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Research Article
18 December 2020

Characterization of the Streptococcus mutans SMU.1703c-SMU.1702c Operon Reveals Its Role in Riboflavin Import and Response to Acid Stress


Streptococcus mutans utilizes numerous metabolite transporters to obtain essential nutrients in the “feast or famine” environment of the human mouth. S. mutans and most other streptococci are considered auxotrophic for several essential vitamins including riboflavin (vitamin B2), which is used to generate key cofactors and to perform numerous cellular redox reactions. Despite the well-known contributions of this vitamin to central metabolism, little is known about how S. mutans obtains and metabolizes B2. The uncharacterized protein SMU.1703c displays high sequence homology to the riboflavin transporter RibU. Deletion of SMU.1703c hindered S. mutans growth in complex and defined medium in the absence of saturating levels of exogenous riboflavin, whereas deletion of cotranscribed SMU.1702c alone had no apparent effect on growth. Expression of SMU.1703c in a Bacillus subtilis riboflavin auxotroph functionally complemented growth in nonsaturating riboflavin conditions. S. mutans was also able to grow on flavin adenine dinucleotide (FAD) or flavin mononucleotide (FMN) in an SMU.1703c-dependent manner. Deletion of SMU.1703c and/or SMU.1702c impacted S. mutans acid stress tolerance, as all mutants showed improved growth at pH 5.5 compared to that of the wild type when medium was supplemented with saturating riboflavin. Cooccurrence of SMU.1703c and SMU.1702c, a hypothetical PAP2 family acid phosphatase gene, appears unique to the streptococci and may suggest a connection of SMU.1702c to the acquisition or metabolism of flavins within this genus. Identification of SMU.1703c as a RibU-like riboflavin transporter furthers our understanding of how S. mutans acquires essential micronutrients within the oral cavity and how this pathogen successfully competes within nutrient-starved oral biofilms.
IMPORTANCE Dental caries form when acid produced by oral bacteria erodes tooth enamel. This process is driven by the fermentative metabolism of cariogenic bacteria, most notably Streptococcus mutans. Nutrient acquisition is key in the competitive oral cavity, and many organisms have evolved various strategies to procure carbon sources or necessary biomolecules. B vitamins, such as riboflavin, which many oral streptococci must scavenge from the oral environment, are necessary for survival within the competitive oral cavity. However, the primary mechanism and proteins involved in this process remain uncharacterized. This study is important because it identifies a key step in S. mutans riboflavin acquisition and cofactor generation, which may enable the development of novel anticaries treatment strategies via selective targeting of metabolite transporters.


Known as a key contributor to dental caries (1, 2), Streptococcus mutans inhabits a complex oral environment composed of over 750 different microbial species (3). The S. mutans genome contains a diverse toolbox of carbohydrate and micronutrient transporters (4), which enable it to adapt and persist during the feast and famine cycle of the human oral cavity. Sugar uptake is primarily mediated by a diverse array of phosphoenolpyruvate-dependent sugar phosphotransferase systems (PTS) (4), importing monosaccharides and disaccharides for use within a lactic acid fermentation-based metabolism. ATP-binding cassette (ABC) transporters, such as the multiple sugar metabolism (Msm) system (5) and MalXFGK (6), also function as oligosaccharide import machinery to maximize carbohydrate uptake. Additionally, recent studies have suggested that S. mutans is able to uptake secondary metabolites, such as pyruvate (7), as a possible means of carbohydrate recovery.
Mechanisms for the import of carbon sources have been well studied in S. mutans, and its genome encodes a range of transporters associated with nonsugar nutrients or inorganic molecules (4). These transport systems range from essential small molecules, such as K+ and Mn2+, to metals, like Fe2+ and Co2+. These systems are key to S. mutans survival and play large roles in physiology. For example, studies of the Trk2 system demonstrated that deletion of this potassium transporter decreased the ability of S. mutans to survive in acidic conditions as well as its ability to form biofilms (8). Furthermore, the complex iron uptake and metabolism system utilized by S. mutans enables its ability to balance acquisition of iron from the environment with prevention of iron toxicity and Fenton chemistry (9, 10).
A relatively recently identified family of ABC transporters, energy coupling factor (ECF) transporters, are primarily associated with the movement of vitamins (11) as well as nickel and cobalt ions (12). The ECF transporter family consists of two groups, group I and group II (11), each consisting of 4 subunits as follows: a substrate binding S subunit, a membrane-bound T subunit, and two ATPases A and A′ (13). Group I transporters are characterized in part by the colocalized organization of all four subunits on the genome. This ECF class has been shown to be involved in the import of metal ions like cobalt (14) or B vitamins like thiamine (B1 [15]) or biotin (B7 [16]). In group II transporters, the T, A, and A′ subunit genes are linked together on the genome in an energy-charging cassette, with genes encoding variant S subunits located elsewhere on the genome. Group II ECF transporters are responsible for transport of many B vitamins, including riboflavin, with specific S subunits for each vitamin molecule (reviewed in references 17 and 18). The different S subunits can each interact and form functional complexes with the shared energy-charging cassette (19) and are unable to transport substrate without the aid of the T, A, and A′ subunits (13). Recent studies dissecting the mechanism of action in ECF-based transport have shown a novel form of substrate release involving the toppling of the substrate-bound S subunit within the membrane (2023). Free S subunits capture their target substrate, inducing a conformational change allowing interaction with the T subunit of the energy-charging cassette. Binding and subsequent ATP hydrolysis powers a “toppling” reaction, shifting the S subunit on to its side and releasing the bound substrate into the cell. Substrate-free S subunits are then released, freeing the energy-charging cassette for binding to another substrate binding unit. ECF S subunits involved in the transport of many B vitamins have been characterized, including thiamine (ThiT [24]), folate/B9 (FolT [22]), pantothenate/B5 (23), biotin (BioY [25]), B12 (CbrT [26]), and riboflavin/B2 (RibU [20, 27]).
S. mutans does not possess any known riboflavin biosynthetic enzymes (4, 28); thus, it must rely on scavenging what it needs from the oral cavity, where free riboflavin in saliva can be found in concentrations ranging from 0.058 to 0.88 μM (29, 30). As the primary niche for S. mutans is within the human mouth, this organism can target the dietary intake of riboflavin by its host as a reliable nutrient source. Once imported into the cell, riboflavin is converted into two cofactors as follows: flavin 5′-monophosphate (FMN) and flavin adenine dinucleotide (FAD). These cofactors are critical to the function of a wide variety of proteins known as flavoproteins. Studies have predicted up to ∼1.5% of total proteins in Staphylococcus aureus and Bacillus subtilis fall within this category (31), with examples of key S. mutans flavoproteins including the NADH oxidase Nox (32) and a key strategy for dealing with a reactive oxide species, superoxide dismutase (33). Riboflavin and its derived cofactors also appear to have a critical role in iron metabolism, as riboflavin biosynthesis has been linked to iron acquisition in Gram-negative bacteria, such as Helicobacter pylori and Campylobacter jejuni (34, 35). The iron-binding protein Dpr (10, 36) in S. mutans is also a protein requiring functional flavin cofactors.
Previous studies have characterized RibU-powered riboflavin import in the lactic acid bacterium Lactococcus lactis (27) as well as other Gram-positive pathogens, such as S. aureus (20, 21, 37), Listeria monocytogenes (38), and B. subtilis (39). However, relatively little investigation has been focused on the process of riboflavin acquisition and how it relates to cell persistence and stress resistance of oral streptococci within dental plaque biofilm. In this current study, we have confirmed and characterized the function of S. mutans SMU.1703c, annotated as a RibU homolog, as a riboflavin transporter. We have also shown that SMU.1702c, annotated as a hypothetical PAP2 family acid phosphatase that is cotranscribed with SMU.1703c, does not appear to be required for S. mutans growth under standard culturing conditions. When cultured in brain heart infusion (BHI) supplemented with saturating riboflavin concentrations, the wild-type strain displayed more sensitivity to acid stress relative to growth in BHI alone, a phenotype that was reversed in the SMU.1703c, SMU.1702c, and double mutant strains. Evidence is also provided that S. mutans growth is sustainable in minimal media containing FMN or FAD in place of riboflavin, a phenotype that is dependent on SMU.1703c. Together, these results suggest that SMU.1703c functions as the primary transporter of riboflavin in S. mutans. In addition, both SMU.1703c and SMU.1702c play a role in response to acid stress, though the mechanism by which this occurs remains unclear.


Expression of S. mutans SMU.1703c in a B. subtilis riboflavin auxotroph restores its ability to uptake riboflavin.

To confirm the RibU function of SMU.1703c, a previously published (40) B. subtilis 168 strain harboring mutations both in ribU and ribB (encoding a protein required for riboflavin biosynthesis) was transformed with the expression vector pHT01 (MoBiTec GmbH) containing the SMU.1703c coding sequence under the control of an isopropyl-β-d-thiogalactopyranoside (IPTG)-inducible promoter. When grown in semidefined SpI medium (41), the plasmid control strain harboring pHT01 showed little to no growth in the absence or presence of 10 μM riboflavin (Fig. 1) over the course of 8 h of aerobic growth at 37°C. As expected, cell growth was observed in this strain with the addition of 100 μM riboflavin (Fig. 1), as this was the saturating amount of this vitamin previously used to sustain growth of the B. subtilis ribB ribU mutant (38, 40). Expression of SMU.1703c had a minimal effect on B. subtilis ribB ribU growth in SpI without riboflavin (Fig. 1) but restored cell growth in media supplemented with 10 μM riboflavin. These data demonstrate that SMU.1703c can functionally complement RibU in a B. subtilis strain missing the ribU gene and, combined with the in silico analyses presented below, confirm that SMU.1703c functions as a RibU riboflavin transport protein.
FIG 1 Functional complementation of a B. subtilis ribU ribB mutant by S. mutans SMU.1703c (RibU). SMU.1703c was expressed in B. subtilis containing deletion of the riboflavin transporter RibU and riboflavin biosynthesis protein RibB. Cells were then grown in SpI medium containing 10 μM, 100 μM, or no riboflavin with cell density measured at 600 nm every hour. Average from n = 3 experiments; error bars represent standard error of the mean (SEM).

SMU.1703c resembles the ECF transporter subunit RibU.

Previous studies of the S. mutans cidAB operon (4244) showed that it is located immediately downstream from an apparent two-gene operon (SMU.1703c-SMU.1702c) (Fig. 2), previously suggested by Northern blotting analysis to be cotranscribed with cidAB (43). The SMU.1703c open reading frame is annotated in UniProt ( as encoding a putative riboflavin transporter and shares an 11-bp overlap with the open reading frame of its predicted operon partner SMU.1702c, annotated as encoding a hypothetical membrane protein of the PAP2 acid phosphatase superfamily. When modeled using the Phyre2 protein fold recognition server (45), the top predicted SMU.1703c structure templates were from the RibU protein of S. aureus TCH60 (13), with 88% amino acid coverage and >90% confidence. This generated structure contains 6 main transmembrane α-helices (see Fig. S1 in the supplemental material) with predicted lengths from 15 to 30 amino acids. Several other templates utilized for SMU.1703c structure generation also surpassed an 80% coverage and 90% confidence threshold, each being a transport protein and most categorized as members of the ECF transporter family. A high level of sequence homology was also observed when the amino acid sequence of SMU.1703c was aligned to other known RibU riboflavin transporters of S. aureus, Thermotoga maritima, L. lactis, and B. subtilis FmnP/RibU (Fig. 3A). When a similarity matrix between every pair of sequences was generated by Clustal Omega (Fig. 3B), the highest amino acid sequence identity was between SMU.1703c and L. lactis RibU at 40.11%, with the lowest between SMU.1703c and B. subtilis FmnP at 26.88%.
FIG 2 The S. mutans SMU.1703c-SMU.1702c locus and immediate genetic neighborhood. The SMU.1703c operon is located on the complement strand downstream of cidAB. Annotations for each open reading frame (ORF) are indicated by arrows.
FIG 3 Amino acid alignment of RibU homologs. (A) S. mutans SMU.1703c, Staphylococcus aureus (SaRibU), Thermotoga maritima (TmRibU), Bacillus subtilis (BsFmnP), and Lactococcus lactis (LlRibU) protein sequences were extracted from the UniProt database and aligned using M-Coffee. Alignment was then scored using BoxShade (ExPASy) with a 0.7 fraction of sequences cutoff for shading. (B) Percent identity matrix of RibU homologs. Identity percentages between the RibU homologs was calculated based on amino acid sequence alignment generated using Clustal Omega under default settings.

SMU.1703c contains a predicted FMN riboswitch within its promoter region and is cotranscribed with SMU.1702c.

Genes involved in riboflavin biosynthesis or salvage often contain an FMN riboswitch within the 5′ untranslated mRNA region. Binding of FMN to this riboswitch causes a conformational change in the secondary structure, inducing formation of a hairpin terminator and/or sequestering the Shine-Dalgarno, which represses transcription and/or translation, respectively (46, 47). To determine if this notable genetic element is also present upstream of the SMU.1703c open reading frame, we used the RNA secondary structure analysis tool Infernal (48) to analyze the S. mutans UA159 genome (GenBank accession number NC_004350.2) for the presence of the FMN riboswitch (Rfam RF00050). Results of this analysis indicated the significant likelihood of this element located 383 bp upstream of the putative SMU.1703c methionine start codon (predicted based on its position 8 bp downstream of a strong putative ribosomal binding consensus sequence) (Fig. 4A). In order to determine if riboflavin concentration impacts transcription of SMU.1703c, real-time PCR was performed on RNA that was isolated from S. mutans defined medium (FMC) cultures containing 0.107 μM (low concentration), 2.13 μM (standard FMC concentration), or 55 μM (saturating) riboflavin. Although large fold changes in SMU.1703c expression were not observed, this analysis showed that as the concentration of riboflavin increased, expression of SMU.1703c decreased (Fig. 4B).
FIG 4 Annotation of SMU.1703c promoter region (A), expression of SMU.1703c in differing riboflavin concentrations (B), and verification of SMU.1703c-SMU.1702c cotranscription (C). (A) The manually predicted start codon (black underline) and upstream consensus ribosomal binding site (RBS; blue) are indicated. 5′ rapid amplification of cDNA ends (RACE) was performed in order to identify the +1 transcriptional start site (red) (74). The −10 element (orange) was then inferred from the +1 site. The presence of an FMN class riboswitch (RF0050; italicized) was predicted via Rfam sequence search and Infernal (48). (B) RNA was isolated from wild-type cells grown to mid-exponential phase (OD600, 0.45 to 0.55) in FMC containing the indicated concentration of riboflavin. qPCR was used to measure the fold change SMU.1703c expression using the Livak method in less and saturating FMC cultures relative to that of full FMC (2.13 μM, calibrator). The gyrB gene was used as a reference gene. Data represent the average from n = 3 or 4 biological replicates; error bars indicate SEM. *, P < 0.05 (Dunn’s method) compared to full FMC culture. (C) S. mutans RNA was extracted from mid-exponential-phase cultures, followed by select cDNA synthesis with a primer targeted to the 3′ end of SMU.1702c. Generated cDNA was then used as a template for a second PCR utilizing the cDNA synthesis primer as well as a forward primer targeted to the 5′ SMU.1703c end. Gel loading: +RT, reverse transcriptase included in cDNA synthesis; −RT, reverse transcriptase not included in cDNA synthesis; H2O, water; gDNA, S. mutans UA159 gDNA.
A 5′ rapid amplification of cDNA ends (RACE) experiment was also performed to identify the +1 transcriptional start site (TSS) of SMU.1703c, which was found to be 324 bp upstream from its start codon (Fig. 4A) in two independently performed and sequenced RACE reactions. This +1 site corresponded to the beginning of the FMN riboswitch site predicted from the Infernal analysis described above. From the TSS, the −10 promoter element was estimated based on sequence similarity (Fig. 4A). In silico scans for a signal sequence did not indicate the presence of either a Sec-dependent or twin arginine translocation (TAT) signal sequence (data not shown).
An 11-base-pair overlap exists between the SMU.1702c start codon and the SMU.1703c stop codon. Therefore, to confirm cotranscription of SMU.1703c and SMU.1702c, PCR was performed on cDNA template synthesized from S. mutans RNA, with a reverse primer targeted to the 3′ end of SMU.1702c and a forward primer designed to amplify the entire SMU.1703c-SMU.1702c genomic region. Amplification of S. mutans genomic DNA (gDNA) template using these primers yielded an expected ∼1.3-kb band (Fig. 4C, rightmost lane). When this primer set was used with the cDNA template, a similarly sized band was amplified (Fig. 4C, leftmost lane), which was calculated to be the same size as the gDNA template band using a standard curve based on the DNA ladder. This amplification was not due to residual genomic contamination of the RNA template for cDNA synthesis, as no amplification was observed in a parallel PCR using RNA as template (Fig. 4C, second lane from left). Taken together, these results verify that SMU.1703c and SMU.1702c are cotranscribed as an operon containing a predicted 5′ end FMN riboswitch.

Deletion of SMU.1703c prevents cell growth without supplementation of excess riboflavin.

To examine the role of SMU.1703c and SMU.1702c in S. mutans physiology, double allelic exchange (49) was performed in order to create ΔSMU.1703c, ΔSMU.1702c, and ΔSMU.1703c ΔSMU.1702c mutant strains. Each mutant strain was subjected to genome resequencing and single nucleotide polymorphism (SNP) analysis (see Tables S1 to S3 in the supplemental material). Compared to our previously sequenced wild-type UA159 strain (44), the ΔSMU.1703c and ΔSMU.1703c ΔSMU.1702c mutants had no extraneous secondary SNPs, while the ΔSMU.1702c mutant had 2 differing SNPs—a change in the 3′ remnant of SMU.1702c and an R13M change in the gene SMU.1088. When struck onto unsupplemented BHI agar, ΔSMU.1702c mutant growth was comparable to that of the wild type (see Fig. S2 in the supplemental material), with no visible decrease in colony size or number. Growth was identical on BHI supplemented with 55 μM riboflavin (BHIrf) (previously used to sustain the growth of an Escherichia coli riboflavin auxotroph [39]) plates as well (Fig. S2), with no defects observed in either the ΔSMU.1702c mutant strain or the wild type. The ΔSMU.1703c and ΔSMU.1703c ΔSMU.1702c mutants, however, showed little to no growth on unsupplemented BHI plates (Fig. S2), with only a small patch of growth seen where the original culture was dropped onto the plate. Growth of each was rescued when struck onto BHIrf (Fig. S2), with cell growth equal to that of the wild type and ΔSMU.1702c mutant strain.
The growth patterns of the wild type and isogenic mutant strains observed on BHI agar plates were paralleled in liquid medium (Fig. 5). When grown in BHI for 24 h at 37°C, wild-type UA159 and the ΔSMU.1702c strain reached a peak optical density at 600 nm (OD600) of ∼0.8 at 5.5 h of growth (early stationary phase), while the ΔSMU.1703c strain and the double mutant supported a reduced level of overall growth, up to an OD600 of ∼0.4 (Fig. 5A). When grown in BHIrf with saturating levels of riboflavin, however, growth of all 4 strains were comparable (Fig. 5B), with a slight increase in lag phase observed for the ΔSMU.1703c mutant strain. Growth was also assessed in a complete defined medium FMC (50) under similar conditions. In this medium, each nutrient component can be individually controlled and manipulated, allowing more defined flavin levels to be studied. Under standard FMC riboflavin concentrations of 2.13 μM, both the wild type and the ΔSMU.1702c mutant grew as expected, reaching a peak OD600 of ∼0.8 with a similar lag phase to that observed during BHI growth (Fig. 5C). In contrast, growth was severely reduced for both the ΔSMU.1703c and double mutant in FMC medium (Fig. 5C) at this concentration of riboflavin, with the final OD600 of both cultures reaching only 0.1. This dramatic reduction in growth was comparable to that observed in all strains when cultured in FMC without any riboflavin (FMCnr, Fig. 5C). It is likely that the small amount of growth observed in these cultures was due to residual carryover of riboflavin from the inoculum, which was prepared from twice-washed phosphate-buffered saline (PBS)-washed cells from overnight FMC cultures supplemented with 55 μM riboflavin (required to support growth of the SMU.1703c and double mutants). Adding a saturating amount (55 μM) of riboflavin to FMC again rescued growth for both the ΔSMU.1703c and double mutants (Fig. 5D), though not quite to the same level as the wild type or ΔSMU.1702c mutant. Increased lag phases were also noted for the ΔSMU.1703c and ΔSMU.1703c ΔSMU.1702c mutants, and slightly reduced early stationary phase cell optical density was observed in each strain compared to that of the wild type (Fig. 5D).
FIG 5 Growth analysis of S. mutans and isogenic mutant panel. (A to D) Cells were grown for 24 h in a Bioscreen C automated growth system (Growth Curves USA) with cell density read at 600 nm every 30 min. Media were BHI (A), BHI plus 55 μM riboflavin (B), FMC (2.13 μM riboflavin) and FMC without riboflavin (FMCnr) (C), and FMC plus 55 μM riboflavin (D). Data are representative of n = 3 independent experiments; error bars indicate SEM. (E and F) Growth of S. mutans and the SMU.1703c-SMU.1702c panel of mutants in FMC with flavin based cofactors as the sole source of flavin(s) was as described above in FMC with 2.13 μM FMN (E), in FMC with 2.13 μM FAD (F), or without any source of flavins. Data are representative of n = 3 independent experiments; error bars indicate SEM.

S. mutans can utilize FMN or FAD as a sole flavin source for growth.

Due to the ability to precisely control the nutrients present within FMC, we next addressed whether flavin 5′-monophosphate (FMN) or flavin adenine dinucleotide (FAD) could support S. mutans growth as the sole flavin source. FMC medium was made with 2.13 μM either FMN or FAD as the only flavin source, the same concentration as riboflavin in standard FMC, and cell density throughout growth was measured as in the previous experiments. The presence of either FMN or FAD as the sole source of flavin(s) was able to support growth of the wild-type strain at levels comparable to standard FMC containing 2.13 μM riboflavin (Fig. 5E and F). Similarly, growth of the ΔSMU.1702c mutant with FAD or FMN matched growth observed in standard FMC with riboflavin (Fig. 5E and F). In the ΔSMU.1703c strain and ΔSMU.1703c ΔSMU.1702c double mutant, FMN and FAD were no longer able to support growth (Fig. 5E and F), a pattern also observed when grown in FMC containing comparable levels of riboflavin and FMC without riboflavin (Fig. 5C, E, and F). Mass spectrometry was additionally utilized to quantify riboflavin contamination within the FAD and FMN chemical stocks that were used to prepare the FMC media used in Fig. 5E and F (see the supplemetnal material). Riboflavin contamination was observed within the FAD and FMN chemicals at levels of 0.049% and 0.651%, respectively, placing the amount of contaminating riboflavin at subpicomolar levels in our FMC medium containing 2.13 μM FMN or FAD.

The SMU.1703c-SMU.1702c operon impacts S. mutans acid stress tolerance.

A key aspect of S. mutans physiology is the ability to adapt to and survive acidic conditions. The UA159 wild type and panel of mutant strains were challenged with acidic conditions at the time of inoculation, and growth was monitored over 24 h at 37°C. Growth of all four strains was comparable in a BHIrf pH of 7.4 (Fig. 6A). However, when grown in BHIrf preadjusted to pH 5.5, a noticeable decline in cell density was observed in the wild type as cells approached stationary growth phase (Fig. 6B). Although a modest decline was also noted in the ΔSMU.1703c mutant, it still had significantly (P < 0.01, Holm-Sidak) increased overall cell growth at 24 h relative to that of the wild type. Cell densities at 24 h of growth for the ΔSMU.1702c strain and double mutant were comparable to those of their counterpart pH 7.4 cultures and significantly (P < 0.01, Holm-Sidak) higher than the wild type grown at pH 5.5 (Fig. 6B). Overall, the ΔSMU.1702c and ΔSMU.1703c ΔSMU.1702c mutants displayed much more robust growth in pH 5.5 medium compared to that of the wild type, as evidenced by their decreased lag phase (Fig. 6B). Curiously, growth in BHI (without riboflavin supplementation) pH 5.5 medium restored the growth inhibition of the wild type that was observed in BHIrf pH 5.5 to levels comparable to those of the ΔSMU.1702c mutant BHI pH 5.5 culture and all mutants grown in BHIrf pH 5.5 (Fig. 6B). The ΔSMU.1703c and ΔSMU.1703c ΔSMU.1702c mutants were not tested in BHI without riboflavin supplementation due to their impaired growth in this medium (Fig. 5A). Viability of exponential-phase S. mutans wild-type and mutant cells grown in BHIrf and then exposed to acid shock (pH 3.5) was also followed over a 90-min period (Fig. 6C). These results indicated similar trends as was observed in the acid tolerance assay (Fig. 6B), but the differences between wild-type and mutant strains were less apparent in this assay. Collectively, these results suggest that excess levels of riboflavin have a negative impact on S. mutans growth under low pH stress and that deletion of SMU.1703c and/or SMU.1702c reverses this effect.
FIG 6 Growth of S. mutans and isogenic mutant panel in BHI preadjusted to pH 7.4 (A) and pH 5.5 (B) and viability upon exposure to acid stress (C). (A) Cells were grown in BHI or BHIrf at pH 7.4 for 24 h at 37°C in a Bioscreen C automated growth system. Optical density at 600 nm (OD600) was measured every 30 min. (B) Cultures were grown in BHI or BHIrf at pH 5.5 as described for panel A. OD600 was measured every 30 min. **, significant difference when comparing the wild type and ΔSMU.1703c mutant, the wild type and ΔSMU.1702c mutant, and the wild type and double mutant (P < 0.01, Holm-Sidak). Data in panels A and B represent the average from n = 3 to 5 experiments; error bars indicate SEM. (C) Cells were grown overnight at 37°C and 5% CO2 in BHIrf and diluted to an OD600 of 0.02 in fresh BHIrf. Following 4 h of growth, cells were harvested and resuspended in 0.1 M glycine buffer adjusted to pH 7 or pH 3.5. CFU were assessed at 20-min intervals. Data represent the average from n = 3 experiments; error bars indicate SEM. *, P = 0.052, Student’s t test, when comparing the wild type and ΔSMU.1703c ΔSMU.1702c mutant at pH 3.5.


Successful and consistent nutrient acquisition within the oral cavity is vital for S. mutans survival and niche establishment. As it cannot synthesize its own riboflavin (vitamin B2), S. mutans is an auxotroph for this essential molecule (4) and must scavenge what it needs from the environment around it. During our previous studies of the cidAB operon (4244, 51) the neighboring operon SMU.1703c-SMU.1702c was shown to possibly be cotranscribed with cidAB under certain conditions. The high amino acid sequence identity of SMU.1703c to well-characterized RibU homologs in other bacteria (Fig. 3A) and the presence of a predicted FMN riboswitch directly upstream of SMU.1703c (Fig. 4A) indicate that SMU.1703c encodes the ECF substrate binding unit RibU in S. mutans. The modest decrease in SMU.1703c expression in saturating levels of riboflavin, combined with a trend toward increased expression in media with lower levels of riboflavin (Fig. 4B), supports the predicted FMN element riboswitch (Fig. 4A) acting in a transcriptional attenuation manner observed in other Gram-positive bacteria (47, 52). However, these data alone do not exclude the possibility that the FMN riboswitch also exerts translational control over SMU.1703c expression in S. mutans, especially since small fold changes in SMU.1703c transcription were observed under our tested conditions. The FMN riboswitch upstream of ribB (involved in riboflavin biosynthesis) in E. coli was shown to have a modest effect on transcription and a stronger effect on translation (53). Since S. mutans is restricted to riboflavin salvage, it is also possible that the expression of SMU.1703c is less tightly controlled by the FMN riboswitch in this organism since it does not need to balance riboflavin biosynthesis and salvage.
The ability of SMU.1703c to functionally complement a B. subtilis ribU ribB mutant (riboflavin auxotroph) (Fig. 1) further supports the activity of SMU.1703c as an RibU riboflavin transporter as does phenotypic analysis of an S. mutans SMU.1703c mutant, which displayed very poor agar plate (see Fig. S2 in the supplemental material) and planktonic growth (Fig. 5A to D) unless the culture medium was supplemented with saturating levels of riboflavin. Interestingly, SMU.1703c is the only annotated riboflavin transporter gene within the S. mutans genome (4). Growth by mutants lacking SMU.1703c in the presence of saturating amounts of riboflavin is most likely driven by passive permeation of riboflavin across the cell membrane (38); however, we cannot definitively exclude the possibility of activity of a novel or uncharacterized vitamin transporter without further experimentation. The essentiality of SMU.1703c has also been recently demonstrated using an in vivo rodent caries model to screen an S. mutans transposon insertion sequencing (Tn-seq) library (54), which revealed that SMU.1703c was essential for survival in this model. Although much remains unknown about the vitamin acquisition and metabolism pathways of organisms within the human oral cavity, this study is a first step in elucidating how this essential process works in a key contributor to dental caries. With the emergence of biogeographical studies of human plaque (55, 56), enhanced understanding of nutrient and vitamin flux may also help to drive further insights on the organization of microorganisms and why specific associations form within the human mouth.
Interestingly, S. mutans can also utilize exogenous FMN and FAD as alternative flavin sources to sustain growth in the absence of riboflavin, and this is dependent on SMU.1703c (Fig. 5E and F). This result was unlikely due to the minimal riboflavin contamination of the FMN and FAD chemical stocks detected by mass spectroscopic analysis (0.049% and 0.651%, respectively, for FMN and FAD), as this would put the contaminating levels of riboflavin in the FMC growth medium at a subpicomolar concentration, which is well below riboflavin concentrations typically found in either complex BHI or defined FMC and is unlikely enough to support robust S. mutans growth. The FMC base medium used for these experiments was also not contaminated with growth-sustaining levels of riboflavin, as only a small amount of S. mutans growth was observed in the absence of FMN or FAD (Fig. 5E and F), comparable to that of FMC cultures lacking supplementation with either flavin. It is also unlikely that riboflavin was generated from FMN and FAD by chemical means. Light driven degradation of FMN at a biologically relevant pH of 5.6 has been shown to generate lumichrome and several forms of lumiflavin including formyllumiflavin but without the regeneration of the base molecule riboflavin (57). Furthermore, acid hydrolysis of FMN to riboflavin is unlikely to occur under the growth conditions used in this study, as this reaction appears to require incubation at 80°C (58). Evidence for the binding of FMN but not FAD by L. lactis RibU has been previously reported (27, 59). Without further experimentation, it is unclear if S. mutans is capable of directly transporting extracellular FMN and/or FAD in an SMU.1703c-dependent manner or if it enzymatically converts one or both flavins to riboflavin prior to transport. For example, enzymes involved in extracellular phosphatase activity recouping riboflavin from FMN and FAD have been described in Saccharomyces cerevisiae (60) and in rat intestinal walls (61), but it is unknown if genes encoding enzymes of similar function are present on the S. mutans genome.
Inundation of oral biofilms with acidic metabolic by-products of both S. mutans and other oral streptococci makes acid stress tolerance vital for persistence and niche establishment by these bacteria (62, 63). Surprisingly, growth of the wild-type strain in BHI with saturating (55 μM) riboflavin at pH 5.5 increased its sensitivity to acid stress, and deletion of SMU.1703c and/or SMU.1702c reversed this phenotype (Fig. 6B). However, when grown in unsupplemented BHI at pH 5.5, no differences in growth were observed between wild type and the ΔSMU.1702c mutant. Furthermore, growth of the wild type in BHI pH 5.5 was similar to growth of the ΔSMU.1702c and ΔSMU.1703c ΔSMU.1702c mutants in BHI pH 5.5 with saturating riboflavin. These results suggest that an excess level of transported riboflavin and/or production of downstream flavin cofactors has a negative impact on the ability of S. mutans to grow in and/or adapt to low pH conditions. Although the mechanism behind this is not known, one possibility is that flooding of wild-type cells with excess riboflavin at low pH disrupts the redox balance and slows growth, which may be alleviated by blocking active riboflavin uptake and/or metabolism by mutation of SMU.1703c and/or SMU.1702c. Annotated as a putative member of the PAP2 acid phosphatase family, SMU.1702c may also have a function in lipid metabolism or membrane alteration as described for other members of this protein family (6466), which may impact the response of S. mutans to acid stress. Remodeling the composition of its membrane fatty acids is a key component of the S. mutans acid tolerance response (ATR) (67). Although the exact protective mechanism is not understood, several studies have shown that S. mutans UA159 alters the composition of fatty acids in the cell membrane during acid stress at a pH of 5. Increases of longer monounsaturated fatty acids during exposure to acid stress (68, 69) have been shown, including shifts away from C14:0 and C16:0 saturated fatty acids toward C18:1 and C20:1 monounsaturated fatty acids (70). Similar results have been observed when comparing clinical carious S. mutans isolates from children at a pH of 7 versus 5 (68). Additionally, a role for cardiolipin in promoting S. mutans acid tolerance has been suggested, as a cardiolipin synthase mutant was more sensitive to acid stress and had a decreased proportion of unsaturated fatty acids (71). Thus, a second possibility is that the increase in acid stress tolerance observed in the ΔSMU.1702c strain and double mutant during growth in medium with excess riboflavin is linked to disruption of SMU.1702c activity, which participates in remodeling of membrane fatty acids. Current and ongoing experiments are exploring this possibility through characterization of the S. mutans ΔSMU.1702c lipidome and how it compares to the wild-type UA159 strain during acid stress.
In summary, the data presented in this study provide the basis for a testable working model of the early stages of S. mutans flavin transport and metabolism (Fig. 7), which is the basis for our ongoing and future research. Extracellular riboflavin is brought into the cell via the function of the RibU homolog SMU.1703c, along with the putative energy-charging cassette encoded by SMU.2148c-SMU.2150c. The flavin cofactors FMN and/or FAD may also enter the cell directly in an SMU.1703c-dependent manner or may be converted to riboflavin prior to transport. Once within the cell, riboflavin can be acted upon by the putative bifunctional riboflavin kinase/FMN adenyltransferase SMU.1143c to generate cofactors FMN and FAD, which are incorporated into the flavoproteome, whose composition may change in response to cellular needs under different metabolic conditions and/or environmental conditions, such as acid stress. Although the role of SMU.1702c in this system remains unknown, the fact that SMU.1703c and SMU.1702c are cotranscribed is suggestive of an as yet unidentified functional linkage. Interestingly, genes encoding RibU homologs in organisms such as B. subtilis (11) and L. lactis (59) are not organized as part of an operon, and the operon structure of ribU and a downstream gene encoding a putative PAP2 phosphatase seem to be a hallmark unique to the streptococci (see Fig. S3 in the supplemental material). With the determination of SMU.1703c as a key cog in the S. mutans vitamin metabolism wheel, future research will aim to identify and confirm the other genes involved in cellular processes linked to riboflavin import and metabolism. This will also further our general understanding of streptococcal vitamin metabolism, which has been understudied compared to that of other bacteria. Understanding this essential process within a key oral pathogen is an important step toward a better comprehension of how the oral microbiome functions and how disease may form within the human mouth.
FIG 7 Working model of riboflavin import and metabolism in S. mutans. Riboflavin is bound and imported via ECF machinery with SMU.1703c as the primary substrate binding S subunit. Sequence analysis predicts the operon containing SMU.2148c, SMU.2149c, and SMU.2150c as the remainder of the ECF energy cassette acting as the transmembrane T subunit and the ATPases A and A′, respectively. Once inside the cell, riboflavin is acted upon by the putative bifunctional riboflavin kinase/FMN adenyltransferase SMU.1143c. This activity provides the vital flavin cofactors FMN and FAD for a variety of cellular processes, including numerous redox reactions and metabolic pathways. SMU.1702c plays an as yet unknown role in this process, possibly in riboflavin import/metabolism or modification of the cell membrane.


Bacterial strains and culture conditions.

All bacterial strains and plasmids used in this study are listed in Table 1. S. mutans UA159 and/or its isogenic SMU.1703c mutant (ΔSMU.1703c::Kmr, nonpolar [NP]), SMU.1702c mutant (ΔSMU.1702c::NPKmr), and double mutant (ΔSMU.1703c ΔSMU.1702c::NPKmr) were routinely cultured on brain heart infusion (BHI) containing 1 mg/ml kanamycin for the mutant strains, along with 55 μM riboflavin for the ΔSMU.1703c and ΔSMU.1703c ΔSMU.1702c mutants. Unless otherwise noted, strains were cultured at 37°C in a 5% CO2 incubator in BHI, BHI supplemented with saturating (55 μM) riboflavin (BHIrf), or complete defined FMC medium (50) with/without riboflavin supplementation at standard (2.13 μM) or saturating (55 μM) concentrations. The B. subtilis ΔribB::Ermr ΔribU::Kmr strain (40) was generously provided by the laboratory of Matthias Mack (Mannheim University of Applied Sciences) and cultured on lysogeny broth (LB) supplemented with 55 μM riboflavin and with 1 μg/ml erythromycin and 5 μg/ml kanamycin in aerobic conditions (37°C, 250 rpm). The B. subtilis ΔribB ΔribU strain harboring the plasmid pHT01 (MoBiTec GmbH, Germany) was cultured as above with the addition of 30 μg/ml chloramphenicol.
TABLE 1 Bacterial strains and plasmids used in this study
Strain or plasmidGenotype or descriptionSource
Streptococcus mutans  
    UA159Wild type 
    ΔSMU.1703c mutantΔSMU.1703c::KmrThis study
    ΔSMU.1702c mutantΔSMU.1702c::KmrThis study
    ΔSMU.1703c ΔSMU.1702c mutantΔSMU.1703c ΔSMU.1702c::KmrThis study
Bacillus subtilis ΔribB ΔribU mutantΔribB::Ermr ΔribU::Kmr40
Escherichia coli DH5αfhuA2 lacΔU169 phoA glnV44 ϕ80' lacZΔM15 gyrA96 recA1 relA1 endA1 thi-1 hsdR1781
    pHT01E. coli to B. subtilis shuttle vector, IPTG-inducible promoter upstream of MCS; Ampr CmrMoBiTec GmbH
    pHT01_1703cpHT01 with SMU.1703c under control of inducible IPTG promoterThis study
    pALH124Source of Kmr cassette for generating S. mutans mutants72

S. mutans genetic manipulations and genome resequencing.

All mutants were created by using the PCR ligation mutagenesis approach (49) in which genes were disrupted with a nonpolar (NP) kanamycin resistance (Kmr) marker that was PCR amplified from plasmid pALH124 (72). Primers used for mutant making are listed in Table S4 in the supplemental material. Transformants were plated onto BHI (ΔSMU.1702c mutant) or BHIrf (ΔSMU.1703c mutant, double mutant), each containing appropriate antibiotic selection. Double crossover recombination was confirmed via PCR and by Sanger sequencing (Genewiz, NJ). Genomic DNA isolation and genome resequencing using the Illumina MiSeq 2 × 300 platform with Illumina v3 sequencing reagents were performed as previously described (44). Single nucleotide polymorphisms (SNPs) and other sequence changes were determined with CLC Genomics workbench (Qiagen) using basic variant detection with a 90% minimum frequency cutoff as described previously (73).

Bioinformatic analyses.

In order to model a predictive structure for SMU.1703c, the Phyre Protein Homology/analogY recognition engine (45) v2.0 was used with the intensive job option. SMU.1703c amino acid sequence with genome-annotated methionine start codon was taken from UniProt (accession number Q8DSR8). To visualize and manipulate protein models, PyMOL v1.7.4.5 Edu (Schrödinger, NY) was used. Amino acid sequences for S. mutans UA159 (SMU.1703c, using manually predicted start codon indicated in Fig. 4A), S. aureus TCH60 (ribU, HMPREF0772_11721), Thermotoga maritima (ribU, TM.1455), B. subtilis 168 (fmnP/ribU, BSU23050), and Lactococcus lactis MG1363 (ribU, LLMG.1195) were retrieved from the National Center for Biotechnology Information (NCBI) or from UniProt. Alignments were then generated using the M-Coffee protein alignment tool (74) followed by shading generated by BoxShade v3.2.1 ( Infernal 1.1.2 (48) was run searching the S. mutans UA159 genome accession NC_004350.2. The genome was scanned for the FMN riboswitch (RFN element) under the Rfam RF00050. The genome neighborhood diagram was generated using the Enzyme Function Initiative genome neighborhood tool (EFI-GNT [75]) using a neighborhood size of 10 and minimal cooccurrence lower limit of 20. An original sequence similarity network for SMU.1703c was generated on the EFI enzyme similarity tool focused on the Pfam family PF07155 under Uni Ref 90 and an E value of 5 with sequence length cutoffs of 145 and 162 min/max, respectively.

Mapping the transcriptional start site of SMU.1703c.

A 5′ rapid amplification of cDNA ends (RACE) was performed as previously described (51). Briefly, total RNA was isolated from S. mutans UA159 cultures grown at 37°C, 5% CO2 in BHI to an OD600 of 0.5 using the RNeasy minikit (Qiagen, Germany) as previously described (76, 77). SMU.1703c cDNA was then generated using the iScript Select cDNA synthesis kit (Bio-Rad, CA, USA) and the primer GSP-1 (see Table S1 in the supplemental material). cDNA was then purified using the Zymo Clean and Concentrator kit (Zymo, CA) and used in a homopolymeric tailing reaction with 2 mM dCTP and 15 units of TdT (Invitrogen). The reaction product was amplified further in several rounds of PCR using nested gene-specific primers, and products were visualized on a 1% agarose gel using gel electrophoresis. Once a suitable amount of DNA was visualized, products were sent for Sanger sequencing (Genewiz). The TSS was then determined from sequencing results by locating the homopolymeric tail and the first nucleotide following it.

Measurement of SMU.1703c expression.

S. mutans UA159 was inoculated to an OD600 of 0.02 in FMC containing 0.107, 2.13, or 55 μM riboflavin. Cultures were grown at 37°C and 5% CO2 until mid-exponential phase was reached (OD600, 0.43 to 0.525), and RNA was isolated from harvested cells as described previously (76, 77). Synthesis of cDNA and quantitative PCR (qPCR) was performed as described previously (78) using the Livak method (2−ΔΔCT) (79) to calculate the relative fold change in SMU.1703c between the calibrator sample (full FMC) and test samples using gyrB as a reference gene. Primers to detect gyrB (gyrB-F/gyrB-R) and SMU.1703c are listed in Table S4.

Cotranscription PCR.

As the coding sequences for SMU.1703c and SMU.1702c overlap, determination of whether or not these genes were cotranscribed was done using a cotranscription PCR (80). S. mutans UA159 cultures were grown as previously stated, and RNA was isolated as described above. cDNA was then generated using the iScript Select cDNA synthesis kit (Bio-Rad) with the primer 3/2_CoTrans_R targeted to the stop codon of SMU.1702c. Reverse transcriptase reactions were also performed without reverse transcriptase enzyme as a control to check for DNA contamination in the RNA sample. PCR amplification was then performed on these samples along with an S. mutans UA159 genomic DNA control, using the primers 3/2_CoTrans_F and 3/2_CoTrans_R for 30 cycles. Products were visualized on a 1% agarose gel using gel electrophoresis.

Functional complementation.

Functional complementation of riboflavin uptake in B. subtilis was performed based on the protocol designed by Hemberger et al. (40). The SMU.1703c coding sequence was amplified from the S. mutans UA159 genome using AccuPrime polymerase (Invitrogen) using the primers 1703_Func_CDS_F and 1703_Func_CDS_R. This sequence was then ligated into the plasmid pHT01 (MoBiTec GmbH) to generate pHT01-1703c, which was then moved into E. coli DH5α by heat shock transformation. Transformants were then screened on appropriate antibiotic plates, and plasmid was Sanger sequenced (Genewiz) to confirm correct insertion of the SMU.1703c coding sequence. Correct pHT01-1703c constructs were then transformed into the B. subtilis ΔribB::Ermr ΔribU::Kmr strain, along with empty pHT01 as a vector control. For growth experiments, both strains were grown 14 to 16 h in tryptic soy broth (TSB) (BD) with 55 μM riboflavin and appropriate antibiotics. Cells were then washed twice in sterile 1× PBS and diluted down to an OD600 of 0.02 in sterile 500-ml Erlenmeyer flasks with 50 ml of SpI medium (41) containing one of the following: 10 μM riboflavin, 100 μM riboflavin (38), or dimethyl sulfoxide (DMSO) (vehicle control). Expression of SMU.1703c was then induced with 1 mM IPTG, and cultures were grown aerobically for 10 h at 37°C and 250 rpm. Cell density was measured every hour at 600 nm.

Growth curves.

Growth of S. mutans UA159 and its isogenic mutants was measured using a Bioscreen C automated growth system (Growth Curves USA). Cells were grown for 16 to 18 h in either BHIrf or FMC (50) supplemented with riboflavin as indicated for each experiment and washed twice in 1× sterile phosphate-buffered saline (PBS). Cells were then diluted to an OD600 in fresh medium as follows: BHI, BHIrf, FMC, FMC supplemented with riboflavin (at 2.13 μM or 55 μM [39] as indicated for each experiment), or FMC without a flavin source. A honeycomb well plate (Growth Curves USA) was then inoculated in a 1:4 (250 μl) well-to-volume ratio, and cell optical density was measured over 24 h at 37°C. The ability of S. mutans UA159 and its isogenic SMU.1703c and/or SMU.1702c mutants to grow using FMN or FAD as a sole flavin source was also measured using a Bioscreen C automated growth system as described above. Cell cultures were grown overnight in FMC medium supplemented with excess riboflavin (55 μM) and then washed twice in 1× sterile PBS. Cells were then diluted to an OD600 of 0.02 in fresh FMC medium with either FMN (Sigma, Germany) or FAD (Sigma) at the standard FMC concentration used for riboflavin (2.13 μM) or without any flavin source at all as a negative control. A honeycomb well plate (Growth Curves USA) was then inoculated as described previously, and cell optical density was measured over 24 h at 37°C.

Acid tolerance assays.

To determine if deletions within the SMU.1703c-SMU.1702c operon affected the ability for S. mutans to tolerate acid stress, two separate assays were performed as follows: a cell growth assay and a cell survival assay. For the cell growth assay, S. mutans UA159 and its isogenic ΔSMU.1703c mutant, ΔSMU.1702c mutant, and double mutant were grown 16 to 18 h in filter-sterilized BHIrf at 37°C with 5% CO2. Cells were then diluted to an OD600 of 0.02 in fresh BHIrf (pH ∼7.4) or BHIrf with an adjusted pH of 5.5. Cell density was then measured over 24 h using a Bioscreen C automated growth system as described above. In a separate experiment, both the wild type and SMU.1702c mutant were grown in BHI (without riboflavin supplementation), pH 7.4, or preadjusted to pH 5.5 as described above. For the cell survival assay, S. mutans UA159 and its isogenic ΔSMU.1703c mutant, ΔSMU.1702c mutant, and double mutant were grown 16 to 18 h in BHIrf at 37°C with 5% CO2. Each strain was then diluted to an OD600 of 0.02 in 10 ml fresh BHIrf and incubated for 4 h at 37°C and 5% CO2. Cultures were split evenly (2 × 5 ml) in 15-ml falcon tubes, and cells were then pelleted via centrifugation at 4,500 rpm for 10 min. Supernatant was removed, and cell pellets were resuspended with 5 ml 0.1 M glycine buffer either at pH 7 or pH 3.5. Aliquots were then removed every 20 min and track plated on square BHIrf agar plates. After incubation at 37°C and 5% CO2 for 48 h, plates were removed and CFU were quantified.

Statistical analyses.

All statistical tests were performed using SigmaPlot 14.0 (Systat Software, CA, USA). Data were tested for normality and equal variance prior to selection of appropriate parametric or nonparametric tests as indicated in each figure legend.

Data availability.

Resequenced genome files were deposited to the NCBI Sequence Read Archive (SRA) under BioProject accession number PRJNA632033.


This work was supported by the National Institutes of Health (NIH)-National Institute of Dental and Craniofacial Research (NIDCR) grant R01 DE025237 (S.-J.A. and subaward to K.C.R.) and in part by a UF graduate school fellowship award and NIH predoctoral fellowship (F31DE029401) to M.E.T.
We thank Ning Zhu, Jin Koh, and Sixue Chen (Proteomics and Mass Spectrometry Facility, Interdisciplinary Center for Biotechnology Research, University of Florida) for performing mass spectrometry methodology optimization and analysis of FAD and FMN to detect contaminating riboflavin. We thank Matthias Mack (Mannheim University of Applied Sciences) for generously providing the B. subtilis ribB ribU mutant (40) and Valerie de Crécy-Lagard (Department of Microbiology and Cell Sciences, University of Florida) for helpful discussions.

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Eriksson L, Lif Holgerson P, Esberg A, Johansson I. 2018. Microbial complexes and caries in 17-year-olds with and without Streptococcus mutans. J Dent Res 97:275–282.
Loesche WJ. 1986. Role of Streptococcus mutans in human dental decay. Microbiol Rev 50:353–380.
Chen T, Yu WH, Izard J, Baranova OV, Lakshmanan A, Dewhirst FE. 2010. The Human Oral Microbiome Database: a web accessible resource for investigating oral microbe taxonomic and genomic information. Database (Oxford) 2010:baq013.
Ajdić D, McShan WM, McLaughlin RE, Savić G, Chang J, Carson MB, Primeaux C, Tian R, Kenton S, Jia H, Lin S, Qian Y, Li S, Zhu H, Najar F, Lai H, White J, Roe BA, Ferretti JJ. 2002. Genome sequence of Streptococcus mutans UA159, a cariogenic dental pathogen. Proc Natl Acad Sci U S A 99:14434–14439.
Tao L, Sutcliffe IC, Russell RR, Ferretti JJ. 1993. Transport of sugars, including sucrose, by the msm transport system of Streptococcus mutans. J Dent Res 72:1386–1390.
Webb AJ, Homer KA, Hosie AH. 2008. Two closely related ABC transporters in Streptococcus mutans are involved in disaccharide and/or oligosaccharide uptake. J Bacteriol 190:168–178.
Ahn S-J, Deep K, Turner ME, Ishkov I, Waters A, Hagen SJ, Rice KC. 2019. Characterization of LrgAB as a stationary phase-specific pyruvate uptake system in Streptococcus mutans. BMC Microbiol 19:223.
Binepal G, Gill K, Crowley P, Cordova M, Brady LJ, Senadheera DB, Cvitkovitch DG. 2016. Trk2 potassium transport system in Streptococcus mutans and its role in potassium homeostasis, biofilm formation, and stress tolerance. J Bacteriol 198:1087–1100.
Ganguly T, Kajfasz JK, Miller JH, Rabinowitz E, Galvão LCC, Rosalen PL, Abranches J, Lemos JA. 2018. Disruption of a novel iron transport system reverses oxidative stress phenotypes of a dpr mutant strain of Streptococcus mutans. J Bacteriol 200:e00062-18.
Yamamoto Y, Poole LB, Hantgan RR, Kamio Y. 2002. An iron-binding protein, Dpr, from Streptococcus mutans prevents iron-dependent hydroxyl radical formation in vitro. J Bacteriol 184:2931–2939.
Rodionov DA, Hebbeln P, Eudes A, ter Beek J, Rodionova IA, Erkens GB, Slotboom DJ, Gelfand MS, Osterman AL, Hanson AD, Eitinger T. 2008. A novel class of modular transporters for vitamins in prokaryotes. J Bacteriol 191:42–51.
Kirsch F, Eitinger T. 2014. Transport of nickel and cobalt ions into bacterial cells by S components of ECF transporters. Biometals 27:653–660.
Zhang P, Wang J, Shi Y. 2010. Structure and mechanism of the S component of a bacterial ECF transporter. Nature 468:717–720.
Bao Z, Qi X, Hong S, Xu K, He F, Zhang M, Chen J, Chao D, Zhao W, Li D, Wang J, Zhang P. 2017. Structure and mechanism of a group-I cobalt energy coupling factor transporter. Cell Res 27:675–687.
Josts I, Almeida Hernandez Y, Andreeva A, Tidow H. 2016. Crystal structure of a group I energy coupling factor vitamin transporter S component in complex with its cognate substrate. Cell Chem Biol 23:827–836.
Finkenwirth F, Kirsch F, Eitinger T. 2017. Complex stability during the transport cycle of a subclass I ECF transporter. Biochemistry 56:4578–4583.
Bousis S, Setyawati I, Diamanti E, Slotboom DJ, Hirsch AKH. 2019. Energy-coupling factor transporters as novel antimicrobial targets. Adv Therap 2:1800066.
Rempel S, Stanek WK, Slotboom DJ. 2019. ECF-type ATP-binding cassette transporters. Annu Rev Biochem 88:551–576.
ter Beek J, Duurkens RH, Erkens GB, Slotboom DJ. 2011. Quaternary structure and functional unit of energy coupling factor (ECF)-type transporters. J Biol Chem 286:5471–5475.
Karpowich NK, Song J, Wang DN. 2016. An aromatic cap seals the substrate binding site in an ECF-type S subunit for riboflavin. J Mol Biol 428:3118–3130.
Karpowich NK, Wang DN. 2013. Assembly and mechanism of a group II ECF transporter. Proc Natl Acad Sci U S A 110:2534–2539.
Swier LJ, Guskov A, Slotboom DJ. 2016. Structural insight in the toppling mechanism of an energy-coupling factor transporter. Nat Commun 7:11072.
Zhang M, Bao Z, Zhao Q, Guo H, Xu K, Wang C, Zhang P. 2014. Structure of a pantothenate transporter and implications for ECF module sharing and energy coupling of group II ECF transporters. Proc Natl Acad Sci U S A 111:18560–18565.
Erkens GB, Slotboom DJ. 2010. Biochemical characterization of ThiT from Lactococcus lactis: a thiamin transporter with picomolar substrate binding affinity. Biochemistry 49:3203–3212.
Finkenwirth F, Sippach M, Landmesser H, Kirsch F, Ogienko A, Grunzel M, Kiesler C, Steinhoff HJ, Schneider E, Eitinger T. 2015. ATP-dependent conformational changes trigger substrate capture and release by an ECF-type biotin transporter. J Biol Chem 290:16929–16942.
Santos JA, Rempel S, Mous ST, Pereira CT, Ter Beek J, de Gier JW, Guskov A, Slotboom DJ. 2018. Functional and structural characterization of an ECF-type ABC transporter for vitamin B12. Elife 7:e35828.
Duurkens RH, Tol MB, Geertsma ER, Permentier HP, Slotboom DJ. 2007. Flavin binding to the high affinity riboflavin transporter RibU. J Biol Chem 282:10380–10386.
Jijakli K, Jensen PA. 2019. Metabolic modeling of Streptococcus mutans reveals complex nutrient requirements of an oral pathogen. mSystems 4:e00529-19.
Sugimoto M, Saruta J, Matsuki C, To M, Onuma H, Kaneko M, Soga T, Tomita M, Tsukinoki K. 2013. Physiological and environmental parameters associated with mass spectrometry-based salivary metabolomic profiles. Metabolomics 9:454–463.
Dame ZT, Aziat F, Mandal R, Krishnamurthy R, Bouatra S, Borzouie S, Guo AC, Sajed T, Deng L, Lin H, Liu P, Dong E, Wishart DS. 2015. The human saliva metabolome. Metabolomics 11:1864–1883.
Macheroux P, Kappes B, Ealick SE. 2011. Flavogenomics–a genomic and structural view of flavin-dependent proteins. FEBS J 278:2625–2634.
Baker JL, Derr AM, Karuppaiah K, MacGilvray ME, Kajfasz JK, Faustoferri RC, Rivera-Ramos I, Bitoun JP, Lemos JA, Wen ZT, Quivey RG. 2014. Streptococcus mutans NADH oxidase lies at the intersection of overlapping regulons controlled by oxygen and NAD+ levels. J Bacteriol 196:2166–2177.
Martin ME, Strachan RC, Aranha H, Evans SL, Salin ML, Welch B, Arceneaux JE, Byers BR. 1984. Oxygen toxicity in Streptococcus mutans: manganese, iron, and superoxide dismutase. J Bacteriol 159:745–749.
Crossley RA, Gaskin DJ, Holmes K, Mulholland F, Wells JM, Kelly DJ, van Vliet AH, Walton NJ. 2007. Riboflavin biosynthesis is associated with assimilatory ferric reduction and iron acquisition by Campylobacter jejuni. Appl Environ Microbiol 73:7819–7825.
Worst DJ, Gerrits MM, Vandenbroucke-Grauls CM, Kusters JG. 1998. Helicobacter pylori ribBA-mediated riboflavin production is involved in iron acquisition. J Bacteriol 180:1473–1479.
Yamamoto Y, Fukui K, Koujin N, Ohya H, Kimura K, Kamio Y. 2004. Regulation of the intracellular free iron pool by Dpr provides oxygen tolerance to Streptococcus mutans. J Bacteriol 186:5997–6002.
Karpowich NK, Song JM, Cocco N, Wang DN. 2015. ATP binding drives substrate capture in an ECF transporter by a release-and-catch mechanism. Nat Struct Mol Biol 22:565–571.
Matern A, Pedrolli D, Großhennig S, Johansson J, Mack M. 2016. Uptake and metabolism of antibiotics roseoflavin and 8-demethyl-8-aminoriboflavin in riboflavin-auxotrophic Listeria monocytogenes. J Bacteriol 198:3233–3243.
Vogl C, Grill S, Schilling O, Stülke J, Mack M, Stolz J. 2007. Characterization of riboflavin (vitamin B2) transport proteins from Bacillus subtilis and Corynebacterium glutamicum. J Bacteriol 189:7367–7375.
Hemberger S, Pedrolli DB, Stolz J, Vogl C, Lehmann M, Mack M. 2011. RibM from Streptomyces davawensis is a riboflavin/roseoflavin transporter and may be useful for the optimization of riboflavin production strains. BMC Biotechnol 11:119.
Erickson RJ, Copeland JC. 1972. Structure and replication of chromosomes in competent cells of Bacillus subtilis. J Bacteriol 109:1075–1084.
Ahn SJ, Rice KC. 2016. Understanding the Streptococcus mutans Cid/Lrg system through CidB function. Appl Environ Microbiol 82:6189–6203.
Ahn SJ, Rice KC, Oleas J, Bayles KW, Burne RA. 2010. The Streptococcus mutans Cid and Lrg systems modulate virulence traits in response to multiple environmental signals. Microbiology (Reading) 156:3136–3147.
Turner ME, Huynh K, Carney OV, Gross D, Carroll RK, Ahn SJ, Rice KC. 2019. Genomic instability of TnSMU2 contributes to Streptococcus mutans biofilm development and competence in a cidB mutant. Microbiologyopen 8:e934.
Kelley LA, Mezulis S, Yates CM, Wass MN, Sternberg MJ. 2015. The Phyre2 web portal for protein modeling, prediction and analysis. Nat Protoc 10:845–858.
Serganov A, Huang L, Patel DJ. 2009. Coenzyme recognition and gene regulation by a flavin mononucleotide riboswitch. Nature 458:233–237.
Vitreschak AG, Rodionov DA, Mironov AA, Gelfand MS. 2002. Regulation of riboflavin biosynthesis and transport genes in bacteria by transcriptional and translational attenuation. Nucleic Acids Res 30:3141–3151.
Nawrocki EP, Eddy SR. 2013. Infernal 1.1: 100-fold faster RNA homology searches. Bioinformatics 29:2933–2935.
Lau PC, Sung CK, Lee JH, Morrison DA, Cvitkovitch DG. 2002. PCR ligation mutagenesis in transformable streptococci: application and efficiency. J Microbiol Methods 49:193–205.
Terleckyj B, Willett NP, Shockman GD. 1975. Growth of several cariogenic strains of oral streptococci in a chemically defined medium. Infect Immun 11:649–655.
Kim HM, Waters A, Turner ME, Rice KC, Ahn SJ. 2019. Regulation of cid and lrg expression by CcpA in Streptococcus mutans. Microbiology (Reading) 165:113–123.
Wang H, Mann PA, Xiao L, Gill C, Galgoci AM, Howe JA, Villafania A, Barbieri CM, Malinverni JC, Sher X, Mayhood T, McCurry MD, Murgolo N, Flattery A, Mack M, Roemer T. 2017. Dual-targeting small-molecule inhibitors of the Staphylococcus aureus FMN riboswitch disrupt riboflavin homeostasis in an infectious setting. Cell Chem Biol 24:576–588.
Pedrolli D, Langer S, Hobl B, Schwarz J, Hashimoto M, Mack M. 2015. The ribB FMN riboswitch from Escherichia coli operates at the transcriptional and translational level and regulates riboflavin biosynthesis. FEBS J 282:3230–3242.
Shields RC, Zeng L, Culp DJ, Burne RA. 2018. Genomewide identification of essential genes and fitness determinants of Streptococcus mutans UA159. mSphere 3:e00031-18.
Kim D, Koo H. 2020. Spatial design of polymicrobial oral biofilm in its native disease state. J Dent Res 99:597–603.
Mark Welch JL, Rossetti BJ, Rieken CW, Dewhirst FE, Borisy GG. 2016. Biogeography of a human oral microbiome at the micron scale. Proc Natl Acad Sci U S A 113:E791–E800.
Holzer W, Shirdel J, Zirak P, Penzkofer A, Hegemann P, Deutzmann R, Hochmuth E. 2005. Photo-induced degradation of some flavins in aqueous solution. Chemical Physics 308:69–78.
Nielsen P, Harksen J, Bacher A. 1985. Hydrolysis and rearrangement reactions of riboflavin phosphates. An explicit kinetic study. Eur J Biochem 152:465–473.
Burgess CM, Slotboom DJ, Geertsma ER, Duurkens RH, Poolman B, van Sinderen D. 2006. The riboflavin transporter RibU in Lactococcus lactis: molecular characterization of gene expression and the transport mechanism. J Bacteriol 188:2752–2760.
Pallotta ML. 2011. Evidence for the presence of a FAD pyrophosphatase and a FMN phosphohydrolase in yeast mitochondria: a possible role in flavin homeostasis. Yeast 28:693–705.
Akiyama T, Selhub J, Rosenberg IH. 1982. FMN phosphatase and FAD pyrophosphatase in rat intestinal brush borders: role in intestinal absorption of dietary riboflavin. J Nutr 112:263–268.
Dashper SG, Reynolds EC. 2000. Effects of organic acid anions on growth, glycolysis, and intracellular pH of oral streptococci. J Dent Res 79:90–96.
de Soet JJ, Nyvad B, Kilian M. 2000. Strain-related acid production by oral streptococci. Caries Res 34:486–490.
Fan J, Jiang D, Zhao Y, Liu J, Zhang XC. 2014. Crystal structure of lipid phosphatase Escherichia coli phosphatidylglycerophosphate phosphatase B. Proc Natl Acad Sci U S A 111:7636–7640.
Gasiorowski E, Auger R, Tian X, Hicham S, Ecobichon C, Roure S, Douglass MV, Trent MS, Mengin-Lecreulx D, Touzé T, Boneca IG. 2019. HupA, the main undecaprenyl pyrophosphate and phosphatidylglycerol phosphate phosphatase in Helicobacter pylori is essential for colonization of the stomach. PLoS Pathog 15:e1007972.
Ghachi ME, Howe N, Auger R, Lambion A, Guiseppi A, Delbrassine F, Manat G, Roure S, Peslier S, Sauvage E, Vogeley L, Rengifo-Gonzalez JC, Charlier P, Mengin-Lecreulx D, Foglino M, Touzé T, Caffrey M, Kerff F. 2017. Crystal structure and biochemical characterization of the transmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus subtilis. Cell Mol Life Sci 74:2319–2332.
Baker JL, Faustoferri RC, Quivey RG. 2017. Acid-adaptive mechanisms of Streptococcus mutans-the more we know, the more we don't. Mol Oral Microbiol 32:107–117.
Bojanich MA, Calderón RO. 2017. Streptococcus mutans membrane lipid composition: virulence factors and structural parameters. Arch Oral Biol 81:74–80.
Fozo EM, Quivey RG. 2004. Shifts in the membrane fatty acid profile of Streptococcus mutans enhance survival in acidic environments. Appl Environ Microbiol 70:929–936.
Quivey RG, Faustoferri R, Monahan K, Marquis R. 2000. Shifts in membrane fatty acid profiles associated with acid adaptation of Streptococcus mutans. FEMS Microbiol Lett 189:89–92.
MacGilvray ME, Lapek JD, Friedman AE, Quivey RG. 2012. Cardiolipin biosynthesis in Streptococcus mutans is regulated in response to external pH. Microbiology (Reading) 158:2133–2143.
Ahn SJ, Wen ZT, Burne RA. 2006. Multilevel control of competence development and stress tolerance in Streptococcus mutans UA159. Infect Immun 74:1631–1642.
Mashruwala AA, Roberts CA, Bhatt S, May KL, Carroll RK, Shaw LN, Boyd JM. 2016. Staphylococcus aureus SufT: an essential iron-sulphur cluster assembly factor in cells experiencing a high-demand for lipoic acid. Mol Microbiol 102:1099–1119.
Moretti S, Armougom F, Wallace IM, Higgins DG, Jongeneel CV, Notredame C. 2007. The M-Coffee web server: a meta-method for computing multiple sequence alignments by combining alternative alignment methods. Nucleic Acids Res 35:W645–W648.
Zallot R, Oberg N, Gerlt JA. 2019. The EFI web resource for genomic enzymology tools: leveraging protein, genome, and metagenome databases to discover novel enzymes and metabolic pathways. Biochemistry 58:4169–4182.
Ahn SJ, Qu MD, Roberts E, Burne RA, Rice KC. 2012. Identification of the Streptococcus mutans LytST two-component regulon reveals its contribution to oxidative stress tolerance. BMC Microbiol 12:187.
Patton TG, Rice KC, Foster MK, Bayles KW. 2005. The Staphylococcus aureus cidC gene encodes a pyruvate oxidase that affects acetate metabolism and cell death in stationary phase. Mol Microbiol 56:1664–1674.
Lewis AM, Rice KC. 2016. Quantitative real-time PCR (qPCR) workflow for analyzing Staphylococcus aureus gene expression. Methods Mol Biol 1373:143–154.
Livak KJ, Schmittgen TD. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods 25:402–408.
Sapp AM, Mogen AB, Almand EA, Rivera FE, Shaw LN, Richardson AR, Rice KC. 2014. Contribution of the nos-pdt operon to virulence phenotypes in methicillin-sensitive Staphylococcus aureus. PLoS One 9:e108868.
Hanahan D. 1983. Studies on transformation of Escherichia coli with plasmids. J Mol Biol 166:557–580.

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Published In

cover image Journal of Bacteriology
Journal of Bacteriology
Volume 203Number 218 December 2020
eLocator: 10.1128/jb.00293-20
Editor: Tina M. Henkin, Ohio State University


Received: 15 May 2020
Accepted: 15 October 2020
Published online: 18 December 2020


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  1. Streptococcus mutans
  2. riboflavin
  3. ECF transporter
  4. acid stress



Matthew E. Turner
Department of Microbiology and Cell Science, Institute of Food and Agricultural Sciences, University of Florida, Gainesville, Florida, USA
Khanh Huynh
Department of Microbiology and Cell Science, Institute of Food and Agricultural Sciences, University of Florida, Gainesville, Florida, USA
Department of Biological Sciences, Ohio University, Athens, Ohio, USA
Sang-Joon Ahn
Department of Oral Biology, College of Dentistry, University of Florida, Gainesville, Florida, USA
Department of Microbiology and Cell Science, Institute of Food and Agricultural Sciences, University of Florida, Gainesville, Florida, USA


Tina M. Henkin
Ohio State University


Address correspondence to Kelly C. Rice, [email protected].

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