while no difference was observed relative to the empty vector strain when overexpressing MogR QN→AA ( Figure 4C).

Overexpression of MogR affects the expression of genes not related to motility functions
Global microarray-based transcriptional profiling was used to identify genes whose expression was affected by MogR overexpression in B. thuringiensis 407 relative to the empty vector control strain. Samples were taken in the early stationary growth phase (3 hours), and a total of 110 genes were identified to be differentially regulated by MogR (FDR-corrected p<0.05), either directly or indirectly. Selected genes are listed in Table 2 (see Supplementary  Table S3 for complete list). Of the differentially expressed genes, 87 were repressed by MogR overexpression, while 23 genes exhibited higher transcriptional levels compared to the control. In total, 21 motility genes were downregulated by MogR overexpression, including flagellin genes flaA and flaB (Supplementary Figure S3). Overexpression of MogR affected genes widely dispersed in the motility loci, including the motAB operon located separately in the chromosome. Interestingly, the microarray analysis revealed that MogR also affected the expression of 89 non-motility-related genes, including six virulence genes which were downregulated. The latter genes included inhA encoding immune inhibitor A, hblA and hblD encoding enterotoxin binding component B and lytic component L1, respectively, sfp encoding a subtilase family serine protease, as well as genes encoding phosphatidyl-choline specific phospholipase C and sphingomyelinase. Also downregulated were five stress-related genes: the groES, groEL and grpE chaperonin genes, sigB encoding the  B alternative sigma factor, and hrcA. The gene encoding the pleiotropic transcriptional regulator NprR (Dubois et al., 2012) was also downregulated. Among the 23 genes upregulated in the MogR overexpression strain were sinI, encoding a protein inducing biofilm formation by direct interaction and sequestration of the biofilm repressor SinR, and a putative capsular polysaccharide biosynthesis protein-encoding gene (BTB_RS26935) corresponding to BC5278 in B. cereus strain ATCC 14579. The genes BC5267 to BC5278 constitute a conserved locus in B. cereus and B. thuringiensis and are homologous to the epsAO locus of B. subtilis involved in the synthesis of the exopolysaccharide component of the biofilm matrix (Branda et al., 2001;Ivanova et al., 2003;Kearns et al., 2005;Fagerlund et al., 2014). Genes in this locus have been found to be implicated in biofilm formation in B. cereus in a pellicle biofilm model (Hayrapetyan et al., 2015;Okshevsky et al., 2017). Another upregulated gene was cbpA, encoding a collagen adhesion protein positively regulated by c-di-GMP (Finke et al., 2019).  Shen et al., 2009). Three putative binding sites were predicted in the intergenic region upstream of the flagellin (flaAB) operon, two potential sites upstream of cheY, while four putative binding sites were predicted in the promoter region of the hbl operon (Lindbäck et al., 1999), all overlapping the -10 promoter element ( Figure 5A). Among the other genes downregulated by MogR overexpression, multiple binding sites were found upstream of inhA, as well as upstream of the biofilm-promoting gene sinI and mogR itself. To investigate whether MogR may affect the expression of candidate genes in B. thuringiensis 407 by directly binding to upstream regulatory regions, we carried out electrophoretic mobility shift assays (EMSA) with a purified maltose-binding protein (MBP)-MogR fusion protein, and purified DNA fragments constituting PCR-amplified candidate regulatory regions ( Figure 5A). Results showed that MogR could bind the promoter regions upstream of both flaAB and the hbl enterotoxin locus in vitro ( Figures 5B,C), while for MogR QN→AA DNA binding was abolished ( Figure 5B), demonstrating the essentiality of the Q119 and N120 residues for DNA-binding. The binding reactions to which competing cold DNA were added (labelled ctrl1 and ctrl2 in Figures 5B,C) indicated that MogR exhibits sequence specific binding to flaAB and hbl upstream regions in vitro.

Cytotoxin production and in vivo virulence are reduced upon MogR overexpression
Western blot analyses were performed to measure the effect of MogR overexpression on the amount of secreted enterotoxins. Hbl, CytK and Nhe enterotoxins were all present but at clearly reduced levels in the culture supernatant from the MogR overexpression strain, compared to that of empty vector and MogR QN→AA control strains ( Figure 6A). To further assess the effect of MogR on virulence capacity, supernatants collected from mutant and overexpression strains grown to exponential phase were tested in an in vitro Vero cell toxicity assay (Lindbäck and Granum, 2006). Supernatants from the MogR overexpressing strain, which synthesized lower amounts of the Hbl, Nhe and CytK toxins ( Figure 6A), were less cytotoxic than those of the empty vector control (p=0,059) and the MogR QN→AA strain ( Figure  6B). In order to more confidently assess the effect of MogR on virulence, an in vivo model was used, where the strains were tested in an oral infection model using Galleria mellonella larvae. In accordance with the in vitro toxicity data, the MogR overexpressing strain was severely attenuated in virulence compared to the empty vector control strain in vivo (p=0.02; Figure 6C).

Biofilm formation is increased upon MogR overexpression in a flaAB negative background
The MogR overexpression strain was compared to the empty vector and MogR QN→AA strains both in glass tube and microtiter plate biofilm assays ( Figure 7A,B). The glass tube assay showed similar development of a biofilm pellicle at the air-liquid surface in all three strains: After 72 hours the pellicle in the MogR overexpression strain was only slightly (14%) thicker than that of the empty vector and MogR QN→AA strains ( Figure 7A). In the microtiter plate assay, no significant effect of MogR overexpression on biofilm formation was observed at 48 or 72 hours, with a slight decrease in biofilm formation at 24 hours compared to MogR QN→AA and empty vector controls ( Figure 7B). Overexpression in a ΔflaAB background however allowed examination of the effect of the MogR protein on biofilm formation, independently of any effect conferred through flagella. Interestingly, in the ΔflaAB host, overexpression of MogR led to a substantial increase in biofilm formation relative to the MogR QN→AA and empty vector controls ( Figure 7C), while biofilm formation in the control strain overexpressing MogR QN→AA was comparable to that of the strain carrying empty vector. Intriguingly, LC-MS/MS quantitation of the biofilm-related second messenger cyclic di-GMP (c-di-GMP) in cellular extracts indicated slightly increased levels of c-di-GMP in the MogR overexpression strain (wild type background; three measurements: 8.38 ng ml -1 ; 6.0 ng ml -1 ; 9.0 ng ml -1 ) compared to the B. thuringiensis 407 empty vector strain (two measurements: < LOD; 6.0 ng ml -1 ) (Limit of detection, LOD: 0.8 ng ml -1 ; Limit of quantitation, LOQ: 3.5 ng ml -1 ).

Discussion
Overexpression of MogR leads to repression of a range of motility genes in B. thuringiensis 407, including the flagellins, rendering the bacteria non-flagellated and non-motile. In addition, amino acids in B. thuringensis MogR which correspond to those shown to constitute DNA backbone-interacting residues in L. monocytogenes MogR are clearly essential for B. thuringiensis MogR DNA binding in vitro. In accordance, overexpression of the mutated variant of MogR restores phenotypes to those of a vector control strain. Collectively these results, and analyses of MogR binding to predicted regulatory sites in vitro, strongly support the role of B. thuringiensis MogR as a DNA-binding transcriptional regulator and that the role of MogR as a key motility regulator in B. thuringiensis is conserved with that in L. monocytogenes. Global gene expression profiling however showed that in B. thuringiensis a large number of non-motility genes were also affected by MogR overexpression, including genes involved in stress response, virulence and biofilm formation. This was paralleled by an observed decrease in cytotoxin production, Vero cell toxicity and in vivo toxicity, indicating that MogR can affect additional phenotypes in B. cereus group bacteria. This is in apparent contrast to L. monocytogenes, where the primary function of MogR during intracellular infection is to repress flagellar motility gene expression, and where deletion of the flagellin gene (flaA) in a mogR deletion strain (producing ΔmogR ΔflaA) restores the LD50 values to wild type levels in a mouse model of infection .
In bacteria, motility and virulence can be subject to coordinated downregulation, e.g. as is observed for intracellular signaling networks involving the second messenger c-di-GMP (Fagerlund et al., 2016). Flagellar motility and virulence appears to be co-regulated in B. cereus group bacteria Ghelardi et al., 2002;Bouillaut et al., 2005;Ghelardi et al., 2007a;Fagerlund et al., 2010;Salvetti et al., 2011;Mazzantini et al., 2016), and a B. cereus strain deleted in flhA, which encodes a component of the flagellum-formation apparatus, is impaired in adhesion (Ramarao and Lereclus, 2006), expression of virulence factors (Ghelardi et al., 2002;Fagerlund et al., 2010), and in cytotoxicity and in vivo virulence independently of PlcR (Bouillaut et al., 2005). Similarly, deletion of the gene encoding the signal recognition particle-like GTPase FlhF, located directly downstream of flhA, resulted in a marked reduction in the number of flagella, and a reduction in secretion of several virulence-related proteins with attenuation of in vivo virulence (Salvetti et al., 2007;Mazzantini et al., 2016;Mazzantini et al., 2020). A link between synthesis of the flagellar apparatus and the production of virulence factors has also been observed for flhA deletion mutants in other bacteria that do not carry a mogR ortholog (Fleiszig et al., 2001;Carrillo et al., 2004). The observed effects of MogR on toxin production and virulence-related phenotypes may thus be conferred through direct regulation, as potentially indicated through in vitro binding to the hbl upstream region, or indirectly through repression of genes in the motility locus, such as e.g. flhA and/or flhF, or both. MogR appears to function independently of the virulence gene activator PlcR, as plcR expression was not affected by MogR overexpression, and only six out of the 28 genes comprising the PlcR regulon were differentially expressed in the microarray analysis. Clearly, the effect on toxin production observed following deletion of flhA or flhF was not replicated in the flaAB deletion mutant, showing that not all deletions in the motility locus affect toxin synthesis and/or transport.
When employing a ΔflaAB genetic background to compare the effect of MogR overexpression on biofilm formation, relative to empty vector and MogR QN→AA overexpression controls, a significant increase in biofilm formation was observed by MogR overexpression. In line with this, the transcriptional profiling experiments showed upregulated expression of the biofilm anti-repressor sinI, and BTB_RS26935 located in a putative polysaccharide locus previously shown to be implicated in the formation of the biofilm matrix in some B. cereus strains (Hayrapetyan et al., 2015;Okshevsky et al., 2017). Moreover, LC-MS/MS quantitation indicated a slight increase in cellular c-di-GMP upon MogR overexpression, which is linked to increased biofilm formation in B. thuringiensis 407 (Fagerlund et al., 2016;Fu et al., 2018). A microarray experiment analyzing the effect of B. thuringiensis 407 overexpressing the c-di-GMP synthesizing protein CdgF relative to an empty vector control strain (Fagerlund et al., 2016;A. Fagerlund, unpublished data deposited in the ArrayExpress database, accession no. E-MTAB-8898) showed that the expression of the biofilm regulators spo0A and sinI were upregulated and the transcriptional regulator codY was downregulated at high levels of c-di-GMP, all paralleling an increase in biofilm formation (Pflughoeft et al., 2011;Lindbäck et al., 2012;Fagerlund et al., 2014). Taken together, these experiments provide strong indication that MogR, directly or indirectly promotes biofilm formation in B. thuringiensis 407, a phenotype in line with the role of the protein in promoting sessility and repressing toxicity.
In L. monocytogenes, MogR activity is controlled by its anti-repressor GmaR (Kamp and Higgins, 2009). GmaR expression is halted when L. monocytogenes experiences 37 °C, which allows MogR to repress flagellar synthesis during human infection. The gmaR gene is not present in most B. cereus group genomes, in line with mogR appearing to be regulated in a growth-dependent but temperature-independent manner in B. thuringiensis 407. It is however interesting to note that a gmaR ortholog is found in at least twelve sequenced B. cereus group strains, including all hitherto characterized strains of the thermotolerant species B. cytotoxicus, perhaps reflecting a need for strains in this phylogenetic group (group VII) to regulate flagellin synthesis at different temperatures. Most B. anthracis strains also harbor a copy of gmaR, which however is truncated and rendered non-functional due to mutation. Uncharacteristically, an apparently full-length gmaR copy was identified in the genome of B. cereus VD184, an environmental soil isolate belonging to phylogenetic cluster IV (Hoton et al., 2009;Jalasvuori et al., 2013), as well as B. cereus strains RU36C, F528-94, MB.22, B4118 and FSL W8-0932, and Bacillus wiedmannii FSL P4-0569, for which optimal growth temperatures are not known.
In conclusion, MogR is a key regulator of motility in B. thuringiensis 407, a function which is probably conserved across the motile species in the B. cereus group. Most importantly however, both microarray experiments, protein expression analyses, biofilm screening results, and the fact that the mogR gene is evolutionary retained in the non-motile B. mycoides, B. pseudomycoides, and the highly virulent B. anthracis, as well as expressed in B. anthracis, speaks to the MogR transcriptional regulator serving additional functions in B. cereus group bacteria, beyond those identified in L. monocytogenes. In this respect it is interesting to note that all attempts to produce a gene deletion mutant in mogR in B. thuringiensis 407 were unsuccessful, and we cannot rule out the possibility of mogR constituting an essential gene in the B. cereus group. From the analyses performed in this study, it is tempting to speculate that MogR may serve as part of a molecular system coordinating motility, virulence, and biofilm formation, and possibly additional phenotypes. Biofilm formation and motility are reciprocally regulated in a number of bacterial species, including B. cereus group bacteria, by the second messenger c-di-GMP (Hengge, 2009;Fagerlund et al., 2016;Jenal et al., 2017;Fu et al., 2018), and several transcriptional regulators in both the B. cereus group and L. monocytogenes are known to have overlapping functions in regulation of virulence and biofilm formation (Hsueh et al., 2006;Lemon et al., 2010;Pflughoeft et al., 2011;Frenzel et al., 2012;Lindbäck et al., 2012;Fagerlund et al., 2014;Slamti et al., 2015;Böhm et al., 2016). It thus appears that, in the B. cereus group, there is substantial cross-regulation between motility, virulence, and biofilm formation, involving a multitude of regulatory networks based on both c-di-GMP and transcription factors (PlcR, MogR, NprR).

Data availability statement
The microarray data comparing overexpression of MogR with the empty vector control is available in the ArrayExpress database (www.ebi.ac.uk/arrayexpress) under accession number E-MTAB-7633.

Author Contributions
OØ and AF contributed to conception and design of the study. VS performed EMSA, motility assays, Western immunoblotting, and biofilm assays. VS and AF performed RT-qPCR. VS, MJ and AF performed cloning and mutagenesis. MJ performed the microarray experiment, SF performed growth experiments, IH performed AFM, TL performed cytotoxicity assays, and CN performed the in vivo toxicity assays. AF performed bioinformatic, microarray, and statistical analyses and NT performed bioinformatic analysis of the subgroup I genomes. VS, OØ, and AF wrote the first draft of the manuscript. TL, IH, and CN wrote sections of the manuscript. All authors contributed to data analysis, read and approved the final manuscript.

Funding
This work was funded by a project grant from the Research Council of Norway to OAØ through the FUGE II Programme (channel 3 grant; project number 183421), the Jahre Foundation, and by internal grants from the Department of Pharmacy, University of Oslo to OAØ.

Conflict of Interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be interpreted as a potential conflict of interest.