TY - JOUR
AU1 - Halter, Mathew C
AU2 - Zahn, James A
AB - Abstract White biotechnology has made a positive impact on the chemical industry by providing safer, more efficient chemical manufacturing processes that have reduced the use of toxic chemicals, harsh reaction conditions, and expensive metal catalysts, which has improved alignment with the principles of Green Chemistry. The genetically-modified (GM) biocatalysts that are utilized in these processes are typically separated from high-value products and then recycled, or eliminated. Elimination routes include disposal in sanitary landfills, incineration, use as a fuel, animal feed, or reuse as an agricultural soil amendment or other value-added products. Elimination routes that have the potential to impact the food chain or environment have been more heavily scrutinized for the fate and persistence of biological products. In this study, we developed and optimized a method for monitoring the degradation of strain-specific DNA markers from a genetically-modified organism (GMO) used for the commercial production of 1,3-propanediol. Laboratory and field tests showed that a marker for heterologous DNA in the GM organism was no longer detectable by end-point polymerase chain reaction (PCR) after 14 days. The half-life of heterologous DNA was increased by 17% (from 42.4 to 49.7 h) after sterilization of the soil from a field plot, which indicated that abiotic factors were important in degradation of DNA under field conditions. There was no evidence for horizontal transfer of DNA target sequences from the GMO to viable organisms present in the soil. Introduction The industrial fermentation of grains or sugars produces large amounts of microbial biomass as a bioresidual, which is often reused or discarded. This bioresidual is rich in organic material, and contains micro- and macronutrients that are essential in plant growth, making it a value-added product for agriculture and animal production. The use of fermentation bioresiduals from ethanol and biodiesel production as a crop fertilizer or animal feed has been well studied [15, 21, 22, 27, 28, 30]. Many large-scale corn processors, brewers and distillers provide low-cost animal feed products to local farmers, and the use of these products for animal production and crops is well documented after decades of use [25]. With the relatively recent introduction of bacterial metabolic engineering to the industrial chemical production industry, there has been increasing debate over the safety and efficacy of fermentation bioresiduals obtained from GM origin. This debate includes the potential for animal, plant, and aquatic toxicity [2], potential impacts in the human food chain [13], and the potential to adversely impact microbial community structure through recombinant DNA transfer [3]. Field trials, therefore, are necessary to characterize and confirm the viability of these biomaterials as a crop fertilizer. We have recently conducted research field trials that have compared heat-inactivated microbial biomass from the production of 1,3-propanediol to synthetic nitrogen phosphorus and potassium (NPK) crop fertilizers. Of particular interest, was the degradation and fate of recombinant deoxyribonucleic acid (DNA) present in the field-applied microbial biomass. As a chemical compound, eukaryotic DNA present in bone has recently been calculated to have a half-life of 521 years [1], but due to the varying polymeric forms taken by long strand DNA, which is capable of forming numerous three-dimensional structures depending on primary sequence [26], the degradation rates of viable DNA in a wide range of environmental conditions depends on many factors. First, it depends on the chosen definition of “viable” DNA. Outside of a homeostatic cell, a single full chromosome (non-plasmid) of DNA has a rapid degradation rate [6], particularly in an environmental and uncontrolled setting. Strand breaks or cross-links, which would normally be repaired by a living cell, would begin to rapidly accumulate due to environmental catalysis [16–18]. But much shorter strands of DNA, strand lengths on the gene-scale, for instance, could be expected to last much longer before accumulating random damage and much shorter strands even more so. Second, the term half-life (in this context) refers to the amount of time required for a chemical to degrade to half its original concentration, and this typically involves a degradation rate that follows a well characterized model of degradation [1]. Long strand DNA, when present in the environment, is exposed to many stresses outside of the expected, random covalent breakages and rearrangements expected of DNA [6, 7, 9, 11]. Enzymatic, heat-shock, and ultraviolet degradation can all be expected to play a role, which would imply that degradation rates may vary from geographic point A to B depending on weather, climate, and season. Local microbial communities will also play a role in decay rates [8, 11]. We developed and optimized a method for monitoring the degradation of strain-specific DNA markers from a genetically-modified organism (GMO) used for the commercial production of 1,3-propanediol. The persistence of DNA in certain soil types has been characterized before by hybridization and transformation assays [23, 24]. To gain a better insight into the amount of time a functional unit of DNA remained in the soil, both end-point and qPCR were used. Because DNA polymerase relies on intact stretches of DNA for replication, its use for successful target amplification implies the presence of intact, viable fragments. Using these two assays, we were able to track the degradation of strain-specific DNA under field and laboratory conditions, characterizing the decay rates of DNA present in microbial biomass applied to farm fields. Materials and methods Composition and viability analyses of the biomass Field trials utilized heat-inactivated microbial biomass produced from a genetically-modified derivative of Escherichia coli K12 (Genbank Accession number U00096.3) that lacked λ-DNA sequence and F plasmid (F−). Prior to transport, five random 1 kg samples of biomass were collected from a 40 cubic yard commercial waste container fitted with a polyethylene liner (0.162 mm or 6 mil thickness). An equal portion of these samples were combined as a composite sample and chemically analyzed according to U.S. EPA SW 846 standard methods 6010B, 7471A, and 1311. In addition to the chemical analyses, a smaller composite of 10 g was collected under aseptic conditions, diluted in 10 g of sterile water, and subjected to viability measurements by standard plate counting on HiCrome™ ECD Agar with MUG (Product# 09142; Sigma Chemical, St. Louis, MO.). Bacterial colonies with fluorescence under UV light (E. coli positive) were subjected to PCR to determine the identity of viable cells using DNA signatures from the recombinant 1, 3-propanediol production strain. This primer pair targeted a synthetic DNA linker sequence region composed of an E. coli-specific target sequence for the forward primer, and a Saccharomyces cerevisiae chromosome V-specific target sequence (GPP2; dl-glycerol-3-phosphate phosphatase) from the synthetic dha regulon [5] for glycerol biosynthesis in the 1,3-propanediol production strain. The primers used to target this sequence were: F: 5′-GGACCACCGCGCTACTG-3′, R: 5′-GCCGCCGGTTGTAAGATCA-3′. Reactions were performed using GoTaq® Green Mastermix (Promega) at a T m of 50 °C for 30 cycles. Land application was performed with heat-treated biomass that contained no measureable viable counts for E. coli. Field plot layout and biomass application Biomass was applied at five application rates (1.0–5.0 short tonnes per acre; 2.24–11.21 tonnes per hectare) during the month of May in an East Tennessee farm field using a Gehl Scavenger II 322 side-discharge manure spreader on a corn field prior to planting, and a fescue feed. Particle distribution ranged from 2 to 80 mm, with an average of 23 mm. Distribution was intermittently uneven due to aggregate formation, and therefore, soil sampling was performed using 1.0 m2 plots, which were marked with string lines. Composite samples were generating by scraping the surface of the 1.0 m2 plot with a flat shovel to a soil depth of <1 mm, and transferring this soil/residue mixture to a 500 mL glass sample container that was returned to the laboratory on dry ice. Crude, large scale field DNA extractions end-point PCR Soil samples were taken immediately before and following biomass application, as well as every 24 h after application. One sample was taken from a new 1.0 m2 plot every day until end-point PCR no longer indicated the presence of target template DNA. The field soil DNA extraction was performed by resuspending 100 mg of soil sample in 1 mL water in a 1.5 mL centrifuge tube. Samples were vortexed for 1 min and centrifuged at 3000×g., then 400 µL of supernatant was removed, and to it, 250 µL of 10% SDS was added. After vortexing for 30 s, 350 µL of 8 M potassium acetate was added to precipitate sample protein. Flocculent was removed by 16,000×g centrifugation for 10 min, and the supernatant was subjected to a standard phenol extraction followed by an isopropyl alcohol precipitation. PCR primers (as described in the section above) composition, and viability analyses of the biomass were used to identify DNA specific to the recombinant production strain. Reactions were performed using GoTaq® Green Mastermix (Promega) at a T m of 50 °C for 30 cycles. Lab-based soil preparation and DNA extraction Soil was collected from the field trial plot site. Six soil samples of 15 g each were dried down to 10 g in a 45 °C incubator (a 5 g water loss). After drying, larger clumps of soil were ground down into a fine powder. Three samples were autoclaved to remove the soil microbiome, and three samples were not. All six samples were then rehydrated with 5 mL of H2O with the DNA suspension, at a 10−3 dilution from a 70 ng/µl DNA prep. Soil DNA was immediately extracted from each of the six samples using the SurePrep™ Soil DNA Isolation Kit (ThermoFisher). These extractions served as time-point zero samples. Extractions were then performed in technical triplicate each day for 7 days. The DNA extracted served as template for qPCR reactions. Absolute quantification of GMO target sequence by qPCR qPCR experimental optimization was performed targeting a synthetic linker region of the recombinant dha regulon [5] cloned into the E. coli production strain. The DNA from the production strain was prepped and quantified in triplicate using the QIAxpert DNA quantifier (Qiagen). Copy number was determined using the equation m = M/N a, where “m” is the mass of a single copy, “M” is molecular weight of a single copy (in Daltons, based on nucleotide sequence), and N a is Avogadro’s constant (6.022 × 1023 molecules/mole). Serial dilutions were then performed on the prepped plasmid, with the original prep containing 132 ng/µl (2.44 × 1010 copies). A SYBR green standard curve was produced using a Rotor gene-Q 5-Plex HRM cycler (Qiagen), graphing threshold cycle (Y-axis) against log (copy number) (X-axis). This primer pair targeted the synthetic GPP2 gene (dl-glycerol-3-phosphate phosphatase) that is part of the synthetic dha regulon [5] in the recombinant 1,3-propanediol production strain. The primer sequences were: F: 5′-GATCATTGGTATTGCCACTACTTTC-3′, R: 5′-CCGCCAACTCTGATGGATT-3′. Melt curve and trend line analysis confirmed that this primer set amplified a single 96 base pair (bp) amplicon with an amplification efficiency of 99.02%. Using the standard curve trend line equation, C t values from reactions involving plasmid DNA isolated from soil samples were used to calculate copy number. Results and discussion Composition analyses of the biomass Bioresiduals generated from the manufacture of 1,3-propanediol are regulated as industrial solid waste. The chemical profile for these bioresiduals closely resemble nonhazardous municipal biosolids, but in contrast, lack measurable viable bacteria and toxic chemicals as defined by the Resource Conservation and Recovery Act (RCRA; 40 CFR Part 261). Table 1 shows the chemical composition of biomass as applied to research corn and fescue plots at application rates from 2.24 to 11.21 tonnes per hectare, and provides a comparison to regulatory limits that are enforced for industrial waste under U.S. EPA under 40 CFR Part 503-Standards for the Use or Disposal of Sewage Sludge, and the toxicity characteristic leaching procedure (TCLP). Soil tests for the research plots identified the soil as having a loam texture (30% sand, 48% silt, and 22% clay), and recommended an application rate of 201.8 kg per hectare for nitrogen (N), 156.9 kg per hectare for phosphate (P2O5) and 156.9 kg per hectare for potassium (K) to support a yellow dent field corn yield of 370.7–432.4 bushels per hectare. At application rates based on nitrogen sufficiency, the analysis of total-N (Table 1) indicated that an application rate of 3.0 tonnes bioresiduals per hectare would be necessary to meet the nitrogen requirement. However, both phosphate and potassium levels in the bioresiduals were below the recommendations at the 3.0 tonne per hectare rate, and required addition phosphate and potash fertilizers to meet recommended rates. Since over 75% of the nitrogen in the bioresiduals was associated with protein and other complex biomolecules, the N-availability was predicted to follow slow release kinetics, and therefore, higher application rates up to 5.0 tonnes per hectare were tested to evaluate nitrogen sufficiency for plant growth. Although no analytes were detected relating to TCLP regulations, two metals, copper and zinc were detected in the bioresiduals and subject to U.S. EPA 40 CFR Part 503 rules. The copper concentration in the bioresiduals for a dry basis was 142-fold lower than the regulatory threshold, while zinc was 107-fold lower than the regulatory threshold. No soil accumulation issues of these essential plant micronutrients are projected based on the application rates used in this study. Chemical composition of 1,3-propanediol bioresiduals compared to regulatory limits, U.S. EPA 40 CFR Part 503 and toxicity characteristic leaching procedure (TCLP) for solid waste Analyte . Units . Minimum detection level, as-produced . Bioresiduals composition, as-applied . Bioresiduals composition, dry basis . TCLP rule, (mg/kga) . 40 CFR part 503 rulea, (mg/kg) . Solids % – 62.2 ± 0.1 100.0 – – Moisture % 0.1 37.8 ± 0.1 0.0 – – Carbon, combustion % 0.01 32.9 ± 0.1 52.9 ± 0.2 – – Hydrogen, combustion % 0.01 8.3 ± 0.2 13.3 ± 0.3 – – Nitrogen, combustion % 0.01 6.71 ± 0.06 10.79 ± 0.10 – – Ammonia (NH3) % 0.01 1.38 ± 0.03 2.22 ± 0.05 – – Nitrate mg/kg 0.55 12.0 ± 0.04 19.3 ± 0.06 – – Sulfur, combustion % 0.01 0.95 ± 0.1 1.53 ± 0.16 – – DNA % 0.0001 1.41 ± 0.13 2.27 ± 0.21 – – Phosphorus % 0.004 0.492 ± 0.093 0.791 ± 0.150 – – Potassium % 0.04 0.257 ± 0.01 0.413 ± 0.016 – – Magnesium % 0.005 0.108 ± 0.003 0.174 ± 0.005 – – Calcium mg/kg 13.4 222 ± 0.11 357 ± 0.18 – – Sodium % 0.002 0.111 ± 0.006 0.178 ± 0.010 – – Chloride mg/kg 23.4 88 ± 5.0 142 ± 8.0 – – Arsenic mg/kg 1.00 ndb ndb 5.0 41 Barium mg/kg 1.00 nd nd 100.0 – Cadmium mg/kg 0.672 nd nd 1.0 39 Cobalt mg/kg 0.93 1.87 ± 0.09 3.01 ± 0.15 – – Copper mg/kg 0.93 6.5 ± 0.2 10.5 ± 0.3 – 1500 Chromium mg/kg 1.00 nd nd 5.0 – Lead mg/kg 1.00 nd nd 5.0 300 Mercury CVAA mg/kg 0.0093 nd nd 0.2 17 Nickel mg/kg 0.90 nd nd – 420 Selenium mg/kg 1.00 nd nd 1.0 100 Silver mg/kg 5.00 nd nd 5.0 – Zinc mg/kg 0.34 16.2 ± 0.4 26.1 ± 0.6 – 2800 Analyte . Units . Minimum detection level, as-produced . Bioresiduals composition, as-applied . Bioresiduals composition, dry basis . TCLP rule, (mg/kga) . 40 CFR part 503 rulea, (mg/kg) . Solids % – 62.2 ± 0.1 100.0 – – Moisture % 0.1 37.8 ± 0.1 0.0 – – Carbon, combustion % 0.01 32.9 ± 0.1 52.9 ± 0.2 – – Hydrogen, combustion % 0.01 8.3 ± 0.2 13.3 ± 0.3 – – Nitrogen, combustion % 0.01 6.71 ± 0.06 10.79 ± 0.10 – – Ammonia (NH3) % 0.01 1.38 ± 0.03 2.22 ± 0.05 – – Nitrate mg/kg 0.55 12.0 ± 0.04 19.3 ± 0.06 – – Sulfur, combustion % 0.01 0.95 ± 0.1 1.53 ± 0.16 – – DNA % 0.0001 1.41 ± 0.13 2.27 ± 0.21 – – Phosphorus % 0.004 0.492 ± 0.093 0.791 ± 0.150 – – Potassium % 0.04 0.257 ± 0.01 0.413 ± 0.016 – – Magnesium % 0.005 0.108 ± 0.003 0.174 ± 0.005 – – Calcium mg/kg 13.4 222 ± 0.11 357 ± 0.18 – – Sodium % 0.002 0.111 ± 0.006 0.178 ± 0.010 – – Chloride mg/kg 23.4 88 ± 5.0 142 ± 8.0 – – Arsenic mg/kg 1.00 ndb ndb 5.0 41 Barium mg/kg 1.00 nd nd 100.0 – Cadmium mg/kg 0.672 nd nd 1.0 39 Cobalt mg/kg 0.93 1.87 ± 0.09 3.01 ± 0.15 – – Copper mg/kg 0.93 6.5 ± 0.2 10.5 ± 0.3 – 1500 Chromium mg/kg 1.00 nd nd 5.0 – Lead mg/kg 1.00 nd nd 5.0 300 Mercury CVAA mg/kg 0.0093 nd nd 0.2 17 Nickel mg/kg 0.90 nd nd – 420 Selenium mg/kg 1.00 nd nd 1.0 100 Silver mg/kg 5.00 nd nd 5.0 – Zinc mg/kg 0.34 16.2 ± 0.4 26.1 ± 0.6 – 2800 Values are reported as the average ± the standard deviation for three individual measurements aLimit value reported as dry weight for the monthly average bNot detected Open in new tab Chemical composition of 1,3-propanediol bioresiduals compared to regulatory limits, U.S. EPA 40 CFR Part 503 and toxicity characteristic leaching procedure (TCLP) for solid waste Analyte . Units . Minimum detection level, as-produced . Bioresiduals composition, as-applied . Bioresiduals composition, dry basis . TCLP rule, (mg/kga) . 40 CFR part 503 rulea, (mg/kg) . Solids % – 62.2 ± 0.1 100.0 – – Moisture % 0.1 37.8 ± 0.1 0.0 – – Carbon, combustion % 0.01 32.9 ± 0.1 52.9 ± 0.2 – – Hydrogen, combustion % 0.01 8.3 ± 0.2 13.3 ± 0.3 – – Nitrogen, combustion % 0.01 6.71 ± 0.06 10.79 ± 0.10 – – Ammonia (NH3) % 0.01 1.38 ± 0.03 2.22 ± 0.05 – – Nitrate mg/kg 0.55 12.0 ± 0.04 19.3 ± 0.06 – – Sulfur, combustion % 0.01 0.95 ± 0.1 1.53 ± 0.16 – – DNA % 0.0001 1.41 ± 0.13 2.27 ± 0.21 – – Phosphorus % 0.004 0.492 ± 0.093 0.791 ± 0.150 – – Potassium % 0.04 0.257 ± 0.01 0.413 ± 0.016 – – Magnesium % 0.005 0.108 ± 0.003 0.174 ± 0.005 – – Calcium mg/kg 13.4 222 ± 0.11 357 ± 0.18 – – Sodium % 0.002 0.111 ± 0.006 0.178 ± 0.010 – – Chloride mg/kg 23.4 88 ± 5.0 142 ± 8.0 – – Arsenic mg/kg 1.00 ndb ndb 5.0 41 Barium mg/kg 1.00 nd nd 100.0 – Cadmium mg/kg 0.672 nd nd 1.0 39 Cobalt mg/kg 0.93 1.87 ± 0.09 3.01 ± 0.15 – – Copper mg/kg 0.93 6.5 ± 0.2 10.5 ± 0.3 – 1500 Chromium mg/kg 1.00 nd nd 5.0 – Lead mg/kg 1.00 nd nd 5.0 300 Mercury CVAA mg/kg 0.0093 nd nd 0.2 17 Nickel mg/kg 0.90 nd nd – 420 Selenium mg/kg 1.00 nd nd 1.0 100 Silver mg/kg 5.00 nd nd 5.0 – Zinc mg/kg 0.34 16.2 ± 0.4 26.1 ± 0.6 – 2800 Analyte . Units . Minimum detection level, as-produced . Bioresiduals composition, as-applied . Bioresiduals composition, dry basis . TCLP rule, (mg/kga) . 40 CFR part 503 rulea, (mg/kg) . Solids % – 62.2 ± 0.1 100.0 – – Moisture % 0.1 37.8 ± 0.1 0.0 – – Carbon, combustion % 0.01 32.9 ± 0.1 52.9 ± 0.2 – – Hydrogen, combustion % 0.01 8.3 ± 0.2 13.3 ± 0.3 – – Nitrogen, combustion % 0.01 6.71 ± 0.06 10.79 ± 0.10 – – Ammonia (NH3) % 0.01 1.38 ± 0.03 2.22 ± 0.05 – – Nitrate mg/kg 0.55 12.0 ± 0.04 19.3 ± 0.06 – – Sulfur, combustion % 0.01 0.95 ± 0.1 1.53 ± 0.16 – – DNA % 0.0001 1.41 ± 0.13 2.27 ± 0.21 – – Phosphorus % 0.004 0.492 ± 0.093 0.791 ± 0.150 – – Potassium % 0.04 0.257 ± 0.01 0.413 ± 0.016 – – Magnesium % 0.005 0.108 ± 0.003 0.174 ± 0.005 – – Calcium mg/kg 13.4 222 ± 0.11 357 ± 0.18 – – Sodium % 0.002 0.111 ± 0.006 0.178 ± 0.010 – – Chloride mg/kg 23.4 88 ± 5.0 142 ± 8.0 – – Arsenic mg/kg 1.00 ndb ndb 5.0 41 Barium mg/kg 1.00 nd nd 100.0 – Cadmium mg/kg 0.672 nd nd 1.0 39 Cobalt mg/kg 0.93 1.87 ± 0.09 3.01 ± 0.15 – – Copper mg/kg 0.93 6.5 ± 0.2 10.5 ± 0.3 – 1500 Chromium mg/kg 1.00 nd nd 5.0 – Lead mg/kg 1.00 nd nd 5.0 300 Mercury CVAA mg/kg 0.0093 nd nd 0.2 17 Nickel mg/kg 0.90 nd nd – 420 Selenium mg/kg 1.00 nd nd 1.0 100 Silver mg/kg 5.00 nd nd 5.0 – Zinc mg/kg 0.34 16.2 ± 0.4 26.1 ± 0.6 – 2800 Values are reported as the average ± the standard deviation for three individual measurements aLimit value reported as dry weight for the monthly average bNot detected Open in new tab DNA degradation rates for field conditions In this study, we have characterized the degradation rates of DNA target sequences from a strain of E. coli used in the manufacture of 1,3-propanediol. Biomass was applied directly to soil on a local East Tennessee corn farm, and DNA persistence was measured directly by end-point PCR. In the field, biomass and DNA degradation is driven by a large number of biotic and abiotic factors, including heat, moisture, UV light, nonenzymatic catalytic degradation, and microbe-mediated degradation. The polymerase chain reaction assay is known to be highly sensitive to very small amounts of template DNA [10], theoretically being capable of amplifying a single copy of target. Figure 1 shows the number of days post-application that target DNA was no longer detectable by the sensitive PCR assay. This data provides a sense of the longest possible amount of time, based on application rate, that production strain DNA was present in the soil samples collected from the field. Intuitively, band strength strongly correlates with biomass application rates, with the 5 tonnes per hectare rates showing the highest intensity. There is also a clear decline in band intensity over time, finally leading to no detectable DNA template after 14 days. Because of the wide array of potential degradation pathways present in the environment and soil for a molecule as large and complex as DNA, it is difficult to claim to have developed a general half-life for all scenarios. End-point PCR is also not conducive to target strand quantitation [12]. With all this being said, it has been clearly displayed that DNA applied to this field of sandy loam soil is degraded beyond PCR detectability (30 cycles) after a period of 14 days (day 14 was the first of 3 consecutive days without banding, data from redundant screening days not included). The target sequence is relatively short (321 bp), increasing the probability of its detection relative to that of a larger, functional sequence on the scope of a gene or chromosome [31]. Therefore, it can be implied that sequences of significantly longer length, which are more ecologically and proprietarily relevant, would have degraded beyond detectability well within the 14 day window. While this data is in contrast to previously reported data [24], it is important to note that these samples were exposed to outdoor field conditions in East Tennessee during the month of May, and not lab conditions. While weather data was not collected, average yearly temperature databases will indicate high temperature, humidity, and UV levels at this time of the year, which is more than likely the root of the deviation. In the context of the purpose of this study, we believe in-field sample collection was a necessary aspect of thoroughly characterizing this process. Fig. 1 Open in new tabDownload slide End-point PCR monitoring of DNA degradation after field application. Application rates (1–5 tonnes per acre) indicated by first number, application rate sub-plots indicated by second number. Negative control (−C) is a sample extracted from the water used in the DNA extraction process. Positive control (+C) is a DNA extraction sample from live production strain. a Samples taken from each sub-plot prior to biomass application. b Samples taken from each sub-plot immediately following biomass application. c Samples taken on day 7. d Samples taken on day 14 DNA degradation rates for laboratory-generated conditions To develop a more accurate description of degradation rates, a more controlled environment was necessary. In-house soil samples (autoclaved or not) were mixed with cleanly prepped production strain plasmid DNA, and degradation was measured over the course of 8 days. Laboratory studies were conducted to evaluate field associated variation in critical variables, including temperature, solar irradiance, moisture, and biotic factors. Autoclaved and non-autoclaved samples were compared to characterize rates of microbial degradation of applied DNA (Fig. 2). qPCR also requires a much higher degree of purity in order for data to be used for quantification. Therefore, extractions were performed using a soil DNA extraction kit to ensure accuracy, efficiency, and repeatability. A standard curve was first prepared for absolute quantification [4, 14, 20, 29]. A log transformation of the standard curve created allowed for plasmid copy number to be calculated directly from threshold cycle (C t). When plotted against time, a significant decrease in plasmid number per day is evident, with average copy numbers decreasing by over a fold in both of the first 2 days. This trend finally reduced in slope after the first 2 days for both sterile and non-sterile samples. DNA adsorption to soil particulate has been demonstrated in the past [11, 19, 23, 24] and this reduction in degradation rates could be an indication of DNA not immediately associated with the protection of soil adsorption from the beginning being degraded more rapidly than that which immediately bound a protective surface. Fig. 2 Open in new tabDownload slide qPCR used to determine half-life of DNA under lab conditions. a Serial dilutions were performed to determine amplification efficiency of the primers. The equation used to determine the efficiency of 99.02% is included. Melt curve analysis indicates a single peak, and therefore a single amplicon. b Log transformation of the serial dilutions and standard curve to determine the copy number present in each reaction sample by Ct. c The period of time of extraction graphed against each measured C t (Lighter squares are sterile soil, and darker diamonds are non-sterile soil). d The period of time graphed against the calculated copy number from each given C t. The fluorescence signals of the non-sterile sample e and the sterile sample f The average copy numbers calculated from the technical triplicates can be used to calculate the half-life of DNA decay in the two different soil environments using the following formula, where h = half-life, t = time in hours, A = final copy number after 7 days, A o = initial copy number immediately after mixing soil and DNA: h=[-tLn2]/LnAAo $$h = [\left( { - t} \right){\text{Ln}}\left( 2 \right)]/\left[ {{\text{Ln}}\left( {\frac{A}{{A_{\text{o}} }}} \right)} \right]$$ The half-lives are listed in Table 1. As was expected, the half-life of DNA in non-sterile soil (42.40 h) is shorter than that in sterile soil (49.67 h), likely due to the presence of nuclease excreting microbes. But the difference in half-life between sterile and non-sterile is much smaller than initially expected, suggesting that microbial degradation of DNA in soil plays a much smaller role in overall degradation than expected (Table 2). The rate limiting step, therefore, seems to be nonenzymatic catalysis and random bond rearrangement/breakage. The variable used in the listed equation for half-life determination, as determined from the C t values Property . Sterile soil . Native soil . Final copy number (A) 210,881 ± 5483 142,054 ± 4260 Initial copy number (A 0) 1,572,841 ± 62,919 1,495,217 ± 31,340 Time in hours (t) 144 144 Half-life (h) 49.7 42.4 Property . Sterile soil . Native soil . Final copy number (A) 210,881 ± 5483 142,054 ± 4260 Initial copy number (A 0) 1,572,841 ± 62,919 1,495,217 ± 31,340 Time in hours (t) 144 144 Half-life (h) 49.7 42.4 Open in new tab The variable used in the listed equation for half-life determination, as determined from the C t values Property . Sterile soil . Native soil . Final copy number (A) 210,881 ± 5483 142,054 ± 4260 Initial copy number (A 0) 1,572,841 ± 62,919 1,495,217 ± 31,340 Time in hours (t) 144 144 Half-life (h) 49.7 42.4 Property . Sterile soil . Native soil . Final copy number (A) 210,881 ± 5483 142,054 ± 4260 Initial copy number (A 0) 1,572,841 ± 62,919 1,495,217 ± 31,340 Time in hours (t) 144 144 Half-life (h) 49.7 42.4 Open in new tab Conclusions Here we have characterized the degradation rates of strain-specific DNA present in biomass from a commercial fermentation process for production of 1,3-propanediol, under laboratory and field conditions. The use of microbial fermentation bioresiduals as an agricultural soil amendment has many advantages including: (1) the potential to improve the life cycle assessment for the biomanufacturing process through capture of biomass carbon from the biomass into soil organic matter and plants; (2) the potential for a more “use the whole buffalo” approach to industrial chemical production, with viability and ecological impacts first being considered. For production companies, the presence of intellectually sensitive DNA sequences in production biomass waste presents a vested interest in ensuring degradation rates are not significantly impacted by the specific molecular profiles of a process-specific biomass waste product. To this end, DNA persistence in the field was monitored by end-point PCR. At multiple application rates, ranging from 1 to 5 tonnes per hectare, it is clear that full degradation of relatively small (321 bp) DNA fragments takes place within 14 days of application. It is also clear that the DNA is not undergoing horizontal transfer within the local, sampled microbial community, as any DNA maintenance would be reflected by persistent banding. The half-life of DNA in sterile and non-sterile soil from the same farm was also measured under controlled lab conditions. Production strain DNA had a half-life of 42.40 h in non-sterile (non-autoclaved) soil, while in sterile soil had a half-life of 49.67 h. Previous studies have indicated much shorter half-lives (28.2, 15.1, and 9.1 h for three different soil types), but these half-lives were based on the transformability of full plasmids extracted from soil [24]. Here, we used qPCR to measure the presence of much shorter fragments of DNA than a full, circular, functional plasmid. It is not surprising, therefore, that the half-lives calculated here are significantly larger, as it will take a ~100 bp fragment much more time to degrade beyond function than it will a circular plasmid of multiple kilobases to degrade beyond function or transformability. This data serves well to begin to characterize the molecular dynamics of industrial fermentation biomass waste applied to agricultural plots as a fertilizer replacement or supplement. Acknowledgements The authors would like to thank the Tennessee Department of Agriculture and the University of Tennessee for assistance in conducting these research field trials. References 1. 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TI - Degradation and half-life of DNA present in biomass from a genetically-modified organism during land application
JF - Journal of Industrial Microbiology and Biotechnology
DO - 10.1007/s10295-016-1876-x
DA - 2017-02-01
UR - https://www.deepdyve.com/lp/oxford-university-press/degradation-and-half-life-of-dna-present-in-biomass-from-a-genetically-05V7sPXiSV
SP - 213
EP - 220
VL - 44
IS - 2
DP - DeepDyve
ER -