FUEL & FUEL SYSTEM MICROBIOLOGY PART 34 – Connecting the Dots, Part 2

Refresher from Part 1: What do Microbiology Test Results Mean?

In January’s Fuel & Fuel System Microbiology article I led with this question and commented that it is actually a double question. In one sense, it is asking: “Do my microbiology test results tell me conclusively whether microbes are damaging my fuel or fuel system?” In another sense, the question means: “Why don’t the results from different fuel microbiology test kits always agree?” I am then asked why often, even when microbiological test data indicate that there is heavy biocontamination present, the fuel does not seem to be affected. In today’s post – the second of three on this topic – I’ll discuss the relationship between microbiological test results and system damage.

Do My Microbiology Test Results Tell Me Conclusively Whether Microbes are Damaging My Fuel System?”

As I wrote, last month, the short answer is no. Keep in mind, all three of these posts about whether detected microbial contamination invariably signals biodeterioration is happening. This is different from the situation in which there are numerous indications of system biodeterioration, but microbiological test results are negative. I’ll revisit that issue in a future post.

Fuel System Biodeterioration

Biodeterioration is any damage caused by organisms. In fuel systems, the most common forms of biodeterioration are biofouling and microbiologically influence corrosion (MIC).

Biofouling is the result of microbes and the slime they produce (i.e., extracellular polymeric substance – EPS – the primary material in biofilms (see Part 15 for a refresher on biofilms) accumulating on system surfaces. When biomass accumulates on filters or screens, it restricts product flow. Figure 1 shows photographs of a dispenser filter, dispenser strainer, and leak detector strainer – each of which has become fouled with biomass.

Biofouling can also cause other problems including valves sticking or failing to close completely. When biofouling accumulates on the surface of an automatic tank gauge’s (ATG’s) water float (Figure 2a) the impact will depend on the biofilm. If the biofilm is filled with gas pockets, the float will be lighter than normal and will float within the fuel – giving a false signal that bottoms-water is present when it is not (Figure 2b). Conversely if the EPS is loaded with rust particles, the water float will be heavier than normal. It will rest on the tank bottom, even when 2 cm to 3 cm bottoms-water as accumulated (Figure 2c)

Biofilms coating vehicle fuel gauges will cause the gauges to give inaccurate readings.

Note that all of these biofilm accumulation zones are on system components. Fuel systems can have substantial bioburdens in tank bottom samples, but no biofouling. Although the possibility of fouling increases with increased bioburden in fuel tank bottom samples, detection of substantial microbial loads in fluid samples doesn’t necessarily mean that fouling has occurred. The only way to know for certain whether biofouling has occurred is by direct inspection of the fuel system components that are likely to become fouled.

Microbiologically influenced corrosion (MIC) includes any from of material damage that is caused either directly or indirectly by microbes. Most commonly, MIC is related to metallic components, but polymeric materials are also susceptible to MIC.

Contamination Detection

Connie Francis recorded Where the Boys Are as the title track for the 1961 movie of the same name. The next several paragraphs could be titled Where the Microbes Are. Microbial contamination is not a fuel property. Unlike fuel properties, the distribution of microbes in fuel systems is non-uniform (heterogeneous). The heterogeneous distribution of microbial contamination makes it difficult to collect a sample that is guaranteed to contain microbes – even if microbial contamination is present in the fuel system.

In my fuel microbiology courses I recount a lecture I heard as an undergraduate. My professor was part of the team tasked with developing a reliable test method for determining whether there was life on Mars. One member of the team suggested using a camera that would scan the horizon for signs of life. The device would scan 15 ° of arc at a time, completing a 360 ° scan each hour. The counterargument – illustrated in figure 3 – was that large life forms (elephants in figure 3) might be present but missed entirely because they continually moved out of the camera’s line of sight.

Fig 3. Not detecting the elephants – a) elephants are to the east while camera is pointing west; b) elephants are to the west while camera is pointing east. A researcher viewing the camera’s photo record would conclude that there are no elephants in the area photographed!

Now consider a 0.5 L (0.13 gal) sample collected from the bottom of a 38,000 (38 m3, 10,000 gal) tank. The sample represents 0.001 % of the total liquid volume in the tank. Similarly, a bottom sample from the bottom of a tank with a 31 m2 (31,000 cm2, 334 ft2) surface area draws in fluid, sludge, and sediment form a 3 cm to 5 cm radius. That represents 0.02 % of the total bottom surface area. Figure 4 illustrates how a two bottoms samples, taken from spots just a few cm apart, can have substantially different bioburdens.

Bottom Line

Fuel system samples used for microbiological testing are meant to be diagnostic – not representative. To be reliably diagnostic, samples must come from locations most likely to harbor microbes. This is can be impractical (if not impossible). Consequently, samples from systems with substantial fouling, MIC, or both can have negligible detectable bioburdens. Conversely, it is not uncommon for systems from which samples have apparently heavy bioburdens to have no biodeterioration symptoms. In Connecting the Dots – Part 3, I’ll write about why test results from different microbiology methods can lead to different conclusions. As I was writing today’s blog I decided to add a Part 4 – the impact of specific microbial activities on the link between bioburden and biodeterioration.

The details

For more details about understanding the relationship between microbiology test data and fuel or fuel system biodeterioration, please contact me at either fredp@biodeterioration.control.com or 01 609.306.5250.

FUEL & FUEL SYSTEM MICROBIOLOGY PART 29

What Does “Viable But Not Culturable” Mean and Why Should I Care?

In microbiology, the term used to describe microbes that appear to be healthy and active by test methods other than culturing is viable but not culturable – VNBC. Since the term first came into vogue in the 1980s, it has always reminded me of the Monty Python skit in which the customer – played by John Cleese – and the shop owner – played by Michael Palin – debate whether the parrot that Mr. Cleese had just bought was dead or simply resting, check it out at The Parrot Sketch.

The viability versus culturability debate

The issue is relevant for two reasons. First, if a fuel or other industrial process fluid system (think heat exchange fluids, metalworking fluids, lubricating oils and hydraulic fluids) is home to a population of microbes that are biodeteriogenic (i.e., causing damage to the fluid, the system, or both) but are not detected by culture testing, the risk of experiencing a failure event can high.

Second, the term VNBC has numerous meanings – depending on researchers’ focus. The varied definitions creates confusion among both microbiologists and others who rely on microbiological test results to drive maintenance decisions.

What does viable mean?

The Online Biology Dictionary defines viable as an adjective meaning (“1) Alive; capable of living, developing, or reproducing, as in a viable cell.” ASTM is a bit more helpful offering several similar definitions. From F2739 Guide for Quantifying Cell Viability within Biomaterial Scaffolds we get: “viable cell, n – a cell capable of metabolic activity that is structurally intact with a functioning cell membrane.” D7463 and E2694 offer: “viable microbial biomass, n – metabolically active (living) microorganisms.” These slight variations all agree that viability relates to a microbe’s ability to:

• function under favorable physical and chemical conditions (more on this in a bit), or
• to survive in an inactive (dormant) state under unfavorable conditions, and
• to become active again once conditions improve.

What does culturable mean?

ASTM defines culturable as an adjective: “microorganisms that proliferate as indicated by the formation of colonies on solid growth media or the development of turbidity in liquid growth media under specific growth conditions.” This definition is used in several ASTM standard test methods, guides, and practices.
When microbes reproduce – i.e., proliferate – go through repeated cycles of division – on a solid or semi-solid medium, after approximately 30 generations (doubling cycles, or generations) they accumulate enough mass to form a visible colony. Thirty generations (230) yields approximately a billion cells. Liquid growth media become visibly turbid once the population density (cells mL-1) reaches approximately one million (106) cells – 20 generations. The duration of a single generation varies among microbial species and growth conditions. At present, known generation times range from 15 min for the fastest proliferating bacteria to >30 days (recent discoveries of deep earth microbes suggest that these microbes might have generation times measured in years or decades – the generation time for humans is approximately 30 years). The key point is that culturable, microbes reproduce in or on growth media under specific environmental conditions.

Before leaving our discussion of culturable lets consider time. Microbes with 15 min generation times will turn broth media turbid in 5h to 6h and form visible colonies on solid media within 8h to 10h. For microbes with a 1h generation time, detection as turbidity or colonies lengthens to 20h and 30h respectively. Many culture-based test protocols state that final observations are to be made after 3-days – sometimes 5-days. Any microbe with a generation time longer than 4h is unlikely to produce a visible colony within 5-days. They will not be detected unless observations are continued for a week or longer. For example, the culture test for sulfate reducing bacterial is not scored negative until after 30-days observation. If you end a culture test at 3-days, are all of the slower growing microbes non-culturable?

What are growth media?

Since the mid-1850s, microbiologists have developed thousands of different recipes designed to support microbial growth and proliferation (recall from an early post that growth refers to the increase in mass, and as noted above, proliferation refers to an increase in numbers). Some growth media are undefined. They are simple recipes made up of extracts from yeasts, soy, and animals. These are the components of media used for the most common culture test: the standard plate count. Other media are prepared from individual chemicals. Their recipes can include more than a dozen ingredients. Solid and semi-solid media include a gelling agent such as agar (extracted from seaweed), gelatin, or silica gel. One of the most frequently referenced microbiological media cookbooks – the Difco Manual – lists more than a thousand recipes. Each of these recipes was developed to detect one or more types of microbes. In addition to the diversity of recipe ingredients, growth media vary in pH, total nutrient concentration (some microbes cannot tolerate more than trace concentrations of nutrients), and salts concentration (ranging from deionized water to brine). The microbes targeted for recovery dictate post-inoculation incubation conditions. Some microbes require an oxygen-free (anoxic) environment. Others require special gas mixtures. Microbes also vary widely on the temperatures at which they will grow. Some only grow at temperatures close to freezing. Others require temperatures closer to boiling.

The growth medium defines the chemical environment and the incubation conditions define the physical environment in which microbes are cultured. No single growth medium is likely to support the proliferation of more than a tiny fraction of the different types of microbes present in an environmental sample. Many microbiologists estimate that <0.1 % of the microbes in a sample will be culturable in a given medium. Similarly, we suspect that for every microbe that has been cultured, there are at least a billion that haven’t.

Is my microbe really dead or simply resting? When conditions are unfavorable to either growth, proliferation or both, many different types of microbes have coping mechanisms. For nearly 200 years, we have recognized that some types of microbes can form endospores – their cell wall chemistry changes and metabolic activity ceases. Only in the past 20 years, we have come to recognize that non-spore-forming microbes can enter into a dormant state that enables them to survive unfavorable conditions for centuries or millennia – becoming metabolically active once conditions once again become favorable. Moreover, in some environments, although the microbes are metabolically active, the rate of their activity is so slow as to be nearly undetectable.

Within some fields of microbiology, VNBC refers to microbes that have been injured due to exposure to a microbicide. Pre-incubation in so-called recovery media – improves their ability to reproduce in or on growth media. In my opinion, this is a very myopic view of VNBC.

Microbial ecologists define VNBC as microbes that are metabolically active or dormant in their home environments but will not growth on the culture media and incubation conditions used to detect them.

I first experienced this phenomenon in the 1970s when I was testing water from oilfield production wells. Using radioactive carbon labelled nutrients to measure metabolic activity, my team routinely found that samples that yielded <1 CFU mL-1 (CFU – colony forming unit: “a viable microorganism or aggregate of viable microorganisms, which proliferate(s) in a culture medium to produce a viable colony.” ASTM E2896) held very active populations. Poisoned controls demonstrated that conversion from radiolabeled acetic acid or glucose to radiolabeled carbon dioxide was from metabolic activity – not from a non-biological (abiotic) process.

This means that culture testing invariably underestimates the microbial population density in tested samples. Conversely, because microbes that were dormant in the environment from which they were sampled can become active once transferred into or onto nutrient media, culture testing can overestimate the presence of metabolically cells. For example, most microbes suspended in fuels or lubricants are dormant, but can become active and form colonies on growth media. These issues do not make culture testing good or bad. Culture testing is still the only practical tool for obtaining microbial isolates that can be used for further testing. Moreover, culture testing is a useful condition monitoring tool if you are tracking changes over time. The more important limitation of culture testing as a condition monitoring tool is the delay between test initiation and the availability of test results. In the days or weeks it takes for microbes to form colonies on growth media, they are also continuing to proliferate in the system from which the sample was collected. This is where real-time (10 min) tests such as ASTM D4012, D7687, and E2694 have a major advantage over culture testing.

For more information on the most strategic use of culture or non-culture microbiology test methods, I invite you to 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.

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).

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.

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.

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):

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).

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.