ACS Symposium Series (ACS Publications)

Chapter 9

Biofilms and Microbiologically Influenced Corrosion in the Petroleum Industry Downloaded via COLUMBIA UNIV on September 3, 2019 at 08:38:21 (UTC). See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.

Damon C. Brown and Raymond J. Turner* Biological Sciences, University of Calgary, 2500 University Drive NW, T2N 1N4 Calgary, Alberta, Canada *E-mail: [email protected].

Microbiologically influenced corrosion (MIC) is a process caused by microbial growth activities. Some MIC is caused by the metabolites whereas other MIC originates from surface-associated microbes (biofilms). MIC causes significant and costly damage in environments ranging from ship hulls to water distribution and cooling systems to oil and gas processing and transportation facilities. Due to the nature of these systems, there are frequently microniche environments with varying concentrations of electron acceptors, such as oxygen and sulfate, which allow for the growth of diverse microbial communities. In systems such as oil and gas pipelines, planktonic cells adhere to surfaces where water accumulates and begin forming biofilms. This chapter provides an overview of the different ways microbes growing as a biofilm can influence corrosion and biofouling in the petroleum distribution industry (pipelines primarily). MIC is believed to occur via two mechanisms: chemical MIC and electrical MIC. This chapter will also reflect on the resistance of MIC biofilm to antiseptic corrosion control inhibitors.

The process by which microbes exist in a biofilm and how these biofilms contribute toward and facilitate corrosion has many names. This process has been referred to in literature as microbiologically influenced/induced corrosion, biocorrosion, and biodegradation, and biofouling. The term induced has fallen out of favor as the microbial mechanism does not (always) initiate corrosion but can accelerate other forms of corrosion. This is discussed in detail later. Additionally, microbially is often used interchangeably with microbiologically, but the abbreviation of microbiologically influenced corrosion (MIC) refers to this process generally. MIC is an issue in many industries, including petroleum production and transportation, water cooling systems, and ship hulls. Generally, any industry where metal and water are in contact has the potential for corrosion, and any of these systems that can support microbial growth is therefore also susceptible to MIC. Here, we will focus primarily on petroleum transportation systems, specifically the role microbes growing as a biofilm play in internal corrosion of these pipelines. © 2019 American Chemical Society

Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

Biofilm Formation Microbial cells exist in two main phenotypic forms of growth: planktonic and sessile. Planktonic cells, or free-floating/free-swimming cells, exist as individual cells and act in an ecological sense as a means of relocation for the microbes into another habitat. In this state, they are often more susceptible to antimicrobial treatment and predation from protozoa (1, 2). Sessile cells, or cells growing in a biofilm, are planktonic cells that have irreversibly attached to a surface and are the predominant form of microbial growth in nature (3). Biofilms are multispecies communities and have been associated with dental plaque, biofouling of water treatment facilities, MIC, and infections from medical implants and devices (3). The formation of biofilms is extensively reviewed elsewhere (4–6) and references therein) and is therefore only described briefly here. As illustrated in Figure 1, biofilm formation and growth consist of five stages: initial attachment, irreversible attachment, biofilm maturation, metabolic stratification, and dispersal. To begin, planktonic cells initiate a reversible attachment to a surface mediated by pili and flagella (Figure 1A). Cells will attach and detach from a surface until appropriate conditions are met, at which point they initiate irreversible attachment. Many different attachment factors have been proposed, including surface associated systems (such as proteinaceous curli fibers, type I fimbriae, and antigen 43 (7)) and enhanced attachment or surface sensing by pili and flagella (8–10). Following irreversible attachment, the cells initiate biofilm formation by secreting the components of a biofilm, namely exopolysaccharides (EPS), extracellular DNA, lipids, and proteins, and are considered sessile (Figure 1B) (11). A key physiological change that occurs once a microbe enters the irreversible state of sessile growth is a significant change of its genetic expression, resulting in a distinct phenotype from planktonic (12, 13). Escherichia coli has been shown to change its genetic expression by 14–38% (14, 15) and Pseudomonas aeruginosa showed a >50% change in proteomic expression in biofilms compared to planktonics (16). Although E. coli and P. aeruginosa are not predominant organisms in classical MIC environments, the drastic change in phenotypes of these model organisms serves as an indicator of the issues that arise from studying planktonic cells as a proxy for the sessile cells. During this shift, the global regulator csgD is upregulated, which leads to the increased expression of curli fimbriae, EPS, and antigens (12). The heterogeneity of the biofilm and its components means it is impractical to target individual components as a means of controlling biofilm production. One system attributed to inducing biofilm formation is quorum sensing through signaling molecules known as autoinducers. However, autoinducers do not have universally received signals, such as in the case of Staphylococcus aureus where the quorum sensing negatively regulates biofilm formation (17). In addition to endogenous signaling molecules and metabolites which trigger population responses, subinhibitory concentrations of antibiotics and biocides have also been shown to induce biofilm formation (18–21). Once formed, the biofilm grows and matures while the cell population increases and diversifies as different species attach and are recruited into the community (Figure 1C). As the population diversifies, metabolic stratification occurs as a result of nutrients being consumed at the outer edge and metabolic end products are released (22, 23). One such example is the consumption of oxygen at the outer edge of P. aeruginosa biofilms, resulting in depleted oxygen concentrations deeper within the biofilm (24). At this stage, the biofilm forms a porous structure, facilitating delivery nutrients, although not preventing the formation of these gradients (Figure 1D) (3, 25). The structure of the biofilm improves nutrient diffusion, but the activity and density of cells in the biofilm still results in 188 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

lower metabolically active or completely inactive subpopulations (known as persisters) (4). As the biofilm reaches maturation, planktonic cells are dispersed from the biofilm and travel through the environment to settle in a different location, restarting the process (Figure 1E). It has been shown the dispersed cells may be individual planktonic cells or small flocs of biofilm-containing aggregates of cells (26).

Figure 1. The steps of biofilm formation and maturation. A) Initial attachment—planktonic cells initiate a reversible attachment to a surface mediated by pili and flagella. B) Irreversible attachment—cells shed their pili and flagella and begin producing components of the biofilm matrix (EPS, DNA, lipids, and proteins). C) Biofilm maturation—cells begin to grow and replicate. Microbial diversity increases as new genera contact the biofilm and are incorporated into the community. D) Metabolic stratification—as the biofilm grows and develops, the community members will create niche environments as nutrients are taken up at the outer edge and metabolic end products are released and diffuse deeper into the biofilm. E) Dispersal—mature biofilms begin shedding mobile planktonic cells into the environment to form new biofilms and begin the cycle again. Biofilms in the Petroleum Industry As mentioned earlier, MIC can affect any environment where microbes can exist in contact with metal and water. In the petroleum industry, biofilms are found in locations where water accumulates or persists long enough for the establishment of a microniche suitable for growth, at which time the microbes can become irreversibly attached. High-velocity systems that never see stagnation or reduction in flow velocity are theoretically immune to biofilm formation. Unfortunately, this is not the case in any pipeline system. The presence of water accumulation points such as dead legs, low flow conditions, shut-ins, or the structural designs that facilitate water condensation, such as the bottom of (sufficiently) inclined pipes, provides niche opportunities for biofilms to initiate. Additionally, in systems with water wet internal surfaces and laminar flow, the fluid velocity near the pipewalls (on the micron scale) will experience near zero velocity. These periods of low or stagnant flow provide adequate conditions for the initiation of a biofilm formation, at which point even after flow is resumed the biofilm will persist, barring physical removal. One important note about the accumulation of water: microbes do not require significant amounts of water to be present (27). In fact, in predominantly oil wet systems, the water entrained on solid particulates can provide sufficient water to disrupt the oil wet surface, which allows microbes, in association with water, to contact the metal surface and yet still feed off the petroleum products. The partitioning of oil hydrocarbons into water at low concentrations (due to solubility) acts as a barrier to protect the cells from hydrocarbon 189 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

toxicity. These compounds can then be used as a food source from the ambient concentration in the water layer. The high cell density and microbial diversity of a biofilm results in the accumulation of metabolic end products, such as organic acids, CO2, and H2S, which can result in significantly different microniche environments within and underneath biofilms compared to the environment. Some of these metabolic end products can be shared between species in syntrophic and symbiotic associations, thus increasing the potential for microbial diversity. Microbial populations in a diverse range of environments have been assessed using 16S ribosomal RNA (rRNA) sequencing and have shown the number of different species can reach up to 50,000 per gram of soil (28), 9700 unique operation taxonomic units (OTUs) in marine sediments (29), a core microbiome of 7522 OTUs in dental plaque (with up to 10,000 OTUs identified) (30), 71–88 phylotypes in petroleum reservoirs (31, 32), and 7–52 in MIC (29, 33). The number of species consistently identified in environmental samples illustrates that biofilms are complex habitats harboring a wide diversity of species and thus vast metabolic potential far beyond what has been replicated in research laboratories. It is assumed that oil reservoirs are the main source of inoculation for petroleum-associated facilities. Therefore, a glance at the commonly identified microbes living in oil reservoirs suggests the range and types of microbes present in the petroleum industry. Using culturing methods, many of the identified reservoir microbes are halophilic, thermophilic, and anaerobic, although not exclusively so (34). The types of microbes found are predominantly fermentative, sulfate-reducing, iron(iii)reducing, and methanogenic (34). Aerobic thermophilic bacteria were also identified near the injection wells, indicating they are consuming the oxygen present in the injection waters when it enters the reservoir but are likely naturally occurring in these environments (34). Molecular techniques (specifically 16S rRNA sequencing) have revealed similar community compositions. Petroleum reservoirs are frequently composed of methanogenic Archaea, Firmicutes, and β‑, δ‑, γ-Proteobacteria (32). A more recent study using samples from a waterflooded reservoir to assess active microbial members confirmed these trends (35). A 16S rRNA analysis identified multiple types of methanogenic Archaea and the bacterial population was predominantly Geobacillus. Others identified belong to aerobic organotrophs, denitrifying, fermentative, iron-reducing, and sulfate/sulfur-reducing bacteria (35).

Microbiologically Influenced Corrosion As with abiotic corrosion, MIC requires the four fundamental elements of a corrosion cell to be present in order for corrosion to occur. These elements are: anode (electron donating region), cathode (electron accepting region), electrolyte (conductive fluid), and an electrical connection between the anode and the cathode. As mentioned earlier, MIC influences corrosion as opposed to the outdated name, microbiologically induced corrosion, which implies the microbes initiate the corrosion. As the new name suggests, the microbes do not initiate corrosion but affect corrosion in multiple ways, which are discussed in detail later. Therefore, with the understanding that the microbes influence corrosion, it is easier to understand their role in affecting the anodic and cathodic elements of a corrosion cell. On the anode side, microbes can create small local anodes underneath their biofilms that can cause rapid corrosion as the size difference between the anode and cathode increases the rate of electron removal. On the cathode side, microbes can cause depolarization, consuming hydrogen from the metal surface and driving corrosion forward. It is important to note 190 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

that not all biofilms are corrosive. In certain cases, biofilms can form a protective barrier that reduces corrosion, as in the case of E. coli and Geobacter sulfurreducens biofilms grown on stainless steel (36, 37). MIC has been studied extensively in laboratory conditions using single species cultures (38–43). These studies have reported maximum corrosion rates of 0.7 mm/year by a pure culture of sulfatereducing bacteria (SRB) on mild steel (44) and 0.8 mm/year by nitrate-reducing bacteria (NRB) on C1018 carbon steel (45) in laboratory conditions. These rates are often considered insignificant for industrial settings. Although mathematically, this implies that a 15 mm thick pipe will corrode through in 15–20 years, defining the lifetime of the system. One of the more problematic issues with MIC is its tendency to cause pitting corrosion, which is corrosion that causes little metal loss but potentially leads to asset failure due to through-wall penetration. The size of the corrosion pit and the small amount of metal loss make MIC-associated corrosion difficult to detect using standard detection methods, such as intelligent pigging. Microbes have three main mechanisms by which they can influence corrosion: chemical microbiologically influenced corrosion (cMIC), electronically microbiologically influenced corrosion (eMIC), and concentration cell units. These mechanisms are visited in more detail below. Chemical Microbiologically Influenced Corrosion The process by which the metabolites of microbial metabolism can influence the rate and process of corrosion is cMIC. Generally, cMIC can be considered an indirect corrosion mechanism. Historically, the most well-studied mechanisms are the ones thought to produce the most aggressive corrosion rates, which are sulfate-reducing prokaryotes (SRP, alternatively referred to as SRB) and, more recently, acid-producing bacteria. As their name suggests, SRP reduce sulfate, producing a spectrum of sulfur species including sulfur, sulfide, and polysulfides. Sulfur is a highly corrosive element but is infrequently produced as it has an intermediarily reduced state, and thus sulfide species are more commonly produced. Sulfide can interact with the metal surface to remove electrons from the cathode, thus allowing corrosion to occur by, in part, physically removing the metal atoms from the surface into a metal sulfide (Figure 2) (46). This process with ferrous ions produces iron sulfide, which precipitates out on the surface along with iron carbonate forming from reactions with CO2. Iron sulfide and iron carbonate scales are conductive, which effectively increases the size of the cathode and facilitates increased corrosion rates because of an increased surface area, allowing more microbes and metabolites to interact with, and effectively remove electrons from, the steel surface (44).

Figure 2. Schematic of chemical reactions associated with cMIC. The acid-producing bacteria are a broad group of microorganisms that produce weak organic acids, such as acetic acid, as their main metabolic end product. These weak organic acids are produced in the biofilm, where they can be concentrated. This produces localized pH values that 191 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

are magnitudes different from that of the bulk fluid. As a comparative example, oral biofilms using Streptococcus mutans, a fermentative bacteria which produces organic acids, can create a pH of 4.5–5.5 at the base of the biofilm compared to a pH of 7 in the bulk fluid (47, 48). These weak acids serve as a source of protons, which interact with the cathodic region of the metal, removing electrons and leading to corrosion. As the lowest pH is concentrated at the base of the structures of the biofilm, it is considered that this can lead to increased pitting corrosion under the biofilm. Electrical Microbiologically Influenced Corrosion Microbes can employ eMIC as a more direct mechanism that facilitates corrosion. In this mechanism, the microbes strip the electrons directly from the metal surface for use in their metabolism but can gain access to the electrons through different mechanisms (Figure 3). The most well-known instances of this are certain SRP (40, 49) and some NRB (50, 51) that are capable of stripping electrons directly from the metal for use in their metabolism. SRP remove the electrons from the insoluble iron surface and conduct them to the inner membrane, where they are used to reduce sulfate. This is a process known as extracellular electron transfer (EET), which generally involves the conduction of electrons to or from an external surface for use in metabolism. In eMIC, iron serves as the electron donor in lieu of hydrocarbons. Thus, EET is promoted under carbon starvation conditions (52).

Figure 3. eMIC highlighting A) electron uptake via microbial nanowires, B) direct electron uptake, and C) conductive iron scale transfer of electrons to microbes throughout the crust. The release of electrons causes solubilization of the iron, which reacts abiotically with hydrogen sulfide and bicarbonate, settling out as iron scales. EET is a cytochrome-dependant process (specifically c-type cytochromes) and is believed to occur via three main pathways: direct cellular contact with the external electron donor (or acceptor), contact through a biological nanowire, and contact through a soluble, extracellular electron shuttling process (45). There is some controversy regarding the actual mechanisms involved in EET, as the distance that needs to be covered by the electron in all instances between the surface and the microbe is vast. It is unclear if the c-type cytochromes are able to directly contact the metal surface and transfer electrons or if soluble electron shuttles such as flavins (e.g., flavin mononucleotide and riboflavin) are required to bridge this gap in all suggested EET pathways (53). It has been observed that increased corrosion rates by the SRP Desulfovibrio vulgaris (54, 55) and the NRB P. aeruginosa (56) are observed following the addition of riboflavin and flavin mononucleotide. While much of the MIC data is focused on SRB, it is becoming more apparent from studies using P. aeruginosa (which is also known 192 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

for its potent ability to form biofilms) that MIC involves much more than a single corrosive species cohabitating in a diverse, innocuous community. Recently, the mechanisms (nanowires, c-type cytochromes, and flavins) behind direct EET have received more interest for their role in processes involving microbial fuel cells (53, 57–59). Overall, these appear to be conversed mechanisms used by different bacteria capable of operating in either direction, depending on the energetic needs of the cell. Concentration Cells Another mechanism of MIC is the creation of concentration cells. While not frequently considered as a driving mechanism of MIC, this mechanism can be caused by any species existing in a biofilm. In fact, there is even the potential of facilitating corrosion as a sterilized biofilm. A concentration cell is effectively formed whenever a difference in electrical potential is created on a metal surface. In aerobic systems with a biofilm, this is most easily seen by observing the oxygen concentrations surrounding and underneath a biofilm. As the aerobic microbes consume the oxygen in solution, the environment under the biofilm, specifically at the biofilm/metal interface, will become more anoxic (45). Correspondingly, the uncovered metal surface surrounding the biofilm will become more cathodic, thus completing the electrochemical corrosion cell (Figure 4). Although oxygen is the simplest example of how a concentration cell is formed, these cells can also be created by the accumulation of other compounds such as soluble metal ions (which interact with negatively charged compounds in biofilms such as DNA, discussed earlier), which become entrapped in the biofilm and concentrate passively, without the need for microbial metabolism to accelerate the process. As such, these metal concentration cells can form in preexisting biofilms without active microbial cells.

Figure 4. Concentration cell depicting A) an oxygen gradient, which forms when active microbes consume oxygen, primarily at the surface, and prevent oxygen from reaching the metal surface and B) a metal concentration cell, which traps solubilized metal ions, preventing contact with the metal surface. The concentration cells create an anodic environment under the biofilm, allowing electrons to flow to the cathodic regions around the biofilm where they are consumed and drive corrosion forward.

Microbial Biocide Resistances Biofilms offer multiple mechanisms for resistances. In a medical context, this includes altering antibiotic targets, preventing entry of the antibiotic into the cell, enzymatic degradation of the antibiotic, and maintaining a low intracellular concentration. Unlike the medical field, the petroleum industry does not employ antibiotics but uses broad-spectrum antiseptics (commonly referred to as biocides) such as glutaraldehyde, quaternary ammonium/cationic compounds, surfactants, and mixes thereof. Due to the nature of the killing mechanism of these compounds, many of the 193 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

resistance mechanisms used against antibiotics (e.g., degradation or altering the target) are not useful against these biocides. Briefly, a clarification between tolerance and resistance should be made. Resistance is an acquired trait that persists beyond the presence of the dangerous chemical as a stable resistant variant. This may be through specific mutations or the acquisition of genes for resistance. Alternatively, tolerance is an insusceptibility that results from physiological changes caused by biofilm-associated growth, including slow growth or nutrient stress. Tolerance is lost as the cells revert back to planktonic form (60). Both resistance and tolerance mechanisms provide protection against antimicrobial biocidal compounds. Under typical pipeline conditions, microbes have developed alternative resistance or tolerance mechanisms. The two most prominent are believed to be the use of the EPS of the biofilm as a diffusion barrier and the use of multidrug resistance efflux pumps (MDREPs). MDREPs are inner membrane proteins that bind a broad range of substrates and extrude them from the cytosol into the periplasm or directly into the environment (when in association with outer membrane proteins, e.g., TolC), maintaining a low intracellular concentration. Tolerant bacteria are not killed by exposure to the biocide but are not in a metabolically active state. When the biocide is removed, these cells become active once again and repopulate the community. These cells are known as persister cells and comprise approximately 1% of a community in mid-log growth phase (61). Table 1. List of MDREPs and Their Nonantibiotic Substrates Superfamily

MDREP Gene

Substrates

Reference

emrE

Benzalkonium, acriflavine, tetraphenylphosphonium, methyl viologen, rhodamine 6G

Bay et al. 2008 (69)

sugE

Ethidium, benzalkonium, cetylpyridinium, cetyltrimethylammonium bromide, tetraphenylphosphonium

He et al. 2011 (70)

qacH

QACs, ethidium

Sundheim et al. 1998 (71)

ABC

lmrA

Ethidium, tetraphenylphosphonium, daunorubicin, several chemotherapy drugs

Van Veen et al. 2000 (72)

RND

acrAB

Sodium dodecyl sulfate, organic solvents, triclosan, chlorhexidine, QACs

Zgurskaya and Nikaido 2000 (73)

MFS

qacA

QACs, acriflavine, diamidines, ethidium

Wassenaar et al. 2015 (74)

mdfA

Benzalkonium, tetraphenylphosphonium

Edgar and Bibi 1997 (75)

norA

Most QACs

Costa et al. 2013 (76)

PACE

aceI, adeABC

Acriflavin, benzalkonium, chlorhexidine

Hassan et al. 2018 (77)

MATE

norM

Acriflavin, benzalkonium, tetraphenylphosphonium

Ogawa et al. 2015 (78)

SMR

194 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

The close proximity, diversity, and cell count of a microbial community within a biofilm offers an excellent opportunity for genetic transfer. Indeed, multiple groups have shown that horizontal transfer (i.e., genetic exchange between cells not through cell replication) occurs at elevated rates within biofilms (62, 63). Many MDREP genes are encoded on mobile gene elements such as transposons, plasmids, insertion sequences, genomic islands, integrons, and integrative and conjugative elements (64–7). These elements have led to the hypothesis that these proteins play a role in increasing the microbes’ tolerance to these antiseptic compounds. Little is understood about how the specificity of these efflux pumps is determined, as they are able to efflux vast ranges of compounds with seemingly little or no structural similarities. Table 1 provides a brief list of representative MDREPs and their nonantibiotic substrates. These MDREPs and their role in biocide tolerance are described elsewhere (68). These genes exist on mobile gene elements, resulting in the potential for their rapid spread through a microbial community during sublethal exposure to specific biocides and an increased microbial tolerance toward the biocides. Eradication of Biofilms In the petroleum industry, pipeline maintenance is typically done using two main methods, and they are often combined to improve efficacy. The two treatment methods are mechanical and chemical. Mechanical cleaning is done using scraping tools sized to the internal diameter of the pipeline known as pigs. The pigs are pushed along the length of the pipeline by the force of the product flow. Many different types of pigs exist, offering different types of scraping edges (e.g., brushes and discs) or equipped with electronic equipment, which can assess the integrity of the internal surface and detect areas of corrosion (intelligent pigging). These electronic pigs (or smart pigs) are not used for integrity assessment instead of cleaning. As such, multiple pigs are typically launched together in trains to clean the internal surface prior to the intelligent pigs’ assessment. It is important to note that even following pigging treatment that scrapes the internal surface, if the biofilm has already formed and occupied a pit, the pigs are unable to remove the biofilm embedded in the pipewall. Additionally, biofilms can exist on the order of microns in thickness and thus can be missed by pigs that are unable to maintain such a tight scraping junction throughout the entire length of the pipe. Chemical treatments and the microbial responses thereof are discussed below. Assessing the microbial population and viability following mitigation treatments is a difficult task. The potential for microbial tolerance toward biocides results in inconsistent results following repeated, identical treatments. The biocides/antiseptics treat the active cells (although rarely to annihilation, as discussed later), but the biofilm thickness remains intact and able to create abiotic concentration cells. Furthermore, the presence of an eradicated biofilm existing on a pipewall of a water system following chlorine treatment showed a twofold faster bacterial accumulation rate than a bare pipewall (79), illustrating how quickly cells can repopulate a treated biofilm. Traditional belief has been that an upregulation of MDREP genes in the presence of a biocide has maintained a sublethal concentration within the cell (80, 81). Mature biofilms (4 or more days old) have been shown to have an increased resistance to glutaraldehyde, attributed to the increased expression of the MDREPs emrA and oprN (82). Microbial tolerance is also attributed to the diffusion barrier of the biofilm toward charged or large-sized compounds and the presence of metabolically inactive cells in the biofilm community (60). Additionally, population resistance has been attributed to biofilms since the additional biofilm components and cells create a reaction-diffusion barrier, effectively shielding cells deeper in the biofilm (83).


As discussed prior, biofilms are a diverse community that have been found to be more resistant to biocides, although the mechanism has not been elucidated (84–86). Recent work has suggested that MDREPs may not be the driving force behind Staph. aureus tolerance toward quaternary ammonium compounds (QACs), but rather a change in membrane composition (87). This remains an area of intense research in many fields, including the medical, food, and petroleum industries. Biocide Corrosion Control Internal surfaces are chemically treated using the previously discussed biocides. In product flow pipelines, oxidizing biocides (e.g., ClO2 and hydrogen peroxide) are not employed as they are corrosive themselves. In the petroleum industry, they are reserved for applications that have short exposure time to metal surfaces, such as water injection systems. As such, nonoxidizing biocides are much more frequently employed in transportation pipelines. Some of the more common nonoxidizing biocides include glutaraldehyde, formaldehyde, tetrakis(hydroxymethyl)phosphonium sulfate, QACs, 2-bromo-2-nitro-1,3-propanediol (bronopol), and 2,2-dibromo-3-nitrilopropionamide (DBNPA). These biocides are applied in proprietary blends that can contain surfactants to improve penetration into biofilms. A thorough review of biocides, their compatibilities, and other considerations for their use is available elsewhere (88). Briefly, glutaraldehyde is a broad-spectrum biocide that cross-links primary amines, such as lysine residues in cell walls. While this effectively kills microbial cells, it also reacts with proteins present in the biofilm, creating more tenacious biofilms by cross-linking the matrix. Tetrakis(hydroxymethyl)phosphonium sulfate is also broad-spectrum and disrupts proteins in the cell membrane and inhibits the activity of lactate dehydrogenase. QACs are cell membrane surface active compounds that disrupt membrane organization but can also bind DNA (89). Bronopol oxidizes thiol groups in proteins and is a slow killing biocide. At acidic pH, bronopol persists for long periods, and at alkaline pH, it will break down into formaldehyde, bromide, and nitrates. Finally, DBNPA is a highly reactive biocide that degrades quickly and is believed to react with cell membrane proteins and the inactivation of enzyme systems. Further complicating the issue of assessing MIC is the use of traditional growth-based assays (known as bug bottles) to determine cell viability following treatment. These methods use bottles with prepared media designed to grow specific microbes of interest, such as SRP and NRB, but can frequently provide false positives and negatives. Additionally, bug bottles are inoculated with planktonic cells from a liquid sample, which is then assumed to represent the sessile population present throughout the entire system. Research has shown that in a continuous flow cell, planktonic cells are rapidly released by biofilm communities as young as 6 h old, even when the flow rate was 58 times the planktonic cell growth rate (90). While there is no debate that biofilms release planktonic cells, there is no evidence to suggest that a consistent, reliable ratio exists between planktonic and sessile cell counts, or even that the planktonic diversity accurately depicts the sessile populations. It is extremely difficult to get precise surface samples from the internal pipewall in practice. Although testing methods such as Robbins devices (91) and other midstream or flush mounted corrosion coupons exist, their location in the pipe ultimately determines how accurately they will represent the microbial community, if at all. For example, does a small, easily accessible slipstream outfitted with a Robbins device accurately represent the microbial population that would develop throughout the pipeline when positioned at the base of a subterranean incline? Likely not. Does the community that attaches to a midstream coupon accurately depict the community that settles 196 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

onto the pipe surface in association with solids and debris? Likely not. Additionally, MIC issues manifest in local niches on the scale of microns to millimeters, so even when the coupons are flush mounted and located in the correct region of the pipe, there is a small likelihood that the coupons will accurately report the extent to which MIC is occurring. Here we did not discuss preventative methods, such as internal linings/coatings to prevent cell attachment and biofilm proliferation, as these methods are not typically employed and prevent the use of mechanical and chemical treatments.

Conclusion Biocide resistance in the petroleum industry is a field of growing interest. To date, no conclusive evidence has been put forward to suggest the mechanism driving microbial tolerance of biocide treatments observed in field operations. As our control over abiotic corrosion increases, the impact of MIC and our awareness thereof grow. It is increasingly obvious that MIC is a complex system and requires a more elegant approach than using broad-spectrum biocides with little or no prescreening or understanding of how the microbial community will respond and adapt to such treatments. The true danger of MIC lies in microbes’ ability to slowly, persistently influence corrosion on a small scale (with respect to metal loss) but still lead to system failure as a result. Our understanding of tolerance toward biocides stems from the medical field and its treatment of biofilm infections and from the food industry and its treatment of surface contaminations, but the obvious differences in the petroleum industry drive the need to develop unique mitigation strategies tailored for the unique conditions of the petroleum industry. Improving our understanding of how individual microbes and the MIC biofilm as a whole influence corrosion is an important fundamental way to improve our handling and transportation of a key economic and growing social issue.

References 1. 2.

3. 4. 5. 6. 7. 8. 9.

Costerton, J. W.; Stewart, P. S; Greenberg, E. P. Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284, 1318–1322. Raghupathi, P. K.; Liu, W.; Sabbe, K.; Houf, K.; Burmølle, M.; Sørensen, S. J. Synergistic Interactions Within a Multispecies Biofilm Enhance Individual Species Protection Against Grazing by a Pelagic Protozoan. Front. Microbiol. 2018, 8, 2649. Flemming, H.-C.; Wingender, J.; Szewzyk, U.; Steinberg, P.; Rice, S. A.; Kjelleberg, S. Biofilms: An Emergent Form of Bacterial Life. Nat. Rev. Microbiol. 2016, 14, 563–575. López, D.; Vlamakis, H.; Kolter, R. Biofilms. Cold Spring Harb. Perspect. Biol. 2010, 2, a000398. Donlan, R. M. Biofilms: Microbial Life on Surfaces. Emerg. Infect. Dis. 2010, 8, 881–90. Costerton, J. W.; Lewandowski, Z.; Caldwell, D. E.; Korber, D. R.; Lappin-Scott, H. M. Microbial Biofilms. Annu. Rev. Microbiol. 1995, 49, 711–745. Beloin, C.; Roux, A.; Ghigo, J. M. Escherichia Coli Biofilms. Curr. Top. Microbiol. Immunol. 2008, 322, 249–89. Friedlander, R. S.; Vogel, N.; Aizenberg, J. Role of Flagella in Adhesion of Escherichia coli to Abiotic Surfaces. Langmuir 2015, 31, 6137–6144. Belas, R. When the Swimming Gets Tough, the Tough Form a Biofilm. Mol. Microbiol. 2013, 90, 1–5. 197 Rathinam and Sani; Introduction to Biofilm Engineering ACS Symposium Series; American Chemical Society: Washington, DC, 2019.

10. Belas, R. Biofilms, Flagella, and Mechanosensing of Surfaces by Bacteria. Trends Microbiol. 2014, 22, 517–527. 11. Besharova, O.; Suchanek, V. M.; Hartmann, R.; Drescher, K.; Sourjik, V. Diversification of Gene Expression During Formation of Static Submerged Biofilms by Escherichia coli. Front. Microbiol. 2016, 7, 1568. 12. White, A. P.; Weljie, A. M.; Apel, D.; Zhang, P.; Shaykhutdinov, R.; Vogel, H. J.; Surette, M. G. A Global Metabolic Shift is Linked to Salmonella Multicellular Development. PLoS One 2010, 5, e11814. 13. Mikkelsen, H.; Duck, Z.; Lilley, K. S.; Welch, M. Interrelationships Between Colonies, Biofilms, and Planktonic Cells of Pseudomonas aeruginosa. J. Bacteriol. 2007, 189, 2411–2416. 14. Prigent-Combaret, C.; Vidal, O.; Dorel, C.; Lejeune, P. Abiotic Surface Sensing and BiofilmDependent Regulation of Gene Expression in Escherichia coli. J. Bacteriol. 1999, 181, 5993–6002. 15. Schembri, M. A.; Kjaergaard, K.; Klemm, P. Global Gene Expression in Escherichia coli Biofilms. Mol. Microbiol. 2003, 48, 253–267. 16. Sauer, K.; Camper, A. K.; Ehrlich, G. D.; Costerton, J. W.; Davies, D. G. Pseudomonas aeruginosa Displays Multiple Phenotypes During Development as a Biofilm. J. Bacteriol. 2002, 184, 1140–1154. 17. Boles, B. R.; Horswill, A. R. agr-Mediated Dispersal of Staphylococcus aureus Biofilms. PLoS Pathog. 2008, 4, e1000052. 18. Hoffman, L. R.; D’Argenio, D. A.; MacCoss, M. J.; Zhang, Z.; Jones, R. A.; Miller, S. I. Aminoglycoside Antibiotics Induce Bacterial Biofilm Formation. Nature 2005, 436, 1171–1175. 19. Yim, G.; Wang, H. H.; Davies, J. Antibiotics as Signalling Molecules. Philos. Trans. R. Soc. B Biol. Sci. 2007, 362, 1195–1200. 20. Tezel, U.; Pavlostathis, S. G. Quaternary Ammonium Disinfectants: Microbial Adaptation, Degradation and Ecology. Curr. Opin. Biotechnol. 2015, 33, 296–304. 21. Pagedar, A.; Singh, J.; Batish, V. K. Adaptation to Benzalkonium Chloride and Ciprofloxacin Affects Biofilm Formation Potential, Efflux Pump and Haemolysin Activity of Escherichia coli of Dairy origin. J. Dairy Res. 2012, 79, 383–389. 22. Spormann, A. M. Physiology of Microbes in Biofilms. Curr. Top. Microbiol. Immunol. 2008, 322, 17–36. 23. Stewart, P. S.; Franklin, M. J. Physiological Heterogeneity in Biofilms. Nat. Rev. Microbiol. 2008, 6, 199–210. 24. Xu, K. D.; Stewart, P. S.; Xia, F.; Huang, C. T.; McFeters, G. A. Spatial Physiological Heterogeneity in Pseudomonas aeruginosa Biofilm is Determined by Oxygen Availability. Appl. Environ. Microbiol. 1998, 64, 4035–4039. 25. Flemming, H.-C.; Wingender, J. The Biofilm Matrix. Nat. Rev. Microbiol. 2010, 8, 623–633. 26. Hall-Stoodley, L.; Costerton, J. W.; Stoodley, P. Bacterial Biofilms: From the Natural Environment to Infectious Diseases. Nat. Rev. Microbiol. 2004, 2, 95–108. 27. Place, T. D.; Holm, M. R.; Cathrea, C.; Ignacz, T. Understanding and Mitigating UnderDeposit Corrosion in Large Diameter Crude Oil Pipelines: A Progress Report. Proceedings of


28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

the 2008 7th International Pipeline Conference, Vol. 2, Calgary, Alberta, Canada, Sept 29–Oct 3, 2008; ASME: Calgary, Alberta, Canada 2008; pp 809–821; IPC2008-64562. Vasileiadis, S.; Puglisi, E.; Arena, M.; Cappa, F.; Cocconcelli, P. S.; Trevisan, M. Soil Bacterial Diversity Screening Using Single 16S rRNA Gene V Regions Coupled with Multi-Million Read Generating Sequencing Technologies. PLoS One 2012, 7, e42671. Zhu, D.; Tanabe, S.-H.; Yang, C.; Zhang, W.; Sun, J. Bacterial Community Composition of South China Sea Sediments Through Pyrosequencing-Based Analysis of 16S rRNA Genes. PLoS One 2013, 8, e78501. Xiao, C.; Ran, S.; Huang, Z.; Liang, J. Bacterial Diversity and Community Structure of Supragingival Plaques in Adults with Dental Health or Caries Revealed by 16S Pyrosequencing. Front. Microbiol. 2016, 7, 1145. Li, H.; Yang, S.-Z.; Mu, B.-Z.; Rong, Z.-F.; Zhang, J. Molecular Phylogenetic Diversity of the Microbial Community Associated with a High-Temperature Petroleum Reservoir at an Offshore Oilfield. FEMS Microbiol. Ecol. 2007, 60, 74–84. Pham, V. D.; Hnatow, L. L.; Zhang, S.; Fallon, R. D.; Jackson, S. C.; Tomb, J.-F.; DeLong, E. F.; Keeler, S. J. Characterizing Microbial Diversity in Production Water from an Alaskan Mesothermic Petroleum Reservoir with Two Independent Molecular Methods. Environ. Microbiol. 2009, 11, 176–187. Jan-Roblero, M.; Romero, M.; Amaya, S.; Le Borgne, J. J. Phylogenetic Characterization of a Corrosive Consortium Isolated from a Sour Gas Pipeline. Appl. Microbiol. Biotechnol. 2004, 64, 862–867. Nazina, T. N.; Grigor’yan, A. A.; Shestakova, N. M.; Babich, T. L.; Ivoilov, V. S.; Feng, Q.; Ni, F.; Wang, J.; She, Y.; Xiang, T.; Luo, Z.; Belyaev, S. S.; Ivanov, M. V. Microbiological Investigations of High-Temperature Horizons of the Kongdian Petroleum Reservoir in Connection with Field Trial of a Biotechnology for Enhancement of Oil Recovery. Microbiology 2007, 76, 287–296. Nazina, T. N.; Shestakova, N. M.; Semenova, E. M.; Korshunova, A. V.; Kostrukova, N. K.; Tourova, T. P.; Min, L.; Feng, Q.; Poltaraus, A. B. Diversity of Metabolically Active Bacteria in Water-Flooded High-Temperature Heavy Oil Reservoir. Front. Microbiol. 2017, 8, 707. Abdoli, L.; Huang, J.; Li, H. Electrochemical Corrosion Behaviors of Aluminum-Based Marine Coatings in the Presence of Escherichia coli Bacterial Biofilm. Mater. Chem. Phys. 2016, 173, 62–69. Mehanna, M.; Basséguy, R.; Délia, M.-L.; Bergel, A. Geobacter sulfurreducens Can Protect 304L Stainless Steel Against Pitting in Conditions of Low Electron Acceptor Concentrations. Electrochem. Commun. 2010, 12, 724–728. Yuan, S. J.; Pehkonen, S. O. Microbiologically Influenced Corrosion of 304 Stainless Steel by Aerobic Pseudomonas NCIMB 2021 Bacteria: AFM and XPS Study. Colloids Surf., B. 2007, 59, 87–99. Jia, R.; Tan, J. L.; Jin, P.; Blackwood, D. J.; Xu, D.; Gu, T. Effects of Biogenic H2S on the Microbiologically Influenced Corrosion of C1018 Carbon Steel by Sulfate Reducing Desulfovibrio vulgaris Biofilm. Corros. Sci. 2018, 130, 1–11.


40. Venzlaff, H.; Enning, D.; Srinivasan, J.; Mayrhofer, K. J. J.; Hassel, A. W.; Widdel, F.; Stratmann, M. Accelerated Cathodic Reaction in Microbial Corrosion of Iron Due to Direct Electron Uptake by Sulfate-Reducing Bacteria. Corros. Sci. 2013, 66, 88–96. 41. Xu, D.; Zhou, E.; Zhao, Y.; Li, H.; Liu, Z.; Zhang, D.; Yang, C.; Lin, H.; Li, X.; Yang, K. Enhanced Resistance of 2205 Cu-Bearing Duplex Stainless Steel Towards Microbiologically Influenced Corrosion by Marine Aerobic Pseudomonas aeruginosa Biofilms. J. Mater. Sci. Technol. 2018, 34, 1325–1336. 42. Liu, H.; Sharma, M.; Wang, J.; Cheng, Y. F.; Liu, H. Microbiologically Influenced Corrosion of 316L Stainless Steel in the Presence of Chlorella vulgaris. Int. Biodeterior. Biodegrad. 2018, 129, 209–216. 43. Liu, H.; Cheng, Y. F. Mechanistic Aspects of Microbially Influenced Corrosion of X52 Pipeline Steel in a Thin Layer of Soil Solution Containing Sulphate-Reducing Bacteria Under Various Gassing Conditions. Corros. Sci. 2018, 133, 178–189. 44. Enning, D.; Venzlaff, H.; Garrelfs, J.; Dinh, H. T.; Meyer, V.; Mayrhofer, K.; Hassel, A. W.; Stratmann, M.; Widdel, F. Marine Sulfate-Reducing Bacteria Cause Serious Corrosion of Iron Under Electroconductive Biogenic Mineral Crust. Environ. Microbiol. 2012, 14, 1772–1787. 45. Li, Y.; Xu, D.; Chen, C.; Li, X.; Jia, R.; Zhang, D.; Sand, W.; Wang, F.; Gu, T. Anaerobic Microbiologically Influenced Corrosion Mechanisms Interpreted Using Bioenergetics and Bioelectrochemistry: A Review. J. Mater. Sci. Technol. 2018, 34, 1713–1718. 46. Ewing, S. P. Electrochemical Studies of the Hydrogen Sulfide Corrosion Mechanism. CORROSION 1955, 11, 51–55. 47. Xiao, J.; Klein, M. I.; Falsetta, M. L.; Lu, B.; Delahunty, C. M.; Yates, J. R.; Heydorn, A.; Koo, H. The Exopolysaccharide Matrix Modulates the Interaction Between 3D Architecture and Virulence of a Mixed-Species Oral Biofilm. PLoS Pathog. 2012, 8, e1002623. 48. Hwang, G.; Liu, Y.; Kim, D.; Sun, V.; Aviles-Reyes, A.; Kajfasz, J. K.; Lemos, J. A.; Koo, H. Simultaneous Spatiotemporal Mapping of In Situ pH and Bacterial Activity Within an Intact 3D Microcolony Structure. Sci. Rep. 2016, 6, 1–11. 49. Beese-Vasbender, P. F.; Nayak, S.; Erbe, A.; Stratmann, M.; Mayrhofer, K. J. J. Electrochemical Characterization of Direct Electron Uptake in Electrical Microbially Influenced Corrosion of Iron by the Lithoautotrophic SRB Desulfopila corrodens Strain IS4. Electrochim. Acta. 2015, 167, 321–329. 50. Xu, D.; Li, Y.; Song, F.; Gu, T. Laboratory Investigation of Microbiologically Influenced Corrosion of C1018 Carbon Steel by Nitrate Reducing Bacterium Bacillus licheniformis. Corros. Sci. 2013, 77, 385–390. 51. Jia, R.; Yang, D.; Xu, J.; Xu, D.; Gu, T. Microbiologically Influenced Corrosion of C1018 Carbon Steel by Nitrate Reducing Pseudomonas aeruginosa Biofilm Under Organic Carbon Starvation. Corros. Sci. 2017, 127, 1–9. 52. Sherar, B. W. A.; Power, I. M.; Keech, P. G.; Mitlin, S.; Southam, G.; Shoesmith, D. W. Characterizing the Effect of Carbon Steel Exposure in Sulfide Containing Solutions to Microbially Induced Corrosion. Corros. Sci. 2011, 53, 955–960. 53. Edwards, M. J.; White, G. F.; Norman, M.; Tome-Fernandez, A.; Ainsworth, E.; Shi, L.; Fredrickson, J. K.; Zachara, J. M.; Butt, J. N.; Richardson, D. J.; Clarke, T. A. Redox Linked


54.

55.

56.

57. 58.

59. 60. 61. 62. 63.

64. 65. 66. 67. 68.

69.

Flavin Sites in Extracellular Decaheme Proteins Involved in Microbe-Mineral Electron Transfer. Sci. Rep. 2015, 5, 11677. Li, H.; Xu, D.; Li, Y.; Feng, H.; Liu, Z.; Li, X.; Gu, T.; Yang, K. Extracellular Electron Transfer Is a Bottleneck in the Microbiologically Influenced Corrosion of C1018 Carbon Steel by the Biofilm of Sulfate-Reducing Bacterium Desulfovibrio vulgaris. PLoS One 2015, 10, e0136183. Zhang, P.; Xu, D.; Li, Y.; Yang, K.; Gu, T. Electron Mediators Accelerate the Microbiologically Influenced Corrosion of 304 Stainless Steel by the Desulfovibrio vulgaris Biofilm. Bioelectrochemistry 2015, 101, 14–21. Jia, R.; Yang, D.; Xu, D.; Gu, T. Electron Transfer Mediators Accelerated the Microbiologically Influence Corrosion Against Carbon Steel by Nitrate Reducing Pseudomonas aeruginosa Biofilm. Bioelectrochemistry 2017, 118, 38–46. Leang, C.; Qian, X.; Mester, T.; Lovley, D. R. Alignment of the C-Type Cytochrome OmcS Along Pili of Geobacter sulfurreducens. Appl. Environ. Microbiol. 2010, 76, 4080–4084. Malvankar, N. S.; Vargas, M.; Nevin, K.; Tremblay, P.-L.; Evans-Lutterodt, K.; Nykypanchuk, D.; Martz, E.; Tuominen, M. T.; Lovley, D. R.; Richard, E.; Brennan, G. Structural Basis for Metallic-Like Conductivity in Microbial Nanowires. MBio 2015, 6, e00084-15. Sure, S.; Ackland, M. L.; Torriero, A. A. J.; Adholeya, A.; Kochar, M. Microbial Nanowires: An Electrifying Tale. Microbiology 2016, 162, 2017–2028. Bridier, A.; Briandet, R.; Thomas, V.; Dubois-Brissonnet, F. Resistance of Bacterial Biofilms to Disinfectants: A Review. Biofouling 2011, 27, 1017–1032. Lewis, K. Multidrug Tolerance of Biofilms and Persister Cells. Curr. Top. Microbiol. Immunol. 2008, 322, 107–131. Savage, V. J.; Chopra, I.; O’Neill, A. J. Staphylococcus aureus Biofilms Promote Horizontal Transfer of Antibiotic Resistance. Antimicrob. Agents Chemother. 2013, 57, 1968–1970. Molin, S.; Tolker-Nielsen, T. Gene Transfer Occurs with Enhanced Efficiency in Biofilms and Induces Enhanced Stabilisation of the Biofilm Structure. Curr. Opin. Biotechnol. 2003, 14, 255–261. Ochman, H.; Lawrence, J. G.; Groisman, E. A. Lateral Gene Transfer and the Nature of Bacterial Innovation. Nature 2000, 405, 299–304. Dzidic, S.; Bedeković, V. Horizontal Gene Transfer-Emerging Multidrug Resistance in Hospital Bacteria. Acta. Pharmacol. Sin. 2003, 24, 519–526. Huddleston, J. R. Horizontal Gene Transfer in the Human Gastrointestinal Tract: Potential Spread of Antibiotic Resistance Genes. Infect. Drug Resist. 2014, 7, 167–176. Poole, K. Mechanisms of Bacterial Biocide and Antibiotic Resistance. J. Appl. Microbiol. 2002, 92, 55S–64S. Brown, D.; Demeter, M.; Turner, R. J. Prevalence of Multidrug Resistance Efflux Pumps (MDREPs) in Environmental Communities. In Microbial Diversity and Infectious Diseases; Das, S., Dash, H. R., Eds.,: Elsevier Inc.: London, 2019; Vol. 1, pp 545–557. Bay, D. C.; Rommens, K. L.; Turner, R. J. Small Multidrug Resistance Proteins: A Multidrug Transporter Family That Continues To Grow. Biochim. Biophys. Acta. – Biomembr. 2008, 1778, 1814–1838.


70. He, G.-X.; Zhang, C.; Crow, R. R.; Thorpe, C.; Chen, H.; Kumar, S.; Tsuchiya, T.; Varela, M. F. SugE, a New Member of the SMR Family of Transporters, Contributes to Antimicrobial Resistance in Enterobacter cloacae. Antimicrob. Agents Chemother. 2011, 55, 3954–3957. 71. Sundheim, G.; Langsrud, S.; Heir, E.; Holck, A. L. Bacterial Resistance to Disinfectants Containing Quaternary Ammonium Compounds. Int. Biodeterior. Biodegrad. 1998, 41, 235–239. 72. van Veen, H. W.; Margolles, A.; Müller, M.; Higgins, C. F.; Konings, W. N. The Homodimeric ATP-Binding Cassette Transporter LmrA Mediates Multidrug Transport by an Alternating Two-Site (Two-Cylinder Engine) Mechanism. EMBO J. 2000, 19, 2503–2514. 73. Zgurskaya, H. I.; Nikaido, H. Multidrug Resistance Mechanisms: Drug Efflux Across Two Membranes. Mol. Microbiol. 2000, 37, 219–225. 74. Wassenaar, T.; Ussery, D.; Nielsen, L.; Ingmer, H. Review and Phylogenetic Analysis of Qac Genes that Reduce Susceptibility to Quaternary Ammonium Compounds in Staphylococcus Species. Eur. J. Microbiol. Immunol. 2015, 5, 44–61. 75. Edgar, R.; Bibi, E. MdfA, an Escherichia coli Multidrug Resistance Protein with an Extraordinarily Broad Spectrum of Drug Recognition. J. Bacteriol. 1997, 179, 2274–2280. 76. Costa, S. S.; Viveiros, M.; Amaral, L.; Couto, I. Multidrug Efflux Pumps in Staphylococcus aureus: an Update. Open Microbiol. J. 2013, 7, 59–71. 77. Hassan, K. A.; Liu, Q.; Elbourne, L. D. H.; Ahmad, I.; Sharples, D.; Naidu, V.; Chan, C. L.; Li, L.; Harborne, S. P. D.; Pokhrel, A.; Postis, V. L. G.; Goldman, A.; Henderson, P. J. F.; Paulsen, I. T. Pacing Across the Membrane: The Novel PACE Family of Efflux Pumps is Widespread in Gram-Negative Pathogens. Res. Microbiol. 2018, 169, 450–454. 78. Ogawa, W.; Minato, Y.; Dodan, H.; Onishi, M.; Tsuchiya, T.; Kuroda, T. Characterization of MATE-Type Multidrug Efflux Pumps from Klebsiella pneumoniae MGH78578. PLoS One 2015, 10, e0121619. 79. Mathieu, L.; Francius, G.; El Zein, R.; Angel, E.; Block, J.-C. Bacterial Repopulation of Drinking Water Pipe Walls After Chlorination. Biofouling 2016, 32, 925–934. 80. Gillis, R. J.; White, K. G.; Choi, K.-H.; Wagner, V. E.; Schweizer, H. P.; Iglewski, B. H. Molecular Basis of Azithromycin-Resistant Pseudomonas aeruginosa Biofilms. Antimicrob. Agents Chemother. 2005, 49, 3858–3867. 81. Kvist, M.; Hancock, V.; Klemm, P. Inactivation of Efflux Pumps Abolishes Bacterial Biofilm Formation. Appl. Environ. Microbiol. 2008, 74, 7376–82. 82. Vikram, A.; Bomberger, J. M.; Bibby, K. J. Efflux as a Glutaraldehyde Resistance Mechanism in Pseudomonas fluorescens and Pseudomonas aeruginosa Biofilms. Antimicrob. Agents Chemother. 2015, 59, 3433–3440. 83. Chapman, J. S. Disinfectant Resistance Mechanisms, Cross-Resistance, and Co-Resistance. Int. Biodeterior. Biodegrad. 2003, 51, 271–276. 84. Simões, M.; Simões, L. C.; Vieira, M. J. Species Association Increases Biofilm Resistance to Chemical and Mechanical Treatments. Water Res. 2009, 43, 229–237. 85. Simões, L. C.; Simões, M.; Vieira, M. J. Influence of the Diversity of Bacterial Isolates from Drinking Water on Resistance of Biofilms to Disinfection. Appl. Environ. Microbiol. 2010, 76, 6673–6679.


86. van der Veen, S.; Abee, T. Mixed Species Biofilms of Listeria monocytogenes and Lactobacillus plantarum Show Enhanced Resistance to Benzalkonium Chloride and Peracetic Acid. Int. J. Food Microbiol. 2011, 144, 421–431. 87. Jennings, M. C.; Forman, M. E.; Duggan, S. M.; Minbiole, K. P. C.; Wuest, W. M. Efflux Pumps Might Not Be the Major Drivers of QAC Resistance in Methicillin-Resistant Staphylococcus aureus. ChemBioChem. 2017, 18, 1573–1577. 88. Downs, H. Selection, Application, and Evaluation of Biocides in the Oil and Gas Industry; NACE Publication 31205-2006-SG; NACE International: Houston, TX, 2006. 89. Gerba, C. P. Quaternary Ammonium Biocides: Efficacy in Application. Appl. Environ. Microbiol. 2015, 81, 464–469. 90. Bester, E.; Wolfaardt, G.; Joubert, L.; Garny, K.; Saftic, S. Planktonic-Cell Yield of a Pseudomonad Biofilm. Appl. Environ. Microbiol. 2005, 71, 7792–7798. 91. Kharazmi, A.; Giwercman, B.; Høiby, N. Robbins Device in Biofilm Research. Methods Enzymol. 1999, 310, 207–215.


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