Archive for the ‘Fuel Microbiology’ Category


FUEL & FUEL SYSTEM MICROBIOLOGY PART 24 – PETROLEUM EQUIPMENT INSTITUTE’S 2018 CONVENTION

The Petroleum Equipment Institute (PEI) held its 2018 convention at the Las Vegas Convention Center from 07 to 10 October 2018. As usual, the PEI convention was held in conjunction with the much larger National Association of Convenience Store (NACS) convention. Today, I’ll focus on a few items that are particularly relevant to fuel and fuel system microbiology. I’m not going to attempt to provide anything approaching an overview of the entire convention. Instead I’ll report and discuss a few statements I heard from speakers during PEI’s Tuesday 09 October education sessions.

Regulatory issues

EPA Regulatory Update – Carolyn Hoskinson, Director of EPA’s Office of Underground Storage Tanks (OUST) and several members of her staff spoke to the current state of affairs regarding UST regulations. Tony Raia reported that with the 13 October 2018 compliance deadline looming, 32 states had updated their UST regulations to harmonize them with the 2015 updated US EPA regulations. Tony identified five state categories:
1. State Program Approval (SPA) States that have completed their updates and which are now in full compliance
2. Non-SPA States that have completed their updates to comply with the 2015 regulations
3. SPA States that have delayed revising their state regulations
4. SPA States that have updates in progress
5. Non-SPA States that have not yet updated their regulations per the US EPA 2015 regulations.
Bottom line is that we are entering a period during which there will be some confusion over compliance.

U.S. EPA UST Enforcement – Mark Barolo – the US EPA OUST official responsible for enforcement – noted that in nearly all cases, individual States were responsible for enforcement. Recognizing the confusion, Mark opined that inspectors were going to address violations on a case-by-case basis. Generally speaking, retailers who had been incompliance, had the required documentation, and demonstrated that they were making good-faith efforts to ensure that they remained in compliance, would experience less enforcement grief than those who have not. Mark’s colleague, Cho Yi Risher noted that the regulations do not prescribe the time permitted for site owners to repair or replace non-compliant equipment. Moreover, the inspections required by the 2015 regulations identify non-compliant equipment. There is no incentive for owners to institute predictive maintenance programs (see my 16 January 2017 post) that would detect failure trends before equipment became non-compliant.

Failure to detect uncontrolled microbial contamination and biodeterioration before they cause valves to seize or tanks and lines to leak is a false economy. A few pennies saved during regulation-mandated inspections can lead to remediation expenses in excess of $0.5 million.
Although for some of us, it’s hard to believe that the UST installed in 1987 – in compliance with the original UST regulations – are now beyond their 30-year warranty life. Discussion during the session’s question and answer period indicated that all stakeholders shared a common interest in ensuring that sites with tanks that were more than 30-years old would be able to continue to operate. I anticipate seeing articles to this issue in PEI Journal in the coming months.

Fuel Quality and Corrosion
Scott Boorse – PEI’s Technical Program Manager; recently retired from a major fuel retailer – made several observations that validated much of what I’ve been discussing in Fuel Microbiology What’s New posts. He suggested that in his experience, 100 % of all retail site fuel systems had some corrosion. He attributed much of this corrosion to the bacterial genus Acetobacter converting ethanol to acetic acid. I am convinced that most of the headspace and spill containment well acid production comes form chemical oxidation of ethanol to acetic acid. Microbes are involved in an estimated 50 % of all system corrosion issues, but – as I’ve written previously – microbes produce a variety of organic acids. These acids can react with chloride, sulfate, and nitrate in fuel-associated water to form organic bases (or salts) and strong, highly corrosive, inorganic acids – hydrochloric, sulfuric, and nitric acids, respectively. Still, Scott was on spot suggesting that UST system corrosion was much more wide-spread than most stakeholders realize.

Rebbeca Moore – GM and chair of the automotive industry’s Top Tier Detergent Gasoline and Diesel Fuel consortia – discussed the importance of fuel quality on engine performance. Top Tier is an auto industry sponsored compliance program intended to go beyond ASTM product specifications typically cited in state regulations. I’ll steer clear of the perennial debates between engine manufactures and petroleum producers that enliven our semi-annual ASTM D02 (Petroleum Products) subcommittee A (Gasoline and Oxygenated Fuels) and E (Burner, Diesel, Non-Aviation Gas Turbine, and Marine Fuels), but Rebbeca made an important point. ASTM specifications are often misused. They are meant to indicate whether a product (i.e., fuel) is fit for use at a single point and place in time (i.e., when and where the sample was collected). The petroleum industry’s infrastructure is vast and complex. Moreover, product ages (if it didn’t it wouldn’t combust so well in engines). Specification tests provide little information about how the product will age during storage.
Rebbeca illustrated the dilemma by listing the typical components of 7,500 gal of in-specification ULSD delivered to UST:
• 1 cup of dirt
• 1 to 2 gallons of water
• Up to 325 gallons of FAME (B5 ULSD is now included in ASTM Specification D975 Diesel Fuel Oils)
• 1 gallon of glycerin
• 5 to 40 gallons of additives
At sites that receive frequent deliveries, these trace amounts of dirt and water add up! Not surprisingly, along with measures that are outside the retail site or fleet owner’s control, Rebecca recommended more aggressive water removal and better dispenser filtration (there is an ongoing debate among stakeholders with some recommending that all dispensers have 5.0 µm, water absorbing filters and others arguing for 100 µm particulate filters). I share Rebecca’s view that all dispensers should have 5.0 µm, water absorbing filters. Marketers who are focused only on low rates and not product quality have argued for eliminating the filtration requirement completely.

Ryan Haerer – of US EPA’s OUST (https://www.epa.gov/ust) – wrapped up the session, sharing a few notable points. First, Ryan reminded attendees that UST regulations apply only to system components that are in contact with the soil. The US EPA does not regulate the condition of internal components that are not in direct contact with the soil (for example, submerged turbine pumps – STP – and their associated hardware). He also explained that under the UST regulations, there is a requirement that system components be compatible with the substance stored. This is likely to become interesting as new products (for example, E15 gasoline or substitution of ethanol with isobutyl alcohol) are introduced into the commercial fuel infrastructure – here I’m using interesting – in the same way it is used in of the phrase: “May you live in interesting times.” (Austen Chamberlain – British Foreign Secretary, 1924 to 1929 wrote that he had been told that this was an ancient Chinese curse, but his claim has never been verified).

Bringing it home

At this point we are enjoying an interesting paradox. Regulators, insurers, and an increasing number of retailers recognize that waiting until fuel systems fail is a problem. However, the system largely provides incentives for site owners to wait until failures have occurred. After failure, insurance covers component replacement costs. In many states, superfund monies cover remediation costs. When site owners invest in predictive maintenance, they only see the costs. Although there are benefits – not the least of which is customer satisfaction and a positive corporate image – they are intangible. How do we break the paradox?

Please share your thoughts on this issue with me at fredp@biodeterioration-control.com. I’ll compile comments and post them anonymously as a future What’s New column.

FUEL & FUEL SYSTEM MICROBIOLOGY PART 21 – PREVENTING MICROBIAL DAMAGE TO FUEL SYSTEMS PART 3; TREATING SYSETMS WITH MICROBICIDES

Fig 1. Microbes in fuel systems and the biocides used to control them – a) fuel (yellow-orange) over bottoms-water (dark blue), with red lines showing where microbes tend to accumulate; b) after treatment with water-soluble biocide (purple stars); c) after treatment with fuel-soluble biocide; d) after treatment with universally-soluble biocide.


Where are the bugs?If you intend to use a biocide to disinfect a fuel system, the first question to ask is: “Where are the bugs?”  Figure 1a shows where microbes tend to be most abundant in fuel systems. The red lines symbolize microbe accumulations in the bottoms sludge and sediment, at the fuel-water interface, and on tank walls.Fuel treatment microbicidesMicrobicides are chemicals that are manufactured and sold for the purpose of killing microbes. They are part of a greater family of chemicals called pesticides or biocides. Formally, microbicides are called antimicrobial pesticides – i.e., they are pesticides that target microbes. I’ll come back to the issue of pesticide registration at the end of the post. For now, I want to split the biocides used for fuel and fuel system treatment into three groups:
  • 1. Water-soluble
  • 2. Fuel-soluble
  • 3. Universally-soluble
Why should I care whether my biocide is water, fuel or universally soluble? Water-soluble biocidesWater soluble biocides fall through the product and dissolve into the water-phase (figure 1b). They are not present in the fuel for a long enough period (see Fuel and Fuel System Microbiology Part 22) to kill microbes either in the fuel-phase or on tank walls exposed to fuel. They can effectively kill microbes in bottoms-water, sludge and sediment, but it is reasonable to ask whether this makes good sense.Typically, when water, sludge, and sediment are vacuumed or drained out of tanks, the wastes are shipped to a biological wastewater treatment plant. Biological wastewater treatment depends on microbes to eat organic molecules to reduce the water’s biochemical oxygen demand (percentage of organic matter that microbes can digest in a five-day period), chemical oxygen demand (percentage of organic matter that is chemically oxidizable – convertible to carbon dioxide), and total petroleum hydrocarbons (TPH).I have never understood the logic for killing microbes that can help the waste treatment process, just before shipping those microbes to waste treatment. Therefore, I have never understood the logic of treating fuels or fuel systems with water soluble biocides.Fuel-soluble biocidesFuel-soluble biocides (figure 1c) mirror the performance of their water-soluble cousins. These products effectively kill microbes in the fuel and can be somewhat effective against microbes growing on tank walls in contact with the fuel. They can also attack microbes living at the fuel-water interface. They do not contact microbes living either on tank bottoms or on those portions of the tank wall that are in contact with water rather than fuel.Universally-soluble biocidesAs their name implies, universally-soluble biocides (figure 1d) can disperse within both the fuel and water phases. Typically, they are fully soluble in fuel and partially soluble in water. Most importantly, they are chemically stable in both phases. As figure 1d illustrates, they can interact with microbes in fuel, in water, at the fuel-water interface, and on all tank surfaces in contact with fuel or water. Consequently, universally-soluble biocides are the most reliable products for disinfecting fuels and fuel systems.RegulationsPesticides are regulated by the U.S. EPA in the USA. Under the U.S. Federal Insecticide, Fungicide and Rodenticide Act (FIFRA – 7 U.S.C. §136 et seq. [1996]), EPA’s Office of Pesticide Programs had direct responsibility for pesticide registration and management. The details of biocide regulations are found in 40 CFR Chapter I, Subchapter E, Parts 152-180.Outside the U.S., the European Union and many individual countries have regulatory agencies responsible for pesticide approval and oversight. The key point here is that microbicides are highly regulated products. Only registered products may be used. Each registered product has one or more approved end-uses (sites). Two US EPA end-uses sites of interest to us are:
  • • For use in treating fuel-associated water, and
  • • For use in fuels.
The language can vary among product labels, but the difference between these two general sites is important. The first site applies to water-soluble biocides. The second one refers to fuel-soluble and universally-soluble biocides. If you are considering a microbicide, read the label carefully and make certain that the product has a use in fuels end-use site.There is another fuel-related regulation: 49 CFR Chapter I, Subchapter C, Part 79 Registration of Fuel and Fuel Additives. The regulations under 49 CFR 79 address the use of fuels and fuel additives. Fuels are comprised of molecules built from carbon (C), hydrogen (H), oxygen (O), nitrogen (N), and sulfur (S). This list of elements has the acronym, CHONS. Fuel additives that contain only CHONS are designated as being substantially similar to fuel (subsim). The American Petroleum Institute (API) has created a consortium of companies who produce subsim products. Once a suitable test method has been developed, the members of the consortium will share the cost of engine emission toxicological tests. Each member’s share will be based on the volume of CHONS product they produce. Recognizing the infinitesimally small volume of fuel treatment microbicides, relative to fuels, API charges microbicide manufacturers a minimal fee for consortium membership.The bottom-line here is that fuel treatment microbicides should have two registrations:
  • 1. A pesticide registration, and
  • 2. A fuel additive registration.
None of the water-treatment microbicides that list a variation on the theme of “for use in treating fuel-associated water” are also registered as fuel additives. Most of the products that list “fuel treatment” as an end-use site, carry both registrations. Just to help confuse users, there are several products that have waivers from the U.S. EPA’s Fuel Programs Manager. These waivers are based on the assumption that the microbicides are used to treat fuel systems rather than fuels, and that none of the product remains dispersed or dissolved in fuel.In part 22, I’ll write about how to use universally-soluble fuel treatment microbicides. In the meantime, if you have questions or comments about today’s post, please contact me at fredp@biodeterioration-control.com.
DisclaimerAs in my previous two post, I’ll open with a disclaimer. Microbes are ubiquitous. There are extraordinarily few habitats on earth where thriving, microbial communities have not been detected. In practical terms, this means that it is unlikely that operators will ever have a completely sterile fuel system or that they will reduce their fuel system biodeterioration risk to zero. Biodeterioration can still occur in the best maintained fuel systems. However, the risk of it occurring in an inadequately maintained system is much more likely.

FUEL & FUEL SYSTEM MICROBIOLOGY PART 20 – PREVENTING MICROBIAL DAMAGE TO FUEL SYSTEMS PART 3; WATER BITS


Fig 1. From Rime of the Ancient Mariner, Samuel Taylor Coleridge, 1798

Water, water everywhere…

Samuel Coleridge’s infamous mariner paid dearly for having killed an albatross (figure 1). Do fuel quality managers and personnel responsible for fuel system integrity pay dearly for underestimating the ability of small (<1 oz; 30 mL) pools of fuel-associated-water left behind after water has been nominally purged from a fuel tank? A water bit is any small volume of water that remains in a fuel tank after dewatering (my personal, technical definition).
In Part 20, I wrote: “No water means no bugs. Is it as easy as all that?” I also explained why the short answer to the question was: “No.” For emphasis, I’ll again share figure 3 from Part 20 (figure 2, here):


Fig 2. Scale: how 2 mm of water appears to a bacterial cell.

A – a 6’6” tall man standing at the base of Mt. Kilimanjaro; B- a bacterial cell “standing” in a pool of water that is >2 mm deep; the ratios between the height of Mt. Kilimanjaro and the man in A, and between the depth of the pool of water and the bacterial cell in B are the approximately the same.

What can we do about these traces of water?
I confess that I am not a big fan of dispersants. When water dispersants are used routinely as fuel additives, the dispersed water can act as a corrosive agent; damaging engine components. However, when used to complete the job started by draining or vacuuming most of the free-water out of a tank, dispersants can be quite effective.
Figure 3 illustrates how dispersants work. Most dispersants are organic molecules that have a polar (charged; water-soluble) head and a non-polar (non-charged; fuel-soluble) tail. When added to fuel over water (figure 3b), they move towards anywhere where fuel contacts water (figure 3c) and trap tiny (typically <1 µm; 0.0004 in dia) fuel droplets. The fuel “sees” only the dispersant’s non-polar tails, so the droplets disperse uniformly throughout the fuel (figure 3d). The dispersed droplets (micelles) get transported with the fuel and evaporate during combustion in the engine cylinder.



Fig 3. Dispersant action: a) fuel over bottoms-water; b) dispersant added to fuel – inset shows dispersant molecule with polar head and non-polar tail; c) dispersant heads and tails align in water an fuel phases, respectively; d) dispersants form micelles with water droplet trapped in center; typical droplet size is < 1 μm dia.
The use of dispersants is controversial. Dispersant manufacturers and marketers focus on dispersant effectiveness in keeping free-water from accumulating in fuel systems. Moreover, under most circumstances, microbes are unlikely to make use of water trapped within dispersant micelles. Conversely, engine manufacturers focus on the potential for dispersed water to corrode and erode injector nozzles; particularly on modern, high-pressure, common-rail diesel engines. Interestingly, there is at least one additive manufacturer that has tried to promote water-emulsion diesel fuels – diesel with a dispersant that enables the fuel to hold as much as 25 % water. As a microbiologist, I was looking forward to investigating microbial contamination problems in systems that handled 25-75 water-in-diesel blends. But that’s another discussion.
In my next blog, I’ll focus on fuel treatment biocides. In the meantime, if you have questions or comments about today’s post, please contact me at fredp@biodeterioration-control.com.

Disclaimer:
Microbes are ubiquitous. There are extraordinarily few habitats on earth where thriving, microbial communities have not been detected. In practical terms, this means that it is unlikely that operators will ever have a completely sterile fuel system or that they will reduce their fuel system biodeterioration risk to zero. Biodeterioration can still occur in the best maintained fuel systems. However, the risk of it occurring in an inadequately maintained system is much more likely.

FUEL & FUEL SYSTEM MICROBIOLOGY PART 19 – PREVENTING MICROBIAL DAMAGE TO FUEL SYSTEMS PART 2; WATER

Disclaimer:

As in my previous post, I’ll open this post with a disclaimer. Microbes are ubiquitous. There are extraordinarily few habitats on earth where thriving, microbial communities have not been detected. In practical terms, this means that it is unlikely that operators will ever have a completely sterile fuel system or that they will reduce their fuel system biodeterioration risk to zero. Biodeterioration can still occur in the best maintained fuel systems. However, the risk of it occurring in an inadequately maintained system is much more likely.

No water means no bugs. Is it as easy as all that?

Best practice for keeping microbes from growing in fuel systems is to keep them dry. You’ll find this guidance in just about every consensus guidance document (for example ASTM D6469) and peer reviewed paper on the subject; including the ones I’ve written. Of course, there is a catch. In today’s post, I’ll write about why keeping fuel systems water-free is much easier said than done.

Water, water everywhere…

Water in fuel systems is typically present in three forms. Dissolved water is made up of individual water molecules and water droplets that are smaller than 1 m diameter (dia; 1 m = 0.00004 in) in fuel. Fuel containing only dissolve water appears to be transparent – no visible haze. The concentration of dissolved water that a fuel product can hold depends on temperature and product chemistry. Water solubility (also called the product’s water tolerance) increases with product temperature. For example, water solubility in E-10 gasoline increases from 0.2 % (by vol.) to 0.5 % as the product temperature increases from 0 C (32 F) to 20 C (68 F). For products typically sold at retail sites, water solubility is greatest in B-5 biodiesel. Water is least soluble in conventional gasoline.

A – Clear and bright gasoline (ASTM D4176 haze rating: 1; B – gasoline with ASTM D4176 haze rating: 6; C – gasoline over bottoms-water; fuel’s ASTM D4176 haze rating: 3.

Once water concentration reaches its saturation limit, individual molecules join together to form dispersed water. Dispersed water droplet diameters range from 1 m dia to  10 m dia. Fuel haze (see ASTM D4176) increases with the number and size of dispersed water droplets. Droplets that are 10 m are heavy enough to settle out of the product and coalesce to form free water. Free-water can accumulate on tank walls, but is most commonly seen as bottoms water. Figure 1 shows three 87 octane gasoline fuel samples: figure 1A has < 0.1 % water (by vol.), figure 1B has an ASTM D4176 haze rating of 6 (i.e., with dispersed water), and figure 1C shows haze rating 3 fuel over free water.

Ethanol-blended gasoline (E-10)

There was a time when competent fuel chemists argued that phase-separation (i.e.: accumulation of bottoms-water) would never happen in E-10 tanks. They believed that because water was more soluble in E-10 than in E-0, there would never be sufficient water accumulation to drive the fuel-water split. The only problem is that they were wrong. Others argued that even if phase-separation did occur, the water-phase would be approximately 60% ethanol. Everyone (except microbes) knows that 60 % ethanol is a good disinfectant. Microbes couldn’t possibly grow in bottoms-water under E-10; except that they do. Most commonly, I do not detect active microbes in E-10 bottoms-water. On occasion, I do. Others have also reported detecting microbes in E-10 bottoms-water. We are not sure what’s going on (i.e., why the bugs are killed by the ethanol), but it is no doubt interesting. Bottom-line: do not assume that underground storage tanks (USTs) containing E-10 do not have bottoms-water or that bottoms-water under E-10 is microbe-free.

Implications

Assume that most UST have some free water. Here’s why:

  • As explained above, depending on the fuel grade, good-quality quality (i.e., fit-for-use) product can as much as 0.5 % dissolved water.
  • It is reasonable to assume that in a fuel carrying 0.01 % (i.e., 100 ppm) water, approximately 10 % of that water will separate from solution while product is in a UST. That 10 % of 100 ppm translates to 10 gal per million gal (a retail site that receives a 7,000-gal delivery every two to three days receives approximately 1 million gallons per year).
  • Based on the previous bullet, approximately 10 gal of water will accumulate, if no other water enters the tank. Figure 2 illustrates this scenario. It shows three-years’ water accumulation in a 10,000 gal (8 ft diameter x 27 ft long underground storage tank; 10 gal = 0.3 in to 0.5 in water).

Figure 2. Underground storage tank (UST); 10,000 gal capacity, with  1 in (30 gal) water.

End and side view of UST that is approximately half filled with product. 1 in of water is approximately 30 gal; barely visible in this schematic. If water does not enter UST by any other means, it can take three years for this volume to accumulate.

End and side view of UST that is approximately half filled with product. 1 in of water  30 gal; barely visible in this schematic. If water does not enter UST by any other means, it can take three years for this volume to accumulate.

Using water-paste on a sounding stick, it should be easy to detect bottoms-water long before 1 in of water has accumulated. In fact, many companies specify that UST should be dewatered whenever 0.5 in or (10 gal to 15 gal) more water is detected. This guidance is based on two key assumptions:

  • 1. USTs rest at the same angle at which they were installed: 1 in per 10 ft grade; with fill-line at low end (figure 3A).
  • 2. UST floors (longitudinal, bottom dead-centerline) are straight (i.e. UST bottoms are flat). In the words of the song from Porgy and Bess: “taint necessarily so.”

There is a problem with these assumptions. USTs are installed on top of backfill. Despite best efforts to fully compact backfill before placing a UST, the tank’s weight – particularly after it has been filled – will cause additional backfill compaction (i.e., settling).

As illustrated in figures 3B and 3C, UST bottoms can settle flat or with the end opposite the fill-line (often the end with the submerged turbine pump – STP) lower than the fill-end. Tanks can also sag (lower in the center than at either end; figure 3D) hog (lower at both ends than in the center; figure 3E), or have numerous peaks and valleys along the longitudinal bottom centerline.

Figure 3. How USTs settle.

A – Typical, planned configuration: UST is lower at fill-end than at turbine end; B- UST is installed to lie flat; C – UST has settled so that turbine end is lower than fill end; D – UST has settled so that center is lower than ends; E – UST has settled so that ends are lower than center. All angles are exaggerated to illustrate settling issues.

Typically, the distance between peaks and valleys are measured in mm (1 mm = 0.04 in), so they are imperceptible to the naked eye. A 1 in per 10 ft incline is impossible to detect without using a bubble level. However, water will flow to low point(s). Long before free-water is detected, a UST is likely to have numerous, small pools of bottoms-water. Each is a great habitat for fuel-system microbes.

Can pools of 1 ounce (30 mL) water be habitats for microbes?

As illustrated in figure 4, to a bacterium a 2 mm deep pool of water (30 mL; 1 oz.) is like an large lake. In figure 4A, a 6 ft 6 in (2 m) tall man is standing at the foot of Mt. Kilimanjaro (19,700 ft – 6,000 m – tall). Figure 4B shows a bacterial cell (0.5 mm dia x 2 m long) “standing” at the bottom of a pool of water that is 2 mm (0.08 in) deep by 6 mm (0.24 in) wide. The relative height of Mt. Kilimanjaro’s peak over the man in figure 4A and the height of the pool of water over the bacterium in figure 4B is the same. In other words, from the perspective of microbes, traces of water that are undetectable to fuel system operators can be like large lakes to microbes; providing mini-habitats for millions of cells.

Figure 4. Scale: how 2 mm of water appears to a bacterial cell.

A – a 6’6” tall man standing at the base of Mt. Kilimanjaro; B- a bacterial cell “standing” in a pool of water that is approximately 2 mm deep; the ratios between the height of Mt. Kilimanjaro and the man in A, and between the depth of the pool of water and the bacterial cell in B are the approximately the same.

Summary:

In summary, water can be present in one or more of three forms in fuel systems:

  • Dissolved
  • Dispersed
  • Free

It is much easier to prescribe keeping tanks water-free than it is to actually eliminate all water.

In my next blog, I’ll focus on options for minimizing water accumulation in fuel systems. In the meantime, if you have questions or comments about today’s post, please contact me at fredp@biodeterioration-control.com

FUEL & FUEL SYSTEM MICROBIOLOGY PART 18 – PREVENTING MICROBIAL DAMAGE TO FUEL SYSTEMS PART 1

Disclaimer:

I’ll open this post with a disclaimer. Microbes are ubiquitous. There are extraordinarily few habitats on earth where thriving, microbial communities have not been detected. In practical terms, this means that it is unlikely that operators will ever have a completely sterile fuel system or that they will reduce their fuel system biodeterioration risk to zero. Biodeterioration can still occur in the best maintained fuel systems. However, the risk of it occurring in an inadequately maintained system is much more likely.

Biodeterioration:

I’ll also take this opportunity to remind readers that biodeterioration (damage caused by organisms) and bioremediation (using microbes or other organisms to degrade or remove toxic or noxious chemicals) are the flip-sides of the biodegradation coin (figure 1).

Figure 1. Just as the two sides of a coin are its obverse (front) and reverse (back) sides, the two sides of biodegradation are bioremediation and biodeterioration.

Microbes degrade fuel quality and fuel system components. In high-turnover retail systems, product deterioration is unlikely. I consider any tank that is refilled at least weekly to be a high-turnover (or high-throughput) system. The time that product spends in the storage tank is too short for degradation to occur. Studies that investigate the rate at which microbes change fuel chemistry, typically show substantial changes after a month or longer. Consequently, fuel in tanks used for emergency generators, or seasonally operated equipment is at greater biodeterioration risk than fuel in retail underground storage tanks (UST), or frequently operated vehicles. That’s all I want to say about fuel biodeterioration for now. I’ll return to the topic in a future blog post.

For now, I’ll focus on fuel system biodeterioration. Most of the damage caused to fuel system components is caused by biofilm communities (see post #16 https://biodeterioration-control.com/2017/11/). Microbes cause damage either directly or indirectly (more on this in a future post). The most obvious indications of biodeterioration are filter plugging and corrosion. Although it’s typically the first indication of a biodeterioration problem, filter plugging is a late symptom. I often compare it to a heart attack; a late – but often first recognized – symptom of coronary disease.

So how do we substantially reduce fuel system biodeterioration risk? Step one is cost-effective condition monitoring (CM). Step two is cost-effective predictive maintenance (PdM; see https://biodeterioration-control.com/microbial-damage-fuel-systems-hard-detect-part-5-predictive-maintenance-pdm/). The former drives the latter. Why do I emphasize cost-effectiveness? As I see it, there is as little justification for investing $10,000 per year to detect problems that might cause $1,000 per year of damage, as there is in refraining from spending $10,000 per year to detect problems that could cost $100,000 per year. I’m not suggesting that any fuel system CM program should cost $10,000 per year. An effective program can cost less than $2,000 per year. My point here is that before setting up a CM/PdM program, stakeholders should invest a bit of time and effort to determine their actual annual biodeterioration-related costs.

Opportunity Cost:

Nearly two decades ago, I first argued that a 10% flow-rate loss at high-traffic, retail sites, can easily translate to more than $100,000 per dispenser per year opportunity cost (Passman, F.J., 1999. “Microbes and Fuel Retailing: The Hidden Costs of Quality.” Nat. Petrol. News 91 [7]: pp: 20-23). My model did not include lost C-store revenues related to customer discontent with their fueling experience. Retailers who have tested my model have invariably been shocked by the huge impact of seemingly minor flow-rate reductions on fuel sales volumes. I’m still trying to understand the psychology behind retailers’ general reluctance to even test my model (for model details, contact me at fredp@biodeterioration-control.com). Bottom line: the return on investment (ROI) for well-designed and executed, CM can easily be >$1,000 return on each $1 invested.

Condition Monitoring:

In Parts 2 through 18 of this series, I’ve written about the details of condition monitoring. I won’t repeat that information here. Instead, I’ll offer a few basic guidelines:

1. Testing hierarchy – CM plans should include two or more tiers. Tests that are easiest and least expensive to perform should be done most frequently (checking UST for bottoms-water accumulation and dispenser flow-rate checks are great examples of Tier 1 tests). Tier 2 tests include bottom-sample visual inspection (optimally, samples should be collected from the fill, automatic tank gauge, and submerged turbine ports). A simple microbiological test (for example ASTM D7687) is indicated whenever the bottom-sample is turbid or when it includes water. When Tier 2 tests indicate that the biodeterioration risk is moderate to high, Tier 3 tests (generally performed by a qualified laboratory) are used to confirm the risk.

2. Test method selection – notwithstanding the examples I mentioned under Testing hierarchy each site owner should develop a CM plan that best meets their needs. You can read my test specific blog posts (or ASTM D6469) for discussions of the benefits and limitations of each test method.

3. Testing frequency – my rule of thumb, after you have determined how often a test parameter is likely to indicate an increased biodeterioration risk, divide that period in three. That gives you the optimal test interval. Testing more often typically translates into greater costs without any real ROI. Testing less frequently increases the risk of having to perform corrective – rather than preventive – maintenance actions.

4. Understanding trends – each of the three previous guidelines depends on a basic understanding of trends. For example, even fuel with an ISO 4406 cleanliness rating of 18/16/13 (see https://www.iso.org/standard/72618.html), will eventually plug dispenser filters. Consequently, dispenser flow-rates will invariably decrease. How many operators know what normal looks like? How many have a control limit? Typically, dispenser filters can process >250,000 gal of fuel before the flow rate will fall below 7 gpm. Replacing fuel filters when the flow-rate is <7 gpm strikes a balance between the opportunity maintenance costs. Knowing whether the flow rate has fallen to <7 gpm after 50,000 gal or 500,000 gal of fuel have been filtered serves to trigger additional sampling and testing. If the amount of fuel filtered before substantial flow reduction occurs is much less than expected, then additional testing is indicated.

Summary:

In summary, microbial contamination control depends on a good CM program that is linked to a good PdM program. A reasonable investment in CM and PdM should be a fraction of the likely cost impact of not having those programs in place. A cost-effective CM program is driven by an understanding of system trends, definition of the methods that will provide the most useful, actionable information; selection of which tests to run; and determination of sampling and testing frequency. In my next blog, I’ll focus on preventive measures. In the meantime, if you have questions or comments about today’s post, please contact me at fredp@biodeterioration-control.com.

 

FUEL & FUEL SYSTEM MICROBIOLOGY PART 17 –TEST METHODS – GENETIC TESTING

In today’s blog, I’ll cover the lastest family of microbiology methods used for testing fuels & fuel associated water. These methods fall under the category genomics – the study of genes. Warning: genetic testing is more technically complex than the methods I’ve described in recent posts. I’ll do my best to keep the language as simple as possible.

Genetic methods have evolved substantially over the past 30 years. They all depend on the polymerase chain reaction (PCR); first reported in 1983. The common steps of all PCR methods include:

   1) Genetic material (deoxyribonucleic acid – DNA) extraction (fig 1).
   2) Heating to separation of the two strands of DNA’s double helix (fig 2) into two single-strands.
   3) Cooling and using an enzyme – polymerase – to convert each single strand back into double stranded DNA (fig 3).
   4) Repeating steps 2 and 3 until there are millions of copies of each of the DNA originally extracted in step 1.
   5) Using analytical tools to: identify the different types of DNA that were extracted from the original sample in step 1.

Fig 1. Bacterial cell lysing and ejecting its cytoplasm

Fig 2. a) DNA molecule ; b) section of DNA being denatured to two single strands.

Fig 3. Denatured DNA (left) coupled with primer and reacted with DNA polymerase to form two new double helices.

When PCR methods were first developed in the mid-1980s, DNA was extracted from colonies that had developed on nutrient agar plates (see Part 12 and fig 4). Early PCR testing revolutionized microbial taxonomy. Microbes that seemed to be closely related because of their appearance and nutrient preferences turned out to be quite distant genetic relatives. Conversely, some microbes that had historically been classified as being members of different groups, were discovered to be nearly identical genetically. However, PCR could only be used to identify microbes that formed colonies.

Fig 4. Bacterial colonies on nutrient agar.

In the 1990s quantitative-PCR (qPCR) methods were developed. In qPCR, messenger-RNA (mRNA) is extracted and used to synthesize complimentary-DNA (cDNA). The PCR process then continues as described in steps 2 through 5. The RNA used for qPCR is typically tagged with a fluorescent dye. A fluorometer is used to measure the DNA concentration as a function of time during repeated cycles of heating and annealing (steps 2 and 3). As shown in fig 5, the time required for the fluorescence to reach a threshold value, can be used to compute the amount of mRNA that was originally extracted from the sample. This, in turn provides an accurate estimate of the population density (i.e., cells/mL) in the sample.

Fig 5. DNA amplification curves: delta Rn is the amount of fluorescence detected and ct is the threshold delta Rn used to compute the DNA concentration in the original sample. The five curves show that the number of PCR cycles needed to reach ct increases as the original DNA concentration decreases.

Four molecules (nucleotides) make up the genetic code (adenosine – A, cytosine – C, guanine – G, and thiamine – T). Each three-nucleotide sequence is the code for a specific amino acid. Thus, long strings of three-letter messages specify the amino acid sequence of enzymes – the cell’s machinery for carrying out all of life’s processes. The total genome of each type of cell (operational taxonomic unit – OTU) is unique. Because each of the four nucleotides – A, C, G, and T – has a unique electrical charge, each OTU’s DNA has a unique net electrical charge. Using a technique called gel electrophoresis, after amplification (step 5) analysts can separate and isolate each type of DNA that was recovered from the original sample (fig 6). They can then sequence the genes and attempt to match the sample’s DNA against a DNA library. The result is a taxonomic profile of the microbes that were present in the original sample.

Fig 6. DNA profiling using gel electrophoresis: a) schematic illustration of process ; b) photograph of gel.

In earlier posts, I’ve referred to Donald Rumsfeld’s “unknown unknowns.” Although qPCR and, the more recently variation called next generation sequencing – NGS, is a powerful tool for studying microbial communities in fuel systems, it is probably not the last word in microbiology testing. True, qPCR detects many types of microbes that are undetectable by historically used culture methods. However, extracting DNA or RNA from cells is as much art as science. Genetic material that isn’t extracted isn’t detected. Additionally, qPCR testing depends on the use of primers – short sections of mRNA selected to either be universal (i.e., include a section of A, C, G, T basis that are believed to be present in all bacteria) or specific (i.e., include a section of genetic coding that is unique to a microbe of specific interest). Consequently, researchers are on a steep learning curve about how to select primers. As task force within ASTM D02.14 has just restarted work on a qPCR standard test method for fuels and fuel associated water. The last attempt stalled when participants could not agree on a consensus DNA extraction protocol. As the new task force makes progress I provide readers with updates. The goal is to develop a method that non-technical folks will be able to use.

In the meantime, please contact me at fredp@biodeterioration-control.com you’d like to learn more about fuel system microbiology or microbiological contamination control.

Footnotes:

Source: http://www.newswise.com/images/uploads/2013/01/9/lysis_cover.jpg.
Source: http://www-nmr.cabm.rutgers.edu/photogallery/proteins/gif/dna.gif.
Source: https://laboratoryinfo.com/wp-content/uploads/2015/07/Polymerase_chain_reaction.svg_.png.
Source: https://laboratoryinfo.com/wp-content/uploads/2015/07/Polymerase_chain_reaction.svg_.png.
Source: https://www.microbiologyinpictures.com/bacteria-photos/escherichia-coli-photos/e.-coli-staphylococcu-aureus-colonie.jpg.
Source: https://media.nature.com/full/nature-assets/leu/journal/v17/n6/images/2402922f5.jpg.
Source: http://science.halleyhosting.com/sci/ibbio/biotech/pics/electrophoresisnotes.gif.
Source: https://78.media.tumblr.com/tumblr_lwthmyKO441qzcf71o1_500.gif.

FUEL & FUEL SYSTEM MICROBIOLOGY PART 15 – TEST METHODS – HOW DO WE DETECT BUGS ON SURFACES?

In my August post (https://biodeterioration-control.com/microbial-damage-fuel-systems-hard-detect-part-14-test-methods-still-microbiological-tests/), I discussed using ASTM D7687 to quantify microbial loads (AKA bioburdens) in liquid samples – fuels and fuel associated water. This post will focus on surface samples.

Generally speaking, microbes tend to be most abundant on surfaces. By some estimates, in any given system, for every microbe floating in the bulk fluid, there are 1,000 to 1,000,000 growing on surfaces. These surface microbes are invariably part of biofilm communities. I’ll discuss biofilms in more detail in a future post. For now, it is sufficient to understand that biofilms are slime-encased microbial communities growing on surfaces (fig 1). It is much easier to grab a fluid sample than a surface sample. Consequently, most fuel system samples – even those intended for microbiology testing – are fluids. However, there are a few fuel system surfaces that can be sampled relatively easily.


Fig 1. Scanning electron micrograph of a mature biofilm. Note its structural complexity. Source http://drandreastevens.com/wp-content/uploads/2016/02/Biofilm-Photo.png


Fig 2. Automatic tank gauge, water float showing slime accumulation (right) and swabbed area (left).

Biofilms tend to develop on automatic tank gauge (ATG) water floats (fig 2). The left side of the water float shown in figure 2 has been swabbed. The right side shows the undisturbed deposit. This deposit includes microbes, their slime, and metal fins (i.e. rust). Most often, I use a swab to collect a sample from a premeasured surface area. If the deposit is > 2 mm (1/8 in) thick, I use a spatula to collect the sample.

The second location I routinely check for microbial contamination is the filter. Figure 3a shows a 76 cm (30 in) filter cartridge. It was one of 16 cartridges in a high-capacity filter housing. However, except for its length, the 76 cm cartridge does not look very different from the filter element inside a typical fuel dispenser filter (fig 3b). To test the filter element for microbial contamination, I first inspect the element visually; looking for slime accumulations or discolored zones. For larger filters, I use an alcohol-disinfected forceps and scissors to cut out a section ( 4 cm x 4 cm; fig 3c), and from that cut out a 1 cm x 2 cm piece of filter medium (fig 3d). For dispenser filters, I cut out a 1 cm x 2 cm piece directly. This is my specimen.

If a dispenser has a screen (fig 4), upstream of the filter I collect either a swab or spatula sample just as I would from the ATG water float.



Fig 3. Fuel filter sampling: a) 60 cm filter element from high-capacity housing; b) dispenser filter element; c) section of filter media taken from element shown in fig 3a; d) 1 cm x 2 cm specimen taken from section shown in fig 3c.


Fig 4. Fuel dispenser prefilter screen partially covered with slime.

Once I’ve collected my surface sample I run LuminUltra Technologies, Ltd, Deposit and Surface Analysis (DSA) test (for more information about the DSA method visit https://www.luminultra.com/dsa/; for a video demonstration, visit https://www.youtube.com/watch?v=VEhpbvtej3E). The method provides me with a rapid, quantitative measure of the bioburden on these fuel system surfaces.

Total ATP concentration ([tATP]) <100 pg/cm2 indicates negligible surface contamination. [tATP] between 100 pg/cm2 and 1,000 pg/cm2 indicates moderate contamination (it’s time for maintenance action). [tATP] ≥ 1,000 pg/cm2 signals that prompt corrective action is needed! If you have weighed out samples, the [tATP] per g threshold levels are the same as those for [tATP] per cm2.

If you’d like to learn more about fuel system surface microbiology, please contact me at fredp@biodeterioration-control.com.

FUEL MICROBIOLOGY NEWS FROM RECENT CONFERENCES

In September, I attended two conferences; each of which included a half-day, fuel microbiology session. Although most of the folks presenting fuel microbiology papers were onboard for both conferences, the information overlap was minor. My overall take home lesson is that when it comes to fuel microbiology, we are all still like the five blind men attempting to describe an elephant (if you are not familiar with this ancient, Indian parable, I invite you to look it up).

Although it has been more than 120 years since the first peer-reviewed paper about fuel biodeterioration was published, there is still much we do not understand. The papers presented at the International Biodeterioration and Biodegradation Society (IBBS17) conference during the week of 04 September and the International Conference on the Stability and Handling of Liquid Fuels (ICSHLF15) the following week shed new light on old questions. At the same time, they highlighted the need for more research.
In a nutshell, in my decades of investigating fuel system biodeterioration, I have often detected substantial microbial communities in tanks that showed no evidence of damage. Just as often, I’ve detected considerable damage in systems that seemed to have negligible microbiological contamination. We might just be getting to the point where we can reasonably investigate why some populations cause damage and others don’t. I’ll get to that in a bit.

For those of you who don’t have the patience or inclination to read this entire post, I’ll start with the highlights:
     1. Anaerobic fuel biodeterioration is an important, but often overlooked component of the overall fuel biodeterioration picture.
     2. Sulfur concentration has no impact of fuel biodegradability.
     3. New test methods, still under development, hold tremendous promise to improving our understanding of fuel and fuel system biodeterioration mechanisms.
     4. Fiber-reinforced-polymer biodeterioration is real.

I had the honor of being the keynote speaker, kicking off the IBBS17 session on fuel microbiology. My presentation focused on just how critical sampling is if microbiology data from fuel systems is going to be either relevant or meaningful. During the session, Prof. Joe Suflita (University of Oklahoma) presented the results of studies he and his team have done on microbiologically influenced corrosion (MIC) caused by anaerobic bacteria (anaerobes are microbes that grow only when there is no oxygen present). His two take-home lessons were:
     1. Anaerobes growing in seawater-ballasted diesel tanks cause MIC; and
     2. The fuel’s sulfur concentration (HSD to ULSD) does not affect, microbial growth, fuel biodeterioration, or MIC risk.

Next, Dr. Oscar Ruiz (Air Force Research Lab – AFRL, Dayton) summarized his recent work on genomic (techniques that profile microbial communities, based on the types of genetic material present and the relative abundance of each unique type of microbe – based on its unique genetic profile – in a sample) and metabolomic (techniques that determine which genes are turned on and which are turned off) testing of fuels and fuel-associated waters. Per my comment earlier in this post, it’s not unusual to detect heavy contamination, but not see evidence of biodeterioration. I suspect that as metabolomic testing becomes more practical to run on lots of samples, we will gain a critical understanding of the triggers that cause some microbial populations to cause damage and other to be benign. As an aside, I’ll note here that understanding these triggers has become a major focus of human and animal disease research. More often than previously understood, we get sick when microbes on which we normally depend turn rogue. The next great leap in microbiology will be to understand what genetic switches are turned on or off. After that, the key will be to learn what triggers these switching actions. I am very excited about the work that Dr. Ruiz is doing at AFRL.

Mr.Graham Hill (ECHA Microbiology, Ltd.) reviewed his Energy Institute sponsored work on the relationship between water and microbial contamination levels in biodiesel blends. Graham and his colleagues looked to the effect of fatty acid methyl esters (FAME) on dissolved, dispersed, and free water. Importantly, they found a critical relationship between dispersed and free water, and bioburdens. Microbial loads did not increase as dissolved water concentration increased. Only once fuel-associated water became biologically available, did bioburden increase. These results weren’t surprising, but it is always great to see hard data that support conventional wisdom.

Prof. Ji-Dong Gu (University of Hong Kong) shared some of the work he had done as a post-doctoral fellow at Harvard, in the early 1990’s. This U.S. Air Force sponsored research is still the most comprehensive study of fiber-reinforced polymer (FRP) biodeterioration that has been published (Prof. Gu has several peer-reviewed papers covering this work; several years ago, the Fiberglass Tank and Pipeline’s attorney demanded that I remove all reference to FRP biodeterioration from the BCA website www.biodeterioration-control.com). Prof. Gu’s research demonstrated that a diverse range of polymers and fibers (including several that had biocides blended into the polymer) were susceptible to biodeterioration. He showed a number of very elegant electron microscope images that illustrated the attack mechanism. He also presented electrical impedance data that demonstrated that FRP lost structural strength as biodeterioration progressed.

Dr. George Dodos (Technical University of Athens) presented data demonstrating that FAME composition affected both the rate and specific nature of biodiesel (B5) biodeterioration. His work built on previous studies that showed similar results. Biodeterioration is more rapid when FAME molecules have more carbon-to-carbon (C=C) double bonds (this is called: degree of unsaturation). Dr. Dodos’ research focused on examining the chemical changes that occurred in different biodiesel blends. Publications of this sort of corroborative research is essential to scientific progress.

Prof. Egemen Aydin (Istanbul University) wrapped up the IBBS17 fuel microbiology session with his paper on the biodeterioration of water-soluble molecules in navy fuels. As many readers know, the refining processes used to produce LSD and ULSD adversely affect fuel lubricity, oxidative stability, and corrosivity. Although they are primarily fuel-soluble, additives used to restore these properties have some water solubility. Consequently, they are nutrients for microbes growing in fuel-associated water. Prof. Aydin’s presentation illustrated how water-soluble fuel molecules can stimulate bioburden development and biodeterioration.

ICSHLF15 followed immediately on the heels of IBBS17. As I noted above, many of the same actors attended and presented papers at both conferences. I’ll only mention the presentations that were unique to the ICSHLF15 fuel microbiology session.
Dr. Giovani Cafi (Conidia Bioscience, Ltd.) presented research being done at Conidia using genetic tools to detect and quantify anaerobic microbes in fuels and fuel associated waters. As I noted, apropos of Prof. Suflita’s IBBS17 presentation, except for sulfate reducing bacteria, historically, anaerobic microbes in fuel systems have been largely overlooked. Dr. Cafi reported that anaerobes are commonly part of the fuel microbiology community. Clearly, more research is needed to better understand what anaerobes are present and how they contribute to both product and system biodeterioration.

Mr. Gareth Williams (EHCA Microbiology, Ltd.) discussed EHCA’s recent investigations in which they compared the results of different fuel microbiology test methods. Not surprisingly, Mr. Williams reported that culture tests do not covary strongly with non-culture tests. As I discussed in my December 2015 blog post (https://biodeterioration-control.com/microbial-damage-fuel-systems-hard-detect-part-3-testing/), any single culture test is unlikely to detect >0.1 % of the different types of microbes present in a fuel or fuel-associated water sample. Put another way, the culture tests typically used for routine condition monitoring are 99 % likely to miss microbes that are present in samples, but won’t grow in the nutrient recipe used to manufacture the culture test kit. Unfortunately, as a manufacturer of culture test kits, ECHA presents methods comparisons as though culture test data represent a gold standard for microbiology testing. Conversely, in my own experience, I have routinely detected heavy microbiological contamination by non-culture methods in samples that appear to be microbiologically clean, based on culture test results. Interestingly, the ECHA data set indicated that ATP data obtained using a method other than ASTM D7687 appeared to have no relationship to other measures of microbial loads.

In addition to the oral presentations there were several noteworthy ICSHLF15 posters that addressed fuel microbiology issues.
Dr. Joan Kelly (Conidia Bioscience, Ltd.) presented the results of a survey that she and her collaborators performed on microbial contamination in U.S. retail site UST. The team collected samples from UST across two states. Not surprisingly (to me), Dr. Kelly’s team detected moderate to high levels of microbial contamination in most of the sampled UST.

Dr. Marlin Vangsness (University of Dayton Research Institute) presented a poster reporting bioburden in bulk storage tanks. Dr. Vangsness reported that most sampled tanks had moderate to heavy microbial contamination. Moreover, he reported that ATP data obtained using ASTM D7463 did not correlate with other microbiological parameters. Having spent 30 years working to separate interferences that had historically made ATP testing unusable for complex, organic chemical rich fluids like fuels and lubricants, I have argued that ASTM D7463 is an unreliable test method. D7463 does not separate water-soluble organic chemicals and salts from microbes in the test specimen. Consequently, ASTM D7463 results are subject to both positive (high values caused by chemical reactions) and negative (low values caused when chemicals in samples capture the light that is generated by the test reaction – see https://biodeterioration-control.com/microbial-damage-fuel-systems-hard-detect-part-14-test-methods-still-microbiological-tests/ – before the light reaches the detector) interferences. Dr. Vangsness’ poster and Mr. Willams’ presentation both corroborate findings that National Research Defense Canada presented at a NATO conference, nearly a decade ago. In contrast to ASTM D7463, ASTM D7687 (see the previous hyperlink) effectively separates interfering chemicals from microbes before extracting ATP.

Speaking of ATP, Ms. Chrysovalanti Tsesmeli, a doctoral candidate at the Technical University of Athens, presented a poster reporting her use of ASTM D7687 and chemical analysis to explore the effects of FAME and hydrogenated vegetable oil (HVO) on marine diesel fuel biodeterioration. Her work showed that HVO-blended fuels were more biostable than FAME-blended fuels.
Last, but not least, Ms. Silvia Bozzi (Chimec S.p.A.) presented a poster that was similar to Dr. Kelley’s. Ms. Bozzi reported on a survey of retail UST in Italy. As in the U.S., the incidence of moderate to high microbial contamination levels in Italy’s retail site UST is considerably greater than generally recognized by site owners and operators.

One of the particularly gratifying aspects of both conferences was the number of young (i.e. under the age of 40) researchers who are investigating fuel microbiology. These young scientists are applying new techniques to ask new questions and to obtain answers that we cannot get using traditional microbiology methods. Moreover, often the young researchers come from non-microbiology disciplines. Because this reflects a multidisciplinary approach to fuel and fuel system biodeterioration it bodes well for the future of fuel microbiology.
Although I didn’t mention any posters or presentations made by Prof. Fatima Bento or her graduate students, I’ll close this blog with a special call out acknowledging the great research being done by this group at Instituto de Ciências Básicas da Saúde, Sao Paulo, Brazil. Few months pass when I don’t have an opportunity to review manuscripts submitted by members of Prof. Bento’s team. They have made many important contributions to our understanding of ULSD and biodiesel biodeterioration. I’d be sorely remiss if I didn’t mention their fine work.

As always, if you’d like to learn more about fuel and fuel system microbiology testing, please contact me at fredp@biodeterioration-control.com.

FUEL & FUEL SYSTEM MICROBIOLOGY PART 13 –TEST METHODS – MORE ON MICROBIOLOGICAL TESTS

In Part 13, I discussed culture testing. One of the points I made was that any given culture test (of which there are >5,000) is unlikely to detect >1 % of all of the microbes present. Before moving on to discuss methods that detect more of the microbes present – in terms of percent detection of each type of microbe and the fraction of the different microbes present that are detectable – I will invoke one of Donald Rumsfeld’s most famous quotes:

“There are known knowns. These are things we know that we know. There are known unknowns. That is to say, there are things that we know we don’t know. But there are also unknown unknowns. There are things we don’t know we don’t know.”

Although, in February 2002, when Secretary of Defense Rumsfeld offered this statement, he was discussing the possibility that Iraq had weapons of mass destruction; he could just as well been talking about microbial contamination condition monitoring. In Part 12’s fig 1, I indicated that genomic testing (you’ll have to wait until Blog Post 15 or 16 for more on genomics) detected a greater proportion of the total microbiome (all of the microbes present in a particular environment) than any other method currently available. However, I also noted that I doubted if current genomic testing detected more than 80% of a given microbiome. This begs the question: “If no method provides a perfect measurement of microbial contamination, which one should I use?”

The perhaps ungratifying answer is: “It depends on your intention.” Let’s start with an illustration. Fig 1 illustrates three ways to take a measurement. You can use a ruler or tape measure to determine an object’s dimensions. If It is a liquid, you can use a measuring cup or graduated cylinder to determine its volume. You can also use a scale to determine its weight. Each of these is a valid measurement, but each provides different information.

Fig 1. Three different ways to measure.

 

It’s the same thing with testing from microbial contamination. Each method that I illustrated in Blog Post 12, figure 1, provides useful information about the microbial population, but each provides different information. If you need to have pure cultures of microbes, on which to do research, culture testing is the most appropriate tool. If, however, you want to quickly determine how heavily contaminated your system is, then one of the chemical microbiology test methods is a better choice.

A chemical microbiology test method is a method that detects specific molecules that are either part of or are produced by microbes. The three chemical microbiology methods illustrated in Fuel Microbiology Part 12 are: catalase activity, adenosine triphosphate concentration, microbial antigen detection.

Today, I’ll write about the catalase test. In the interest of full disclosure, in the early 1980’s, after a University of Houston graduate student developed the HMB catalase test method (www.biotechintl.com), I did most of the method validation for a variety of industrial applications. I also developed ancillary HMB tests to verify that the test results were due to microbes. Starting in 1982, and for the next 27 years, the HMB was my primary field test for detecting and quantifying microbial contamination in industrial fluid systems.

The catalase test is based on the reaction between the enzyme catalase and hydrogen peroxide. Catalase is the enzyme that made life in an oxygen-rich atmosphere possible. Cells that grow in normal air (aerobes) produce hydrogen peroxide as part of their energy metabolism. Catalase converts that hydrogen peroxide into water and oxygen. What makes the HMB test quantitative are its two primary components: a patented, electronic pressure gauge (figure 2a) and a stoppered reaction tube (figure 2b).

Fig 2. HMB catalase test system. a) pressure measurement device; b) stoppered reaction tube

 

The HMB pressure gauge is unique because there’s very little volume between its probe and its sensor.

The stoppered reaction tube provides a fixed volume, so that headspace pressure increases as the concentration of oxygen gas increases within that space (the head space is the space between the top of the liquid and bottom of the stopper).

To run the test, add a standard sample volume (typically either 3 mL or 10 mL) to a reaction tube, and then add concentrated hydrogen peroxide (one drop – = 0.05 mL – per mL of sample). Quickly replace the tube’s stopper (it is a septum cap that re-seals itself after it has been pierced with a needle) and briefly vent the tube. This ensures that the headspace pressure is 0 psig when the reaction starts. If there are aerobic microbes in the sample, they will race to convert the hydrogen peroxide to water and oxygen gas, before the hydrogen peroxide kills them. In the meantime, as oxygen is produced, it accumulates in the reaction tube’s head space. The universal gas law teaches that if temperature and volume are constant, the pressure in an enclosed space is proportional to the concentration of gas in that space. Simply put: the more catalase enzyme in the sample, the more oxygen in the headspace; the more oxygen the greater the pressure increase (fig 3). The reaction runs its course in <15 min. At 15 min, stick the reaction tube with the needle that’s attached to the pressure gauge (fig 1a) and read the psig. The psig reading at 15 min is proportional to the microbial contamination load. Correlation between culture test data and HMB catalase test data is generally very strong.

Fig 3. Catalase reaction with hydrogen peroxide in reaction tube. a) negligible contamination = negligible oxygen accumulation = negligible pressure increase in headspace; b) heavy contamination = substantial oxygen accumulation = large pressure increase in headspace.

 

However, the HMB test has its limitations. First: it only detects organisms that have the catalase enzyme. This excludes all anaerobes (microbes that only grow in oxygen-free environments) and aerobes that don’t have a complete catalase enzyme. Second: dissolved iron reacts with hydrogen peroxide to release oxygen gas. Samples with dissolved oxygen will appear to have microbial contamination. Third: at ∼25 psig the pressure is sufficient to launch the reaction tube’s stopper. The noise can be disconcerting and flying stoppers can be eye hazards. Moreover, the foam pouring over the reaction tube’s wall creates a mess. When microbiological contamination is negligible, it generates <1.5 psig pressure. Heavily contaminated samples (many bottoms-water samples) will foam over before the reaction tube’s stopper can be put in place (have you ever seen the reaction when sulfuric acid is poured over a sugar cube; fig 4?). When this occurs, the sample must be diluted to get a quantitative test result. On the few occasions when curiosity has compelled me to get a quantitative answer, after observing a violent reaction in the original sample, I’ve found that the actual psig was 20,000 to 30,000 psig (yes, I had to dilute samples 10,000 to 50,000-fold in order to get a psig reading). Normally, either being unable to get the stopper onto the tube, or having the stopper launch before the end of the 15 min test period, provide the information I need to determine that the sample is heavily contaminated.

Fig 4. Column of sugar charcoal formed after adding sulfuric acid to sugar. The reaction is violent and exothermic (give off lots of heat).

 

Earlier, I mentioned that I had developed ancillary tests for the HMB catalase test. One is used to determine if dissolved iron is producing a false positive result. The other is used to inactivate any enzymes in the sample. When testing unknown samples (i.e.: I don’t know whether they sample is likely to have dissolved iron), I run four tests: hydrogen peroxide (H2O2) only, H2O2 + a chelating reagent (prevents the dissolve iron reaction), H2O2 + a poison (inhibits catalase activity), and H2O2 + chelating reagent + poison (serves as a background control). The H2O2 result tells me if there is a contamination issue. If the chelating reagent reduces the psig by >90 %, then the psig observed in the H2O2 only test is due to dissolve iron. Similarly, if the chelator has no effect but the poison reduces the psig by >90 %, then the psig observed in the H2O2 only test is due to microbes. If both the chelator and poison are needed to reduce the psig by >90 %, then the sample has substantial concentrations of dissolved iron and microbial contamination.

With all of these limitations, why use the HMB test? The truth is, for those 27 years during which I relied on it, the HMB test was the best test available for my specific objectives: to be able to obtain a sample and obtain reasonably reliable, quantitative microbiological data, quickly (15 min), near the point of sampling. These days, I compare the test method to early portable phones and so-called laptop computers (the former weighed in at > 10 lb., and the latter at > 20 lb.) At the time they were introduced, they did their respective jobs better than anything else available. I hope that you are now wondering: What test replaced the HMB test? That will be the topic of Part 14. Stay tuned…

In the meantime, if you’d like to learn more about fuel and fuel system microbiology testing, please contact me at fredp@biodeterioration-control.com.

FUEL & FUEL SYSTEM MICROBIOLOGY PART 12 –TEST METHODS – MICROBIOLOGICAL TESTS

Since November, this series has progressed through fuel system sampling, sample handling and non-microbiological tests used to detect biodeterioration. This post, and the three to follow, will cover microbiological testing.

Let’s take another look at the figure (fig 1) that accompanied Part 3 (December 2016):

Fig 1. Ability of different microbiological test method to detect all microbes present in a microbiome.

The largest circle represents the total microbiome – all the microbes present in a particular environment. The parameters portrayed in the figure are not exhaustive. For example, the figure does not include direct counts: the use of a microscope to examination samples and count the number of microbial cells per microscope field (the area visible when looking through a microscope’s lenses). Nor does it include tests that measure the concentration of building block molecules such as proteins, carbohydrates, or fatty acid methyl esters (FAME). Direct counting is labor intensive. Moreover, it can be difficult to distinguish between microbes and microbe-size inanimate particles. Finally, there is considerable debate about whether direct counting includes both live and dead cells. Although, theoretically, direct count methods detect 100% of the total microbiome, direct counting is rarely used in practice.

Culture testing is currently the most commonly used tool for determining bioburdens in fuel and fuel associated water samples. Culture testing depends on microbes captured in a sample to be able to reproduce (proliferate) either in or on the growth media used to perform the test. The growth media can be either solid (ASTM Practice D7469), semisolid (ASTM D7978), or liquid (for example: LiquiCult Test Kits – LiquiCult is a trademark of MCE, Inc.; http://www.metalchem.com/liqui-cult.html). First developed in the late 19th century to detect disease causing microbes, culture testing is now often used without any real understanding of its real purpose or its limitations.

To produce a visible colony (mass of cells), a microbe must reproduce. A generation is the time required for a population to double: for one cell to become two; two to become four, etc. A visible colony has at least 1 billion cells. It takes 29 generations to get from one cell to a billion cells (fig 2).

Fig 2. Microbe proliferation from individual cell to visible colony.

 

To reproduce, a microbe must have the right nutrients and environmental conditions. There are more than 5,000 different recipes for microbiological growth media. Each one is optimized for the nutrient requirements of specific types of microbes. No individual type of microbe will grow on all media. Additionally, different microbes have unique preferences for growth conditions (atmosphere with oxygen present versus oxygen-free atmosphere; acidic, neutral, or alkaline environment; cold, temperate, or hot – >40 °C/104 °F; etc.). Consequently, the 1% recovery estimated in fig 1 doesn’t reflect the detection power of all test method combined. It reflects the sensitivity (actually: insensitivity) of any individual culture method. If an analyst ran thousands (perhaps millions) of different combinations of growth media and conditions, the combined results might detect 50% to 60% of the total microbiome population. There are still many microbes that we do not know how to culture.

In addition to the selective effects of any combination of growth medium and incubation conditions, time affects culture test sensitivity. Known microbe generation times range from 15 min for the fastest growing bacteria to 30 days for the slowest. The fastest growing microbes can proliferate from single cells to visible colonies in less than a day. A microbe with a 4h generation time needs nearly five days to from a visible colony, and one with a 30-day generation time needs nearly 2.4 years! Most commercial test kits recommend observing colonies daily, for up to three days. Any microbe with a generation time longer than 2h is unlikely to be detected. Analysts testing samples contaminated with microbes that have generation times of >2h will incorrectly conclude that the samples are uncontaminated.

Notwithstanding these limitations, culture testing has been used with reasonable success for more than a century. It remains the only tool available for obtaining pure cultures on which to do additional testing. Consequently, the take home message is not to dismiss culture testing. Rather it is to recognize that culture testing has specific uses. Obtaining an estimate of total levels of microbial contamination (i.e.: bioburdens) is not one of them. In the next several blog posts, we’ll look at tools better suited for that purpose.

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