A commercially available UVLED flow-through device, operating at 40 mJ/cm2, was examined for biofilm control on irrigation pipe material fed by wastewater effluent. Biofouling was monitored through total coliform counts, crystal violet (CV) staining, and ATP analyses. A UV fluence of 40 mJ/cm2 at 280 nm retarded biofilm formation; however, complete biofilm prevention by UV treatment was not achieved despite a high inactivation of planktonic cells. After 5 days of the study, the total coliform and CV biofilm quantification assays between the UV-treated and control bioreactor coupons were not statistically different. The total coliform counts indicated a stable biofilm cell concentration was reached; the CV assay showed biofilm biomass accumulation with time. The ATP results revealed higher coupon ATP on the UV-treated coupons than the control coupons by day 5. The results provoke an interesting discussion surrounding the contribution of viable cells, represented by total coliforms, and extracellular polymeric substance (EPS) to total biofilm biomass. This study also highlighted a need for further investigation into the relationship between ATP responses and complex UV-stress responses of diverse microbial communities as opposed to pure bacteria cultures.

  • A UV fluence of 40 mJ/cm2 at 280 nm slows biofilm formation on irrigation pipe material.

  • Bioreactor effluent total coliform concentrations and crystal violet coupon accumulation were not statistically different between UV-treated and control bioreactors after being fed wastewater effluent for five days.

  • ATP results revealed higher ATP on UV-treated coupons than control coupons after 5 days in bioreactors fed by wastewater effluent.

Water reuse is a promising water management strategy to alleviate water scarcity currently affecting every continent (UN-Water 2018). Water sources for potential reuse include stormwater, industry process water, and municipal wastewater (US EPA 2019). Uses for reuse water include environmental restoration, groundwater recharge, potable reuse, and agricultural irrigation. Wastewater reuse for agriculture is particularly promising; for instance, Israel recycles 87% of its wastewater for irrigation of both food and non-food crops (Marin et al. 2017). However, irrigating with wastewater effluent comes with challenges including biofouling of drip lines and other infrastructure meant to transport water.

Biofouling, or biofilm accumulation, is the unwanted deposition of microorganisms and sticky extracellular polymeric substances (EPS) on surfaces (Flemming 2002). Biofilms are established in five stages: (1) attachment of planktonic cells to a surface, (2) attached bacteria divide and EPS is excreted, (3) the EPS cell mixture expands, (4) the biofilm matures, and (5) the mature biofilm releases planktonic bacteria back into the environment (Flemming et al. 2007). Biofilms play a major role in many water reclamation and reuse strategies like membrane bioreactors and filtration systems used in specific reuse applications (Bishop 2007). The higher concentration of nutrients, organic matter, and microorganisms in wastewater effluent can aggravate the biofouling of water distribution systems, like drip irrigation lines (Tarchitzky et al. 2013).

Strategies for biofouling control during the distribution of effluent include filtration, flushing, and chlorination. Filtration is an important factor in preventing biofilm formation as it removes biofouling microorganisms, organic particulates, and some nutrients that facilitate their growth. Most manufacturers of drip irrigation systems recommend filtration for particulates; however, in the case of effluent, filter clogging is a big concern in addition to the clogging of the drip irrigation lines (Ravina et al. 1997). Regardless, filtration is still an important step during irrigation with reuse water to prevent immediate clogging of emitters by large particles (Adin & Sacks 1991).

Flushing is the process of increasing the water velocity in an irrigation system so the flow hydraulic shear force rapidly sheds attached biofilms while also slowing down the clogging frequently caused by the shredded biofilm falling inside the emitter (Li et al. 2015). Flushing studies have identified a method to effectively slow down emitter clogging in a reclaimed water drip irrigation system, and the longer period it was applied, the better controlling was observed (Li et al. 2015, 2019). However, increasing water velocity causes an immediate increase in bacteria numbers as the increase in shear stress resuspends biofilms (Lehtola et al. 2006). In one study examining the effects of flushing during irrigation with reclaimed water, lateral flushing failed to completely solve the emitter clogging problem and pairing flushing with additional measures was recommended (Li et al. 2015).

Chlorination is considered the most effective method of controlling biofouling during irrigation, but it has drawbacks (Song et al. 2017). Aside from being hazardous to humans, one study demonstrated adverse soil health effects when chlorine was applied in high concentrations during short-term use which is the recommended treatment for irrigation lines (Song et al. 2019). When chlorine and chloramine react with organic matter, like that found in wastewater effluent, it forms hazardous disinfection by-products (DBPs) (Doederer et al. 2014). Although the impact of DBPs on soil and crop health is not well studied, their ability to persist in aquatic environments has been reported (Rostad 2002).

UV disinfection is a common and effective treatment technology for drinking water, wastewater, and reclaimed water globally (Song et al. 2016; Nguyen et al. 2019). Recent advancements in UV light emitting diodes (UVLEDs) have allowed for compact, electrically efficient, and customizable point-of-use UV disinfection options. This includes the integration of UVLEDs into devices like toothbrush holders, coffee makers, and water coolers (Linden et al. 2019). UVLEDs have demonstrated an ability to treat wastewater effluent to US EPA guidelines for irrigation of processed food crops and non-food crops (Nguyen et al. 2019). LEDs are particularly well suited for the intermittent flow applications of agricultural irrigation due to their small size, no required warmup time, and ability to turn on/off without negatively impacting the lifetime and performance of the device (Song et al. 2016; Chen et al. 2017). Despite this, the impact of UVLED disinfection on biofouling during irrigation with wastewater effluent has not been studied.

The goal of this study was to examine a commercially available UVLED flow-through device for biofouling control on drip irrigation line material during the distribution of wastewater effluent. The effects of the UVLED device on biofilm growth in a bioreactor fed by effluent were tested through direct quantification assays (total coliforms) and indirect quantification assays (crystal violet (CV), adenosine triphosphate (ATP)). Before implementing the UVLED device for biofouling mitigation studies, a series of challenge tests were performed to establish a relationship between operating flow rate and fluence for varying water qualities in the UV LED flow-through devices. bacteriophage was chosen as the challenge organism because of its ability to survive high fluence levels, providing countable concentrations for the low-flow rate/high-fluence testing conditions (Malayeri et al. 2016). The log reduction values (LRV) of bacteriophage were determined in three water matrices described later with varying UV transmittances (UVT) at 254 nm (93, 80, and 70%).

Collimated beam experiments

Microbial stocks

bacteriophage

bacteriophage stock (ATCC 23631-B1) was propagated by GAP EnviroMicrobial Services Ltd (London, Ontario, Canada). An Escherichia coli K-12 (ATCC 23631) host culture was made in tryptic soy broth (TSB) with shaking at 35 °C. Once the E. coli reached early log-phase growth (OD600: 0.1–1.0), 1 mL of 1 × 1010 plaque forming units (PFU)/mL stock was added. After overnight incubation with shaking, the stock was centrifuged to remove cellular debris, leaving the phage in the supernatant. A concentration of approximately 1 × 1010 PFU/mL was achieved. Stocks were stored at 4 °C.

E. coli Famp

E. coli Famp (ATCC #700891) was used as the bacteriophage stock host for challenge testing. The Famp was separately streak-plated onto sterile nutrient broth (NB) (BD Difco™) agar plates and incubated for 20 h at 37 °C. One isolated colony was selectively removed with a sterile inoculation loop and suspended in 25 mL sterile NB for overnight incubation for 20 h at 37 °C with shaking (180 rpm). The overnight culture was then centrifuged at 5,000 rpm for 10 min. The supernatant was removed, and the pellet was resuspended in sterile NB. Twenty percent (by volume) of glycerol (100% v/v, Microlytic) was added to produce a final concentration of 9 × 108 colony forming units (CFU)/mL E. coli Famp. Stocks were stored at −80 °C.

Test water

Three water matrices with varying UV transmittances (UVT254nm) (93, 80, and 70%) were used for characterizing the UV devices for fluence delivered as a function of flow rate. bacteriophage stock was serially diluted into dechlorinated tap water to achieve a test water concentration of approximately 5 × 106 PFU/mL, resulting in a 93–95% UVT at 254 nm. UV transmittance was adjusted for the 80 and 70% UVT254nm waters using a mix of SuperHume (UAS of America, Lake Panasoffkee, FL, USA) and vanillin according to NSF/ANSI 55 standards. Dechlorination of test water was verified with a Hach DPD Free Chlorine colorimetric test.

Enumeration

US EPA 1602 method and Standard Methods 9224 B.3 were followed with minor adjustments including the addition of a streptomycin/ampicillin antibiotic to prevent microbial contamination (Bridgewater et al. 2017; Sholtes & Linden 2019). The bacterial host was E. coli Famp (ATCC #700891). Enumeration of was conducted using a combination of spread plate and spot plate methods and each sample was plated in triplicate. For each dilution in spread plating, 1 mL of sample was applied to soft-agar plates. For each dilution in spot plating, 20 μL of sample was serially diluted into 180 μL of phosphate-buffered saline (PBS) and five spots, 10 μL each, were added to the soft-agar plates with a multichannel pipette. After the sample was absorbed into the agar, plates were inverted and incubated for 20 h at 37 °C. Plaques were counted and concentrations were expressed in PFU/mL. For each experiment, the PBS dilution water, dechlorinated tap water, and pre-wash purified water were plated to test for contamination. Microbial reduction at each dose was expressed as log10 (N0/Nx). N0 represents the initial concentration in PFU/mL in the control sample without UV exposure. Nx represents the concentration in PFU/mL after exposure to the various experimental flow rates.

Bench-scale set-up

Bench-scale collimated beam tests were performed with a PearlBeam UVLED System (AquiSense Technologies, Erlanger, Kentucky) and commercial low pressure (LP) lamps (15 W Sankyo Denki, Japan) to create reduction equivalent fluence (REF) equations for the flow-through UVLED devices. The UVLED collimated beam experiments followed Sholtes & Linden (2019) whereas the LP experiments followed Bolton & Linden (2003). The peak and weighted average wavelength of the UVLED over a 200–300 nm spectrum were 281.5 and 283.5 nm, respectively, measured using a Maya 2000 Pro spectrometer (Ocean Insight, Dunedin, FL). Incident irradiance (mW/cm2) was measured with a radiometer (IL2400, SED240, International Light, Newburyport, Massachusetts) at the calibration factors corresponding to the weighted average wavelength (283.5 nm) for the polychromatic UVLED lamps and 254 nm for the monochromatic LP lamps. The calibrated plane of the detector was placed directly under the UVLED at the same height as the water surface of the irradiated samples. Stirred suspensions of 10 mL (0.55 cm sample depth) were irradiated in 5.4 cm diameter dishes at a 15 cm distance from the UVLED and 41 cm from the LP lamps. The UV irradiance was measured before and after the exposures with a calibrated radiometer to ensure there was no decay in lamp output over the exposure periods and less than a 5% difference between final and initial irradiance as recommended in the US EPA UV Disinfection Guidance Manual (US EPA 2006). The petri factor was >0.99 for all UVLED exposures. UV fluence is a product of average irradiance and time; therefore, pre-determined fluences were achieved by manipulating exposure time once average irradiance was determined. All experiments were performed in duplicate, and each duplicate sample had a replicate measurement. Samples were exposed in random order over the target fluence levels.

Flow-Through experiments

The PearlAqua MicroTM 9C UVLED flow-through units (AquiSense Technologies, Erlanger, Kentucky) were set up in accordance with manufacturer instructions. Greater than five reactor and effluent tube volumes of laboratory-grade deionized (DI) water were pumped through the entire system as a rinse. A sample of this DI rinse water was taken to test for contamination from previous testing in the system line prior to each experiment.

The test water was contained in a glass bottle, connected to integrated tubing, and a peristaltic pump for additional flow control. An untreated sample was taken from the glass bottle after sufficient mixing to quantify the starting concentration. To start experimentation, the peristaltic water pump (MasterFlex L/S 7518-62) was turned on. Once the water was flowing through the UVLED system, a sample was taken before the UVLED device was turned on to test for microbial decay due to non-UV factors. Upon changing the system flow rate, five times the reactor and effluent tube volume were allowed to flush the system before further experimentation. Samples from influent and effluent were taken in duplicate.

Once the relationship between fluence and LRV was established in the collimated beam experiments, the reduction equivalent fluences (REF254nm and REF280nm) for various flow rates were back calculated based on the LRVs in the flow-through units. Similar to Nguyen et al. (2019), a REF at 280 nm (as opposed to 254 nm) was chosen to investigate the wavelength-specific UV-induced stress experienced by microorganisms in the 280 nm UVLED flow-through device. However, a benchmark to 254 nm was included as it is helpful to compare to LP UV applications.

Wastewater effluent sample collection

Secondary wastewater effluent was collected at the City of Boulder Water Resource Recovery Facility after biological nutrient removal and prior to UV disinfection. Water was analyzed according to Standard Methods for the Examination of Water and Wastewater, Section 1060B (Bridgewater et al. 2017). The average water quality parameters measured were pH (7.48), temperature (21.3 °C), dissolved organic carbon (DOC) (6.83 mg-C/L), and UVT254nm (72.7%). The secondary effluent was used unfiltered.

Bioreactor set-up

Native wastewater effluent biofilms were grown on high-density polycarbonate coupons in a Centers for Disease Control and Prevention (CDC) biofilm reactor (model CBR 90-1, Biosurface Technologies Corp., Bozeman, MT). High-density polycarbonate coupons were chosen to mimic the adhesion surface found in drip irrigation lines. Two bioreactors, one non-UV control reactor and a UV-treated reactor, were operated for 5 days under semi-batch conditions with effluent replenished after day 1 and day 3. Internal stirring speed of the biofilm reactor, 60 rpm, was chosen to mimic the conditions of a concurrent pilot study at Technion-Israel Institute of Technology, considered the retention time of a standard drip irrigation line and shear stress experienced through a drip emitter. The bioreactors were kept in dark conditions to prevent any influence from bacterial photorepair following UV, similar to the conditions of a water transmission pipe or drip irrigation line. Flow rate was based off the REF280nm equations for the UVLED device and used to achieve the target fluence level, considering the operating limits of the inlet tubing (MasterFlex L/S 16). The experimental design is illustrated in Figure 1.
Figure 1

Flow-through schematic for the 5-day biofilm studies. Reuse water (1) is pumped by a peristaltic pump (2) through the UVLED flow-through device (3) into the CDC bioreactor (4) which drains into the waste container (5).

Figure 1

Flow-through schematic for the 5-day biofilm studies. Reuse water (1) is pumped by a peristaltic pump (2) through the UVLED flow-through device (3) into the CDC bioreactor (4) which drains into the waste container (5).

Close modal

Enumeration

Total coliforms by plate count

Total coliforms were enumerated through membrane filtration on m-ENDO agar LES (NutriSelect™ Plus). Each fouled coupon was rinsed in 30 mL PBS for 20 s to remove planktonic cells then immersed in a 10 mL PBS solution. A vortex-sonication series was used to disaggregate biofilm from the coupons. The UltraSonic cleaner (Branson 8210, 50/60 Hz) was degassed for 5 min prior to testing. Samples were then sonicated for 30 s, vortexed for 30 s, with each step being repeated three times. Serial dilutions of 10 mL were passed through 0.45 μm membrane filters and the filters were placed on the surface of m-ENDO agar plates, filter side up. Plates were incubated at 35 ± 0.5 °C for 24 ± 2.0 h. Coliforms were identified by red colonies with a golden-green metallic sheen.

ATP bioluminescence

ATP bioluminescence is an indirect quantification method using ATP as a proxy marker which infers biofilm viability and biomass. ATP is a nucleoside triphosphate which acts as the primary energy source in all organisms, making it a strong indication of biofilm viability and biomass (Wilson et al. 2017). Coupons were rinsed in 30 mL PBS to remove extracellular ATP then immersed in 2 mL PBS. Coupons were then subjected to 30 s vortexing followed by 5 min of sonication. Samples were then left to incubate at 37 °C for 4 h to diminish any ATP increase due to stress induced by UV irradiation (Rauch et al. 2019). Intracellular ATP was quantified using LuminUltra's standard Quench-Gone Aqueous (QGA) Test (LuminUltra Technologies Ltd, New Brunswick, Canada).

CV assay

Gram staining is a common and optimized indirect biofilm quantification method (Wilson et al. 2017). CV is a basic trianiline dye that permeates cell membranes in both Gram-positive and -negative cells regardless of inactivation or cell lysis. The dye leaves cells violet in color, and after a decolorization step with an ethanol solution, the biofilm can be quantified via spectroscopy. The CV assay was conducted according to Charlton (2008). A calibration curve was established between CV concentration and optical density, and a linear relationship was found. After treatment, coupons were immersed in a 2 mL 0.3% CV solution for 90 min followed by a PBS rinse and a subsequent DI rinse. Coupons were allowed to air dry, then transferred to a 2 mL 95% ethanol solution. Further dye solubilization was achieved by a 30-s vortex followed by 5 min of sonication. In total, 125 μL of sample were added to a 96-well plate and absorbance was measured at 540 nm. Five replicate measurements were taken per sample. Background absorbance was determined with clean coupons taken through the same stain and rinse steps.

The amount of biofilm remaining on the coupons was expressed as percent reduction (PR) where Cav is the average absorbance of a coupon in the non-UV control reactor, Bav represents the background absorbance as represented by the average absorbance of a sterile coupon, and Tcoupon is the absorbance of a coupon within the UV-treated reactor at 540 nm.
formula

Data analysis/statistics

Analysis of variance (ANOVA) full factorial experiments were performed to examine the statistical significance of experimental effects as included in the supplemental information. Model adequacy was verified by checking the three assumptions required for an ANOVA analysis: (1) equal variances, (2) residual normality, and (3) independent data.

Collimated beam results

The F-specific RNA bacteriophage was chosen as the biological indicator for flow-through UVLED device challenge testing because it shows first-order rate kinetics at high UV fluences (>40 mJ/cm2) and can capture the high inactivation rates produced under slow flow rates. It is a commonly used for flow-through UV device challenge testing and is the NSF/ANSI 55 challenge organism for UV device challenge testing (Jenny et al. 2014; Oguma et al. 2016a, 2016b; NSF/ANSI 55 2019). An LP 254 nm exposure was performed to ensure quality control of the microbial stocks and comparable results to other QB LP literature values. In addition to the LP 254 nm exposures, 280 nm UVLEDs were chosen to illuminate the response of microorganisms inactivated in the UVLED flow-through device also manufactured with identical 280 nm LED chips.

The dose response of in three varying UVT254nm waters is presented in Figure 2. As expected, the response is the same between UVTs for both UV devices as the fluence calculations correct for the UV transmittance of the water sample (Bolton & Linden 2003). LP 254 nm produces higher first-order rate kinetics in with an inactivation rate constant of 0.0728 cm2/mJ, similar to the results from previous studies (Aoyagi et al. 2011; Oguma et al. 2013; Jenny et al. 2014; Rattanakul et al. 2014; Beck et al. 2015). The 280 nm UVLED produced a lower inactivation rate constant of 0.0410 cm2/kJ, consistent with previous studies (Aoyagi et al. 2011; Oguma et al. 2016a, 2016b).
Figure 2

bacteriophage log reduction values (LRV) (log10(N0/Nx)) over three UVT254nm waters in response to UV irradiation dose from both polychromatic UVLEDs emitting at a peak of 280 nm and a monochromatic 254 nm LP lamp.

Figure 2

bacteriophage log reduction values (LRV) (log10(N0/Nx)) over three UVT254nm waters in response to UV irradiation dose from both polychromatic UVLEDs emitting at a peak of 280 nm and a monochromatic 254 nm LP lamp.

Close modal

Flow-through results

The flow-through results (Figure 3) display a tailing behavior in flow rates over 1 L/min. Tailing has also been observed in other UVLED flow-through reactor studies indicating lower inactivation efficiencies at higher flow rates (Oguma et al. 2013). Declining disinfection performances with increasing flow rate is expected as a higher flow rate corresponds to a shorter residence time, and shorter LED exposure (and lower dose), within the reactor (Barstow et al. 2014). A previous study examining flow-through UVLEDs for domestic wastewater disinfection for agricultural reuse found that the REF280nm values were high enough (>50 mJ/cm2) to meet water reuse guidelines for agriculture of both processed food crops and non-food crops (US EPA 2012; Nguyen et al. 2019). The results from this study illustrate that similarly high REF280s at low-flow rates (<0.05 L/min) can be achieved; however, the REF will decrease at higher flow rates especially for low UVT waters.
Figure 3

reduction equivalent fluence values at 280 nm (REF280nm) left and 254 nm (REF254nm) right for the flow-through UVLED device tested with three challenge waters. The 93% UVT254nm water was unaltered while the 80 and 70% UVT254nm waters were adjusted with UV absorbing compounds.

Figure 3

reduction equivalent fluence values at 280 nm (REF280nm) left and 254 nm (REF254nm) right for the flow-through UVLED device tested with three challenge waters. The 93% UVT254nm water was unaltered while the 80 and 70% UVT254nm waters were adjusted with UV absorbing compounds.

Close modal
Higher reduction equivalent fluences were reached for the 93% UVT254nm water, with lower values in the 80 and 70% UVT254nm waters for both the REF280nm and REF254nm. The lower germicidal performance for 80 and 70% UVT254nm waters was anticipated as low UVT indicates a lower proportion of light is transmitted through the water sample. At a flow rate of 2 L/min, the REF280nm values were 34, 15, and 12 mJ/cm2 for the 93, 80, and 70% UVT254nm waters, respectively. At the same flow rate of 2 L/min, the REF254nm values were 19, 9, and 7 mJ/cm2 for the 93, 80, and 70% UVT254nm waters, respectively. The 22 mJ/cm2 REF280nm difference and the 12 mJ/cm2 REF254nm difference between the 93 and 70% UVT254nm waters suggest challenges for UVLED performance in wastewater reuse when higher fluence levels are required. Most effluents fall between 70 and 80% UVT254nm while most UVLED flow-through devices are developed for drinking-water applications (93% UVT254nm and higher). This highlights an interesting juxtaposition between the intended use of commercially available UVLEDs (for drinking water in this case) versus the available and growing applications for the devices, and a need to design devices specifically for wastewater applications. A non-linear regression analysis was performed to develop a relationship between the LRV, flow rate in L/min, and UVT254nm resulting in an adjusted regression coefficient of 0.99954. The regression analysis of fit is included in the supplemental information. A is the regression constant while alpha and beta are regression correction factors for UVT and flow rate, respectively.
formula

Using the model equation, at 1 L/min, there is a 0.961 LRV increase when UVT254nm increases from 60 to 93%. At smaller flow rates of 0.1 L/min, this difference is larger, with a 2.647 LRV increase when UVT254nm increases from 60 to 93%. The model results can be paired with the inactivation rate constants (slopes) found in Figure 2 to convert QB LRV to REF280nm and REF254nm. Therefore, the model can be used to contextualize the benefits of technologies that may impact water UV transmittance and the resulting benefit to REF. For example, at a flow rate of 0.5 L/min, a UVLED device paired with a pre-filter that increases UVT254nm from 65 to 80% would result in an additional REF254nm of roughly 10 mJ/cm2.

The primary objective of this study was to test the UVLED device's ability to inactivate planktonic microorganisms in waters with UVT ranging from 70 to 93%. A target REF280nm for water reuse at 40 mJ/cm2, corresponding to a REF254nm of 24 mJ/cm2, was chosen to investigate biofouling control in the CDC bioreactor. 40 mJ/cm2 REF280nm, or 24 mJ/cm2 REF254nm, were obtained using a 0.16 L/min flow rate in the UVT254nm 72.7 ± 2.5% water matrix. The following quantification methods describe the results from the flow-through investigations.

Direct quantification methods

Biofilm quantification can be measured through direct and indirect methods. Direct biofilm quantification methods rely on direct observations for quantification of parameters, for example, the number of cells or total biofilm volume (Wilson et al. 2017). Inferring biofilm quantity through viable cell numbers should be supported with other assays as many factors affect biofilms carrying capacity of cells. Carrying capacity is defined as the maximum potential population size of a given landscape is capable of supporting and is a common attribute used to describe population dynamics (Stilling 1996). Biofilm cell carrying capacity is dependent on; but not limited to, carbon and oxygen depletion, shear stress, temperature, and iron availability (Madigan et al. 1949). These factors also regulate the rate at which biofilms shed planktonic cells.

Total coliforms enumeration
The UV-treated and control bioreactors had 166 (102.22) and 3,240 (103.51) TC/coupon, respectively, on day 1 (Figure 4). Each coupon has a diameter of 12.7 mm. The total surface area exposed for biofouling included the top and bottom of the coupon as the side was embedded in the coupon holder, making the total sampled surface area roughly 2.5 cm2. Normalizing the total coliform growth per square centimeter, the day 1 UV-treated and control bioreactors had 66.1 (101.82) and 1,290 (103.11) TC/cm2, respectively. The 1.29 log per coupon or square centimeter difference indicates that the UV-treated bioreactor had less biofouling initially. Referencing the dose response of in Figure 2, a 280 nm fluence of 40 mJ/cm2 produced a similar log reduction value (1.64 LRV) as the coupons on day 1 (1.29 LRV). This indicates that initially, biofilm formation is inhibited at similar levels to the inactivation of planktonic cells achieved by the UVLED device.
Figure 4

Total coliform concentrations expressed as log TC per coupon (left axis) and log TC per mL of bioreactor effluent (right axis) over a 5-day period. Vertical error bars represent standard deviation.

Figure 4

Total coliform concentrations expressed as log TC per coupon (left axis) and log TC per mL of bioreactor effluent (right axis) over a 5-day period. Vertical error bars represent standard deviation.

Close modal

After day 2, the control and UV-treated coupons approach similar concentrations of 104 TC/coupon and 103.6 TC/cm2, respectively. Similar observations were reported in previous studies (Wenjun & Wenjun 2009; Conrad 2018; Sperle et al. 2020), which showed UV exposures on planktonic cells that only delayed their biofilm formation in a continuous system. One limitation of this investigation is that only sub-lethal UV fluence was used to treat the planktonic cells before biofilm formation. Less viable cells were available for inoculation on the surface in the early stage of biofilm development. But, once the biofilm structure is formed, it may start capturing the incoming planktonic cells and utilizing the nutrients to grow and eventually reach a similar steady-state as the biofilms without UV pretreatment. Long-term biofilm inhibition by UV pretreatment is possible if the UV fluence is high enough to achieve intensive suppression or even elimination of viable planktonic cells. Another method employing UVLEDs for biofilm control is through direct UV exposure orienting the UVLEDs directly above the forming biofilm structure. Previous studies suggest that direct UV exposure can effectively inactivate pre-formed biofilms on surfaces (Gora et al. 2019; Ma et al. 2022) or inhibit biofilm formation (Torkzadeh et al. 2021), achieving a maximum reduction of more than 2 log (99%). In addition, recent studies (Lanzarini-Lopes et al. 2020; Zhao et al. 2023) demonstrated that UVLEDs coupled with side-emitting optical fibers could be an alternative configuration for effective biofilm control via direct UV exposure, providing greater surface area coverage, especially in systems with complex geometries like drip irrigation systems.

An important distinction not investigated in this study is whether the viable planktonic cells are growing within the bioreactor, then adhering to the coupons and forming biofilms, or if the planktonic cells immediately form biofilms then release more planktonic cells as biofilm maturity is reached. To achieve a REF280nm of 40 mJ/cm2, the UVLED device was operated at a specific flow rate corresponding to a 22-min hydraulic residence time within the bioreactors. Under favorable conditions, bacterial doubling time can be less than 20 min (Chalor et al. 2012). This suggests that planktonic cell concentrations could be increasing within the bioreactor independent of biofilm adhesion and maturity.

To understand how the UVLED device may change microbial conditions within the bioreactor with an influent flow rate of 0.16 L/min, the ambient bioreactor effluent coliform concentrations were measured on day 5. A full-factorial ANOVA analysis examining the effects of time in days and reactor (control or UV treated) on effluent coliform revealed that the day 5 effluent coliform values were not statistically different between the non-UV and UV-treated bioreactor (p = 0.65). This supports the previous conclusion that any viable cells surviving UV disinfection could potentially colonize the bioreactor. Interestingly, the cell carrying capacity of the coupons (∼104 CFU/coupon or ∼103.6 CFU/cm2) is similar to the planktonic cell concentration of the wastewater effluent within the bioreactors (∼103.25 CFU/mL). The data may suggest the planktonic cell concentration environment is influencing the carrying capacity of the biofilms, or the opposite, the biofilm may be influencing the planktonic cell concentration in the bioreactor. If biofilm maturity is reached rapidly, planktonic cell dispersion from the mature biofilm may influence the planktonic cell concentration in the bioreactor effluent.

Indirect quantification methods

In addition to direct quantification, biofilms can be quantified indirectly through the use of a proxy marker, such as CV or metabolites. There is a general assumption that proxy markers are directly related to cell concentration within the biofilm (Azeredo et al. 2017; Wilson et al. 2017). However, these indirect biofilm quantification techniques are often dependent on metabolic function and biomolecule production that may not affect cell concentration. This study followed the recommendation provided by Wilson et al. (2017) to pair indirect quantification methods with a direct quantification method.

CV staining

CV staining was first described by Christensen et al. (1985) and is now one of the most optimized microbiological methods for the identification and visualization of bacteria (Wilson et al. 2017). A CV assay was performed on bioreactor coupons on days 1, 3, and 5 of the experiment. Although Gram-positive and Gram-negative cells can be differentiated via microscopy after CV staining, this was not examined. It should be noted that CV stains cell membranes regardless of cell viability and is therefore a quantification of biomass and not viable cell concentration (Wilson et al. 2017). To accurately capture the amount of EPS, a prerinse step was performed prior to CV staining to attempt to ‘wash’ planktonic cells from the coupon biofilm.

The CV, represented by optical density at 540 nm, accumulating on the coupons over the 5-day experiment appears to increase slightly over time with no statistical difference between the control and UV-treated samples (Figure 5). A full-factorial ANOVA analysis examining the effects of time in days and reactor (control or UV treated) on coupon optical density confirmed that biofilms did continue to accumulate with time (p = 0.033) over the 5 days and there was no significant difference between the optical density of coupons from the control versus UV-treated bioreactors across the period measurements that were taken. The results contrast the total coliform findings presented in Figure 4, which found no biofouling increase by the measure of bacterial colony-forming units after day 2. A possible explanation is the ability of CV to stain dead or lysed cell materials contained in the EPS or staining the EPS itself, which would not be removed during the wash step (Flemming & Wingender 2010). In this scenario, the dead/lysed cell material would contribute more CV absorbance to the optical density readings suggesting higher biofouling than a viable cell assay, like total coliforms. This hypothesis is supported by McSwain et al. (2005) who observed significant effects by cell lysis and contamination by dead biomass in EPS leading to different and opposing conclusions in biofilm quantification assays. Future studies may benefit from an EPS-specific quantification assay, such as a ruthenium red dye specific to carbohydrates or scanning electron microscopy (SEM) which detects the presence of EPS (Figueroa & Silverstein 1989; Azeredo et al. 2017). This would allow independent observations of the contribution of viable cells and EPS to the total biofilm biomass, provided by the CV assay. An additional factor contributing to the difference between coupon total coliform cell counts and CV biofilm accumulation following day 2 may be the wastewater effluent replenishment step occurring on day 3. Effluent replenishment may impact the nutritional composition and bacterial diversity of the source water though these water quality parameters were not quantified in this study. It is important to note that both dead bacterial cells and EPS play a significant role in biofouling by increasing hydraulic resistance (Herzberg et al. 2009). An assay targeting the whole biofilm structure, such as CV staining, may be more representative of biofouling than a direct quantification of viable biofilm cells. Regardless, both total coliform and CV assays indicate that an REF280nm of 40 mJ/cm2 retarded the rate, but did not fully prevent biofouling in CDC bioreactors fed by unfiltered wastewater effluent.
Figure 5

Optical density (OD540nm) presented graphically and numerically at 540 nm for the crystal violet solutions derived from the decolorization of crystal violet-stained control and UV-treated coupons. Standard deviations are represented by the vertical error bars and parenthesis.

Figure 5

Optical density (OD540nm) presented graphically and numerically at 540 nm for the crystal violet solutions derived from the decolorization of crystal violet-stained control and UV-treated coupons. Standard deviations are represented by the vertical error bars and parenthesis.

Close modal
ATP bioluminescence

Due to resource limitations, the ATP analysis was only performed on day 5 to provide an understanding of the overall bioactivity after 5 days of biofilm development. ATP was sampled from the bioreactor coupons as well as bioreactor effluent. UV prevents cells from replicating but does not immediately inhibit ATP production in the cell after inactivation. To observe differences between UV and non-UV-treated samples, the coupon and bioreactor effluent samples were incubated for 4 h post-sampling at 37 °C to allow for replication of active cells (Rauch et al. 2019). Allowing cells to undergo replication cycles post-UV provides resolution for the ATP assay between treated and non-UV-treated samples that would not be detectable immediately following UV disinfection.

On day 5, the UV-treated bioreactor appears to have slightly higher coupon ATP (54.8 pg ATP/mm2) than the control coupons (42.9 pg ATP/mm2) (Figure 6). A similar trend is observed for the bioreactor effluents; the UV-treated effluent has higher ATP (1,678 pg ATP/mL) than the control effluent (1,014 pg ATP/mL). Previous studies have reported higher ATP values in E. coli following UV (Villaverde & Barbe 1986). However, these results are contrary to the total coliform (Figure 4) and CV (Figure 5) results, which did not see statistically different results between UV and non-UV-treated bioreactors by day 5.
Figure 6

Day 5 adenosine triphosphate (ATP) values for the coupons [pg ATP/mm2] and effluent [pg ATP/mL] of the control and UV-treated bioreactors.

Figure 6

Day 5 adenosine triphosphate (ATP) values for the coupons [pg ATP/mm2] and effluent [pg ATP/mL] of the control and UV-treated bioreactors.

Close modal

Rauch et al. (2019) developed an ATP biomass recovery method for E. coli and wastewater communities following UV exposure. Adoption of the method has been successfully reproduced in studies examining the effects of UV on pure cell cultures ATP (Gora et al. 2019). Miller et al. (2020) found that a 2-h incubation after UV exposure was enough to diminish background noise for ATP measurements at a pilot-scale direct potable reuse facility, and higher sensitivity was achieved from ATP than flow cytometry following UV advanced oxidation processes (UV-AOP). Hammes et al. (2010) was able to calculate an average ATP-per-cell value (1.75 × 10−10 nmol/cell) from microbial samples taken from a variety of aquatic environments including drinking water, groundwater, river water, and wastewater effluent. Although the study standardized ATP production per cell, the authors also recognized the high ATP heterogeneity of microbial communities of different samples and variations in ATP extraction and analysis procedures when different natural water microbial communities are analyzed (Wilson et al. 1981; Schneider & Gourse 2004; Eydal & Pedersen 2007; Hammes et al. 2010). Rauch et al. (2019) also recognized that variations in microbial communities (like slow-growing organisms) may complicate the method and indicated the importance of the incoming water matrix on the outcome of the biomass recovery-ATP test. The ATP results presented in this study suggest that the 4-h 37 °C incubation was not sufficient at suppressing heightened ATP production in microbial communities post-UV. Future studies should build upon the recommendations made by Rauch et al. (2019) and examine the relationship between UV irradiation and natural microbial community ATP, as opposed to pure cell cultures, over time frames beyond 4 h.

A different explanation for the ATP results on day 5 is the high level of variability characteristic of biofilms and their quantification. Many coliform-specific biofilm studies have noted this stochastic behavior including a study examining coliform retention and biofouling within irrigation pipes (Shelton et al. 2013) and a study investigating drinking-water systems plagued by coliform regrowth (Camper et al. 1996). Through an 18-month survey of 31 water systems in North America, LeChevallier et al. (1996) concluded that the occurrence of coliforms is dependent upon complex interactions between chemical, physical, operational, and engineering parameters. This reinforces the recommendation made by Wilson et al. (2017), to draw upon multiple biofilm assessment methods that include both direct and indirect techniques to improve understanding and knowledge surrounding biofilms. Table 1 provides an overview of the biofilm quantification methods used in this study including the quantification target, quantification time, and high-level pros and cons of each technique.

Table 1

Comparison of biofilm quantification methods used in this study

Total coliformCrystal violetATP
Quantification target Viable coliform bacteria cells All bacterial cells (live and dead)
EPS 
Biological activity 
Quantification time 24 h 3 h 1 h 
Pros Culture-based method that only targets viable cells, which is sensitive to UV treatment. Representative to biofouling Fast 
Cons Slow
May not be representative of biofouling 
Indirect assay based on colorimetric quantification with low accuracy Delayed response to UV treatment 
Total coliformCrystal violetATP
Quantification target Viable coliform bacteria cells All bacterial cells (live and dead)
EPS 
Biological activity 
Quantification time 24 h 3 h 1 h 
Pros Culture-based method that only targets viable cells, which is sensitive to UV treatment. Representative to biofouling Fast 
Cons Slow
May not be representative of biofouling 
Indirect assay based on colorimetric quantification with low accuracy Delayed response to UV treatment 

Challenge testing was conducted for a UVLED flow-through device using three test waters of varying UVT254nm (93, 80, 70%). The study concluded that the UVLED device performed similarly to other UV flow-through devices discussed in the literature, with tailing of inactivation performance at higher flow rates and decreased performance for low UVT waters (Oguma et al. 2013; Barstow et al. 2014). When operating with 70% UVT254nm effluent at low (<0.036 L/min) flow rates, the UVLED device produced fluences >40 mJ/cm2 corresponding to the 2012 US EPA Water Reuse Guidelines for the use of reclaimed water for irrigation of food crops (nondetectable fecal coliforms), and processed/non-food crops (<200 fecal coliforms/100 mL) (US EPA 2012). For disinfected tertiary recycled water intended for agriculture, California Title 22 reuse regulations allow disinfection methods that, when combined with filtration, produce 5-log inactivation of PFU of F-specific bacteriophage MS-2, poliovirus, or a similar virus (Title 22 Code of Regulations). According to NWRI analysis of UV fluence at 254 nm and MS2 inactivation, a 5-log removal of MS2 corresponds to a fluence of 121 mJ/cm2 (NWRI 2012). Using the regression analysis presented in Figure 3, the UVLED system examined for this study would need to operate at 0.003 L/min to produce a REF254nm of 121 mJ/cm2 for a 70% UVT254nm water matrix (assuming a worst-case scenario where the pre-filter provided no virus removal). For reference, drip irrigation row flow rates are generally in the range of 0.8 L/min per 100 feet of row length (Burt 2008). This level of fluence necessitates an evaluation or potentially UV LED device such as a PearlAqua Deca (AquiSense Technologies, Erlanger, Kentucky) with higher UV output or potentially pairing UV treatment with pretreatments, like filtration, that improve the UV transmittance of the distributed reuse water.

The UVLED flow-through device was then assessed for its ability to prevent biofouling in a CDC bioreactor fed by unfiltered wastewater effluent operating at a 40 mJ/cm2 REF280nm. This study demonstrated UVLED disinfection retarded biofouling in a bioreactor fed by unfiltered wastewater effluent. However, by day 5, both non-UV and UV-treated bioreactors approached 104 CFU/coupon as well as 103.75 coliforms per mL of bioreactor effluent. Examining higher reduction equivalent fluences or more extensive pretreatments, such as filtration, would provide a meaningful contribution to this body of literature surrounding biofouling control by UVLEDs. Understanding the interaction between planktonic cells and sessile cells (i.e., whether the planktonic cell concentration is influencing biofouling; or if a rapidly formed mature biofilm is dispersing planktonic cells) would also help advance the conclusions drawn in this study. Overall, UV pretreatment at REF280nm of 40 mJ/cm2 may slow but will not ultimately eliminate biofouling in drip irrigation lines. The impact on drip emitters also may differ due to the hydrodynamics of the water flow through the emitters, which was not tested herein but is addressed in a companion study as part of this larger body of funded research.

Analysis of biological activity after 5 days of biofilm development using ATP did not align with the quantification of biofilm biomass using enumeration assays. This can be attributed to either (1) the high variability known to exist within biofilm behavior and resulting quantification or (2) known challenges in applying ATP assays to UV disinfection and possible insufficient incubation conditions following UV. Natural microbial communities in wastewater will vary geographically and may exhibit varying physiological UV-stress responses depending on treatment conditions. Future studies examining the relationship between ATP production in mixed cell cultures or microbial communities post-UV should be conducted to advance the understanding of microbial response to UV-induced stress.

All relevant data are included in the paper or its Supplementary Information.

The authors declare there is no conflict.

Adin
A.
&
Sacks
M.
1991
Dripper-clogging factors in wastewater irrigation
.
Journal of Irrigation and Drainage Engineering
117
(
6
),
813
826
.
https://doi.org/10.1061/(ASCE)0733-9437(1991)117:6(813)
.
Aoyagi
Y.
,
Takeuchi
M.
,
Yoshida
K.
,
Kurouchi
M.
,
Yasui
N.
,
Kamiko
N.
,
Araki
T.
&
Nanishi
Y.
2011
Inactivation of bacterial viruses in water using deep ultraviolet semiconductor light-emitting diode
.
Journal of Environmental Engineering
137
(
12
),
1215
1218
.
https://doi.org/10.1061/(ASCE)EE.1943-7870.0000442
.
Azeredo
J.
,
Azevedo
N.
,
Briandet
R.
,
Cerca
N.
,
Coenye
T.
,
Costa
A.
&
Desvaux
M.
2017
Critical review on biofilm methods
.
Critical Reviews in Microbiology
43
(
3
),
313
351
.
https://doi.org/10.1080/1040841X.2016.1208146
.
Barstow
C. K.
,
Dotson
A. D.
&
Linden
K. G.
2014
Assessing point-of-use ultraviolet disinfection for safe water in urban developing communities
.
Journal of Water and Health
12
(
4
),
663
669
.
https://doi.org/10.2166/wh.2014.223
.
Beck
S. E.
,
Wright
H. B.
,
Hargy
T. M.
,
Larason
T. C.
&
Linden
K. G.
2015
Action spectra for validation of pathogen disinfection in medium-pressure ultraviolet (UV) systems
.
Water Research
70
,
27
37
.
https://doi.org/10.1016/j.watres.2014.11.028
.
Bishop
P. L.
2007
The role of biofilms in water reclamation and reuse
.
Water Science and Technology: A Journal of the International Association on Water Pollution Research
55
(
1–2
),
19
26
.
https://doi.org/10.2166/wst.2007.005
.
Bolton
J. R.
&
Linden
K. G.
2003
Standardization of methods for fluence (UV dose) determination in bench-scale UV experiments
.
Journal of Environmental Engineering
129
(
3
),
209
215
.
https://doi.org/10.1061/(ASCE)0733-9372(2003)129:3(209)
.
Bridgewater
L. L.
,
Baird
R. B.
,
Eaton
A. D.
&
Rice
E. W.
&
American Public Health Association, American Water Works Association, and Water Environment Federation
2017
Standard Methods for the Examination of Water and Wastewater
, 23rd edn.
American Public Health Association
,
Washington, DC
.
Burt
C.
2008
Avoiding Common Problems with Drip Tape
.
ITRC Report No. R 08-003. California Polytechnic State University Irrigation Training and Research Center (ITRC)
.
Camper
A. K.
,
Jones
W. L.
&
Hayes
J. T.
1996
Effect of growth conditions and substratum composition on the persistence of coliforms in mixed-population biofilms
.
Applied and Environmental Microbiology
62
(
11
),
4014
4018
.
Chalor
J.
,
Kamolchanok
S.
,
Supatpong
M.
&
Ratana
J.
2012
Treatment of formaldehyde-containing wastewater using membrane bioreactor
.
Journal of Environmental Engineering
138
(
3
),
265
271
.
https://doi.org/10.1061/(ASCE)EE.1943-7870.0000430
.
Chen
J.
,
Loeb
S.
&
Kim
J.
2017
LED revolution: Fundamentals and prospects for UV disinfection applications
.
Environmental Science: Water Research & Technology
3
(
2
),
188
202
.
https://doi.org/10.1039/C6EW00241B
.
Christensen
G. D.
,
Simpson
W. A.
,
Younger
J. J.
,
Baddour
L. M.
,
Barrett
F. F.
,
Melton
D. M.
&
Beachey
E. H.
1985
Adherence of coagulase-negative staphylococci to plastic tissue culture plates: A quantitative model for the adherence of staphylococci to medical devices
.
Journal of Clinical Microbiology
22
(
6
),
996
1006
.
Conrad, S.
2018
Evaluation of continuous, low-intensity ultraviolet irradiation for biofilm prevention. Graduate Theses & Non-Theses. 194. https://digitalcommons.mtech.edu/grad_rsch/194
.
Doederer
K.
,
Gernjak
W.
,
Weinberg
H. S.
&
Farré
M. J.
2014
Factors affecting the formation of disinfection by-products during chlorination and chloramination of secondary effluent for the production of high quality recycled water
.
Water Research
48
,
218
228
.
https://doi.org/10.1016/j.watres.2013.09.034
.
Eydal
H. S.
&
Pedersen
K.
2007
Use of an ATP assay to determine viable microbial biomass in fennoscandian shield groundwater from depths of 3–1000m
.
Journal of Microbiological Methods
70
(
2
),
363
373
.
https://doi.org/10.1016/j.mimet.2007.05.012
.
Figueroa
L. A.
&
Silverstein
J.
1989
Ruthenium red adsorption method for measurement of extracellular polysaccharides in sludge flocs
.
Biotechnology and Bioengineering
33
(
8
),
941
947
.
https://doi.org/10.1002/bit.260330802
.
Flemming
H.
2002
Biofouling in water systems – cases, causes and countermeasures
.
Applied Microbiology and Biotechnology
59
(
6
),
629
640
.
https://doi.org/10.1007/s00253-002-1066-9
.
Flemming
H.
&
Wingender
J.
2010
The biofilm matrix
.
Nature Reviews Microbiology
8
(
9
),
623
633
.
https://doi.org/10.1038/nrmicro2415
.
Flemming
H.
,
Neu
T. R.
&
Wozniak
D. J.
2007
The EPS matrix: The ‘House of biofilm cells’
.
Journal of Bacteriology
189
(
22
),
7945
7947
.
https://doi.org/10.1128/JB.00858-07
.
Gora
S. L.
,
Rauch
K. D.
,
Ontiveros
C. C.
,
Stoddart
A. K.
&
Gagnon
G. A.
2019
Inactivation of biofilm-bound Pseudomonas aeruginosa bacteria using UVC light emitting diodes (UVC LEDs)
.
Water Research
151
,
193
202
.
https://doi.org/10.1016/j.watres.2018.12.021
.
Hammes
F.
,
Goldschmidt
F.
,
Vital
M.
,
Wang
Y.
&
Egli
T.
2010
Measurement and interpretation of microbial adenosine tri-phosphate (ATP) in aquatic environments
.
Water Research
44
(
13
),
3915
3923
.
https://doi.org/10.1016/j.watres.2010.04.015
.
Herzberg
M.
,
Kang
S.
&
Elimelech
M.
2009
Role of extracellular polymeric substances (EPS) in biofouling of reverse osmosis membranes
.
Environmental Science & Technology
43
(
12
),
4393
4398
.
Jenny
R. M.
,
Simmons
O. D.
,
Shatalov
M.
&
Ducoste
J. J.
2014
Modeling a continuous flow ultraviolet light emitting diode reactor using computational fluid dynamics
.
Chemical Engineering Science
116
,
524
535
.
https://doi.org/10.1016/j.ces.2014.05.020
.
Lanzarini-Lopes
M.
,
Zhao
Z.
,
Perreault
F.
,
Garcia-Segura
S.
&
Westerhoff
P.
2020
Germicidal glowsticks: Side-emitting optical fibers inhibit Pseudomonas aeruginosa and Escherichia coli on surfaces
.
Water Research
184
,
116191
.
https://doi.org/10.1016/j.watres.2020.116191
.
LeChevallier
M. W.
,
Welch
N. J.
&
Smith
D. B.
1996
Full-scale studies of factors related to coliform regrowth in drinking water
.
Applied and Environmental Microbiology
62
(
7
),
2201
2211
.
Lehtola
M. J.
,
Laxander
M.
,
Miettinen
I. T.
,
Hirvonen
A.
,
Vartiainen
T.
&
Martikainen
P. J.
2006
The effects of changing water flow velocity on the formation of biofilms and water quality in pilot distribution system consisting of copper or polyethylene pipes
.
Water Research
40
(
11
),
2151
2160
.
https://doi.org/10.1016/j.watres.2006.04.010
.
Li
Y.
,
Song
P.
,
Pei
Y.
&
Feng
J.
2015
Effects of lateral flushing on emitter clogging and biofilm components in drip irrigation systems with reclaimed water
.
Irrigation Science; Heidelberg
33
(
3
),
235
245
.
http://dx.doi.org.colorado.idm.oclc.org/10.1007/s00271-015-0462-8
.
Li
Q.
,
Song
P.
,
Zhou
B.
,
Xiao
Y.
,
Muhammad
T.
,
Liu
Z.
,
Zhou
H.
&
Li
Y.
2019
Mechanism of intermittent fluctuated water pressure on emitter clogging substances formation in drip irrigation system utilizing high sediment water
.
Agricultural Water Management
215
,
16
24
.
https://doi.org/10.1016/j.agwat.2019.01.010
.
Linden
K. G.
,
Hull
N.
&
Speight
V.
2019
Thinking outside the treatment plant: UV for water distribution system disinfection
.
Accounts of Chemical Research
52
(
5
),
1226
1233
.
https://doi.org/10.1021/acs.accounts.9b00060
.
Ma
B.
,
Seyedi
S.
,
Wells
E.
,
McCarthy
D.
,
Crosbie
N.
&
Linden
K. G.
2022
Inactivation of biofilm-bound bacterial cells using irradiation across UVC wavelengths
.
Water Research
217
,
118379
.
https://doi.org/10.1016/j.watres.2022.118379
.
Madigan
M.
,
Martinko
J.
,
Bender
K.
,
Buckley
K.
&
Stahl
D.
1949
Brock Biology of Microorganisms
, 14th edn.
Pearson
,
London
.
Malayeri
A.
,
Mohseni
M.
,
BCairns
B.
&
Bolton
J.
2016
Fluence (UV dose) required to achieve incremental log inactivation of bacteria, protozoa, viruses and algae
.
IUVA News
18
,
4
6
.
Marin
P.
,
Tal
S.
,
Yeres
J.
&
Ringskog
K.
2017
Water Management in Israel : Key Innovations and Lessons Learned for Water Scarce Countries
.
Water Global Practice
.
McSwain
B. S.
,
Irvine
R. L.
,
Hausner
M.
&
Wilderer
P. A.
2005
Composition and distribution of extracellular polymeric substances in aerobic flocs and granular sludge
.
Applied and Environmental Microbiology
71
(
2
),
1051
1057
.
https://doi.org/10.1128/AEM.71.2.1051-1057.2005
.
Miller
S.
,
Rodriguez
R.
&
Nelson
K.
2020
Removal and growth of microorganisms across treatment and simulated distribution at a pilot-scale direct potable reuse facility
.
Environmental Science: Water Research & Technology
6
(
5
),
1370
1387
.
https://doi.org/10.1039/C9EW01087D
.
NSF/ANSI 55
2019
NSF International Standard/ American National Standard
.
NSF International
,
Ann Arbor
.
NWRI
2012
Ultraviolet Disinfection Guidelines for Drinking Water and Water Reuse
.
National Water Research Institute (NWRI)
,
Fountain Valley
.
Oguma
K.
,
Kita
R.
,
Sakai
H.
,
Murakami
M.
&
Takizawa
S.
2013
Application of UV light emitting diodes to batch and flow-through water disinfection systems
.
Desalination
328
,
24
30
.
https://doi.org/10.1016/j.desal.2013.08.014
.
Oguma
K.
,
Kita
R.
&
Takizawa
S.
2016a
Effects of arrangement of UV light-emitting diodes on the inactivation efficiency of microorganisms in water
.
Photochemistry and Photobiology
92
(
2
),
314
317
.
https://doi.org/10.1111/php.12571
.
Oguma
K.
,
Rattanakul
S.
&
Bolton
J. R.
2016b
Application of UV light–emitting diodes to adenovirus in water
.
Journal of Environmental Engineering
142
(
3
),
04015082
.
https://doi.org/10.1061/(ASCE)EE.1943-7870.0001061
.
Rattanakul
S.
,
Oguma
K.
,
Sakai
H.
&
Takizawa
S.
2014
Inactivation of viruses by combination processes of UV and chlorine
.
Journal of Water and Environment Technology
12
(
6
),
511
523
.
https://doi.org/10.2965/jwet.2014.511
.
Rauch
K. D.
,
Mackie
A. L.
,
Middleton
B.
,
Xie
X.
&
Gagnon
G. A.
2019
Biomass recovery method for adenosine triphosphate (ATP) quantification following UV disinfection
.
Ozone: Science & Engineering
41
(
2
),
146
155
.
https://doi.org/10.1080/01919512.2018.1518127
.
Ravina
I.
,
Paz
E.
,
Sofer
Z.
,
Marm
A.
,
Schischa
A.
,
Sagi
G.
,
Yechialy
Z.
&
Lev
Y.
1997
Control of clogging in drip irrigation with stored treated municipal sewage effluent
.
Agricultural Water Management
33
(
2
),
127
137
.
https://doi.org/10.1016/S0378-3774(96)01286-3
.
Rostad
C.
2002
Fate of disinfection by-products in the subsurface
. In:
U.S. Geological Survey Artificial Recharge Workshop
.
U.S. Geological Survey
,
Sacramento, CA
.
Schneider
D. A.
&
Gourse
R. L.
2004
Relationship between growth rate and ATP concentration in Escherichia coli
.
Journal of Biological Chemistry
279
(
9
),
8262
8268
.
https://doi.org/10.1074/jbc.M311996200
.
Shelton
D. R.
,
Kiefer
L. A.
,
Pachepsky
Y. A.
,
Blaustein
R. A.
&
Martinez
G.
2013
Coliform retention and release in biofilms formed on new and weathered irrigation pipes
.
Irrigation Science
31
(
5
),
971
981
.
https://doi.org/10.1007/s00271-012-0373-x
.
Sholtes
K.
&
Linden
K. G.
2019
Pulsed and continuous light UV LED: Microbial inactivation, electrical, and time efficiency
.
Water Research
165
,
114965
.
https://doi.org/10.1016/j.watres.2019.114965
.
Song
K.
,
Mohseni
M.
&
Taghipour
F.
2016
Application of ultraviolet light-emitting diodes (UV-LEDs) for water disinfection: A review
.
Water Research
94
,
341
349
.
https://doi.org/10.1016/j.watres.2016.03.003
.
Song
P.
,
Li
Y.
,
Zhou
B.
,
Zhou
C.
,
Zhang
Z.
&
Li
J.
2017
Controlling mechanism of chlorination on emitter bio-clogging for drip irrigation using reclaimed water
.
Agricultural Water Management
184
,
36
45
.
https://doi.org/10.1016/j.agwat.2016.12.017
.
Song
P.
,
Feng
G.
,
Brooks
J.
,
Zhou
B.
,
Zhou
H.
,
Zhao
Z.
&
Li
Y.
2019
Environmental risk of chlorine-controlled clogging in drip irrigation system using reclaimed water: The perspective of soil health
.
Journal of Cleaner Production
232
,
1452
1464
.
https://doi.org/10.1016/j.jclepro.2019.06.050
.
Stilling
P. D.
1996
Ecology
, 4th edn.
Prentice Hall, Inc
,
Upper Saddle River, NJ
.
Tarchitzky
J.
,
Rimon
A.
,
Kenig
E.
,
Dosoretz
C.
&
Chen
Y.
2013
Biological and chemical fouling in drip irrigation systems utilizing treated wastewater
.
Irrigation Science
31
,
1277
1288
.
https://doi.org/10.1007/s00271-013-0406-0
.
Title 22 Code of Regulations
.
State Water Resources Control Board
.
California Water Boards
.
Torkzadeh
H.
,
Zodrow
K. R.
,
Bridges
W. C.
&
Cates
E. L.
2021
Quantification and modeling of the response of surface biofilm growth to continuous low intensity UVC irradiation
.
Water Research
193
,
116895
.
https://doi.org/10.1016/j.watres.2021.116895
.
UN-Water
2018
Water Scarcity
.
Available from: https://www.unwater.org/water-facts/scarcity/ (accessed 4 August 2023)
US EPA
2006
Ultraviolet Disinfection Guidance Manual for the Final Long Term 2 Enhanced Surface Water Treatment Rule
.
US EPA
,
Washington D.C
US EPA
2012
Water Reuse Guidelines
.
United States Environmental Protection Agency, Office of Water
.
US EPA, OW
2019
Basic Information About Water Reuse
.
US EPA
.
August 13, 2019. Available from: https://www.epa.gov/waterreuse/basic-information-about-water-reuse (accessed 4 August 2023)
Villaverde
A. G.
&
Barbe
J.
1986
ATP production after ultraviolet irradiation in Escherichia coli
.
Current Microbiology
14
(
1
),
31
34
.
https://doi.org/10.1007/BF01568098
.
Wilson
C.
,
Stevenson
H. L.
&
Chrzanowski
T. H.
1981
The contribution of bacteria to the total adenosine triphosphate extracted from the microbiota in the water of a salt-marsh creek
.
Journal of Experimental Marine Biology and Ecology
50
(
2–3
),
183
195
.
https://doi.org/10.1016/0022-0981(81)90049-6
.
Wilson
C.
,
Lukowicz
R.
,
Merchant
S.
,
Valquier-Flynn
H.
,
Caballero
J.
,
Sandoval
J.
&
Okuom
M.
2017
Quantitative and qualitative assessment methods for biofilm growth: A mini-review
.
Research & Reviews. Journal of Engineering and Technology
6
(
4
).
Zhao
Z.
,
Luo
Y.
,
Wang
T.
,
Sinha
S.
,
Ling
L.
,
Rittmann
B.
,
Alvarez
P.
,
Perreault
F.
&
Westerhoff
P.
2023
Phenotypic and transcriptional responses of Pseudomonas aeruginosa biofilms to UV – C irradiation via side-emitting optical fibers: Implications for biofouling control
.
Environmental Science & Technology
.
https://doi.org/10.1021/acs.est.3c04658
.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Licence (CC BY 4.0), which permits copying, adaptation and redistribution, provided the original work is properly cited (http://creativecommons.org/licenses/by/4.0/).

Supplementary data