CsrA-Mediated Translational Activation of ymdA Expression in Escherichia coli

The Csr system of E. coli controls gene expression and physiology on a global scale. CsrA protein, the central component of this system, represses translation initiation of numerous genes by binding to target transcripts, thereby competing with ribosome binding. Variations of this mechanism are so common that CsrA is sometimes called a translational repressor. Although CsrA-mediated activation mechanisms have been elucidated in which bound CsrA inhibits RNA degradation, no translation activation mechanism has been defined. Here, we demonstrate that CsrA binding to two sites in the 5′ untranslated leader of ymdA mRNA activates translation by destabilizing a structure that otherwise prevents ribosome binding. The extensive role of CsrA in activating gene expression suggests the common occurrence of similar activation mechanisms.

aminoglycoside antibiotic apramycin (27). A ymdA knockout strain had no significant effect on biofilm formation, but it exhibited twofold increased resistance to apramycin (27).
In this study, we determined that CsrA activates ymdA translation by binding to two sites in the ymdA leader RNA, one of which is present in a structure that sequesters the SD sequence. We show that upon CsrA binding, this hairpin is destabilized, and the SD sequence becomes single stranded and available for ribosome binding. These findings provide direct evidence for CsrA-mediated translational activation of ymdA expression.

RESULTS
CsrA binds to two sites in the ymdA leader transcript causing destabilization of a SD-sequestering hairpin. Previous CLIP-seq results indicated that CsrA binds to the leader region of the ymdA transcript (6). Four GGA motifs were identified within 90 nucleotides (nt) of the translation initiation codon; GGA is a critical component of a CsrA binding site (Fig. 1). Quantitative gel mobility shift assays were performed to investigate the interaction of CsrA with ymdA leader RNA containing all four GGA motifs (Ϫ100 to Ϫ1 relative to the start of ymdA translation). A distinct band indicative of bound CsrA was observed between 8 and 64 nM CsrA, with a second shift appearing at 125 nM CsrA and above ( Fig. 2A). A nonlinear least-squares analysis of this data yielded a K d (dissociation constant) value of 48 Ϯ 3 nM CsrA, indicating that CsrA binds to the ymdA transcript with moderate affinity. While the stoichiometry of each bound state was not investigated, it is likely that the first bound state (B1) is a single CsrA dimer bound to a single transcript, while the second bound state (B2) may contain two CsrA dimers bound to the same transcript. The specificity of this interaction was assessed by competitive gel mobility shift assays in the presence of specific (ymdA) or nonspecific (phoB) unlabeled RNA competitors in 10-and 100-fold excess to the radiolabeled ymdA transcript. With a CsrA concentration of 250 nM, unlabeled ymdA RNA competed for binding of CsrA to the radiolabeled ymdA transcript, whereas the nonspecific competitor did not, indicating that the interaction of CsrA with the ymdA RNA is specific (Fig. 2B).
CsrA-ymdA RNA footprint experiments were performed to identify CsrA binding sites. RNase T1, which cleaves RNA following single-stranded G residues, was used as the probe. Since we identified four potential CsrA binding sites in the ymdA leader region (Fig. 1), all four of the GGA sequences were present in the RNA used in this analysis. Bound CsrA protected the G residues in two GGA motifs from RNase T1 cleavage, indicating that these GGA motifs are components of authentic CsrA binding sites, which we refer to as BS1 and BS2 from hereon ( Fig. 3A and B). Cleavage of the G residues in BS2 was much less efficient than in BS1 even in the absence of CsrA ( Fig. 3A and B). This observation is consistent with an RNA secondary structure that partially sequesters BS2 in a hairpin (Fig. 3C). We did not observe CsrA-dependent protection of the other two GGA sequences, including the GGA motif that overlaps the ymdA SD sequence ( Fig. 3A and B). Of particular importance, RNase T1 cleavage of the G residues in the ymdA SD sequence actually increased in the presence of bound CsrA. Footprint experiments using a transcript containing a BS2 mutation (GGA to AGA) indicated that CsrA is capable of binding to BS1 in the absence of BS2 ( Fig. 3D and E). From these data, we conclude that CsrA binds to BS1 and BS2, which destabilizes the ymdA SDsequestering hairpin.
CsrA activates ymdA translation by promoting 30S ribosomal subunit binding. Primer extension inhibition (toeprint) experiments were performed to test whether bound CsrA affects ribosome binding. In this assay, reverse transcriptase will stop just downstream of a bound protein or stable RNA secondary structure. Two bands at positions ϩ12U and ϩA37 were observed in all experimental lanes (Fig. 4, lanes 2 to 7). RNA structure predictions with Mfold indicate that both of these bands correspond to the base of short RNA hairpins. In the absence of bound CsrA or 30S ribosomal subunits, two adjacent RNA structure-dependent toeprints were identified at positions Ϫ11A and Ϫ12G (Fig. 4A, lane 2), which are located at the base of the SD-sequestering hairpin (Fig. 3C), indicating that this hairpin forms in the absence of bound CsrA. Two adjacent CsrA-dependent toeprints were observed at nucleotides Ϫ24C and Ϫ25A (Fig. 4A, lane 3), which is at the 3= boundary of BS2 (Fig. 3C). To identify the position of bound 30S ribosomal subunit and whether binding was dependent on the presence of CsrA, a ribosome toeprint was also performed. In the absence of CsrA, a smear of reverse transcriptase stops was identified along the 3= stem of the SD-sequestering hairpin (Fig. 4A, lane 5). This could be a consequence of the 30S ribosomal subunit attempting to access the sequestered SD sequence but failing to make the sequence-specific contacts required for productive binding. Importantly, CsrA-dependent 30S ribosomal subunit toeprints were observed at positions 16U and 18A; the toeprint at 16U is the expected position 15 nt downstream from A of the ymdA start codon (Fig. 4A, compare lanes 5 to 7) (19)(20)(21). These results indicate that bound CsrA promotes 30S ribosomal subunit binding by destabilizing the ymdA SD-sequestering hairpin. One intriguing result that we cannot explain is the absence of bands beyond ϩ12U in the lane containing bound ribosomes (Fig. 4, lane 7). Since the ϩ12U band corresponds to transcripts that did not have a bound 30S ribosomal subunit, we would have expected to see all of the longer bands observed in the control lane (Fig. 4, lane 2).
We next utilized the in vitro coupled transcription-translation PURExpress system to determine whether CsrA activates ymdA translation. Three different plasmids carrying a ymdA'-'lacZ translational fusion driven by identical T7 RNA polymerase (RNAP) promoters were used in this analysis. One plasmid contained the WT ymdA leader sequence, while the other two contained a mutation in BS1 (GGA to AGA) or BS2 (GGA to AGA). The expression level of the WT fusion increased with increasing CsrA concentrations CsrA activates ymdA expression posttranscriptionally in vivo. To determine whether CsrA activates ymdA expression in vivo, we monitored expression of a chromosomally integrated P ymdA -ymdA'-'lacZ translational fusion from mid-exponential to early stationary-phase growth in WT and CsrA-deficient (csrA::kan) strains. This fusion contained sequences between Ϫ300 to ϩ4 relative to the start of ymdA translation. The csrA::kan allele contains a transposon insertion following the 50th codon of CsrA's 61-amino-acid coding sequence, resulting in a 62-amino-acid fusion protein that retains ϳ12% of the RNA binding affinity of WT CsrA (24). Expression of the WT translational fusion was 10-to 30-fold higher in the WT strain throughout growth, indicating that CsrA activates ymdA expression in vivo ( Fig. 5A to C). Similar experiments were performed with a translational fusion in which the ymdA leader sequence from Ϫ211 to Ϫ132 was deleted such that it contained only the leader region used in our footprint analysis (Fig. 3). With the exception of a small decrease in expression during exponential-phase growth, the deletion fusion exhibited expression characteristics identical to those of the full-length WT translational fusion including 10-to 30-fold higher expression in the WT strain (Fig. 5C). These results indicate that the leader region used for our in vitro footprint studies is sufficient for CsrA-dependent activation of ymdA expression. When experiments were performed with a P ymdA -ymdA-lacZ transcriptional fusion in which the ymdA leader region is absent, expression levels were similar between the WT and csrA::kan strains, indicating that CsrA-dependent activation is not due to an indirect effect on transcription initiation ( Fig. 5A and D). We next tested a P lacUV5 -ymdA'-'lacZ leader fusion in which the ymdA promoter was replaced with a promoter that is unaffected by CsrA (4). Therefore, any effect of CsrA would occur after transcription initiation. As was observed with the P ymdA -ymdA'-'lacZ translational fusion, expression of the leader fusion was 10-to 20-fold higher in the WT strain throughout growth ( Fig. 5A and E). Taken together with our toeprint and in vitro translation results, we conclude that CsrA directly activates ymdA expression posttranscriptionally.
Since our footprinting and in vitro translation results identified BS1 and BS2 as critical CsrA binding sites, we tested the effect of single nucleotide substitutions in BS1 (GGA to AGA) and BS2 (GGA to AGA) in the context of the P ymdA -ymdA'-'lacZ translational fusion. Both of the individual binding site mutations resulted in the complete loss of the CsrA-mediated activation (Fig. 5F). We also tested whether a GGA-to-GAA mutation in the upstream GGA motif (Fig. 1, GGA*) affected ymdA expression. This mutation, which is predicted to maintain WT-like RNA secondary structure, had no effect on CsrA-mediated activation, and therefore, this sequence was ruled out as a CsrA binding site ( Fig. 3C and 5F).
CsrA stabilizes the ymdA transcript. Previous RNA-seq data indicated that ymdA RNA levels were 9-fold higher in the WT strain compared to the csrA::kan strain (6). Since our expression results excluded indirect effects of CsrA on ymdA transcription (Fig. 5) and because the stability of a transcript can be influenced by translation efficiency, we performed quantitative reverse transcriptase PCR (qRT-PCR) to analyze ymdA mRNA levels in WT and csrA::kan strains. Depending on the stage of growth, ymdA RNA levels were 3-to 7-fold higher in the WT strain (Fig. 6A). Consistent with these results, the mRNA half-life in mid-exponential-phase cultures was threefold greater in the WT strain (Fig. 6B). Whether the stability is a direct effect of CsrA binding or due to CsrAdependent translational activation was not investigated.
YmdA represses biofilm formation. A previous study reported that overexpression of ymdA inhibited biofilm formation, whereas a ymdA knockout strain had no significant effect on biofilm formation (27). Thus, we generated an unmarked deletion of ymdA and tested the effect of this mutation on biofilm formation. The ΔymdA strain resulted in a modest 45% increase in biofilm (Fig. 7). Since CsrA activates ymdA expression, we CsrA-Mediated Translational Activation ® reasoned that loss of activation by introducing a BS1 mutation would lead to increased biofilm. Thus, we generated a markerless BS1 mutation using CRISPR. We found that this mutation resulted in a small but reproducible increase in biofilm; however, a Student's t test did not support statistical significance of this effect. Complementation tests of ΔymdA with a plasmid clone of ymdA (pYmdA) versus an empty vector control failed to decrease biofilm formation (data not shown).

DISCUSSION
CsrA is a conserved global regulatory protein that binds to several hundred mRNAs in E. coli (6). CsrA represses translation by a variety of related mechanisms in which CsrA binds near the translation initiation region and/or SD sequence of target transcripts, thereby blocking 30S ribosome binding (1,2,(19)(20)(21). While translational repression is a well-characterized mechanism in bacteria, few examples of translational activation have been reported. Two prior studies provided evidence that CsrA (RsmA) is capable of activating translation, although the underlying molecular mechanisms remain unresolved (25,26). In this study, we elucidated the first CsrA-mediated translational activation mechanism. CsrA was already shown to activate flhDC expression by blocking

FIG 7
YmdA represses biofilm formation. Biofilm assays were performed in 96-well plates at 37°C in LB medium following the procedure described previously (30). Biofilm formation was quantified as crystal violet staining (A 630 ) over growth (A 600 ). Each column represents the average of results from three independent experiments (number of biological replicates n ϭ 14 total), with the error bars representing the standard errors of the means and asterisks representing the P value (**, P ϭ 0.01; ns, not significant). WT, MG1655; BS1, GGA-to-AAG mutation (PLB2839); ΔymdA, unmarked deletion of ymdA (AP3080).
RNase E access to the transcript (24). In addition, CsrA participates in Rho-dependent transcription attenuation mechanism (22). Thus, CsrA does not function solely as a translational repressor as has been suggested in the literature.
CsrA activates translation of ymdA by binding to two sites in the leader region of ymdA mRNA, one of which is partially sequestered in an RNA hairpin that also fully sequesters the SD sequence (Fig. 3C). CsrA binding to BS1 and BS2 ( Fig. 1 and 3C) destabilizes the ymdA SD-sequestering hairpin, leading to translational activation. RNA-seq data in WT and csrA::kan strains revealed a higher abundance of the ymdA transcript in the WT strain (6). We found that CsrA-dependent stabilization of ymdA mRNA contributes to that effect.
The function of YmdA is unknown, but a previous study reported that overexpression of ymdA inhibited biofilm formation, although a ymdA knockout strain had little to no effect on biofilm formation (27). We found that deletion of ymdA slightly, but reproducibly, increased biofilm formation (Fig. 7). Although introduction of a BS1 mutation that eliminates CsrA-mediated activation in vivo exhibited a small but reproducible increase in biofilm, this difference was not statistically significant. Furthermore, the effect of ΔymdA on biofilm was not complemented in trans. Perhaps the effects of ymdA and CsrA-mediated activation of ymdA on translation would be more substantial under different growth conditions than those used in our studies.
The connection between the E. coli Csr system and inhibition of biofilm formation is well documented (21,22,28,29). CsrA represses biofilm formation by inhibiting synthesis and secretion of the biofilm adhesin molecule poly-␤-1,6-N-acetyl-Dglucosamine (PGA) (21). CsrA regulates PGA levels by repressing expression of the pgaABCD operon, which encodes the cellular machinery required for synthesis, covalent modification, and secretion of PGA (30)(31)(32). CsrA represses translation initiation of the pgaA transcript by binding to a site overlapping the SD sequence and competing with the 30S ribosome (21). CsrA also participates in Rho-dependent termination of the pgaA transcript by destabilizing an RNA secondary structure that sequesters a Rho utilization (rut) site, resulting in transcript termination (22). This mechanism provides another example for how CsrA can destabilize RNA structure. CsrA also indirectly represses pgaABCD expression by repressing translation of nhaR, a transcriptional activator of the pgaABCD operon and GGDEF domain proteins that synthesize cyclic-di-GMP, which is the allosteric activator of PGA synthesis (28,29). Taken together, CsrA-mediated activation of ymdA translation might provide another connection between the Csr system and biofilm formation in E. coli, although the weak effects of ymdA itself on biofilm under our growth conditions leave this open to question. The location of ymdA immediately downstream from the csgBAC operon encoding genes involved in the production of curli fimbriae, which mediate an alternative pathway for E. coli biofilm formation (33), allows the possibility that ymdA may affect biofilm formation via this alternative pathway.

MATERIALS AND METHODS
Bacterial strains and plasmids. All E. coli strains used in this study are listed in Table 1. The E. coli strain S17-1 pir ϩ (34) was used for conditional replication, integration, and modular (CRIM)-based plasmid construction (35). The plasmids pLFT, pLFX, and pUV5 (4) were used to construct the translational, transcriptional, and leader fusions, respectively. Plasmid pAR2 contains a P ymdA -ymdA'-'lacZ translational fusion (Ϫ300 to ϩ4 relative to the ymdA start codon cloned into the PstI and BamHI sites of pLFT). Plasmid pYH370 is identical to pAR2 except that nucleotides Ϫ211 to Ϫ132 were deleted from the ymdA leader region using the QuikChange protocol (Agilent Technologies). Plasmid pAR3 contains a P ymdA -ymdA-lacZ transcriptional fusion (Ϫ300 to Ϫ221 cloned into the PstI and EcoRI sites of pLFX). Plasmid pAR4 contains a P lacUV5 -ymdA'-'lacZ leader fusion (Ϫ217 to ϩ4 cloned into the EcoRI and BamHI sites of pUV5) such that the promoter region of ymdA was replaced with the lacUV5 promoter. Plasmid pAR25 contains the P T7 -ymdA'-'lacZ translational fusion (Ϫ212 to ϩ4 cloned into the PstI and BamHI sites of pLFT). Mutations in the CsrA binding sites BS1 (GGA to AGA) and BS2 (GGA to AGA) and in the predicted site GGA* (GGA to GAA) in the context of the P ymdA -ymdA'-'lacZ and P T7 -ymdA'-'lacZ translational fusions were introduced using the QuikChange protocol. These plasmids contain a ymdA'-'lacZ translational fusion with the ymdA promoter or a T7 RNAP promoter driving transcription. WT and mutant fusions were integrated into the chromosomal att site of E. coli strains CF7789 and TRCF7789 as described previously (35).
CsrA-Mediated Translational Activation ® Strain PLB2839 contains a scarless GGA-to-AAG mutation in BS1. This mutation was engineered using the no-SCAR (scarless Cas9-assisted recombineering) system described previously (36). The correct mutation was confirmed by DNA sequencing. Strain AP3080 contains an unmarked ymdA deletion. This strain was constructed by first replacing the wild-type ymdA coding region with a kanamycin resistance gene using Red recombinase as described previously (37). The antibiotic resistance gene was subsequently removed using Flp recombinase (38), and the ymdA deletion was confirmed by PCR.
␤-Galactosidase assay. Bacterial cultures containing lacZ fusions were grown at 37°C in Luria-Bertani (LB) broth supplemented with 100 g/ml ampicillin and 50 g/ml kanamycin for csrA::kan strains. Cells were harvested at various points throughout growth. ␤-Galactosidase activity was measured as described previously (19).
Gel mobility shift assay. Quantitative gel mobility shift assays followed a published procedure (19). His-tagged CsrA (CsrA-H 6 ) was purified as described previously (39). WT and mutant RNAs (Ϫ100 to ϩ1 relative to the ymdA start codon) were synthesized with the RNAMaxx kit (Agilent Technologies) using PCR-generated DNA templates. Gel-purified RNA was dephosphorylated and then 5= end labeled using T4 polynucleotide kinase (New England Biolabs) and [␥-32 P]ATP. Labeled RNAs were renatured by heating for 1 min at 90°C followed by slow cooling to room temperature. Binding reaction mixtures (10 l) contained 0.1 nM labeled RNA, 10 mM Tris-HCl (pH 7.5), 10 mM MgCl 2 , 100 mM KCl, 40 ng of yeast tRNA, 7.5% glycerol, 0.1 mg/ml xylene cyanol, and various concentrations of purified CsrA-H 6 . Reaction mixtures were incubated for 30 min at 37°C to allow CsrA-RNA complex formation, and then samples were fractionated on a 10% nondenaturing polyacrylamide gel. Free and bound RNA species were visualized with a Typhoon 8600 variable-mode phosphorimager (GE Healthcare Life Sciences). CsrA-RNA interactions were quantified as described previously (19).
Footprint assay. CsrA-ymdA RNA footprint assays followed a published procedure (19). WT and BS2 mutant (GGA to AGA) labeled ymdA RNA (nt Ϫ127 to ϩ1 relative to the ymdA start codon) was labeled as described above for the gel mobility shift assay. The reaction mixtures were identical to those in the gel shift assay except that the concentration of labeled RNA was raised to 2 nM, and 1 g of acetylated bovine serum albumin (BSA) was added to each reaction mixture. Reaction mixtures were incubated for 30 min at 37°C to allow CsrA-RNA complex formation, then RNase T1 (0.02 U) was added, and incubation was continued for 15 min at 37°C. Reactions were stopped by adding 10 l of gel loading buffer (95% formamide, 0.025% sodium dodecyl sulfate (SDS), 20 mM EDTA, 0.025% bromophenol blue, 0.025% xylene cyanol). Samples were heated for 5 min at 90°C and then fractionated through 6% sequencing gels. Cleavage patterns were examined using a phosphorimager and quantified using semiautomated footprinting analysis software (SAFA) (40).
Toeprint assay. Primer extension inhibition (toeprint) assays followed a published procedure (19). Gel-purified ymdA RNA (150 nM) extending from Ϫ79 to ϩ106 relative to the ymdA start codon was hybridized to a 5=-end-labeled DNA oligonucleotide (150 nM) complementary to the 3= end of the RNA in Tris-EDTA (TE) buffer (pH 8) by heating for 3 min at 85°C followed by a slow cooling to room temperature. Toeprint reaction mixtures (10 l) contained 2 l of the hybridization mixture (30 nM final concentration), 375 M each deoxynucleoside triphosphate (dNTP), 10 mM dithiothreitol, and Super- script III (SSIII) reverse transcriptase buffer. CsrA-His 6 (1.5 M), tRNA fMet (10 M), and/or 30S ribosomal subunits (5 M) were added as indicated. Prior to addition, 30S ribosomal subunits were activated by incubation for 15 min at 37°C. Mixtures of the hybridization reaction and CsrA were incubated for 20 min at 37°C to allow for CsrA-RNA complex formation. tRNA fMet was then added, and incubation was continued for 5 min at 37°C. 30S ribosomal subunits were then added, and incubation was continued for 10 min at 37°C. SSIII (2 U) was added, and incubation was continued for 15 min at 37°C. Reactions were stopped by the addition of 10 l of gel loading buffer. Samples were fractionated through standard 6% sequencing gels. Toeprint patterns were visualized with a phosphorimager. Coupled transcription-translation assay. In vitro coupled transcription-translation assays were carried out with the PURExpress kit (New England BioLabs). Plasmids contained the P T7 -ymdA'-'lacZ fusion with either the WT ymdA sequence or a mutation in BS1 or BS2 (described above). A similar P T7 -pnp'-'lacZ translational fusion was used as a negative control (41). These plasmids were used as the templates for coupled transcription-translation reactions following a published procedure (41). Reaction mixtures containing 20 nM plasmid DNA template and various amounts of His-tagged CsrA were incubated for 2 h at 30°C. ␤-Galactosidase activity was monitored according to the manufacturer's instructions.
mRNA half-life analysis. E. coli MG1655 (WT) and TRMG1655 (csrA::kan) were grown in LB at 37°C to exponential phase, diluted to an optical density at 600 nm (OD 600 ) of 0.01 in fresh LB, and then grown to mid᎑exponential phase (OD 600 of 0.5). Rifampicin was added to a final concentration of 200 g/ml to stop transcription initiation. One-milliliter aliquots were removed at 0, 1, 2, 4, 8, and 16 min following rifampicin addition and immediately mixed with 0.125 ml of stop solution (10% phenolϪ90% ethanol [vol/vol]). Total cellular RNA was isolated using hot phenol-chloroform extraction followed by ethanol precipitation. Genomic DNA was digested by treating 20 g of nucleic acid with 2 U of Turbo DNase (Thermo Fisher Scientific), and RNA was purified from these reactions with phenol-chloroform extraction followed by ethanol precipitation. The overall integrity of the RNA was assessed by observing rRNA following 2% denaturing formaldehyde agarose gel electrophoresis in the presence of ethidium bromide and imaged using a ChemiDoc XRSϩ system (Bio-Rad). ymdA mRNA levels were detected by qRT᎑PCR and plotted on semilog decay curves using Prism (GraphPad).
Quantitative reverse transcriptase PCR. Quantitative reverse transcriptase PCR (qRT-PCR) was conducted using the iTaq Universal SYBR green one-step kit (Bio-Rad) and an iCycler iQ5 real-time PCR detection system (Bio-Rad) according to the manufacturer's instructions as described previously (42). The primer sequences used for this analysis were 5=-CTCTCTTATGCTCGGCAGTTT-3= and 5=-ACATGCCGGTTC CACAAT-3=. Reaction mixtures (10 l) contained 200 ng of RNA or standard DNA, 300 nM each primer, iScript reverse transcriptase, and 1ϫ iTaq Universal SYBR green reaction mix. Reaction mixtures were incubated for 10 min of reverse transcription (RT) at 50°C, 1 min of RT inactivation at 90°C, followed by 45 cycles of denaturation for 10 s at 95°C and annealing/extension for 20 s at 60°C. Following amplification, melting curve analysis was used to verify the specificity of the PCR product according to a single melting temperature (T m ). Melting curve analysis consisted of incubation for 1 min at 95°C, 1 min at 60°C, followed by 70 steps in which the temperature was increased to 95°C at a rate of 0.5°C/10 s/step. The ymdA mRNA concentrations were determined relative to a DNA standard curve for the PCR products using iQ5 optical system software version 2.1 (Bio-Rad) and were normalized to 16S rRNA levels.
Biofilm assay. Biofilm assays of WT (MG1655) and ΔymdA mutant strains, with or without complementation of ymdA via ASKA plasmid clone pYmdA versus the empty vector pCtrl (27), were performed in 96-well plates at 37°C in LB medium following the procedure described previously (30). Growth (A 600 ) and crystal violet staining (A 630 ) was measured using a BioTek Synergy HT plate reader, and biofilm formation was quantified as crystal violet staining (A 630 ) over growth (A 600 ).