Environmental Toxicology and Chemistry, Vol. 33, No. 9, pp. 1996–2003, 2014 # 2014 SETAC Printed in the USA
ACTIVITY AND ECOLOGICAL IMPLICATIONS OF MAIZE-EXPRESSED TRANSGENIC ENDO-1,4-b-D-GLUCANASE IN AGRICULTURAL SOILS ADAM J. KENNY and JEFFREY D. WOLT* Department of Agronomy, Iowa State University, Ames, Iowa, United States (Submitted 24 March 2014; Returned for Revision 14 May 2014; Accepted 23 May 2014) Abstract: Plant expression of thermostable endoglucanase (E1) has been proposed for improved conversion of lignocellulose to ethanol for fuel production. Residues of E1-expressing maize may affect ecological services (e.g., C mineralization and biogeochemical cycling) on soils where they occur. Therefore, the activity of residual E1 was investigated using soils amended with bacterial and plant-solubilized E1 compared with soil endogenous activity and residual activity from a mesostable cellulase (Aspergillus and Trichoderma spp.). An optimized analytical method involving a carboxymethyl cellulose substrate and dinitrosalicylic acid detection effectively assayed endoglucanase activity in amended and unamended soils and was used for determining E1 activity in 3 representative soils. The effect of E1 on soil carbon mineralization was determined by comparing CO2 evolution from soils amended with transgenic E1-expressing and wild-type maize tissue. Extraction and recovery of the mesostable comparator, bacterial E1, and plant-soluble E1 showed nearly complete loss of exogenous endoglucanase activity within a 24-h period. Carbon mineralization indicated no significant difference between soils amended with either the transgenic E1 or wild-type maize tissue. These results indicate that maize residues expressing up to 30 mg E1/g tissue negligibly affect soil endoglucanase activity and CO2 respiration for representative soils where transgenic E1 maize may be grown. Environ Toxicol Chem 2014;33:1996–2003. # 2014 SETAC Keywords: Acidothermus cellulolyticus
Genetically engineered
Dissipation
Thermostable
Cellulase
Endoglucanase
One strategy to reduce costs for cellulosic ethanol production is to genetically transform the source biofeedstock for better utilization of the cellulosic material [6,7]. The genetic engineering of bioenergy feedstocks has been explored for a variety of cellulosic crops—Zea mays (maize), Panicum virgatum (switchgrass), and Miscanthus spp.—to improve processing through in planta expression of bioprocessing enzymes [8–13] and through altering the lignin content and composition [14–17]. Improved cellulose conversion efficiencies for maize stover can be achieved through in planta production of bioprocessing enzymes so that, following biomass pretreatment and cell lysis, plant-made cellulases become active and cleave cellulosic polymers into fermentable sugars. A case in point is highly thermostable endo-1,4-b-D-glucanase from Acidothermus cellulolyticus (E1; pdb:1ECE) that has been expressed at high levels in maize green tissue [13]. With an optimal temperature of 83 8C and with marked stability in both acidic and basic environments, E1-endoglucanase has been an attractive target for expression in cellulosic biofeedstock crops [1,18,19]. We estimate that where genetically engineered maize expressing E1 is grown, it will add 60 g ha1 to 120 g ha1 annually of E1 in postharvest residues to receiving soils, depending on the nature of expression (Table 1). This stands in stark contrast to typical agroecosystems in which plants are considered minor contributors to soil enzyme activity through residue decomposition and root exudation [20]. Therefore, widescale agricultural release of E1-expressing maize will require environmental assessment to better understand the potential impact of residues left in the field [21]. Especially important in this regard is the evaluation of effects on ecosystem services (e.g., carbon mineralization and biogeochemical cycles). It has been well documented that enzymes in the soil environment are direct biological instigators of elemental cycling and indicators of overall soil tilth [22–26]. The importance of soil glycoside
INTRODUCTION
Maize cellulosic residues represent a major potential biofeedstock source for biofuel production [1,2]. Current ethanol production strategies using cellulosic feedstocks are inefficient because, in order for carbon polymer sugars to become available for processing into ethanol, the biofeedstock must undergo dilute acid pretreatment at temperatures generally in excess of 75 8C to eradicate the recalcitrant lignin and hemicellulose layers that shield the carbon substrate. Pretreatment is followed with exogenous cellulase addition to release fermentable glucose sugars. Cellulases are biocatalytic systems made up of endoglucanases (Enzyme Commission number 3.2.1.4), exoglucanases (Enzyme Commission number 3.2.1.91), and cellobiases (Enzyme Commission number 3.2.1.21) that work synergistically to depolymerize cellulose. The currently accepted mechanism of cellulase action begins with endoglucanase hydrolyzing internal 1,4 glycosidic bonds within amorphous regions to produce new chain ends. Exoglucanase (e.g., cellobiohydrolase) attacks chain ends to yield glucose or cellobiose. Cellobiase completes any residual breakdown needed and prevents product inhibition from cellobiose [3]. Thermostability of the added cellulases is highly desired in the biofuel conversion process because these enzymes are subjected to high temperatures and variable pH. Additionally, saccharification at higher temperatures can prevent batch contamination, increase solution viscosity for improved mixing, and better solubilize cellulose substrates [4]. Addition of exogenous cellulase enzymes after pretreatment accounts for approximately one-third to one-half of the entire production cost for cellulosederived ethanol [5]. * Address correspondence to [email protected] Published online 26 May 2014 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/etc.2645 1996
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Table 1. Estimated environmental concentration (EEC) for thermostable endoglucanase (E1) in soil from postharvest residues of E1-expressing maize Assumptions
Rationale
11.4 Mg grain ha1 1:1 grain to stover biomass ratio 0.33 postharvest fraction of residual biomass left on field 3.8 Mg residual biomass ha1 (b) 2 103 Mg soil ha1 (m) 17–33 g Mg1 residual biomass (a) EEC ¼ ab/m ¼ 0.032–0.063 g E1 Mg1 soil 60–120 g E1 ha1
Iowa average maize yield, 2009 (National Agricultural Statistics Service [42]) Perlack et al. [43] For erosion control and nutrient sustainability (Graham et al. [44]) Incorporation to 15 cm depth, typical soil bulk density 1.33 g cm3 (White [45], p. 73) Range in mitochondrial expression in green plant tissue (Mei et al. [13])
hydrolases cannot be overemphasized because the soil carbon pool is more than 3 times that of the atmosphere pool and 4.5 times that of the biotic pool [27]. Furthermore, E1, which was originally isolated from hot springs, has thermostable properties that may confer increased stability when released to agricultural soils during postharvest residue decomposition [28]. The novel expression, release to a foreign environment, and potential effects to ecosystem services require better knowledge of E1 fate in agricultural soil. Enzyme function and stability have been linked to numerous soil physicochemical properties, especially clay and organic matter content. Clay and humic substances are thought to protect enzymes/proteins from degradation because of their enhanced stability via steric hindrance and conformational changes on organomineral sorption that make them less susceptible to proteolysis and physicochemical denaturing [29]. Ironically, the sorption process can also alter the tertiary structure of proteins to the extent that they are no longer biologically active; however, there is also evidence that activity may be retained after sorption. Regardless of the fate of enzymes functionality, once adsorbed to soil particles they are generally considered to be irreversibly bound [29]. As a first step toward understanding residue effects of E1expressing maize, we discriminated activity of endoglucanases from various sources (bacterial E1, E1-maize soluble protein, endogenous soil enzymes, and mesostable cellulase) when amended to representative agricultural soils. In addition, carbon dioxide (CO2) evolution was used to compare whole-soil respiration rates for unamended soils and for soils amended with either transgenic (E1) or nontransgenic (wild-type) maize tissue. The CO2 evolution study provides indirect data on activity from whole tissue–derived E1 that has been amended to soils and indicates whether E1 presence in soil affects mineralization of labile carbon. MATERIALS AND METHODS
Test materials
Bacterial E1 from Streptomyces lividans was provided by W.S. Adney (National Renewable Energy Laboratory). Soluble bacterial E1 preparations were used for all analyses. The mesostable cellulase comparator (Aspergillus and Trichoderma spp.) was obtained from MP Biomedical. The bacterial E1-endoglucanase and mesostable comparator enzymes were characterized to determine the temperature and pH optima using the carboxymethyl cellulose and dinitrosalicylic acid analytical technique (see Endoglucanase assay). Dry tissue (leaves plus stems) of transformed (E1) and untransformed (wild-type) maize (V5 stage) was obtained from M. Sticklen (Michigan State University). Transgenic tissues were from T2 maize representing a cross between mitochondria l expressing E1-endoglucanase lines (1.5-E1-10b 1.5-E1-10c)
[13]. Nontransgenic wild type near isoline maize grown under the same conditions as the E1 maize was used as a control. On receipt, tissues were pulverized in liquid nitrogen, screened to pass 1.7-mm mesh openings, and stored at –20 8C until used. Total carbon and nitrogen for subsamples was determined by combustion and infrared detection. Plant total soluble protein was obtained by genogrinding (SPEX CertiPrep) 100 mg of the previously milled tissue in 1 mL of extraction buffer (50 mM sodium acetate [NaAc], 10 mM ethylenediaminetetraacetic acid, 0.1% Triton X-100, pH 5). Crude soluble extracts were centrifuged, and an aliquot was subjected to ammonium-sulfate precipitation using an initial saturation concentration of 100% (0 8C). Crude extracts and ammonium sulfate solution were combined at 0.4:1 ratio (70% final saturation concentration), briefly vortexed, and then stored in a –20 8C freezer for 1 h before centrifugation at 20 800 g for 5 min. The protein pellet formed was reconstituted to an equivalent of 0.2 g original tissue/mL. Activity of plant-soluble protein from E1 and wild-type maize was determined and compared against activity from E1 standards to estimate expression levels (see Endoglucanase assay). All stock enzyme materials were stored concentrated in 1% bovine serum albumin and 50 mM NaAc (pH 5) in polypropylene vials. Test systems
All preliminary work was performed using an air-dried Nicollet loam surface soil (superactive, mesic Aquic Hapludolls). Definitive studies utilized surface soils from the Hanlon– Spillville–Coland alluvial catena (superactive, mesic Cumulic Hapludolls) that were collected fresh, immediately sieved to pass 2-mm mesh openings, and maintained field-moist at a workable consistency in the dark at 4 8C prior to use. These soils were used within 60 d of sampling. Physicochemical characteristics of the test soils are summarized in Table 2. Endoglucanase assay
A number of optimization trials were undertaken to evaluate extraction and detection procedures to allow for discrimination of endoglucanase activity from test materials and test systems (A.J. Kenny, 2012, Master’s thesis). Endoglucanase activity was determined through assay of glucose release from carboxymethyl cellulose substrate [13,18,30,31]. Glucose monomers released by carboxymethyl cellulose hydrolysis from endoglucanase were directly measured using a reducing sugar assay solution [32]. For characterization of test materials, dinitrosalicylic acid reagent solution was mixed 1:1 (400 mL total) with 2% carboxymethyl cellulose solution (pH 5) and heated to 95 8C for 5 min. The colored solution was centrifuged at 20 800 g for 1 min, and a 200-mL aliquot was immediately measured and compared with control samples at 540 nm in microplate wells using a BioTek PowerWave spectrophotometer. Sodium azide (NaN3) was added as a microbial inhibitor during the hydrolysis
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A.J. Kenny and J.D. Wolt Table 2. Physicochemical properties of test soils Water content (%)d
Texture (%) Series Nicollet Hanlon Spillville Coland
Textural class
Sand
Silt
Clay
Organic matter (%)a
pHb
CECc (cmol kg1)
33 kPa
100 kPa
1500 kPa
Loam Sandy loam Silt loam Loam
48 56 24 34
34 28 50 40
18 16 26 26
2.9 1.9 3.4 3.7
6.7 7.6 6.8 7.0
16.8 14.7 22.1 23.9
16.5 14.2 24.7 23.4
— 12.6 17.3 18.1
11.52 8.61 14.73 15.69
a
Loss on ignition. Soil–water extract, 1:1. c Cation exchange capacity (CEC), sodium acetate method (US Environmental Protection Agency 9081). d Pressure plate measurements for 33 kPa and 1500 kPa, 100 kPa estimated using pedotransfer functions (van Genuchten [46]). b
phase of the assay [33] to allow for accumulation of released glucose. For soil assays, a 0.5-g (oven-dry equivalent) soil sample was extracted for 5 min using bead-beating (genogrinding at 100 strokes min1) in the presence of 600 mL of the extraction buffer (50 mM NaAc, 10 mM ethylenediaminetetraacetic acid, 0.01% Triton X-100, pH 5 [34]) and then centrifuged for 5 min at 20 800 g. Aliquots of the supernatant (300 mL) were incubated with 300 mL of 2% carboxymethyl cellulose substrate solution containing 0.05% NaN3. Incubation was conducted at pH 5 and temperatures of 40 8C or 65 8C for the mesostable comparator and E1, respectively, based on the previously determined pH and temperature optima. Recovery of accumulated glucose was determined with the dinitrosalicylic acid reagent using a 200-mL aliquot of the reaction solution, 200 mL of dinitrosalicylic acid reagent, incubating at 95 8C for 5 min, and measuring spectrophotometrically at 540 nm. E1 activity in amended soils
Soil microcosm studies were used to determine exogenous enzyme activity. Test systems utilized 0.5-g soil samples (ovendry equivalent) in 2-mL microcentrifuge tubes. Test materials (enzymes) were diluted from stock and applied to surface soil samples at environmentally relevant concentrations. Soils were amended with 300 ng/g, 60 ng/g, and 80 ng/g oven dry equivalent soil of the mesostable comparator, bacterial E1, and plant-soluble E1, respectively. These levels of fortification were previously determined to represent equivalent amounts of source enzyme activity so that each treatment hydrolyzed carboxymethyl cellulose to release glucose at relatively equal rates. Plant-soluble E1 represented a loading equivalent to 160 g ha1, only slightly above the high-end estimate of E1 that would be released to the soil environment under typical conditions (Table 1). Preliminary studies used the air-dried Nicollet soil and mesostable comparator enzyme and were confirmed using bacterial and plant-soluble E1. For definitive studies, field-moist Hanlon, Spillville, and Coland soils were fortified with either bacterial or plant-soluble E1, while control soils received no enzyme. Following fortification, soils were mixed by spatula, moisture was adjusted with deionized water, and the samples were incubated aerated in the dark at a constant temperature (Ambient Temperature Incubator BOD50A16; Thermo-Scientific). For definitive studies, the base water potential and temperature used were 100 kPa and 25 8C, respectively. The 100 kPa moisture for the Hanlon, Spillville, and Coland soils was 12.6%, 17.3%, and 18.1%, respectively (Table 2). Each study included 6 sampling time points extending over approximately 2.5 half-lives of the test materials as determined on the basis of the preliminary study. At each sampling time
triplicate vials for each treatment were sampled for analysis of enzyme activity. CO2 evolution
Mineralization effects from E1-endoglucanase amended to field-moist Hanlon, Spillville, and Coland soils were determined for soils treated with E1 or wild-type maize tissue or unamended using a fully factorial design in 3 replications. For amended samples, 20 mg of ground tissue was mixed into 10 g soil (ovendry equivalent), representing an environmental loading of 120 g E1 ha1, whereas unamended samples were simply 10 g soil. Amended soils were added to 120-mL bottles, adjusted to 100 kPa water potential with the addition of deionized water, sealed with rubber stoppers, and incubated at 25 8C. Gas samples were analyzed with a LI-7000 CO2/H2O LI-COR infrared gas analyzer (LI-COR Biotechnology). At each sampling, stoppers were pierced with a syringe, and headspace was well mixed before 0.4 mL of gas was withdrawn and injected into the LI-COR analyzer, which had been previously calibrated against CO2 standards. Rubber stoppers were then removed, and headspace was replaced with laboratory air under slight positive pressure. Stoppers were replaced, and a second 0.4-mL aliquot was withdrawn for analysis. Change in CO2 was standardized for the 2 samples to reflect CO2 released per gram of soil. Samples were taken 14 times during the 45-d duration of the study. Instrumentation was calibrated before every use, and soil moisture was readjusted to 100 kPa twice over the course of the study. Statistical analysis
Comparisons presented graphically throughout represent the 95% confidence intervals calculated on the basis of triplicate analysis, unless otherwise indicated. An empirical biphasic decay model, f(x) ¼ ae–bx þ ce–dx, was used to characterize the change in extractable enzyme activity with time [35], where the loss of activity, f(x), over time, x in hours, was determined by the 2 apparent first-order rate constants b and d, describing loss from 2 differentially acting pools of enzyme, a and c, respectively. Data fitting used SigmaPlot ver 10 software (Systat Software), where the relative percentages of a and c were constrained such that c ¼ 100 a, whereas b and d were constrained to values >0. For CO2 evolution studies, a least squares full factorial method was used to examine interactions between variables (time, replication, treatment, and soil). All cross-terms indicated significant interactions; therefore, Tukey’s honestly significant difference test was applied to determine significant effects of treatment soil at the a ¼ 0.05 level using JMP V.10 (SAS).
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Environ Toxicol Chem 33, 2014
A
retained high activity across a much broader range (pH 5–9; Figure 1B). Both E1 and wild-type maize tissue had similar C:N ratios, but total protein content was higher for the E1 tissue, whereas soluble protein content was higher for the wild-type compared with the E1 tissue (Table 3). The differences in total and soluble protein concentrations were slight and, given the favorable C:N ratios, would not markedly contribute to differences in mineralization. Transgenic maize was positive for E1 activity; expression was 0.55% of total soluble protein, representing 5 mg g1 tissue. Wild-type maize, however, exhibited no measurable E1 activity (Table 3). Combining the present expression data with that of Mei and coworkers [13], a refined range for the estimated environmental concentration of 10 mg E1 g1 to 60 mg E1 g1 soil (17–120 g ha1 soil loading) was calculated (Table 1).
Enzyme activity (relative %)
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Q10 = 1.9
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Figure 1. Enzyme activity of bacterial sources of thermostable endoglucanase (E1) and the mesostable comparator as affected by (A) temperature and (B) pH. Each data point represents the average of triplicates 95% confidence interval. Q10 ¼ temperature coefficient.
RESULTS
Enzyme characterization
When incubated at pH 5, bacterial E1 exhibited steadily increasing activity when measured over a temperature range of 15 8C to 60 8C, consistent with the reported temperature optima of 83 8C [30]. The mesostable comparator exhibited a temperature optimum of 40 8C under similar conditions (Figure 1A). Relative to the mesostable comparator, the bacterial E1 had a substantially lower activity at temperatures reasonably anticipated for surface soil under maize residue cover [36]. Both enzymes displayed optimal activity at pH 5 when determined at temperature optima of 40 8C and 65 8C for the mesostable comparator and bacterial E1, respectively; however, bacterial E1
In preliminary studies in which soluble enzyme test materials were amended to the air-dried Nicollet soil, the bacterial E1 and mesostable comparator exhibited similar patterns of activity, although the mesostable comparator showed a slightly more rapid rate of initial decline (Figure 2). Plant-soluble E1, however, exhibited a large pool of slowly dissipating endoglucanase, with approximately 50% of initial activity remaining after 25 h. In definitive studies, when E1 from bacterial and plant-soluble sources was added to fresh soil microcosms, loss of activity was more rapid (Figure 3). Although differences among the fresh soils were not evident, there was a distinctly different trend in initial decline in activity for the plant-soluble versus bacterial source of E1. On the basis of preliminary model fitting, the size of the fast-reacting pool of soil E1 for all soils was fixed at 75% and 50% of initial activity for the plant-soluble and bacterial E1, respectively. Initial decline from this fast-reacting pool was more rapid for the plantsoluble E1 compared with bacterial E1; but in all cases, relative E1 activity declined to Coland > Hanlon (Figure 4).
Plant soluble E1 Mesostable comparator Bacterial E1
100
Plant soluble E1 f(x) = 32e-3.5x + 68e-0.012x r2 = 0.98
80
60
DISCUSSION
Bacterial E1 -0.56x -0.087x + 58e f(x) = 42e r2 = 0.98 Mesostable comparator f(x) = 67e-0.70x + 33e-0.091x 2 r = 0.92
40
20
There were no evident effects of exogenous E1 addition to soils in terms of either the retention of extractable activity or total CO2 evolved in microcosm studies. The 3 soils used in the definitive study were chosen to represent sites where E1 maize would be grown and differed largely in texture and organic matter content within an alluvial catena characteristic of Midwestern production fields. In Iowa, the majority of soils that support maize production have 3% to 5% organic matter, neutral to slightly acidic pH, and 20% to 30% clay content [37]. The Spillville and Coland soils are characteristic of these soils, whereas the Hanlon soil has relatively lower organic matter (1.9%) and clay content (16%) and relatively higher pH (7.6). If E1 were persistent in Iowa soils, its long-term activity would likely be more evident in soils such as Hanlon because of its
0 0
5
10
15
20
25
30
Time (hours)
Figure 2. Comparative loss of endoglucanase activity from Nicollet loam surface soil amended with either bacterial endoglucanase (E1), plant-soluble E1, or mesostable comparator. Each data point represents the average of triplicates 95% confidence interval.
Plant-soluble E1
A
100
Bacterial E1
B
f(x) = 75e-4.30x+ 25e-0.02x r2 = 0.92
80
f(x) = 50e-1.01x+ 50e-0.06x r2 = 0.97
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Enzyme activity (% of initial)
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f(x) = 75e-4.93x+ 25e-0.02x r2 = 0.89
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f(x) = 50e-2.30x+ 50e-0.06x r2 = 0.93
60 40 20
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E
100
F
f(x) = 75e-4.58x+ 25e-0.05x r2 = 0.95
80
f(x) = 50e-1.86x+ 50e-0.04x r2 = 0.97
60 40 20
Spillville 0 0
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0
5
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Time (h) Figure 3. Comparative loss of endoglucanase activity from field-moist Hanlon sandy loam (A,B), Coland loam (C,D), and Spillville silt loam (E,F) surface soils amended with either plant-soluble E1 (A,C,E) or bacterial E1 (B,D,F). Each data point represents the average of triplicates 95% confidence interval.
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A Control Wild-type maize E1-expressing maize
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-1
Cumulative CO2-C evolution (mg kg soil)
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Incubation time (d) Figure 4. Cumulative CO2-C release from field-moist Spillville silt loam (A), Coland loam (B), Hanlon sandy loam (C) surface soils amended with endoglucanase (E1)-expressing or wild-type maize tissue and for unamended controls. Data points are the average of 3 replications 95% confidence interval.
lower reactivity and especially its high pH, which would favor E1 activity over that of endogenous soil enzymes. In the present study, maize tissue expressing E1 in mitochondria was utilized because it shows good promise as a plant expression system for bioprocessing (M. Sticklen, Michigan State University, East Lansing, MI, USA, personal communication). Levels of E1 expression in our test material (5 ng mg1 tissue) were within the range of E1 maize expression in the literature (3–33 ng mg1 tissue) [13,18]. Levels of fortification with various test materials were designed to demonstrate nominally equivalent levels of endoglucanase activity equivalent to plant residue loadings providing 60 ng E1 g1 soil (120 g ha1), which approximates the level that would be found in postharvest residues from E1 maize production. The mesostable comparator originated from
2001
organisms that are ubiquitous in soil (Aspergillus and Trichoderma spp.) and was used to examine the dissipation of representative cellulases in the soil environment. Because of the specificity of carboxymethyl cellulose in the enzyme assay used, the activity measured for the mesostable comparator represented endoglucanase activity from the mix of cellulase enzymes. Because the fortification of soils with the mesostable comparator resulted in a suite of cellulases being added, the comparison to E1-fortified soils is inexact but does serve to compare the relative differences in endoglucanase activities from the mesostable and hyperthermostable sources. The process by which exogenous enzyme activity is lost from soil depends on the overall microbial activity, the relative amounts of clay and humic substances present, and the stability of the particular protein involved [38–40]. The selection of incubation temperatures (40 8C or 65 8C) in the endoglucanase assays allowed us to discriminate between activity of mesostable endoglucanase and thermostable E1 because of differences in temperature optima. Results from these studies show that the activity of bacterial and plant-soluble E1 is rapidly lost within a matter of hours after addition to soil. As with proteins in general, endoglucanases released to the environment will be either rapidly adsorbed to soil particulates or degraded by microbes and proteases [29]. It is possible that E1 becomes bound to colloidal material and retains activity; however, any residual effects were not evident over the short term of the present study. The endoglucanase activity detected when E1 was amended to these soils was no different from that of the mesostable comparator. Direct measurement of plant E1 persistence from maize incorporated into soil microcosms was not possible because of the high glucose background from tissue. However, there was no observed effect on whole carbon mineralization from soils amended with E1 maize, and this is consistent with the very limited duration of activity observed in studies with plantsoluble E1. Additionally, the high level of glucose present in these tissue-amended soils may have resulted in feedback inhibition to stall enzymatic conversion [41]. In all, these data suggest that plant exogenous E1 introduced to soil will have no short-term effect on carbon cycling. The present research utilized maize expressing E1-endoglucanase in mitochondria, but there are many alternative sites of in planta expression and enzyme compartmentalization as well as differing specific suites of enzymes that may be plantexpressed [19], which may affect the nature of activity and carbon cycling effect in soils. In addition, we investigated representative Midwestern US soils for maize production, but there may also be unique soil types that require special consideration relative to the impact of E1 maize residues. For example, soils with high pH, low organic matter and clay content, or low overall microbial activity may have longer tissue residence times, less reactivity, and greater overall effects than shown in the present study. Under conditions in which soil temperatures are substantially higher than investigated in the present study, changes in enzyme activity could be evident. SUMMARY
Clean, renewable energy will be needed in the near future to address dwindling petroleum reserves and mitigate anthropogenically induced climate change from CO2. To meet these objectives in a timely manner, maize has been genetically engineered to produce a bioprocessing enzyme that may reduce the economic and energy costs associated with lignocellulosic ethanol conversion. The novel expression of thermostable
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E1-endoglucanase in maize green tissue requires better understanding of the fate and implications of plant-expressed E1 in representative soil environments. In the United States, harvestable stover for biofuel production is likely to come from highly productive Midwestern soils. Based on residue stover left on fields after harvest, E1 from maize is conservatively anticipated to release 120 g ha1 into representative soils. Rapid decline in endoglucanase activity recovered from soil extracts showed limited activity of exogenous E1 in soil. Subsequent measurements of CO2 evolution for soils amended with E1-maize tissue confirmed no effect of endogenous E1 on short-term carbon mineralization. Together, these activity and CO2 evolution studies show that maize expressing up to 30 g mg1 of E1 (120 g ha1 environmental load) did not contribute significant endoglucanase activity in representative soils. However, the transformed maize used in the present study expressed only 1 component of the cellulase complex, endoglucanase. Maize that expresses the entire suite of cellulase enzymes is under development, and therefore, future studies may benefit from the development and incorporation of rapid immunoassays (e.g., enzyme-linked immunosorbent assays) to allow for more sensitive testing for effects to carbon cycling on agricultural soils expected to receive exogenous loadings of novel enzymes. Acknowledgment—M. Sticklen (Michigan State University) and M. Himmel and W. Adney (National Renewable Energy Lab) provided E1 sources and supporting materials. This project was supported by Biotechnology Risk Assessment Program competitive grant 2008-33522-04758 from the US Department of Agriculture, National Institute of Food and Agriculture.
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A.J. Kenny and J.D. Wolt 13. Mei C, Park S-H, Sabzikar R, Qi C, Ransom C, Sticklen M. 2009. Greentissue–specific production of a microbial endocellulase in maize (Zea mays L) endoplasmic reticulum and mitochondria converts cellulose into fermentable sugars. J Chem Technol Biotechnol 84:689–695. 14. Hu W-J, Harding SA, Lung J, Popko JL, Ralph J, Stokke DD, Tsai C-J, Chiang VL. 1999. Repression of lignin biosynthesis promotes cellulose accumulation and growth in transgenic trees. Nat Biotechnol 17: 808–812. 15. Anterola AM, Lewis NG. 2002. Trends in lignin modification: A comprehensive analysis of the effects of genetic manipulations/ manipulations on lignification and vascular integrity. Phytochemistry 61:221–294. 16. Boerjan W, Ralph J, Baucher M. 2003. Lignin biosynthesis. Annu Rev Plant Biol 54:519–546. 17. Alizadeh H, Teymouri F, Gilbert TI, Dale BE. 2005. Pretreatment of switchgrass by ammonia fiber explosion (AFEX). Appl Biochem Biotechnol 121–124:1133–1141. 18. Brunecky R, Selig MJ, Vinzant TB, Himmel ME, Lee D, Blaylock MJ, Decker SR. 2011. In-planta expression of A. cellulolyticus Cel5A endocellulase reduces cell wall recalcitrance in tobacco and maize. Biotechnology for Biofuels 4:1–4. 19. Park S-H, Ransom C, Mei C, Sabzikar R, Qi C, Chundawat S, Dale B, Sticklen M. 2011. The quest for alternatives to microbial cellulase mix production: Corn stover-produced heterologous multi-cellulases readily deconstruct lignocellulosic biomass into fermentable sugars. J Chem Technol Biotechnol 86:633–641. 20. Skujins JJ. 1978. History of abiotic soil enzyme research. In Burns RC, ed, Soil Enzymes. Academic, New York, NY, USA, pp 1–49. 21. Wolt JD. 2009. Advancing environmental risk assessment for transgenic biofeedstock crops. Biotechnology for Biofuels 2:27. 22. Burns RG. 1986. Interaction of enzymes with soil mineral and organic colloids. In Huang PM, Schnitzer M, eds, Interactions of Soil Minerals with Natural Organics and Microbes. Special Publication 17. Soil Science Society of America, Madison, WI, pp 429–451. 23. Nannipieri P. 1994. The potential use of soil enzymes as indicators of productivity, sustainability and pollution. In Pankhurst CE, Doube BM, Gupta VVSR, Grace PR, eds, Soil Biota Management in Sustainable Farming Systems. CSIRO, East Melbourne, Australia, pp 238–244. 24. Dick RP, Breakwell DP, Turco RF. 1996. Soil enzyme activities and biodiversity measurements as integrative microbiological indicators. In Doran JW, Jones AJ, eds, Methods for Assessing Soil Quality. Soil Science Society of America, Madison, WI, pp 247–271. 25. Naseby DC, Lynch JM. 1997. Functional impact of genetically modified micro-organisms on the soil ecosystem. In Zelikoff JT, Schepers J, Lynch JM, eds, Ecotoxicology: Responses, Biomarkers and Risk Assessment. SOS Publications, Fair Haven, NJ, USA, pp 419–442. 26. Nielsen MN, Winding A. 2002. Microorganisms as indicators of soil health. Technical report 388. National Environmental Research Institute, Aarhus, Denmark. 27. Lal R. 2004. Soil carbon sequestration impacts on global climate change and food security. Science 304:1623–1626. 28. Jaenicke R, Bohm G. 1998. The stability of proteins in extreme environments. Curr Opin Struct Biol 8:738–748. 29. Nielsen KM, Calamai L, Pietramellara G. 2006. Stabilization of extracellular DNA and proteins by transient binding to various soil components. In Nannipieri P, Smalla K, eds, Nucleic Acids and Proteins in Soil. Soil Biology 8. Springer-Verlag, Berlin, Germany, pp 141–157. 30. Himmel ME, Adney WS, Baker JO, Elander R, McMillan JD, Nieves RA, Sheehan JJ, Thomas SR, Vinzant TB, Zhang M. 1997. Advanced bioethanol production technologies: A perspective. In Saha BC, Woodward J, eds, Fuels and Chemicals from Biomass. ACS Symposium Series 666. American Chemical Society, Washington, DC, pp 2–45. 31. Dai Z, Hooker BS, Anderson DB, Thomas SR. 2000. Expression of Acidothermus cellulolyticus endoglucanase E1 in transgenic tobacco: Biochemical characteristics and physiological effects. Transgenic Res 9:43–54. 32. Miller GL. 1959. Use of a dinitrosalicylic acid reagent for determination of reducing sugar. Anal Chem 31:426–428. 33. Trevors JT. 1996. Sterilization and inhibition of microbial activity in soil. J Microbiol Methods 26:53–59. 34. Deng SP, Tabatabai MA. 1994. Cellulase activity of soils. Soil Biol Biochem 10:1347–1354. 35. Wolt JD, Nelson HP Jr, Cleveland CB, van Wesenbeeck IJ. 2001. Biodegradation kinetics for pesticide exposure assessment. Rev Environ Contam Toxicol 169:123–164. 36. Gupta SK, Radke JK, Larson WE. 1981. Predicting temperature of bare and residue covered soils with and without a corn crop. Soil Science Society of America Proceedings 45:405–412.
Maize-expressed transgenic endoglucanase in soils 37. Miller GA, Fenton TE, Oneal BR, Tiffany BJ, Burras CL. 2010. Iowa Soil Survey and Digital Soil Data (ISPAID), Ver 7.3. Iowa State University, Ames, IA, USA. 38. Icoz I, Stotzky G. 2008. Fate and effects of insect-resistant Bt crops in soil ecosystems. Soil Biol Biochem 40:559–586. 39. Kosaki H, Coats JR, Wang K, Wolt JD. 2008. Persistence and degradation of maize-expressed vaccine protein, Escherichia coli heatlabile enterotoxin subunit B, in soil and water. Environ Toxicol Chem 27:1227–1236. 40. Prihoda KR, Coats JR. 2008. Fate of Bacillus thuringiensis (Bt) Cry3Bb1 protein in a soil microcosm. Chemosphere 73:1102–1107. 41. Andric P, Meyer A, Jensen P, Dam-Johansen K. 2010. Reactor design for minimizing product inhibition during enzymatic lignocellulose hydrolysis: I. Significance and mechanism of cellobiose and glucose inhibition on cellulolytic enzymes. Biotechnol Adv 28:308–324.
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42. National Agricultural Statistics Service. 2011. Crop Production. US Department of Agriculture, Washington, DC. 43. Perlack RD, Wright LL, Turhollow AF, Graham RL, Stokes BJ, Erbach DC. 2005. Biomass as feedstock for a bioenergy and bioproducts industry: The technical feasibility of a billion-ton annual supply. ORNL/TM-2005/66. Oak Ridge National Laboratory, Oak Ridge, TN, USA. 44. Graham RL, Nelson R, Sheehan J, Perlack RD, Wright LL. 2007. Current and potential U.S. corn stover supplies. Agron J 99: 1–11. 45. White RE. 1979. Introduction to the Principles and Practice of Soil Science. Blackwell, Oxford, UK. 46. van Genuchten MT. 1980. A closed-form equation for predicting the hydraulic conductivity of unsaturated soils. Soil Sci Soc Am J 44: 892–898.
ACTIVITY AND ECOLOGICAL IMPLICATIONS OF MAIZE-EXPRESSED TRANSGENIC ENDO-1,4-b-D-GLUCANASE IN AGRICULTURAL SOILS ADAM J. KENNY and JEFFREY D. WOLT* Department of Agronomy, Iowa State University, Ames, Iowa, United States (Submitted 24 March 2014; Returned for Revision 14 May 2014; Accepted 23 May 2014) Abstract: Plant expression of thermostable endoglucanase (E1) has been proposed for improved conversion of lignocellulose to ethanol for fuel production. Residues of E1-expressing maize may affect ecological services (e.g., C mineralization and biogeochemical cycling) on soils where they occur. Therefore, the activity of residual E1 was investigated using soils amended with bacterial and plant-solubilized E1 compared with soil endogenous activity and residual activity from a mesostable cellulase (Aspergillus and Trichoderma spp.). An optimized analytical method involving a carboxymethyl cellulose substrate and dinitrosalicylic acid detection effectively assayed endoglucanase activity in amended and unamended soils and was used for determining E1 activity in 3 representative soils. The effect of E1 on soil carbon mineralization was determined by comparing CO2 evolution from soils amended with transgenic E1-expressing and wild-type maize tissue. Extraction and recovery of the mesostable comparator, bacterial E1, and plant-soluble E1 showed nearly complete loss of exogenous endoglucanase activity within a 24-h period. Carbon mineralization indicated no significant difference between soils amended with either the transgenic E1 or wild-type maize tissue. These results indicate that maize residues expressing up to 30 mg E1/g tissue negligibly affect soil endoglucanase activity and CO2 respiration for representative soils where transgenic E1 maize may be grown. Environ Toxicol Chem 2014;33:1996–2003. # 2014 SETAC Keywords: Acidothermus cellulolyticus
Genetically engineered
Dissipation
Thermostable
Cellulase
Endoglucanase
One strategy to reduce costs for cellulosic ethanol production is to genetically transform the source biofeedstock for better utilization of the cellulosic material [6,7]. The genetic engineering of bioenergy feedstocks has been explored for a variety of cellulosic crops—Zea mays (maize), Panicum virgatum (switchgrass), and Miscanthus spp.—to improve processing through in planta expression of bioprocessing enzymes [8–13] and through altering the lignin content and composition [14–17]. Improved cellulose conversion efficiencies for maize stover can be achieved through in planta production of bioprocessing enzymes so that, following biomass pretreatment and cell lysis, plant-made cellulases become active and cleave cellulosic polymers into fermentable sugars. A case in point is highly thermostable endo-1,4-b-D-glucanase from Acidothermus cellulolyticus (E1; pdb:1ECE) that has been expressed at high levels in maize green tissue [13]. With an optimal temperature of 83 8C and with marked stability in both acidic and basic environments, E1-endoglucanase has been an attractive target for expression in cellulosic biofeedstock crops [1,18,19]. We estimate that where genetically engineered maize expressing E1 is grown, it will add 60 g ha1 to 120 g ha1 annually of E1 in postharvest residues to receiving soils, depending on the nature of expression (Table 1). This stands in stark contrast to typical agroecosystems in which plants are considered minor contributors to soil enzyme activity through residue decomposition and root exudation [20]. Therefore, widescale agricultural release of E1-expressing maize will require environmental assessment to better understand the potential impact of residues left in the field [21]. Especially important in this regard is the evaluation of effects on ecosystem services (e.g., carbon mineralization and biogeochemical cycles). It has been well documented that enzymes in the soil environment are direct biological instigators of elemental cycling and indicators of overall soil tilth [22–26]. The importance of soil glycoside
INTRODUCTION
Maize cellulosic residues represent a major potential biofeedstock source for biofuel production [1,2]. Current ethanol production strategies using cellulosic feedstocks are inefficient because, in order for carbon polymer sugars to become available for processing into ethanol, the biofeedstock must undergo dilute acid pretreatment at temperatures generally in excess of 75 8C to eradicate the recalcitrant lignin and hemicellulose layers that shield the carbon substrate. Pretreatment is followed with exogenous cellulase addition to release fermentable glucose sugars. Cellulases are biocatalytic systems made up of endoglucanases (Enzyme Commission number 3.2.1.4), exoglucanases (Enzyme Commission number 3.2.1.91), and cellobiases (Enzyme Commission number 3.2.1.21) that work synergistically to depolymerize cellulose. The currently accepted mechanism of cellulase action begins with endoglucanase hydrolyzing internal 1,4 glycosidic bonds within amorphous regions to produce new chain ends. Exoglucanase (e.g., cellobiohydrolase) attacks chain ends to yield glucose or cellobiose. Cellobiase completes any residual breakdown needed and prevents product inhibition from cellobiose [3]. Thermostability of the added cellulases is highly desired in the biofuel conversion process because these enzymes are subjected to high temperatures and variable pH. Additionally, saccharification at higher temperatures can prevent batch contamination, increase solution viscosity for improved mixing, and better solubilize cellulose substrates [4]. Addition of exogenous cellulase enzymes after pretreatment accounts for approximately one-third to one-half of the entire production cost for cellulosederived ethanol [5]. * Address correspondence to [email protected] Published online 26 May 2014 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/etc.2645 1996
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Table 1. Estimated environmental concentration (EEC) for thermostable endoglucanase (E1) in soil from postharvest residues of E1-expressing maize Assumptions
Rationale
11.4 Mg grain ha1 1:1 grain to stover biomass ratio 0.33 postharvest fraction of residual biomass left on field 3.8 Mg residual biomass ha1 (b) 2 103 Mg soil ha1 (m) 17–33 g Mg1 residual biomass (a) EEC ¼ ab/m ¼ 0.032–0.063 g E1 Mg1 soil 60–120 g E1 ha1
Iowa average maize yield, 2009 (National Agricultural Statistics Service [42]) Perlack et al. [43] For erosion control and nutrient sustainability (Graham et al. [44]) Incorporation to 15 cm depth, typical soil bulk density 1.33 g cm3 (White [45], p. 73) Range in mitochondrial expression in green plant tissue (Mei et al. [13])
hydrolases cannot be overemphasized because the soil carbon pool is more than 3 times that of the atmosphere pool and 4.5 times that of the biotic pool [27]. Furthermore, E1, which was originally isolated from hot springs, has thermostable properties that may confer increased stability when released to agricultural soils during postharvest residue decomposition [28]. The novel expression, release to a foreign environment, and potential effects to ecosystem services require better knowledge of E1 fate in agricultural soil. Enzyme function and stability have been linked to numerous soil physicochemical properties, especially clay and organic matter content. Clay and humic substances are thought to protect enzymes/proteins from degradation because of their enhanced stability via steric hindrance and conformational changes on organomineral sorption that make them less susceptible to proteolysis and physicochemical denaturing [29]. Ironically, the sorption process can also alter the tertiary structure of proteins to the extent that they are no longer biologically active; however, there is also evidence that activity may be retained after sorption. Regardless of the fate of enzymes functionality, once adsorbed to soil particles they are generally considered to be irreversibly bound [29]. As a first step toward understanding residue effects of E1expressing maize, we discriminated activity of endoglucanases from various sources (bacterial E1, E1-maize soluble protein, endogenous soil enzymes, and mesostable cellulase) when amended to representative agricultural soils. In addition, carbon dioxide (CO2) evolution was used to compare whole-soil respiration rates for unamended soils and for soils amended with either transgenic (E1) or nontransgenic (wild-type) maize tissue. The CO2 evolution study provides indirect data on activity from whole tissue–derived E1 that has been amended to soils and indicates whether E1 presence in soil affects mineralization of labile carbon. MATERIALS AND METHODS
Test materials
Bacterial E1 from Streptomyces lividans was provided by W.S. Adney (National Renewable Energy Laboratory). Soluble bacterial E1 preparations were used for all analyses. The mesostable cellulase comparator (Aspergillus and Trichoderma spp.) was obtained from MP Biomedical. The bacterial E1-endoglucanase and mesostable comparator enzymes were characterized to determine the temperature and pH optima using the carboxymethyl cellulose and dinitrosalicylic acid analytical technique (see Endoglucanase assay). Dry tissue (leaves plus stems) of transformed (E1) and untransformed (wild-type) maize (V5 stage) was obtained from M. Sticklen (Michigan State University). Transgenic tissues were from T2 maize representing a cross between mitochondria l expressing E1-endoglucanase lines (1.5-E1-10b 1.5-E1-10c)
[13]. Nontransgenic wild type near isoline maize grown under the same conditions as the E1 maize was used as a control. On receipt, tissues were pulverized in liquid nitrogen, screened to pass 1.7-mm mesh openings, and stored at –20 8C until used. Total carbon and nitrogen for subsamples was determined by combustion and infrared detection. Plant total soluble protein was obtained by genogrinding (SPEX CertiPrep) 100 mg of the previously milled tissue in 1 mL of extraction buffer (50 mM sodium acetate [NaAc], 10 mM ethylenediaminetetraacetic acid, 0.1% Triton X-100, pH 5). Crude soluble extracts were centrifuged, and an aliquot was subjected to ammonium-sulfate precipitation using an initial saturation concentration of 100% (0 8C). Crude extracts and ammonium sulfate solution were combined at 0.4:1 ratio (70% final saturation concentration), briefly vortexed, and then stored in a –20 8C freezer for 1 h before centrifugation at 20 800 g for 5 min. The protein pellet formed was reconstituted to an equivalent of 0.2 g original tissue/mL. Activity of plant-soluble protein from E1 and wild-type maize was determined and compared against activity from E1 standards to estimate expression levels (see Endoglucanase assay). All stock enzyme materials were stored concentrated in 1% bovine serum albumin and 50 mM NaAc (pH 5) in polypropylene vials. Test systems
All preliminary work was performed using an air-dried Nicollet loam surface soil (superactive, mesic Aquic Hapludolls). Definitive studies utilized surface soils from the Hanlon– Spillville–Coland alluvial catena (superactive, mesic Cumulic Hapludolls) that were collected fresh, immediately sieved to pass 2-mm mesh openings, and maintained field-moist at a workable consistency in the dark at 4 8C prior to use. These soils were used within 60 d of sampling. Physicochemical characteristics of the test soils are summarized in Table 2. Endoglucanase assay
A number of optimization trials were undertaken to evaluate extraction and detection procedures to allow for discrimination of endoglucanase activity from test materials and test systems (A.J. Kenny, 2012, Master’s thesis). Endoglucanase activity was determined through assay of glucose release from carboxymethyl cellulose substrate [13,18,30,31]. Glucose monomers released by carboxymethyl cellulose hydrolysis from endoglucanase were directly measured using a reducing sugar assay solution [32]. For characterization of test materials, dinitrosalicylic acid reagent solution was mixed 1:1 (400 mL total) with 2% carboxymethyl cellulose solution (pH 5) and heated to 95 8C for 5 min. The colored solution was centrifuged at 20 800 g for 1 min, and a 200-mL aliquot was immediately measured and compared with control samples at 540 nm in microplate wells using a BioTek PowerWave spectrophotometer. Sodium azide (NaN3) was added as a microbial inhibitor during the hydrolysis
1998
Environ Toxicol Chem 33, 2014
A.J. Kenny and J.D. Wolt Table 2. Physicochemical properties of test soils Water content (%)d
Texture (%) Series Nicollet Hanlon Spillville Coland
Textural class
Sand
Silt
Clay
Organic matter (%)a
pHb
CECc (cmol kg1)
33 kPa
100 kPa
1500 kPa
Loam Sandy loam Silt loam Loam
48 56 24 34
34 28 50 40
18 16 26 26
2.9 1.9 3.4 3.7
6.7 7.6 6.8 7.0
16.8 14.7 22.1 23.9
16.5 14.2 24.7 23.4
— 12.6 17.3 18.1
11.52 8.61 14.73 15.69
a
Loss on ignition. Soil–water extract, 1:1. c Cation exchange capacity (CEC), sodium acetate method (US Environmental Protection Agency 9081). d Pressure plate measurements for 33 kPa and 1500 kPa, 100 kPa estimated using pedotransfer functions (van Genuchten [46]). b
phase of the assay [33] to allow for accumulation of released glucose. For soil assays, a 0.5-g (oven-dry equivalent) soil sample was extracted for 5 min using bead-beating (genogrinding at 100 strokes min1) in the presence of 600 mL of the extraction buffer (50 mM NaAc, 10 mM ethylenediaminetetraacetic acid, 0.01% Triton X-100, pH 5 [34]) and then centrifuged for 5 min at 20 800 g. Aliquots of the supernatant (300 mL) were incubated with 300 mL of 2% carboxymethyl cellulose substrate solution containing 0.05% NaN3. Incubation was conducted at pH 5 and temperatures of 40 8C or 65 8C for the mesostable comparator and E1, respectively, based on the previously determined pH and temperature optima. Recovery of accumulated glucose was determined with the dinitrosalicylic acid reagent using a 200-mL aliquot of the reaction solution, 200 mL of dinitrosalicylic acid reagent, incubating at 95 8C for 5 min, and measuring spectrophotometrically at 540 nm. E1 activity in amended soils
Soil microcosm studies were used to determine exogenous enzyme activity. Test systems utilized 0.5-g soil samples (ovendry equivalent) in 2-mL microcentrifuge tubes. Test materials (enzymes) were diluted from stock and applied to surface soil samples at environmentally relevant concentrations. Soils were amended with 300 ng/g, 60 ng/g, and 80 ng/g oven dry equivalent soil of the mesostable comparator, bacterial E1, and plant-soluble E1, respectively. These levels of fortification were previously determined to represent equivalent amounts of source enzyme activity so that each treatment hydrolyzed carboxymethyl cellulose to release glucose at relatively equal rates. Plant-soluble E1 represented a loading equivalent to 160 g ha1, only slightly above the high-end estimate of E1 that would be released to the soil environment under typical conditions (Table 1). Preliminary studies used the air-dried Nicollet soil and mesostable comparator enzyme and were confirmed using bacterial and plant-soluble E1. For definitive studies, field-moist Hanlon, Spillville, and Coland soils were fortified with either bacterial or plant-soluble E1, while control soils received no enzyme. Following fortification, soils were mixed by spatula, moisture was adjusted with deionized water, and the samples were incubated aerated in the dark at a constant temperature (Ambient Temperature Incubator BOD50A16; Thermo-Scientific). For definitive studies, the base water potential and temperature used were 100 kPa and 25 8C, respectively. The 100 kPa moisture for the Hanlon, Spillville, and Coland soils was 12.6%, 17.3%, and 18.1%, respectively (Table 2). Each study included 6 sampling time points extending over approximately 2.5 half-lives of the test materials as determined on the basis of the preliminary study. At each sampling time
triplicate vials for each treatment were sampled for analysis of enzyme activity. CO2 evolution
Mineralization effects from E1-endoglucanase amended to field-moist Hanlon, Spillville, and Coland soils were determined for soils treated with E1 or wild-type maize tissue or unamended using a fully factorial design in 3 replications. For amended samples, 20 mg of ground tissue was mixed into 10 g soil (ovendry equivalent), representing an environmental loading of 120 g E1 ha1, whereas unamended samples were simply 10 g soil. Amended soils were added to 120-mL bottles, adjusted to 100 kPa water potential with the addition of deionized water, sealed with rubber stoppers, and incubated at 25 8C. Gas samples were analyzed with a LI-7000 CO2/H2O LI-COR infrared gas analyzer (LI-COR Biotechnology). At each sampling, stoppers were pierced with a syringe, and headspace was well mixed before 0.4 mL of gas was withdrawn and injected into the LI-COR analyzer, which had been previously calibrated against CO2 standards. Rubber stoppers were then removed, and headspace was replaced with laboratory air under slight positive pressure. Stoppers were replaced, and a second 0.4-mL aliquot was withdrawn for analysis. Change in CO2 was standardized for the 2 samples to reflect CO2 released per gram of soil. Samples were taken 14 times during the 45-d duration of the study. Instrumentation was calibrated before every use, and soil moisture was readjusted to 100 kPa twice over the course of the study. Statistical analysis
Comparisons presented graphically throughout represent the 95% confidence intervals calculated on the basis of triplicate analysis, unless otherwise indicated. An empirical biphasic decay model, f(x) ¼ ae–bx þ ce–dx, was used to characterize the change in extractable enzyme activity with time [35], where the loss of activity, f(x), over time, x in hours, was determined by the 2 apparent first-order rate constants b and d, describing loss from 2 differentially acting pools of enzyme, a and c, respectively. Data fitting used SigmaPlot ver 10 software (Systat Software), where the relative percentages of a and c were constrained such that c ¼ 100 a, whereas b and d were constrained to values >0. For CO2 evolution studies, a least squares full factorial method was used to examine interactions between variables (time, replication, treatment, and soil). All cross-terms indicated significant interactions; therefore, Tukey’s honestly significant difference test was applied to determine significant effects of treatment soil at the a ¼ 0.05 level using JMP V.10 (SAS).
Maize-expressed transgenic endoglucanase in soils
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A
retained high activity across a much broader range (pH 5–9; Figure 1B). Both E1 and wild-type maize tissue had similar C:N ratios, but total protein content was higher for the E1 tissue, whereas soluble protein content was higher for the wild-type compared with the E1 tissue (Table 3). The differences in total and soluble protein concentrations were slight and, given the favorable C:N ratios, would not markedly contribute to differences in mineralization. Transgenic maize was positive for E1 activity; expression was 0.55% of total soluble protein, representing 5 mg g1 tissue. Wild-type maize, however, exhibited no measurable E1 activity (Table 3). Combining the present expression data with that of Mei and coworkers [13], a refined range for the estimated environmental concentration of 10 mg E1 g1 to 60 mg E1 g1 soil (17–120 g ha1 soil loading) was calculated (Table 1).
Enzyme activity (relative %)
100
Q10 = 1.9
80
60
40
Q10 = 1.4 20
0 10
20
30
40
50
60
o
Temperature ( C)
B
Enzyme activity in soil
100
Enzyme activity (relative %)
1999
80
60
40
20 Thermostable E1 Mesostable comparator 0 3
4
5
6
7
8
9
pH
Figure 1. Enzyme activity of bacterial sources of thermostable endoglucanase (E1) and the mesostable comparator as affected by (A) temperature and (B) pH. Each data point represents the average of triplicates 95% confidence interval. Q10 ¼ temperature coefficient.
RESULTS
Enzyme characterization
When incubated at pH 5, bacterial E1 exhibited steadily increasing activity when measured over a temperature range of 15 8C to 60 8C, consistent with the reported temperature optima of 83 8C [30]. The mesostable comparator exhibited a temperature optimum of 40 8C under similar conditions (Figure 1A). Relative to the mesostable comparator, the bacterial E1 had a substantially lower activity at temperatures reasonably anticipated for surface soil under maize residue cover [36]. Both enzymes displayed optimal activity at pH 5 when determined at temperature optima of 40 8C and 65 8C for the mesostable comparator and bacterial E1, respectively; however, bacterial E1
In preliminary studies in which soluble enzyme test materials were amended to the air-dried Nicollet soil, the bacterial E1 and mesostable comparator exhibited similar patterns of activity, although the mesostable comparator showed a slightly more rapid rate of initial decline (Figure 2). Plant-soluble E1, however, exhibited a large pool of slowly dissipating endoglucanase, with approximately 50% of initial activity remaining after 25 h. In definitive studies, when E1 from bacterial and plant-soluble sources was added to fresh soil microcosms, loss of activity was more rapid (Figure 3). Although differences among the fresh soils were not evident, there was a distinctly different trend in initial decline in activity for the plant-soluble versus bacterial source of E1. On the basis of preliminary model fitting, the size of the fast-reacting pool of soil E1 for all soils was fixed at 75% and 50% of initial activity for the plant-soluble and bacterial E1, respectively. Initial decline from this fast-reacting pool was more rapid for the plantsoluble E1 compared with bacterial E1; but in all cases, relative E1 activity declined to Coland > Hanlon (Figure 4).
Plant soluble E1 Mesostable comparator Bacterial E1
100
Plant soluble E1 f(x) = 32e-3.5x + 68e-0.012x r2 = 0.98
80
60
DISCUSSION
Bacterial E1 -0.56x -0.087x + 58e f(x) = 42e r2 = 0.98 Mesostable comparator f(x) = 67e-0.70x + 33e-0.091x 2 r = 0.92
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There were no evident effects of exogenous E1 addition to soils in terms of either the retention of extractable activity or total CO2 evolved in microcosm studies. The 3 soils used in the definitive study were chosen to represent sites where E1 maize would be grown and differed largely in texture and organic matter content within an alluvial catena characteristic of Midwestern production fields. In Iowa, the majority of soils that support maize production have 3% to 5% organic matter, neutral to slightly acidic pH, and 20% to 30% clay content [37]. The Spillville and Coland soils are characteristic of these soils, whereas the Hanlon soil has relatively lower organic matter (1.9%) and clay content (16%) and relatively higher pH (7.6). If E1 were persistent in Iowa soils, its long-term activity would likely be more evident in soils such as Hanlon because of its
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Figure 2. Comparative loss of endoglucanase activity from Nicollet loam surface soil amended with either bacterial endoglucanase (E1), plant-soluble E1, or mesostable comparator. Each data point represents the average of triplicates 95% confidence interval.
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Time (h) Figure 3. Comparative loss of endoglucanase activity from field-moist Hanlon sandy loam (A,B), Coland loam (C,D), and Spillville silt loam (E,F) surface soils amended with either plant-soluble E1 (A,C,E) or bacterial E1 (B,D,F). Each data point represents the average of triplicates 95% confidence interval.
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Incubation time (d) Figure 4. Cumulative CO2-C release from field-moist Spillville silt loam (A), Coland loam (B), Hanlon sandy loam (C) surface soils amended with endoglucanase (E1)-expressing or wild-type maize tissue and for unamended controls. Data points are the average of 3 replications 95% confidence interval.
lower reactivity and especially its high pH, which would favor E1 activity over that of endogenous soil enzymes. In the present study, maize tissue expressing E1 in mitochondria was utilized because it shows good promise as a plant expression system for bioprocessing (M. Sticklen, Michigan State University, East Lansing, MI, USA, personal communication). Levels of E1 expression in our test material (5 ng mg1 tissue) were within the range of E1 maize expression in the literature (3–33 ng mg1 tissue) [13,18]. Levels of fortification with various test materials were designed to demonstrate nominally equivalent levels of endoglucanase activity equivalent to plant residue loadings providing 60 ng E1 g1 soil (120 g ha1), which approximates the level that would be found in postharvest residues from E1 maize production. The mesostable comparator originated from
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organisms that are ubiquitous in soil (Aspergillus and Trichoderma spp.) and was used to examine the dissipation of representative cellulases in the soil environment. Because of the specificity of carboxymethyl cellulose in the enzyme assay used, the activity measured for the mesostable comparator represented endoglucanase activity from the mix of cellulase enzymes. Because the fortification of soils with the mesostable comparator resulted in a suite of cellulases being added, the comparison to E1-fortified soils is inexact but does serve to compare the relative differences in endoglucanase activities from the mesostable and hyperthermostable sources. The process by which exogenous enzyme activity is lost from soil depends on the overall microbial activity, the relative amounts of clay and humic substances present, and the stability of the particular protein involved [38–40]. The selection of incubation temperatures (40 8C or 65 8C) in the endoglucanase assays allowed us to discriminate between activity of mesostable endoglucanase and thermostable E1 because of differences in temperature optima. Results from these studies show that the activity of bacterial and plant-soluble E1 is rapidly lost within a matter of hours after addition to soil. As with proteins in general, endoglucanases released to the environment will be either rapidly adsorbed to soil particulates or degraded by microbes and proteases [29]. It is possible that E1 becomes bound to colloidal material and retains activity; however, any residual effects were not evident over the short term of the present study. The endoglucanase activity detected when E1 was amended to these soils was no different from that of the mesostable comparator. Direct measurement of plant E1 persistence from maize incorporated into soil microcosms was not possible because of the high glucose background from tissue. However, there was no observed effect on whole carbon mineralization from soils amended with E1 maize, and this is consistent with the very limited duration of activity observed in studies with plantsoluble E1. Additionally, the high level of glucose present in these tissue-amended soils may have resulted in feedback inhibition to stall enzymatic conversion [41]. In all, these data suggest that plant exogenous E1 introduced to soil will have no short-term effect on carbon cycling. The present research utilized maize expressing E1-endoglucanase in mitochondria, but there are many alternative sites of in planta expression and enzyme compartmentalization as well as differing specific suites of enzymes that may be plantexpressed [19], which may affect the nature of activity and carbon cycling effect in soils. In addition, we investigated representative Midwestern US soils for maize production, but there may also be unique soil types that require special consideration relative to the impact of E1 maize residues. For example, soils with high pH, low organic matter and clay content, or low overall microbial activity may have longer tissue residence times, less reactivity, and greater overall effects than shown in the present study. Under conditions in which soil temperatures are substantially higher than investigated in the present study, changes in enzyme activity could be evident. SUMMARY
Clean, renewable energy will be needed in the near future to address dwindling petroleum reserves and mitigate anthropogenically induced climate change from CO2. To meet these objectives in a timely manner, maize has been genetically engineered to produce a bioprocessing enzyme that may reduce the economic and energy costs associated with lignocellulosic ethanol conversion. The novel expression of thermostable
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E1-endoglucanase in maize green tissue requires better understanding of the fate and implications of plant-expressed E1 in representative soil environments. In the United States, harvestable stover for biofuel production is likely to come from highly productive Midwestern soils. Based on residue stover left on fields after harvest, E1 from maize is conservatively anticipated to release 120 g ha1 into representative soils. Rapid decline in endoglucanase activity recovered from soil extracts showed limited activity of exogenous E1 in soil. Subsequent measurements of CO2 evolution for soils amended with E1-maize tissue confirmed no effect of endogenous E1 on short-term carbon mineralization. Together, these activity and CO2 evolution studies show that maize expressing up to 30 g mg1 of E1 (120 g ha1 environmental load) did not contribute significant endoglucanase activity in representative soils. However, the transformed maize used in the present study expressed only 1 component of the cellulase complex, endoglucanase. Maize that expresses the entire suite of cellulase enzymes is under development, and therefore, future studies may benefit from the development and incorporation of rapid immunoassays (e.g., enzyme-linked immunosorbent assays) to allow for more sensitive testing for effects to carbon cycling on agricultural soils expected to receive exogenous loadings of novel enzymes. Acknowledgment—M. Sticklen (Michigan State University) and M. Himmel and W. Adney (National Renewable Energy Lab) provided E1 sources and supporting materials. This project was supported by Biotechnology Risk Assessment Program competitive grant 2008-33522-04758 from the US Department of Agriculture, National Institute of Food and Agriculture.
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