Methods

Fluorescence Tools Adapted for Real-Time Monitoring of the Behaviors of Streptococcus Species

R. C. Shields, J. R. Kaspar, K. Lee, S. A. M. Underhill, R. A. Burne
Donald W. Schaffner, Editor
DOI: 10.1128/AEM.00620-19

ABSTRACT

Tagging of bacteria with fluorescent proteins has become an essential component of modern microbiology. Fluorescent proteins can be used to monitor gene expression and biofilm growth and to visualize host-pathogen interactions. Here, we developed a collection of fluorescent protein reporter plasmids for Streptococcus mutans UA159 and other oral streptococci. Using superfolder green fluorescent protein (sfGFP) as a reporter for transcriptional activity, we were able to characterize four strong constitutive promoters in S. mutans. These promoter-sfgfp fusions worked both for single-copy chromosomal integration and on a multicopy plasmid, with the latter being segregationally stable in the absence of selective pressure under the conditions tested. We successfully labeled S. mutans UA159, Streptococcus gordonii DL1, and Streptococcus sp. strain A12 with sfGFP, DsRed-Express2 (red), and citrine (yellow). To test these plasmids under more challenging conditions, we performed mixed-species biofilm experiments and separated fluorescent populations using fluorescence-activated cell sorting (FACS). This allowed us to visualize two streptococci at a time and quantify the amounts of each species simultaneously. These fluorescent reporter plasmids add to the genetic toolbox available for the study of oral streptococci.

IMPORTANCE Oral streptococci are the most abundant bacteria in the mouth and have a major influence on oral health and disease. In this study, we designed and optimized the expression of fluorescent proteins in Streptococcus mutans and other oral streptococci. We monitored the levels of expression and noise (the variability in fluorescence across the population). We then created several fluorescent protein delivery systems (green, yellow, and red) for use in oral streptococci. The data show that we can monitor bacterial growth and interactions in situ, differentiating between different bacteria growing in biofilms, the natural state of the organisms in the human mouth. These new tools will allow researchers to study these bacteria in novel ways to create more effective diagnostic and therapeutic tools for ubiquitous infectious diseases.

INTRODUCTION

Green fluorescent protein (GFP) is widely used to explore bacterial behaviors (1). GFP has been used in studies of protein localization and gene expression, for examining spatial arrangements in biofilms, and for visualizing bacteria in host cells. The utility of GFP was first described in 1994 when Chalfie et al. (2) identified the gfp gene in the jellyfish (Aequorea victoria). Over the last 20 years, a number of fluorescent protein (FP) variants have been described, including optimized versions of GFP (e.g., superfolder GFP [sfGFP]) and homologs of GFP that emit light at different wavelengths (e.g., DsRed) (3). Besides altering the protein, there are a number of factors that can influence the fluorescence intensity. These include transcription rates, mRNA stability, and codon bias (4–6). To achieve stable and high levels of GFP fluorescence in an organism, optimized reporter systems have to be designed (7). Once reporter systems have been designed, they can be used to visualize the organism in diverse experimental models (8).

Approximately 35% of the human population has untreated caries in permanent teeth, leading to significant reductions in quality of life through pain, disfigurement, and difficulty eating (9). Dental caries, which involves the loss of tooth mineral, is caused by a compositional change in the oral microbiome (10). Streptococcus mutans is the organism that is most consistently associated with carious lesions and has multiple virulence traits that contribute to its ability to initiate and worsen carious lesions (11). Its interactions with other oral bacteria, including health-associated streptococci, are important for determining the health status of a person’s teeth (12). Despite the medical importance of these microorganisms, there is a lack of molecular tools available for their study, in contrast to the number of molecular tools available for the study of the more intensively studied model organisms. This includes fluorescence-based tools that could be used for the study of microbial interactions, host-microbe interactions, single-cell behaviors, and biofilm dynamics. Recent examples of fluorescent protein tagging in oral streptococci include the work of Vickerman et al. (13), who tagged eight oral streptococci with mTFP1 and mCherry, and Shabayek and Spellerberg (14), who used the GFP derivatives enhanced GFP (EGFP) and Sirius to fluorescently tag oral streptococci. Further refinements of the already available tools are required to realize the full potential of fluorescence tagging in oral bacteria. Refinements include increasing expression of the fluorescent proteins, testing different delivery systems and their impact on fluorescence noise, and measuring the utility of fluorescent proteins in more complex experimental systems.

Here, we developed plasmid vectors that carry fluorescent protein markers to allow for the visualization of S. mutans and other oral streptococci. We tested four constitutively active promoters fused to a superfolder gfp (sfgfp) gene as a way to compare promoter strength. The intensity and noise of these promoters delivered in different vectors were determined at the single-cell level. Next, we designed, tested, and confirmed the effectiveness of sfgfp, DsRed-Express2, and citrine gene expression from plasmid vectors in S. mutans, Streptococcus gordonii DL1, and Streptococcus sp. strain A12. Finally, we show that these plasmids can be used to visually discriminate oral streptococci in coculture biofilms.

RESULTS

Ranking the strength of four constitutive promoters in S. mutans.The first goal of this project was to construct a plasmid that would allow for the chromosomal integration of a constitutively expressed fluorescent protein. To accomplish this, synthetic constructs were cloned into the integration vector pPMZ (P, phnA; M, mtlA; Z, lacZ) (15, 16). This plasmid contains DNA sequences flanking the cloning sites that allow for the integration of the desired DNAs into the phnA-mtlA locus, which is required only for mannitol metabolism and which has no effect on the fitness of S. mutans in the absence of mannitol. To maximize the expression of gfp, we designed four constructs with four different constitutive promoters, P3, P23, Pldh, and Pveg. P3 has prokaryotic consensus −35 (TTGACA) and −10 (TATAAT) boxes and is a strong constitutive promoter in Streptococcus pneumoniae (17). P23 is a lactococcal promoter (Lactococcus lactis subsp. cremoris) with strong constitutive activity in S. mutans (18, 19). Pveg is a constitutive promoter originally identified in Bacillus subtilis that has strong expression in a variety of bacterial hosts (20). Pldh is the promoter of the gene encoding the lactate dehydrogenase (ldh, SMu.1115) of S. mutans and has been used as a constitutive promoter in S. mutans (21). The sequences of the promoters are shown in Fig. 1A, with the −35, −10, +1 (the transcription initiation site), and Shine-Dalgarno (SD) sequences being highlighted in bold. For the Pveg and P23 promoters, we used an optimized ribosomal binding site (RBS) that contains a consensus SD (AGGAGG) sequence (22); this RBS displayed maximal translational efficiency in B. subtilis (22). For P3, the original RBS sequence described by Sorg et al. was retained (17), as was the native RBS of the ldh gene. Next, we selected a superfolder GFP (sfGFP) that was codon optimized for Bacillus subtilis but that was also highly expressed in S. pneumoniae (7). sfGFP matures rapidly (6 min in Escherichia coli) and is highly stable, making it a useful fluorescent reporter (23). The sequences of each promoter-sfgfp fusion are available in the supplemental material. For ease of cloning, we used a gene synthesis service provided by Integrated DNA Technologies (IDT). Sequences were synthesized and provided in a plasmid cloning vector. SacI and SphI were used to excise the constructs from the IDT vector, with subsequent cloning into the pPMZ vector. These vectors were then transformed into competent S. mutans cells.

FIG 1
FIG 1

Fluorescence intensities of the four promoter-gfp fusion constructs. (A) Promoter sequences are shown, with the −35, −10, +1 (transcription initiation site [TIS]), Shine-Dalgarno (ribosome binding site [RBS]), and sfgfp ATG sequences being indicated in bold. The promoter sequences were fused to the superfolder gfp gene, and these constructs were cloned into plasmids. To the best of our knowledge, for S. mutans Pldh the TIS has not been experimentally validated. (B) Western blot detection of sfGFP using an anti-sfGFP antibody against whole-cell extracts collected at an OD600 of 0.5. (C) Fluorescence intensity (circles) and growth (lines) measured simultaneously using a microplate assay. (D) Single-cell fluorescence microscopy of S. mutans/pPM_Pveg-sfgfp. sfGFP fluorescence was detected by excitation at 488 nm, and emission was collected using a 525-nm (±25-nm)-band-pass filter. A 63× oil objective lens was used to acquire the image.

The levels of GFP expression for each individual construct were measured using immunodetection and a fluorescent plate reader. For Western blotting experiments, we used an anti-sfGFP polyclonal antibody (Sigma-Aldrich) against whole-cell lysates prepared from cells grown to the mid-exponential phase (optical density at 600 nm [OD600] = 0.5). We were able to detect sfGFP at this time point across all four constructs, with no evidence of degradation of sfGFP in the form of lower-molecular-weight immunoreactive products (Fig. 1B). Using a plate reader, we were able to monitor the strength of the fluorescence signals during the growth of S. mutans. For each construct, the absolute levels of GFP increased as the optical density of the population increased, consistent with promoter activity being constitutive. The fluorescence signals peaked during the transition from late exponential to early stationary phase. In this assay, fluorescence levels were the highest for Pveg-sfgfp (33,000 relative fluorescence units [RFU]) and the lowest for Pldh-sfgfp (15,000 RFU) (Fig. 1C). None of the strains exhibited any significant growth defects compared to wild-type S. mutans. Lastly, using fluorescence microscopy, S. mutans/Pveg-sfgfp cells could readily be visualized as green ovococci, the typical morphology of S. mutans (Fig. 1D).

Next, we cloned each promoter-sfgfp construct into the E. coli-streptococcal shuttle vector pDL278 (24). An advantage of this vector is that it can replicate in multiple streptococcal species. The vector was constructed with a pVA380-1 backbone. Plasmid pVA380-1 was originally isolated from Streptococcus ferus and replicates via rolling-circle replication (RCR) (24–26). Plasmids that replicate via RCR can suffer from segregational instability, which has been attributed to the reliance of replication on single-stranded DNA intermediates, and the formation of linear high-molecular-weight (HMW) plasmid multimers (27). Plasmid-based fluorescence systems may not be useful if selective pressure is required to avoid segregational instability. Addition of antibiotics (to increase plasmid stability) would limit the usefulness of GFP plasmids, as antibiotics are known to modify the physiology of bacteria in ways that alter important behaviors, including biofilm formation (28). To test for plasmid stability in the absence of spectinomycin (pDL278 carries aad9), we grew pDL278 carrying Pveg-sfgfp in rich medium (brain heart infusion [BHI] medium) for approximately 50 generations. The strains were subcultured (1:1,000) into fresh rich medium for a total of 5 serial passages, each of which was incubated for 12 h. Prior to each passage, cells from the cultures were plated onto BHI agar with or without spectinomycin. After 48 h of incubation, the numbers of CFU were counted and the total numbers of cells on selective and nonselective media were compared. Based on this plasmid stability assay, it was observed that pDL278 carrying Pveg-sfgfp was retained over the entire course of the experiment, as there were no significant differences (P = 0.6749) between the total number of cells that grew on BHI agar and the total number of cells that grew on BHI-spectinomycin agar (Fig. 2).

FIG 2
FIG 2

Stability of pDL278_Pveg-sfgfp in the presence and absence of spectinomycin. The numbers of spectinomycin-resistant S. mutans pDL278_Pveg-sfgfp cells cultured in the presence (A) or absence (B) of spectinomycin were compared. S. mutans cells harboring pDL278_Pveg-sfgfp were diluted 1:1,000 into BHI and cultured for 12 h. After 12 h the total numbers of S. mutans cells (blue) and spectinomycin-resistant S. mutans cells (red) were determined via serial dilution and agar plating. This assay was repeated five times for approximately 50 generations.

Confident that the GFP plasmids were segregationally stable under the conditions tested, we next tested the strength of GFP expression of each individual construct using immunodetection and plate reader measurements. As with the integrated reporters, Western blotting experiments confirmed the expression of sfGFP, with no visible degradation of the protein, in the mid-exponential phase of growth (Fig. 3A). Next, sfgfp reporter strains were cultured in a microplate reader with fluorescence, and the optical density was recorded for 16 h. The comparative strengths of the fluorescence signals matched those of the integrated constructs, with Pveg giving the strongest signal (330,000 RFU) and Pldh giving the weakest signal (150,000 RFU) (Fig. 3B).

FIG 3
FIG 3

Characterization of plasmid-borne promoter-sfgfp constructs in S. mutans. (A) Western blot detection of sfGFP using an anti-sfGFP antibody against whole-cell extracts collected at an OD600 of 0.5. (B) Fluorescence intensity (circles) and growth (lines) measured simultaneously using a microplate assay.

Measuring the signal heterogeneity of the two GFP systems.While these GFP constructs could be used for visualizing S. mutans, we were also interested in synthetic biology applications from two perspectives: (i) using the sfgfp variants as readouts for promoter activity and (ii) using the constitutive promoters for overexpression of proteins or small RNAs. In both cases, it was important to know how heterogeneous the signals were, as experimental outcomes can be altered if promoters and/or the type of construct (integrated versus plasmid borne) may influence the signal-to-noise ratios in the system(s). Therefore, constructs were grown to an OD600 of 0.5, aliquots of the cultures were deposited on a microscope slide and covered with a coverslip, and the sfGFP intensity in single cells was measured for >1,000 cells per sample (see Materials and Methods). The data for these experiments were plotted as histograms (Fig. 4), with the number of cells per bin being provided on the x axis and the sfGFP intensity being provided on the y axis. When plotted in this manner, it was possible to observe the range of sfGFP fluorescence across the population. It was clear from these data that the range for sfGFP fluorescence was much larger for plasmid-borne sfgfp constructs than for constructs integrated into the chromosome (Fig. 4). Next, we calculated the Fano factor (σp2/<p>) for each construct, which provides a measure of the variance (square of the standard deviation [σp2]) over the mean value of expression (<p>) and can be used to measure gene expression noise (29). Phenotypic noise (Fano factor) was increased when the constructs were plasmid-borne (Table 1). Noise was also increased with increased promoter activity. For example, the Fano factor for the integrated version of Pveg-sfgfp was almost 8 times that for the Pldh-sfgfp integrated construct.

FIG 4
FIG 4

Single-cell fluorescence intensities of integrated and plasmid-borne constructs. For each promoter-sfgfp fusion, the distribution of fluorescence strength (x axis) is plotted as a histogram, with the results for both integrated and plasmid-borne constructs being shown on the same graph. sfGFP intensities are higher and more widely distributed for the plasmid-borne constructs than for the integrated constructs.

View this table:
TABLE 1

Fano factor measurements of single-cell sfGFP heterogeneity

Fluorescent protein constructs for use in oral streptococci.We next wanted to build several plasmid-based constructs to aid the visualization of other oral streptococci, choosing the E. coli-streptococcal shuttle vector pDL278 in anticipation that it would replicate and be stable in a variety of streptococci. We also selected the P23 promoter with the optimized RBS sequence, along with three different fluorescent protein sequences to incorporate sfGFP, DsRed-Express2 (30), and a yellow fluorescent protein variant, citrine, that is more resistant to acid quenching (31). Both the DsRed-Express2 and citrine gene sequences were downloaded from the SnapGene software website, codon optimized for use in S. mutans using the IDT codon optimization tool, synthesized in combination with the P23 promoter, and cloned into pDL278, similarly to the sfGFP constructs. The vectors pDL278_P23-sfgfp, pDL278_P23-DsRed-Express2, and pDL278_P23-citrine were then transformed into three different streptococci: S. mutans UA159; Streptococcus gordonii DL1; and Streptococcus A12, which is phylogenetically situated between Streptococcus australis and Streptococcus parasanguinis (32).

We detected fluorescence signals from all three constructs in a microplate reader during an 18-h incubation, as well as by fluorescence microscopy (Fig. 5). From these observations, we noted a strain-specific diversity in the fluorescence intensity, depending on the reporter construct tested. For example, S. gordonii displayed higher levels of the sfGFP signal than S. mutans (1.5-fold greater) and strain A12 (4.4-fold greater) (Fig. 5A and B), whereas S. mutans produced a higher citrine signal than S. gordonii (2.3-fold increase) (Fig. 5A and B). In fact, A12 failed to express the P23-citrine construct under the conditions tested. The signal production patterns of DsRed-Express2 for the three strains were similar to those of sfGFP (Fig. 5A, B, and C). The growth of both S. mutans and S. gordonii was not substantially reduced by the reporters in either defined medium (Fig. 5D and E) or rich medium (see Fig. S1A and B in the supplemental material). The reporters, most notably, pDL278_P23-citrine, did slow the growth of Streptococcus A12 (Fig. 5F). Citrine expression in Streptococcus A12 appeared to be toxic to the organism, and this might explain the lack of yellow fluorescence detected during microscopy in this organism (Fig. 5I). The stability of the pDL278_P23-citrine plasmid in Streptococcus A12 is also discussed below.

FIG 5
FIG 5

Detection of fluorescence among different oral Streptococcus spp. in monoculture. The robustness of fluorescent reporter plasmids was evaluated in S. mutans UA159 (top row), S. gordonii DL1 (middle row), and Streptococcus A12 (bottom row). For each species, we measured the maximum fluorescence intensity of reporter plasmids using a Synergy 2 multimode plate reader after 18 h of growth (A, S. mutans; B, S. gordonii; C, Streptococcus A12) (A.U., arbitrary units). Growth measurements in defined medium were taken from the reporter strains using a Bioscreen C automated growth curve analysis system (D, S. mutans; E, S. gordonii; F, Streptococcus A12). Colored circles indicate strain backgrounds, as follows: wild-type strains in gray, pDL278_P23-sfgfp strains in green, pDL278_P23-citrine strains in yellow, and pDL278_P23-DsRed-Express2 strains in red. Fluorescence was also visualized using a confocal microscope, once the cell cultures had reached stationary phase (OD600, ∼0.7 to 0.8) in all three species (G, S. mutans; H, S. gordonii; I, Streptococcus A12). Images are overlays of the bright-field and fluorescence channels. We did not detect yellow fluorescence for Streptococcus A12/pDL278_P23-citrine.

S. mutans UA159 and S. gordonii DL1 are model organisms for oral streptococcal research and are lab adapted, whereas Streptococcus A12 is a novel streptococcal isolate that has not been highly passaged. We wanted to determine if aberrant plasmid stability of pDL278 in Streptococcus A12 might be responsible for the lower fluorescence signal from our constructed vectors. For these experiments, we also included Streptococcus sanguinis SK150, which returned spectinomycin-resistant colonies after transformation with our vectors, but a fluorescent signal could not be detected for S. sanguinis SK150 when assayed, nor could fluorescence be observed by microscopy. Our sampling over a total of approximately 50 generations for S. mutans UA159, S. gordonii DL1, S. sanguinis SK150, and Streptococcus A12 verified that the pDL278_P23-DsRed-Express2 plasmid was stable in all four organisms under the conditions tested, as the numbers of CFU were similar on BHI and BHI-spectinomycin agar at all time points (Fig. S2A). When the presence of pDL278 after 50 generations was checked by PCR using pDL278-specific primers, we found that only a faint band was present in S. sanguinis SK150, even at the beginning of our plasmid stability experiment (time zero) (Fig. S2B). For Streptococcus A12, we could not confirm the presence of pDL278_P23-citrine, even though the strain had become spectinomycin resistant (Fig. S3). This is the most likely reason why no citrine could be detected in Streptococcus A12 in either our microplate reader or our microscopy experiments (Fig. 5G and I). These observations indicate that these particular constructs may not be suitable for all Streptococcus species. Therefore, constructs should be rigorously tested, and changes to the vector, promoter, or fluorescent reporter may be necessary prior to use in an isolate of interest.

Application of the fluorescent tools for biofilm studies.As we were able to detect fluorescence in all three species tested for both the sfGFP and DsRed-Express2 constructs, we next moved to test whether we would be able to detect sufficiently strong signals in a mixed-species biofilm model for future biofilm studies. We chose to examine three different mixed-species biofilms grown over the course of 72 h, with imaging being performed every 24 h: S. mutans/pDL278_P23-sfgfp and S. mutans/pDL278_P23-DsRed-Express2 (S. mutans-only control biofilms), S. mutans/pDL278_P23-sfgfp and S. gordonii/pDL278_P23-DsRed-Express2, and S. mutans/pDL278_P23-sfgfp and Streptococcus A12/pDL278_P23-DsRed-Express2. For this experiment, biofilms were grown in ibidi μ-Slide 8-well chamber slides using chemically defined medium (CDM) supplemented with 20 mM glucose and 5 mM sucrose. After 24 h of incubation, the growth medium of the biofilm was replaced every 12 h (Fig. S4). Biofilms were analyzed by confocal microscopy in situ with no washing or medium replacement so as not to disrupt the biofilm structure.

Over the course of 72 h, we were able to visualize both bacterial species within the growing biofilms (Fig. 6). All three species tested were able to produce a strong, stable signal for imaging over the course of the experiment. Using this technique, we observed differences in biofilm architecture between our three different dual-species groups, especially the distribution of bacteria within the growing biofilms. Microcolony formation was observed within our S. mutans-only control biofilm (Fig. 6A). S. mutans cells at 24 h were visualized within microcolonies and did not completely coat the surface of the glass slides, leaving uncolonized sections of the substratum. Over the course of 72 h, these microcolonies grew in size to 20 to 30 μm in diameter. In contrast, no large microcolony formation for S. mutans was observed when it was cocultured either with S. gordonii (Fig. 6B) or with Streptococcus A12 (Fig. 6C). We found that the distribution of either S. gordonii or Streptococcus A12 differed when it was cocultured with S. mutans. S. gordonii was found to cover much of the surface at 24 h, with S. mutans being found in only small microcolonies. The S. mutans microcolonies eventually expanded over the course of the experiment, with S. gordonii ceding surface area. This was not the case in dual-species biofilms with Streptococcus A12. At 24 h, S. mutans predominately covered the substratum. We noted that Streptococcus A12 was visualized alongside of, but not within, the S. mutans microcolonies at early time points. Streptococcus A12 became associated with the sides of the S. mutans microcolonies as they expanded over time but was not found within the microcolonies. Analysis of these images by the use of the Comstat2 program confirmed these observations, as we found the biomass to decrease over time for both of our S. gordonii DL1 and Streptococcus A12 mixed-species biofilms but to increase in our S. mutans-only control biofilm (Fig. 6D). Similarly, the maximal thickness increased within our S. mutans single-species biofilm over time, whereas the thickness only moderately changed in our S. gordonii DL1 or Streptococcus A12 biofilm group (Fig. 6E). We were also able to obtain mixed-species biofilm images from citrine- and DsRed-tagged bacteria (Fig. S5). The excitation and emission spectra of sfGFP and citrine overlap, which can make these proteins hard to detect with optimal brightness in the same experiment. However, a multispecies biofilm of bacteria colored by GFP, yellow fluorescent protein (YFP), and red fluorescent protein (RFP) might be possible with a confocal microscope that is capable of spectral imaging and linear unmixing.

FIG 6
FIG 6

Images of dual-species biofilms. (A to C) Selected 3D reconstructions of maximum-intensity z-section confocal microcopy images of dual-species biofilms of either S. mutans/pDL278_P23-sfgfp and S. mutans/pDL278_P23-DsRed-Express2 (control biofilm) (A), S. mutans/pDL278_P23-sfgfp and S. gordonii/pDL278_P23-DsRed-Express2 (B), and S. mutans/pDL278_P23-sfgfp and Streptococcus A12/pDL278_P23-DsRed-Express2 (C) at the 24, 48-, and 72-h time points. Images are of 128-μm sections of the fluorescence range within the biofilm collected at 1-μm intervals using a 63× oil objective lens (numerical aperture, 1.40). Biofilms were grown in chemically defined medium (CDM) supplemented with 20 mM glucose and 5 mM sucrose. At 24 h and every 12 h thereafter the spent medium was replaced with fresh medium for the length of the experiment. The biofilm images from time course experiment were reconstructed with Imaris software (6.4.0) using shadow projections. (D and E) The biofilm images from the time course experiment were analyzed by the Comstat2 (v2.1) program for biomass (D) and maximum thickness (E). Green line, S. mutans/pDL278_P23-sfgfp and S. mutans/pDL278_P23-DsRed-Express2 dual-species biofilms (control biofilm); orange line, S. mutans/pDL278_P23-sfgfp and S. gordonii/pDL278_P23-DsRed-Express2 dual-species biofilms; blue line, S. mutans/pDL278_P23-sfgfp and Streptococcus A12/pDL278_P23-DsRed-Express2 dual-species biofilms.

Another attractive option for research studies is the direct quantification of bacteria within the biofilm using flow cytometry for detection of these fluorescent reporters rather than using more traditional and labor-intensive methods, such as enumeration of the CFU on selective media. The use of flow cytometry can save time, as data collection is extremely rapid and can be more cost-effective because of the reduction in the amount of supplies, e.g., agar and plastics, used. We analyzed our 24-h dual-species biofilms by flow cytometry to determine if our constructed reporters would be viable for such an approach (Fig. 7). Indeed, a clear separation between our sfGFP- and DsRed-producing strains could be observed, such that we were able to determine the proportion of each species within our biofilm samples. Additionally, the total number of cells within the population that were analyzed as containing either sfGFP or DsRed was >99%, suggesting that the constructs and fluorescent proteins were stable enough during biofilm growth to allow for accurate counting. Our S. mutans cocultured control biofilm contained a roughly equal proportion of sfGFP and DsRed, as expected (Fig. 7A). However, the proportions of S. gordonii DL1 (Fig. 7B) and Streptococcus A12 (Fig. 7C) were different when they were grown with S. mutans: S. gordonii DL1 cells represented 83% of the biofilm cells at 24 h when S. gordonii DL1 was grown with S. mutans, whereas Streptococcus A12 cells comprised only 2% of the total cells measured at the same time point. These data correlate with what was observed by confocal microscopy at 24 h. In all, these observations highlight the feasibility of the use of the constructed fluorescent reporter strains for biofilm research not only to reveal differences in biofilm architecture or spatial arrangement when coculturing different streptococcal species together but also to allow direct quantification by flow cytometry.

FIG 7
FIG 7

Quantification of dual-species biofilms by flow cytometry. Histogram of cell counts from flow cytometry analysis of the pDL278_P23-DsRed-Express2 reporter on the left and a colored dot plot of both the pDL278_P23-DsRed-Express2 and pDL278_P23-sfgfp fluorescence intensity on the right of the following biofilms grown for 24 h: S. mutans/pDL278_P23-sfgfp and S. mutans/pDL278_P23-DsRed-Express2 (control biofilm) (A), S. mutans/pDL278_P23-sfgfp and S. gordonii/pDL278_P23-DsRed-Express2 (B), and S. mutans/pDL278_P23-sfgfp and Streptococcus A12/pDL278_P23-DsRed-Express2 (C). The green and red colored markers in the histogram denote the DsRed-negative and DsRed-positive cell populations that were used to determine the percentage of each within the sample. A total of 50,000 cells were counted in three independent replicates for each experiment.

DISCUSSION

With this study we aimed to provide new fluorescence visualization tools for S. mutans and other oral streptococci. We also used this study as an opportunity to test the relative strength of four constitutive promoters and determine how consistently these vectors produced fluorescence across a population of cells (noise). Lastly, we used these fluorescence tools to label oral streptococci so that they could be differentiated by using microscopy in a coculture biofilm system. Collectively, we achieved the construction of fluorescent reporter plasmids that are conformationally and segregationally stable with high FP expression for use in a broad range of oral streptococci and that should be valuable for gathering information on bacterial biofilm behaviors and interspecies interactions.

All four of the promoters tested in this study were constitutive and gave measurable levels of gene expression in S. mutans. Of the four, the veg promoter (Pveg) of B. subtilis, which included an optimized SD sequence, was the strongest. In a previous study by Biswas et al. (19), Pveg was also shown to have high activity in S. mutans. The other three promoters, P23, P3, and Pldh, were also active but showed moderately lower levels of transcriptional activity (approximately half the fluorescence intensity) compared with that yielded by the Pveg constructs. For synthetic biology applications, it would be useful to characterize weakly, moderately, and highly expressed promoters in S. mutans. The promoters that we tested here exhibit moderate to high levels of gene expression, but the platforms that we have developed could be used to screen for weaker or stronger promoters in a more comprehensive approach. It is important to note that optimal expression for one protein (e.g., GFP) might not correlate with that for another because of differences in 5′-end mRNA folding affecting mRNA stability and translation initiation (33). This is an especially important consideration here, as we did not use the same RBS for each promoter-sfgfp fusion. Therefore, it is difficult to determine which part of the promoter/5′ untranslated region is responsible for the changes in sfGFP levels across the four constructs. Despite these limitations, the data should be valuable in selecting which promoter-FP construct would be most helpful for particular applications in S. mutans.

The location of the promoter-sfgfp constructs, whether integrated in the chromosome or on a multicopy plasmid, did not alter the promoter strength rankings (Pveg displayed the highest activity). There was a measurable increase in the signal intensity and noise for the plasmid constructs compared with those for the integrated versions. Plasmids are typically present in multiple copies per cell, and this is the most likely explanation for the increased sfGFP fluorescence compared with that achieved with single-copy genome integration. pDL278 is based on the streptococcal plasmid pVA380-1, which has a plasmid copy number of ∼25 (34). The wider distribution of sfGFP fluorescence between cells in a population for pDL278 constructs is not a concern for strain marking. However, it could add stochasticity to experiments where it is not needed (e.g., promoter-reporter or overexpression studies). Increased population heterogeneity is most likely associated with variability of the actual plasmid copy number per cell, which could be influenced by segregational instability (e.g., random transmission to daughter cells or the formation of HMW multimers) or structural issues (e.g., replicative infidelity or illegitimate recombination, leading to plasmid rearrangements) (27, 35). Our single-cell analysis should aid researchers in selecting the optimal vectors and/or promoters for synthetic biology applications in S. mutans.

Having determined the relative expression from the promoter-RBS combinations and compared the noise levels between cells in a population from chromosomal and extrachromosomal sfGFP constructs in S. mutans, we were next able to use this knowledge to create various GFP, RFP, and YFP reporters in S. mutans and other oral streptococci. We selected the E. coli-streptococcal shuttle vector pDL278 as the backbone for reporter constructs because it replicates in oral streptococci and cloning can be performed in E. coli (24). While these constructs worked well in most of the strains under the conditions tested, we did observe species- and gene-specific stability issues. Segregational and/or conformational plasmid stability differences between bacterial species or gene inserts have been observed by others (36). However, we anticipate that the constructs will function well across the majority of transformable oral streptococci and will allow timely fluorescent protein tagging of strains of interest. The reporter methods developed here offer several advantages compared to other techniques, including the imaging of bacteria without addition of exogenous dyes (e.g., LIVE/DEAD staining or fluorescent in situ hybridization), live cell imaging, visualization of biofilm cells without disruption of the biofilm structure, and straightforward dual labeling of strains in coculture experiments. Fluorescent reporter systems also have some weaknesses. Fluorescent proteins are sensitive to certain environmental conditions (e.g., acidic conditions or low oxygen concentrations), and they can photobleach during extended excitation periods (the rate is protein specific) (37). It would be of interest to test these reporters in more diverse environments to determine their activities under more conditions that could conceivably be encountered in a human host.

Using our newly designed reporters, we were able to visualize two Streptococcus spp. simultaneously within biofilms. There is a renewed interest in the study of microbial interactions within different microbiome communities that colonize the human body. One recent example is the exploration of the community structure within supragingival dental plaque (38). Of interest are the interactions between the early colonizers of the oral biofilm, consisting largely of Streptococcus and Actinomyces species that efficiently attach to the salivary pellicle. An inverse association exists between the abundance of these commensal species and that of cariogenic bacteria, such as mutans group streptococci. An increase in the proportions of cariogenic bacteria under acidic conditions promotes the lower diversity of organisms within the biofilms and demineralization of the tooth enamel (12). Exploring the interactions between these health-associated commensals and acidogenic bacteria is critical for our understanding of the pathogenic potential of S. mutans and for the development of novel therapeutic interventions, e.g., probiotic strains that antagonize the growth of S. mutans through multiple pathways (32, 39). The construction and use of the various streptococcal fluorescent strains described here should allow for better insight into the behaviors of multispecies biofilms.

In summary, this study provides new tools that should enable researchers to shed light on critical traits of S. mutans and microbe-microbe interactions. Our detailed investigation of several constitutive promoters should serve as a basis for the correct selection of tools for synthetic biology applications in S. mutans. In addition, the plasmid-borne fluorescence reporters will allow the timely tagging of streptococci and be beneficial in various experimental systems.

MATERIALS AND METHODS

Strains and growth conditions.The strains of Streptococcus mutans and other Streptococcus spp. (Table 2) were cultured in brain heart infusion (BHI) broth (Difco) or chemically defined medium (FMC) (40) supplemented with either 25 mM glucose (for planktonic growth) or 20 mM glucose and 5 mM sucrose (for growth as biofilms). Streptococci were grown in a 5% CO2 aerobic environment at 37°C, unless stated otherwise. Strains of Escherichia coli (Table 2) were routinely grown in lysogeny broth (LB) with slight modifications (Lennox LB; 10 g/liter tryptone, 5 g/liter yeast extract, 5 g/liter NaCl) at 37°C with aeration. The following antibiotics were added to the growth media at the indicated concentrations: kanamycin, 1.0 mg/ml for S. mutans and 50 μg/ml for E. coli; spectinomycin, 1.0 mg/ml for S. mutans and 50 μg/ml for E. coli; and ampicillin, 100 μg/ml for E. coli. The strains and plasmids are listed in Table 2.

View this table:
TABLE 2

Strains and plasmids used in this study

Construction of plasmids.Each construct was created using gene synthesis (Integrated DNA Technologies) and restriction enzyme cloning. Previously published promoter sequences were combined with an sfgfp sequence (which is codon optimized for Bacillus subtilis but which has a good intensity in Streptococcus pneumoniae). At the 5′ end of the sequence we included a SacI recognition site, and at the 3′ end we included an SphI recognition site. These restriction sites facilitated the cloning into the multicopy E. coli-streptococcal shuttle plasmid pDL278 (24) and the integration plasmid pPMZ (16). For pPMZ, restriction digestion removed the PaguR promoter and the lacZ gene. Cloning into these plasmids was carried out in NEB 10-beta competent E. coli (New England BioLabs). After cloning, the plasmids were purified using a QIAprep spin miniprep kit (Qiagen) and transformed into competent S. mutans UA159 with selection on the appropriate antibiotic (for pDL278, spectinomycin; for pPMZ, kanamycin). For the DsRed-Express2 and citrine constructs, a similar protocol was followed, except that the sfGFP sequence was replaced with the sequence for DsRed2-Express2 or citrine.

Microtiter plate assays.For monitoring of cell density and GFP fluorescence over time, S. mutans or strains of commensal streptococci were processed as follows. Strains were cultured to an OD600 of 0.5 and then diluted 1:100 into FMC with glucose as the sole carbohydrate source. In quadruplicate, 200 μl of the cultures was placed in dark-sided, 96-well microtiter plates. For each fluorescent strain, a control that harbored the plasmid or integrated cassette without the gene for the fluorescent protein was used to allow subtraction of the background fluorescence of the cells and/or medium components. After loading of the cultures, the 96-well plate was placed in a Synergy HT microtiter plate reader (BioTek). The fluorescence and optical density (absorbance at 600 nm) were measured at 30-min intervals using Gen5 software (BioTek). The fluorescence settings were as follows: for GFP, excitation at 485 nm, emission at 525 nm, and a sensitivity of 65; for DsRed-Express2, excitation at 560 nm, emission at 590 nm, and a sensitivity of 65; and for citrine, excitation at 510 nm, emission at 530 nm, and a sensitivity of 65. Data readings were collected, and the background fluorescence or OD600 was subtracted prior to data visualization using GraphPad Prism (v7) software (GraphPad Software).

Western blotting.The bacterial strains were grown in FMC at 37°C to an OD600 of 0.5. Cells were harvested by centrifugation at 3,500 × g for 10 min, spent medium was discarded, and cell pellets were stored overnight at −20°C. On the following day, the pellets were resuspended in 100 μl lysis buffer (60 mM Tris, pH 6.8, 2% sodium dodecyl sulfate [SDS]) and transferred to a screw-cap microcentrifuge tube that contained 100 μl 0.1-mm ice-cold glass beads. Samples were homogenized in a bead beater for 30 s three times with 5-min intervals on ice. The samples were then centrifuged at 8,000 × g for 10 min at 4°C. The supernatants, which contained the S. mutans proteins, were carefully removed and placed into a fresh 1.5-ml microcentrifuge tube. The protein concentration was determined using a bicinchoninic acid (BCA) assay following the supplier’s protocol (Pierce), with purified bovine serum albumin used as the standard. Ten micrograms of each lysate was diluted in 4× SDS loading buffer, and the mixture was boiled for 10 min. Protein samples were separated by SDS-polyacrylamide gel electrophoresis (PAGE) and then transferred to a polyvinylidene difluoride (PVDF) membrane using a Trans-Blot Turbo transfer system (Bio-Rad). Green fluorescent protein was detected using a polyclonal anti-GFP antibody (1:5,000 dilution; Millipore Sigma) and a goat-anti-rabbit immunoglobulin G (IgG) antibody (1:5,000 dilution; SeraCare Life Sciences, USA). Western blot signals were detected using a SuperSignalWest Pico chemiluminescent substrate kit (Thermo Fisher Scientific) and visualized with a FluorChem 8900 imaging system (Alpha Innotech, USA).

Plasmid stability assay.Streptococcal strains carrying derivatives of pDL278 were cultured in BHI with or without spectinomycin for 12 h. Every 12 h (for a total of 5 passages, or approximately 50 generations), the strains were subcultured 1:1,000 into fresh medium. At each passage time point, an aliquot from the 12-h culture was serially diluted onto BHI agar with and without selection (spectinomycin). After 48 h, the colonies on these plates were enumerated and the numbers of CFU in the presence or absence of selective pressure were calculated. Experiments were repeated at least three times.

Single-cell GFP measurements.S. mutans cells were washed twice in phosphate-buffered saline (PBS), pH 7.2, from overnight cultures and diluted 20-fold into FMC growth medium. At an OD600 of 0.5, the culture was gently sonicated using a Fisher Scientific FB120 sonic dismembrator probe in order to dechain the cells, and then 4 μl of each sample was pipetted onto a glass coverslip and imaged on a Nikon TE2000-U inverted phase-contrast microscope using a Nikon C-FL GFP HC HISN zero-shift filter cube with a Nikon Intensilight mercury arc lamp source for GFP fluorescence excitation and detection. Images were collected on a Photometrics Prime CMOS camera and analyzed by a previously described method (41).

Biofilm time course imaging and analysis.To begin the biofilm experiments, overnight cultures of selected strains grown in BHI medium with appropriate antibiotics were washed and diluted 1:20 into fresh CDM (42) with glucose as the sole carbohydrate source and were then grown to mid-exponential phase (OD600 = 0.5). An aliquot (10 μl) of each strain was added to 1 ml of CDM supplemented with 20 mM glucose and 5 mM sucrose, and 350 μl of these mixtures was used to inoculate one chamber of an ibidi μ-Slide 8-well chamber slide (catalog number 80826; ibidi GmbH). CDM is used for these experiments because it is more strongly buffered than FMC. The use of CDM rather than FMC reduces the impact of the acidification of the medium on fluorescent protein stability and diminishes the antagonistic capabilities of S. mutans against commensal organisms. The samples were incubated at 37°C in a 5% CO2 aerobic atmosphere, and the time of inoculation was represented as time zero. Every 24 h (times of 24, 48, and 72 h), the biofilms were removed from the incubator and imaged by confocal microscopy (see Fig. S2 in the supplemental material). At 24 h and every 12 h thereafter, the spent medium was gently removed from the biofilms and replenished with fresh medium. Biofilm images were acquired using a spinning-disk confocal system connected to a Leica DM IRB inverted fluorescence microscope equipped with a Photometrics cascade-cooled electron-multiplying charge-coupled-device (EMCCD) camera. GFP fluorescence was detected by excitation at 488 nm, and emission was collected using a 525-nm (±25-nm)-band-pass filter. Detection of DsRed-Express2 fluorescence (RFP) was performed using a 545-nm excitation laser and a 590-nm (±25-nm)-band-pass filter. All z-sections were collected at 1-μm intervals using a 63× oil objective lens (numerical aperture, 1.40). Image acquisition and processing were performed using VoxCell software (VisiTech International). The three-dimensional (3D) reconstruction of selected biofilm images was completed using Imaris software (v6.4.0; Bitplane), and analysis was completed using the Comstat2 program (v2.1; www.comstat.dk) (43). For each biofilm and time point, five images were acquired from different parts of the biofilm and used for image analysis.

Flow cytometry.Dual-species bacterial biofilms were grown for 24 h before being harvested, washed, and resuspended in 1× PBS. Cells were sonicated in a water bath sonicator for 4 intervals of 30 s each while resting on ice in 5-ml polystyrene round-bottom tubes. Samples were analyzed using an LSR II (BD Biosciences) flow cytometer. Forward and side scatter signals were set stringently to allow sorting of single cells. In total, 5 × 104 cells were counted from each event at a maximum rate of 2 × 103 cells per second, and each experiment was performed in triplicate. Data were acquired for unstained cells and single-color-positive controls so that the data collection parameters and compensation could be properly set. The data were collected using FACSDiva software (BD Biosciences) and analyzed with FCS Express (v4) software (De Novo Software). Gating for quadrant analysis was selected by using a dot density plot with forward and side scatter, with the gates being set to capture the densest section of the plot. x- and y-axis data represent logarithmic scales of fluorescence intensity (arbitrary units).

Addgene availability.The following plasmids are stored on Addgene, the not-for-profit plasmid repository: pPM_Pveg-sfgfp (Addgene number 121503; Pveg-sfgfp integration vector for S. mutans), pDL278_P23-sfgfp (Addgene number 121504; P23-sfgfp E. coli-streptococcal shuttle vector), pDL278_P23-DsRed-Express2 (Addgene number 121505; P23-DsRed-Express2 E. coli-streptococcal shuttle vector), and pDL278_P23-citrine (Addgene number 121506; P23-citrine E. coli-streptococcal shuttle vector).

Accession number(s).The DNA sequences of each synthetic construct are deposited at NCBI under accession numbers MK301203 for Pveg-gfp, MK301204 for P23-gfp, MK301205 for Pldh-sfgfp, MK301206 for P3-sfgfp, MK301207 for P23-DsRed-Express2, and MK301208 for P23-citrine. These sequences are also available in the supplemental material.

ACKNOWLEDGMENTS

The research reported in this publication was supported by the National Institute of Dental and Craniofacial Research of the National Institutes of Health under award numbers R01 DE013239, R01 DE025832, R01 DE023339, T90 DE21990, F32 DE028469, and F30 DE028184.

We declare that there are no potential conflicts of interest.

FOOTNOTES

    • Received 14 March 2019.
    • Accepted 11 May 2019.
    • Accepted manuscript posted online 17 May 2019.

  • Supplemental material for this article may be found at https://doi.org/10.1128/AEM.00620-19.

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KEYWORDS

Streptococcus mutans
biofilm
fluorescence microscopy
green fluorescent protein
oral streptococci

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