Pipeline pigging sludge collected during a routine cleaning of a transmission pipeline transporting crude oil served as an inoculum to enrich for SRM (Gieg et al., 2020). Approximately 1 g of the pigging sludge was weighed into sterile serum bottles, and 50 mL of freshly prepared anoxic and sterile CSBK (Coleville Synthetic Brine medium K; Callbeck et al., 2013) medium containing 10 mM sulfate (added as sodium sulfate) and 3 mM volatile fatty acids (VFA: acetate, propionate and butyrate) were added. The serum bottles were stoppered with butyl rubber stoppers and sealed with aluminum crimps. The bottles were incubated at 30 °C, and sulfide concentrations were monitored to confirm sulfate-reducing activity. After a few transfers, propionate (13.3 mM, added as sodium propionate) was added as the sole carbon source (as separate experiments showed that this substrate supported faster growth than the VFA mixture; not shown). The culture was repeatedly transferred in CSBK medium containing 10 mM sulfate and 13.3 mM propionate to maintain its sulfate-reducing activity for use in all the experiments described here. The SRM enrichment culture was always freshly transferred and monitored for sulfate-reducing activity before transferring a second time for inoculation of flow cell experiments. Following the initial enrichment, the sulfate-reducing culture was maintained at room temperature (20–22 °C) and all experiments described herein were performed at this temperature.

The activity of planktonic and biofilm SRM cultures were determined by monitoring dissolved sulfide using a spectrophotometric sulfide assay (Cline, 1969). In some cases, sulfate concentrations were also measured by ion chromatography using procedures described by Okpala et al. (2017). Planktonic cultures were aseptically sampled using sterile, N₂-flushed needles and syringes with 10 μL of sampled fluid immediately added to 200 μL of 0.1 M zinc acetate to trap dissolved sulfide. During the operation of the flow cell experiments, direct sampling from the effluent tubing occurred daily during medium introduction, before the end of biocide treatment, and during medium re-introduction. To limit volatilization of sulfide into the atmosphere, effluent tubing was placed into a microcentrifuge tube containing 5 μL of 1 M ZnCl₂ to trap sulfide. A minimum of 0.5 mL effluent was collected. Sulfide concentrations were measured immediately after this sample collection.

To estimate the active number of cells in the SRM planktonic culture inoculated into the flow cell system, intracellular ATP concentrations were measured using a commercially available ATP test kit according to the manufacturer’s instructions (LuminUltra Technologies, Fredericton, NB). Briefly, samples were filtered, followed by cell lysis to release the ATP. The ATP and luciferin-luciferase reagent were mixed and measured on a luminometer (PhotonMaster, LuminUltra Technologies, Fredericton, NB). The conversion of RLUs, the initial output from the luminometer, to final units of microbial equivalents (ME) of ATP per unit of volume (mL) was done. Volume of sample, background readings, and standard of reagents were accounted for.

Sodium nitroprusside dihydrate was obtained from Sigma-Aldrich (Oakville, ON) and was freshly prepared before each use as a 10 mM (3,000 ppm) sterile anoxic stock solution by dissolving 0.015 g SNP in 5 mL deionized water. ADBAC was obtained as a commercial product from SUEZ North America. A 10,000 ppm ADBAC stock solution was anoxically prepared by dissolving 1 g ADBAC into 98 mL N₂-flushed water. Biocide stock solutions were then diluted to the desired concentrations into CSBK medium for use in the flow cell experiments.

The active SRM enrichment culture was used to determine the effects of different doses of SNP and ADBAC on planktonic sulfate-reducing cells. Sterile, anoxic CSBK medium containing 13.3 mM propionate and 10 mM sulfate was inoculated with 10% vol/vol of the enrichment culture. The sulfate and sulfide concentrations were monitored to determine the approximate mid-log phase of the culture (the point at which about half of the added sulfate has been reduced to sulfide). For the SNP tests, 20 mL of the mid-log phase culture were transferred with a N₂-flushed syringe into sealed 60 mL sterile serum bottles containing a N₂/CO₂ (90/10) headspace. The planktonic cells were immediately challenged with 0, 7.5, 15, 30, 75, or 150 ppm SNP (in triplicate) based on effective doses previously reported by Fida et al. (2018). All treatments were performed in triplicate. After the treatment, the cultures were monitored for 20 days for sulfide (from each replicate bottle) and sulfate (in a single bottle for each treatment) to ascertain the impact on sulfate-reducing activity. The ADBAC dosing tests were performed similarly but were amended with 0, 10, 50, or 100 ppm ADBAC, performed in triplicate. These doses were chosen to span a similar range as those tested for SNP. Sulfide concentrations were monitored over a period of 20 days, followed by a 10% transfer of each treatment inoculated into fresh CSBK medium. Sulfide concentrations of the newly transferred treatments were monitored for an additional 10 days to determine whether any viable (active) cells remained.

After biocide treatment or the re-introduction of CSBK medium to biofilms, the FilmTracer LIVE/DEAD Biofilm Viability kit (Thermo Fisher Scientific, Ottawa, ON) containing the two nucleic acid stains SYTO 9 and propidium iodide (PI), was used to differentially stain “live” or “dead” (membrane-compromised) cells, respectively. The staining procedure was performed according to the manufacturer’s instructions by slowly injecting the appropriate solutions into the flow cell. Stained biofilms were then immediately analyzed using confocal or two-photon microscopy. Different microscopes and data visualization systems were used for examining SNP- and ADBAC-treated biofilms based on availability and health precautions in place when the experiments were done (the SNP work was done when restrictions were in place due to the Covid-19 pandemic; the ADBAC work was conducted after restrictions were removed).

For the SNP-treated biofilm experiments carried out during the Covid restrictions, images were acquired using a Zeiss Axio Imager Z1 confocal laser scanning microscope LSM510 NLO with a 40x/0.8 NA objective (Zeiss, Inc., Germany), as described by Moo et al. (2013). The microscope used a 488 nm Argon laser to excite fluorescence, with a 520/20 green bandpass filter for Channel 0 and a 680/60 red bandpass filter for Channel 1. Z-stacks were acquired at 1-micron intervals and ImageJ (Schneider et al., 2012) was used to review and visualize the imaging data. Biofilms from the untreated (control), 300 ppm SNP, and 750 ppm SNP tests were visualized using this microscopy approach.

More recently, ADBAC-treated biofilm experiments were imaged separately on a multiphoton microscope (Bergamo 2, Thorlabs, USA) using ThorImageLS 3.2. Two-photon microscopy was used rather than standard confocal laser scanning microscopy due to its superior penetration into the thick biofilm sample (Neu et al., 2002; Vroom et al., 1999). Three random fields of view were imaged per condition (control, 50, 100, 250, and 500 ppm ADBAC). Fluorescence of SYTO 9 and PI was simultaneously excited via two-photon excitation using 920 nm from a Tiberius Ti: Saph laser (Thorlabs, USA). To separate the SYTO 9 and PI signals, the fluorescence emission was directed first to a primary dichroic (705 nm long pass), and then to a pair of GaASP photomultiplier tubes, each equipped with a bandpass filter (525/50; SYTO and 607/70; PI).

Large fields of view, corresponding to 640 by 640 μm were acquired using a 20×/1.0 NA (Olympus, Japan) objective. Z-stacks were acquired at 2-micron intervals, and images were recorded until where the fluorescence emission was weak and almost non-detectable. After acquisition, the image stacks were visualized using FIJI (Schneider et al., 2012) as well as Nikon Elements Analysis 4.60 software. Image quantification was carried out using Cell Profiler 4.28 (Stirling et al., 2021) by comparing the relative signal of the SYTO 9 and PI channels for every condition First, each fluorescence channel was segmented into foreground (positive signal corresponding to biofilm) and background using the FindPrimaryObjects module in CellProfiler. Settings applied included global Otsu using 3-class thresholding where only the highest intensity class was considered as foreground. The automated segmentation was verified by visual inspection. Objects were filtered so that only objects representing more than 3 pixels (corresponding to X by Y microns) were analyzed; this was to minimize measuring spurious fluctuations due to shot and other noise as fluorescence emission. The detected objects were then merged, and the fluorescence intensity in the SYTO 9 and PI images were measured for each image in each Z-stack. The integrated intensity, representing the total intensity signal over all the detected regions, was measured for each channel.

Biofilms removed from the glass slides within each flow cell were processed for amplicon sequencing. Prior to DNA extraction, biofilm samples were centrifuged for 10 min at 14,000 g, and resulting pellets were resuspended in 1 mL medium. Samples were then split into two 0.5 mL samples - one for treatment with propidium monoazide (PMA) (Biotium, Fremont, CA, USA) and one remaining untreated (NPMA). PMA is a membrane impermeant dye, known to bind to DNA only in compromised cells, preventing its extraction, thus DNA from PMA-treated samples can be a better reflection of living, intact cells in any given sample (Nocker et al., 2007). For PMA-treated samples, 1.25 μL PMA were added in a dark room. Samples were covered in tinfoil and placed on a rocker for 10 min to allow for PMA to penetrate cells with compromised membranes. Samples were then placed under a halogen light (500 W) on a bed of ice for 10 min and rotated every 2 min to allow for crosslinking of PMA to DNA in cells with compromised membranes, preventing its amplification during PCR. PMA-treated samples were centrifuged at 5000 g for 10 min, then resuspended in 200 μL of phosphate buffer prior to DNA extraction. Samples were stored at −20 °C until DNA extraction could be performed.

Genomic DNA was extracted using the FastDNA™ SPIN Kit for Soil (MP Biomedicals) according to the manufacturer’s protocols. DNA was quantified by fluorometry (Qubit, Invitrogen). The V4-V5 region of the 16S rRNA gene was amplified using a 2-step PCR protocol as described by Rachel and Gieg (2020). Amplicons were further processed and sequenced on an Illumina MiSeq (v2 300 cycle) at the Centre for Health Genomics and Informatics (University of Calgary, Canada). Resulting sequences (raw reads) were processed through a series of quality control steps to remove chimeras, etc. through a DADA2 pipeline in R (Callahan et al., 2016). The R code/pipeline for sequence data processing and visualization can be found at.¹ Supplementary Table 1 shows the number of raw reads and quality-controlled reads obtained for the sequencing results shown in this study. Quality reads were assigned as amplicon sequence variants (ASVs) to the lowest taxonomic level possible against the SIVLA 138. 2 database (Quast et al., 2012), and ASV plots were created using graphing functions in R studio. Amplicon sequencing data is available in GenBank (NCBI) under BioProject accession number PRJNA1284394.

A sulfate-reducing enrichment culture derived from pipeline pigging solids was used as the source of inoculum for all biofilm experiments. The culture was regularly maintained on propionate as the carbon and energy source and sulfate as the electron acceptor. Prior to each use as an inoculum for a biofilm experiment, the culture was transferred into fresh medium, and sulfide was monitored to ensure that it was active before it was transferred again during mid-log phase (when 4–6 mM sulfide was produced) to serve as the starting culture for inoculating the biofilms. We used the ATP assay to estimate the number of active cells being transferred and used for biofilm inoculation when sulfide values were at 4–6 mM, which typically took 5 to 7 days of incubation. Supplementary Figure 2 shows that during mid-log phase (e.g., at 6 days), the culture contained an average of 3.5 × 10⁷ active cells per mL. As the flow cells were inoculated with 2 mL of culture, all flow cells were inoculated with approximately 7 × 10⁷ cells for biofilm establishment.

The SRM planktonic enrichment used as the inoculum was sequenced twice (in late 2020 and early 2023), showing that the planktonic community was comprised of Desulfobulbus as the most abundant sulfate-reducer, with a lower relative abundance of Desulfomicrobium (Supplementary Figure 3). Desulfobulbus is a known propionate-utilizing sulfate-reducing bacterium (El Houari et al., 2017), aligning with the maintenance of this culture on propionate as the carbon and energy source. Desulfobulbus sp. are known to incompletely oxidize propionate to acetate and CO₂ that can be used by other taxa in communities (Ozuolmez et al., 2020). Fermentative bacteria such as Sphaerochaeta and Proteiniphilum, along with the acetogen Acetobacterium also consistently made up the community. Changes in some of the dominant taxa in the culture showed that the community shifted over time as it was transferred and maintained over 3 years on propionate and sulfate. Regardless of these community shifts, the culture continued to demonstrate sulfate-reducing activity so it could reliably be used to establish sulfidogenic biofilms over the time that this research was conducted.

A previous study reported that SNP added at doses as low as 15 to 30 ppm (depending on cell biomass) was effective in preventing sulfidogenic activity using a mixed SRM planktonic culture enriched from oilfield produced waters (Fida et al., 2018). This finding was confirmed in the present study using a different SRM enrichment obtained from pipeline pigging sludge, wherein SNP added at concentrations of 7.5 ppm or higher inhibited sulfide production and sulfate consumption compared to an untreated control (Supplementary Figure 4).

To determine whether similar effects of low SNP doses could also effectively treat existing SRM biofilms, flow cells were established to create active sulfate-reducing biofilms that could be challenged with different concentrations of SNP. The initial flow cell experiment tested concentrations of SNP ranging from 15 to 150 ppm (Figure 3A) alongside an untreated control. Following flow cell inoculation, active biofilms were established in the flow cells within approximately 5 days, after which medium flow was initiated in the system. This action initially lowered sulfide concentrations (due to dilution with medium) but sulfide production increased within the 5 days following the flow regime. It should be noted that prior to biocide dosing, these biofilms that were established in separate flow cells are biological replicates (n = 5), all showing similar trends in sulfide production (Figure 3A; Supplementary Figure 1). At day 11, SNP was added at the various concentrations for approximately 10 h, after which sulfide levels dropped in all SNP-treated biofilms; this was not observed in the control (untreated) biofilm (Figure 3A). After the period of SNP treatment, fresh CSBK was then added to each flow cell, and by the following day (approximately 24 h later), sulfide concentrations increased (rebounded) in all treated biofilms, showing that SNP doses ranging from 15 to 150 ppm were only transiently effective. This effect was also observed when these SNP concentrations were tested in separate flow cell experiments (Supplementary Figure 5), confirming the results. Untreated biofilms, and those treated with 30 or 75 ppm SNP were analyzed using 16S rRNA gene sequencing, which revealed that these biofilms were comprised primarily of Desulfobulbus, with lower relative abundances of Desulfomicrobium and other taxa (Figure 4A), like the community composition of the planktonic culture used as the inoculum for the flow cells (Supplementary Figure 3). PMA-treated samples did not show substantive differences compared to the non-PMA treated samples for the control or the 30 ppm SNP-treated biofilms. For the 75 ppm SNP-treated biofilm samples treated with PMA, Rhizobium became enriched in contrast to the non-PMA treated biofilms which was dominated by Desulfobulbus like the other samples (Figure 4A).

Since concentrations of SNP up to 150 ppm were only transiently inhibitory to existing SRM biofilms, a second flow cell experiment was carried out wherein established biofilms were untreated, or treated with 300 or 750 ppm SNP to determine whether these higher doses would result in more permanent inhibition of sulfidogenic activity in the biofilms. Like the lower-dose SNP experiment, replicate sulfide-producing biofilms were successfully established (n = 3, showing similar trends in sulfide production, Supplementary Figure 1), then treated on day 12 of flow cell operation (Figure 3B) for approximately 10 h with these higher SNP doses. Similar to the lower dose flow cell experiment, sulfide levels initially dropped in the SNP-treated biofilms but not in the untreated biofilm (Figure 3B). Following replenishment with fresh CSBK medium after this treatment time, sulfide concentrations again increased within 1 day, showing that these higher concentrations of SNP were also only transiently effective at inhibiting sulfide production (Figure 3B). Sequencing of the biofilms following the flow experiment showed that Desulfobulbus remained as the dominant sulfate-reducer in all samples (PMA-treated or not treated), but in this case, sequences identified as Pseudomonas were present as the second most abundant taxon (Figure 4B). There were few differences in the PMA-treated or non-PMA-treated samples from the 300 ppm or 750 ppm SNP-treated biofilms, but the control biofilm (untreated) showed different relative abundances of Pseudomonas which was present only in the PMA-treated sample (Figure 4B).

Replicate biofilm flow cells from this higher dose SNP experiment were also established to determine the efficacy of biocide penetration into the biofilms, as visualized by confocal laser scanning microscopy. A visual comparison of data sets recorded using identical settings reveals differences in the fluorescence emission from the SYTO 9 (live or viable; membrane-intact) and PI (dead or non-viable; membrane-compromised) channels. Using the LIVE/DEAD stain, an increased green signal indicates more “live” (membrane-intact) cells, while an increased red signal indicates more “dead”, or membrane-compromised cells (Tawakoli et al., 2013). As seen in Figure 5, control (untreated) biofilms show a primarily green signal, suggesting predominantly live (membrane-intact) cells while the 300 ppm-treated biofilm shows a mixture of green and red signal, suggesting the presence of both live and membrane-compromised cells. The 750 ppm-treated biofilm shows mostly membrane-compromised cells, showing effective SNP penetration into the biofilm. However, some green signals in the image suggests that some membrane-intact cells remain in the biofilm even after 750 ppm treatment (Figure 5). An increase in sulfide production soon after the biocide pressure was removed suggests that sulfate-reducing cells can recover after exposure to this higher SNP dose (Figure 3B).

ADBAC is well-known to be effective for treating microbial cells in a variety of applications and sectors (Boyce, 2023; Jennings et al., 2015; Merchel Piovesan Pereira and Tagkopoulos, 2019). To verify this efficacy, ADBAC was added to the planktonic SRM culture. Several replicates of this culture were established and demonstrated active sulfide production by 3 days of incubation. On day 3, various concentrations of ADBAC (0, 10, 50, or 100 ppm) were added to the active SRM culture and sulfide was monitored. As seen in Supplementary Figure 6, all tested concentrations of ADBAC inhibited sulfate-reducing activity, as indicated by a continuous decrease in sulfide levels compared to the untreated control. After 20 days of incubation, cultures were then transferred to fresh CSBK medium to determine whether removing the biocide pressure would allow for a resurgence in sulfate-reducing activity. Only the untreated control continued to show sulfide production - this activity was unable to re-establish from all the ADBAC-treated cultures (Supplementary Figure 6). These results show that doses as low as 10 ppm ADBAC were effective at inhibiting sulfide production in the planktonic SRM culture.

Like the SNP tests, biofilms were established in flow cell systems to determine the doses of ADBAC that would inhibit sulfide production from existing sulfate-reducing biofilms. For ADBAC, concentrations of 25 (not shown), 50, 100, 250, and 500 ppm were tested against established biofilms, with different biofilm analyses conducted at the end points of three experiments. Note that for each flow cell experiment conducted, 5 biofilms were established representing biological replicates that were then each treated with a different concentration of ADBAC (Supplementary Figure 1). Further, each experiment was repeated twice to determine reproducibility of the results (Supplementary Figures 7, 8 show the results of repeat experiments). For the first experiment, biofilms were established, treated with varying concentrations of ADBAC, and then the flow cells were opened soon after to determine microbial community composition. In the 5 replicate biofilms, sulfide production reached expected values (e.g., mid-log phase) by day 13 (Figure 6A). At this time, ADBAC was added to the biofilms at various concentrations for 14 h. Sulfide concentrations were measured immediately thereafter, and the flow cells were opened to collect the biofilms for microbial community analysis. In this experiment, sulfide concentrations dropped when all doses of ADBAC were applied compared to the untreated control, where sulfide concentrations continued to increase (Figure 6A). A replicate experiment subsequently conducted showed similar results (Supplementary Figure 7). 16S rRNA gene analysis of the various biofilms showed that Pseudomonas were of the highest relative abundance in the control (untreated) biofilms (75–80% relative abundance) with lower relative abundances of Desulfobulbus and other taxa (Figure 7A). However, when ADBAC was added at 50 or 100 ppm doses, Desulfobulbus became dominant (with or without PMA treatment). When 250 ppm ADBAC was added to the biofilms, the PMA-treated sample showed an increase in relative abundance of Pseudomonas compared to Desulfobulbus, while the opposite was true when PMA was not added (Figure 7A). DNA could not be extracted from the biofilm samples treated with 500 ppm ADBAC and then treated with PMA suggesting that only membrane-compromised/dead cells were present.

For the second flow-cell experiment testing ADBAC, the aim was to assess whether biofilms treated with ADBAC could resume sulfide production after ADBAC was removed from the system. An initial flow cell experiment was conducted to assess the reproducibility of sulfide production from all 5 biofilms, and the resumption of sulfide production following ADBAC treatment. Following ADBAC treatment, the biofilm treated with 50 ppm ADBAC resumed sulfide production, while all other concentrations inhibited further sulfide production (Supplementary Figure 8). As microbial community analysis was not carried out for this initial experiment, a second similar experiment was carried out that also assessed microbial community composition (Figures 6, 7). Like the first experiment, sulfide production, though variable across all biofilms, continuously increased following the introduction of CSBK medium (Figure 6B; Supplementary Figure 1). On day 46, each biofilm was then treated for 14 h with ADBAC at different doses. The temporary decrease in sulfide production observed on the day of ADBAC treatment was likely due to its dilution caused by adding medium, but sulfide production levels increased after the introduction of medium. Biocide addition resulted in a decrease of sulfide concentrations when ADBAC was added, similar to the previous experiment (Figure 6A, Supplementary Figure 7). To observe whether sulfate-reducing activity of treated biofilms could re-establish after treatment, CSBK medium was re-introduced for 10 days. In the 50 ppm ADBAC-treated biofilm, sulfide rebounded quickly, in a similar manner to the untreated control, showing that this concentration was ineffective at permanently inhibiting sulfide production from this biofilm, also previously seen (Supplementary Figure 8). The 100 ppm ADBAC treatment also showed a sulfide rebound, but in a slightly delayed manner (Figure 6B); this result differed from the previous experiment that showed no sulfide production following 100 ppm treatment (Supplementary Figure 8). In contrast, both the 250 ppm and 500 ppm biocide treated biofilms did not show sulfide production within 10 days after medium re-introduction, showing that these concentrations of ADBAC were comparatively effective at inhibiting sulfate-reducing activity and confirming the results of the replicate flow cell experiment (Supplementary Figure 8). After 10 days of medium flow, the flow cells were opened, and the biofilms were sampled for microbial community analysis by 16S rRNA gene sequencing. In this biofilm experiment, Desulfobulbus was the most abundant taxon, followed by Desulfomicrobium, as seen in the untreated biofilms (Figure 7B). When 50 ppm ADBAC were added, the community showed a decrease in the relative abundance in Desulfobulbus and an increase in Desulfomicrobium, which was even more pronounced when 250 ppm ADBAC were added (in the PMA-treated samples). Like the previous experiment, DNA could not be extracted from the PMA-treated biofilm sample retrieved from the 500 ppm ADBAC treatment (Figure 7B).

A final flow cell experiment was established in an identical manner to obtain samples for microscopy analysis (Supplementary Figure 7). Supplementary Figure 1 shows the results of the sulfide production from the 5 replicate biofilms prior to ADBAC addition, again showing a general upward trend of sulfide production despite variability amongst each biological replicate. Following the treatment of each flow cell with different concentrations of ADBAC, medium was allowed to flow through each biofilm for 30 min then biofilms in the flow cells were stained with the LIVE/DEAD Biofilm Viability kit for analysis by two-photon microscopy. Figure 8 illustrates representative 3D visualizations of the imaging data, with green representing SYTO 9 (live) and magenta representing PI (green and magenta were chosen as they are more visually accessible for people with color vision deficiencies). Cross-sectional views (edge on view of the Z-stack) are shown, allowing qualitative inspection of the profile of the various biofilms.

The LIVE/DEAD kit should reflect the relative abundance of viable cells in the total population. We noted that the overall SYTO 9 fluorescence decreased when the samples were treated with biocide. Figure 8A provides representative 3D visualizations for comparison across conditions. Each channel is displayed using identical settings for brightness and contrast. The untreated and 50 ppm ADBAC-treated biofilms emit primarily in the SYTO 9 channel indicating live cells (shown as green) and corresponding closely to the flow cell observations (Figure 6). ADBAC added at 100 ppm yielded signal primarily in the SYTO 9 channel as well, but also showed discrete regions where fluorescence was higher in the PI channel (non-viable, shown as magenta), aligning with the sulfide measurements showing more of a delay in the resurgence of sulfate-reducing activity. Finally, the 250 and 500 ppm treatments showed little emission in the SYTO 9 channel (live, detected as green), aligning with the flow cell data showing no sulfide activity rebounding following treatment with these ADBAC concentrations.

To compare the relative SYTO 9 and PI fluorescence emission for each condition, the ratio of the integrated intensity in the SYTO 9 channel was then divided by the sum of the integrated intensity of the SYTO 9 and PI channels. This normalization was required because the overall intensity in both the STYO 9 and PI channels decreased in the treated samples. The resulting ratio varied between 0 and 1, with 1 being the case when all the intensity is derived from the SYTO 9 channel only and conversely, when all the signal is derived from PI signal only.

Figure 8B illustrates a plot of the ratio of the integrated intensity (0–1) by z-slice (z-stack 20 to 100, with 20 being closer to the coverslip). A plot of the ratios for each conditions demonstrates a systematic trend, with a decreasing ratio when ADBAC was applied. A decreasing ratio implies that the SYTO 9 fluorescence emission is decreasing relative to PI emission, as would be expected when the cells become less viable. As shown in Figure 8B, the control and 50 ppm as well as the 250 ppm and 500 ppm results track closely, while the 100 ppm is intermediate between these two extremes. Thus, both the qualitative and quantitative imaging results from the ADBAC biofilm experiments align closely with the results of the biological assays.