Soil Phospholipid Fatty Acid (PLFA) Analysis! The Not So New Kid on the Block
We routinely monitor the orchard for insect damage and disease symptoms, take soil samples to monitor the pH, and collect annual leaf samples to check the nutrient levels of the orchard. As some orchard managers have ventured into the Regenerative Ag realm, they have added additional monitoring techniques such as the Haney Soil Test, Solvita soil respiration, water infiltration, and the PLFA test. The PLFA (Phospholipid Fatty Acid) test seems to be creating the most confusion with growers, so I thought we would spend some time reviewing the history and basics of the test.
While the PLFA technique is new to a lot of farmers, especially with the increase of its use as new land management systems (regenerative, sustainable, agroforestry, silvopasture, etc.) have been implemented in the last few years, the contemporary PLFA analysis we currently use has been in development for over six decades.
Quideau et al. (2016) provide a good synopsis of the evolution. The present test traces back to the Bligh and Dyer method (Bligh and Dyer, 1959) and has undergone several modifications over the years. David White and colleagues at the University of Tennessee did a lot of the early heavy lifting, modifying buffers used and improving the gas chromatography analysis, including enhancement of peak identification and quantification. After a couple of years working as a post-doc under White, Anders Tunlid returned to Lund University in Sweden and started collaborative work with Erland Bååth (also at Lund) and Åsa Frostegård (Norwegian University of Life Science), which led to several more important modifications, such as their use of multivariate statistical techniques in exploring and presenting the data. Also, their use of citrate buffer instead of phosphate buffer led to increases in the extracted amount of lipid phosphate. Frostegård’s paper (1993) on the influence of liming on soil microbial communities, along with the detailed methodology by White & Ringelberg (1998), provided the foundation for an explosion in the use of the PLFA technique to investigate fundamental questions in soil science in the 2000s. Since then, further refinements of the method by Firestone and colleagues (2011) have included adding an internal GC standard (C10:0) and C19:0 as a surrogate standard to improve quantification.
Now that we better understand the history, let’s get back to the basics. The first thing we have to realize is the soil isn’t an inert growing medium but a home to billions of bacteria, fungi, and other microbes that make up the foundation of a healthy symbiotic ecosystem. All of these microbes have cells surrounded by a biological membrane that protects the interior of the cell from the external environment. These membranes generally comprise predominantly phospholipids but contain other lipids, cholesterol, and proteins. A phospholipid has a hydrophilic “head” containing a phosphate group with two hydrophobic “tails” derived from fatty acids. It is the composition of these fatty acid tails that provide the basis of the PLFA test. Even though phospholipids occur in many different lifeforms (plant roots, fungi, bacteria, protists, etc.), the fatty acid side chains between differing lifeforms can be unique and serve as a biomarker for a specific group of lifeforms.
A typical PLFA report will generally provide values for total living microbial biomass, gram-positive bacteria, gram-negative bacteria, arbuscular mycorrhizae, saprophytes, protozoa (predators), and undifferentiated microorganisms. Let’s quickly look at some of the microorganisms we expect to find in a pecan orchard’s soil. Starting with bacteria, when a “Gram” staining agent is used in the lab, bacteria can be separated into “gram-negative” or “gram-positive” categories. The staining agent attaches to the bacteria’s cell walls. Gram-negative bacteria are generally the smallest bacteria and are sensitive to drought and water stress. Gram-positive bacteria tend to be larger with thicker cell walls and resist water stress; their outside cell wall surface is negatively charged.
Gram-negative bacteria are a major component of the soil and provide a positive nutrition impact on plants by increasing phosphate solubility, producing ionophore compounds that increase the uptake of iron or other minerals, and possibly producing antifungal compounds. One of the more important gram-negative bacteria is Rhizobium, which forms an association with leguminous plants such as clover, vetch, and alfalfa. Rhizobium forms nodules on the plant roots and converts atmospheric nitrogen to a plant-available form. In return, the plant supplies carbon in the form of simple sugars.
Gram-positive bacteria. Actinobacteria, are found in various environments, including soil, freshwater, and marine sites. Actinomycetes are a special group of bacteria that have several similarities with fungi, such as the growth of mycelia in a filamentous and branched pattern. They are active in the decomposition of organic matter, breaking down some of the toughest dead animal, fungal, and plant polymers. They produce a wide range of hydrolytic enzymes needed to decompose organic residues. You recognize their presence whenever you pass a tilled field or stroll through a pasture after a rain. That rich earthy smell is geosmin, an organic compound produced by this group of bacteria. Actinomycetes are involved in forming stable humus, leading to enhanced soil structure, increased nutrient storage, and improved water retention. Other common genera of soil Actinobacteria include Rhodoccus, Nocardia, Corynebacterium, and Streptomyces (renowned for its prolific production of antibiotics). Other common Gram-positive bacteria would include aerobic Bacillus species, especially those related to B. cereus and B. subtilis. These may sound familiar to some of you because selected strains of Bacillus are used as the active ingredient of some biofungicides labeled on pecan, including Serenade OPTI (Bacillus subtilis), LifeGard WP (Bacillus mycoides), and Double Nickel 55 WDG (Bacillus amyloliquefaciens).
Fungi—Arbuscular mycorrhizae and saprophytes. Mycorrhizae are divided into several types depending on how they physically interface with the host plant. The ectomycorrhizae grow on the surface layers of the roots and have a Hartig net of hyphae that physically extends into the root but between the cells. This type is most commonly associated with trees. Another group of mycorrhizae is the endomycorrhizae that penetrate the cell wall of root cells and are generally associated with grasses, row crops, vegetables, and shrubs. Arbuscular mycorrhizal (AMF) fungi are a type of endomycorrhizal fungi. In these mycorrhizae, the fungi form arbuscules that penetrate root cells and are the site of the metabolic exchanges of phosphorus, nitrogen compounds, and other minerals between the fungus and the host plant.
Although AMF are found in many soils, they are not free-living in the soil. They are obligate symbiotic fungi and can form an association with about 80% of all vascular plant families. Regarding the PLFA analysis, the AMF hyphae in the root form lipid materials, which then are transported to the hyphae that extend into the soil from the root, which means they can be detected in a soil sample. Fungal monounsaturated biomarkers include C16:1ω5 as an indicator of arbuscular mycorrhizal fungi (AMF), C18:1ω9 is commonly used for saprophytic fungi, and ectomycorrhizae typically contain C16:1ω9.
Typically, most trees will only form an association with one type of mycorrhiza, either endomycorrhiza or ectomycorrhiza. However, research has shown that pecan can associate with both types. It appears to prefer ectomycorrhiza, with colonization by arbuscular mycorrhiza usually only occurring on less than one-third of the root system. A question that still must be answered is, “Does this dual association provide pecan trees an advantage in soil exploration for nutrients, and is this one of the reasons pecans can expand into geographically different environments so easily?”
Protozoa. Microorganisms in soil are intensively grazed by a variety of soil animals with protozoa, forming the most abundant and ubiquitous grazers of bacteria. Protozoa are classified into three subgroups: ciliates, flagellates, and amoebae. As bacteria decompose organic matter, they retain the nutrients in their bodies. These nutrients are not released until the bacteria die or protozoa capture them. Protozoans do not require all the nutrients contained in the bacteria, and about 30% of the bacterial nitrogen is excreted as ammonium by protozoa and thereby made available for plant uptake. With enhanced nitrogen supply, plant growth increases and the growing plants allocate more carbon into the rhizosphere, thereby fuelling rhizosphere bacteria. Thus, protozoans likely benefit from increased plant nitrogen uptake as it fosters the growth of bacterial prey via enhanced root exudation.
Undifferentiated microorganisms. This category will usually be the highest portion of your total living biomass, often greater than 50% of the total. This is generally so high because the diversity of soil microorganisms is enormous, and most species are still unidentified. A few grams of soil is home to billions of microorganisms, including archaea, bacteria, fungi, and protozoa. Archaean microbes are involved in nutrient recycling (carbon, nitrogen, sulfur) in soils but differ from bacteria and fungi because they do not contain PLFAs. Consequently, the PLFA protocol is probably underestimating the total living microbial biomass because it can’t detect archaeal communities in soil.
So now that we have discussed some of the microorganisms you will find in the soil in your orchard let’s get back to discussing the PLFA test. As mentioned earlier, the basic premise is that the phospholipids of distinct groups of bacteria and fungi contain various somewhat unusual fatty acids that can serve as useful biomarkers for such groups. Microorganisms produce PLFAs of different chain lengths and compositions as a means to maintain cell-membrane integrity and cellular function in response to their immediate environmental conditions. The PLFAs in the soil at the time of collection are considered a snapshot of the viable, living organisms present. Phospholipid fatty acids are synthesized quickly as microbes grow. Still, as microorganisms die, their PLFAs are rapidly decomposed in an aerobic, active microbial community and do not accumulate in the soil. So, the PLFAs analyzed in the sample are considered to be from only living microorganisms, and the total PLFA content equals the total microbial biomass in the soil sample.
The PLFA analysis is widely used to estimate the total microbial biomass and identify broad changes in the composition of the microbial community in the soil. It has been successfully applied in characterizing soil response to land management changes, including forest clear-cutting, liming, and reclamation, as well as disturbances such as fire, contamination by metals and hydrocarbons, and insect outbreak. It has been used to characterize the structural composition of microbial communities found in a wide range of environments, ranging from forests to prairies and agricultural fields. This recent surge of interest in PLFAs, evident from the increase in peer-reviewed journal articles and references on the subject, has increased concern that some researchers may be assigning PLFAs to a specific microbial group when in fact those specific PLFAs are present in numerous other life forms.
In soils containing a diverse microbial population, PLFA analysis cannot be used to identify individual bacterial and fungal species, but it can provide information on different segments of the microbial communities found in soils. However, to minimize and avoid misinterpretation of PLFA data, careful data screening must be done because some PLFAs found in the soil microbial community are also present in single and multicellular eukaryotic organisms, such as plant roots, algae, and soil animals. For instance, the tri-unsaturated C18:3ω 6,9,12 has been used as a biomarker for fungi in some labs; however, C18:3ω6c can be found in other eukaryotic organisms, including plants (roots in the sample) and algae, although it is typically not found in bacteria. It is important to note that plant tissue (active roots, cover-crop residue, incorporated green manures) share similar phospholipids as bacteria and fungi. Sieving and tweezers can be used to reduce the amount of plant debris, thereby reducing these interferences to a minimal level.
While contamination can be problematic, PLFA tests can still yield meaningful data, particularly on long-term management comparisons with similarly derived samples run in the same laboratory. Take note of the last two words of the previous sentence: same laboratory. Because not all labs use the same biomarkers, it is important to select a lab and continue to use them over several years. Switching back and forth between different labs may not allow you to accurately compare results.
PLFA analysis can provide information about the overall structure of the microbial community. It can provide a biological index of overall soil quality and serve as a quantitative indicator of soil response to land management and environmental stresses. It allows the tracking of change over time following the implementation of a new management system. As we discussed earlier, the PLFA is a snapshot of community structure and abundance at the moment the sample was collected. But the microbial population can be influenced by numerous environmental conditions, including soil pH, temperature, and soil moisture content. These communities are also impacted by soil type, organic matter, soil disturbance, crop rotations, cover crop selection, and applied pesticides. And while producers can use PLFA to compare agricultural management techniques with respect to overall better soil health and fertility, currently we do not have baseline values for biological testing as we do with soil and foliar nutrient analysis.