The Metabolism of Exogenously Provided Atrazine by the

Chapter 11

The Metabolism of Exogenously Provided Atrazine

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by the Ectomycorrhizal Fungus Hebeloma crustuliniforme and the Host Plant Pinus ponderosa J. L. Gaskin and J. Fletcher Department of Botany and Microbiology, University of Oklahoma, Norman, OK 73019

The mineralization of atrazine by mycorrhizal fungi was examined. The percent mineralization was determined by comparing the amount of CO evolved from the plant and plant-fungus systems when measured amounts of C-labeled atrazine were provided under axenic conditions in specially designed exposure chambers. The percent mineralization was 0.1 and 0.3, respectively, for the plant and plant-fungus systems. The study demonstrated that an ectomycorrhizal fungus in association with its host plant increased the metabolism of exogenously provided atrazine above that of the plant by itself, thereby supporting the proposed use of plant-fungal systems in bioremediation of contaminated soils. 14

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14

The ability of ectomycorrhizae to degrade xenobiotic compounds has been studied with pure cultures examined under laboratory conditions. Donnelly et al. ( 1) screened several mycorrhizal fungi for their ability to degrade two aromatic herbicides, 2,4-D and atrazine. In a 30-day study with l ^ C labeled atrazine and 2,4-D, it was shown that the rates of degradation as measured by CO2 evolution varied depending on the species of fungus and herbicide tested (1). In general atrazine was degraded at a higher rate than 2,4-D by the fungi tested. Gautieria crispa, was the most active dégrader of atrazine; whereas Oidiodendron griseum, an ericoid mycorrhizal fungus, was most active against 2,4-D. Maximum mineralization of atrazine occurred between the third and fifth day, while it occurred between the fifth and tenth day for 2,4-D. Robideaux et al. (2) found that 10 of 11 ectomycorrhizal fungi tested removed the herbicide hexazinone from liquid medium. The amount of hexazinone removed depended on the species tested and the C / N ratio in the medium. In a study by Donnelly and Fletcher (3), examination of the ability of 21 different pure cultures of ectomycorrhizal fungi to metabolize 19 different polychlorinated biphenyl (PCB) congeners, showed that 14 of the fungi metabolized PCB's. These findings with pure cultures suggest that ectomycorrhizae are capable of degrading organic pollutants, but it is not certain that they retain this property when in association with roots of a host plant. The objective of this study was to examine the ability of roots colonized with ectomycorrhizal fungi to metabolize atrazine.

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© 1997 American Chemical Society

In Phytoremediation of Soil and Water Contaminants; Kruger, E., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1997.

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Methods Ponderosa pine seeds were sterilized in 30% hydrogen peroxide for 55 min., rinsed with sterile water, and placed on solid nutrient medium. When seeds had germinated and grown to approximately 3 cm in length they were examined for microbial contamination, and uncontaminated seedlings were transferred to sterile pouches (Figure 1). The pouches used were plant culture made by Agristar Inc. (Sealy, TX) The bags were modified by inserting paper pads removed from seed germination pouches available from Vaughn's Seed Company (Downers Grove, IL). Although Vaughn plastic polyester pouches have been used previously in ectomycorrhizal work (4) we found the Agristar bags to be preferred because they had much higher rates of gas exchange, and could also be autoclaved. Pouches were prepared as shown in Figure 1. Ten ml of Hoagland's solution were added to each pouch. A glass disposable pipette with a foil cap was clipped on the inside of each pouch for future watering. Two fiberglass pads were used as spacers to prevent the sides of the bag from collapsing and thereby permit air circulation. The assembled pouches were autoclaved, permitted to cool to room temperature, and sterile seedlings were carefully inserted to insure that the root and pouch remained axenic. The plant/pouches were placed in a 22 °C growth chamber and received 14 h of 300-400 pEinsteins of illumination every 24 hr. Sterile water (approx. 5 ml) was added aseptically every three days through the pipette attached to each pouch (Figure 1). Hoaglands solution (10%) was added every two weeks instead of water. Hebeloma crustuliniforme obtained from Paul Rygiewicz, (Environmental Research Laboratory, Corvallis, Oregon) was grown in petri dishes on mycorrhizal solid medium, which contained in one liter of media: glucose 10 g, NH4H2PO4 0.7 mg, KH2PO4 0.4 g, MgS04 0.5 g, CaCl2 100 mg, Fe-citrate 5 mg, MnS04 5 mg, ZnS04 4.4 mg, yeast extract 50 mg, agar IS g(l). Ponderosa pines were inoculated with fungal plugs after short roots appeared from lateral roots, the stage of root development considered to be most receptive to colonization (4,5). Inoculum plugs were prepared by removing 4 mm cork borings from the outer edge of 4 week old solid cultures and incubating them in liquid medium until the fungal plug was engulfed in new mycelium. The plugs were then used to inoculate the pines by placing a plug next to a group of short roots, typically comprised of 3-4 roots (Figure 1). Following inoculation, the plant roots and fungi were examined every three days for fungal growth and developmental changes, such as growth toward roots, formation of the mantle, and the appearance of extramatrical hyphae. When a root had been colonized, the inoculum plug was removed and a filter paper disc laden with l^C-atrazine carefully placed in individual pouches so that it was in contact (covered) with a dense region of extramatrical hyphae and a few branch roots. The ^C-laden discs were prepared by asceptically transferring 0.5 ml of ^Catrazine stock solutions (2 pCi-ml ) with a specific activity of 19.1 pCi-pmole" onto autoclaved filter paper discs. The transfers were made with a syringe fitted with a 0.2 pm Acrodisc filter. The discs were left in the transfer hood until the solvent carrier evaporated from the discs. Following addition of the ^C-discs, each pouch was wrapped around a small piece of PVC pipe cylinder (4x10 cm), and a rubber band was used to fit and hold the pouch to the cylinder thereby securing the position of the l^C-disc inside the pouch. Pouches prepared in this manner were then placed into the incubation chambers (Figure 2). The laden discs placed over roots in the pouches imbibed approximately 0.5ml of H2O giving an exposure conentration of ΙΟΟμΜ atrazine. The metabolism of l^C-atrazine was determined by comparing the release of C 0 2 from pouches containing: plant, plant-fungus (mycorrhizae), and no living w a s

-1

1 4


1


f!

Paper Towel

Short Root Lateral Root

Fungal Plug

Disposable Pipet

I POUCH I

liter Disk

Figure 1. Diagrammatic representation of the inoculation of ponderosa pine with Hebeloma crustuliniforme. 1) Insertion of the fungus plug in proximity to short roots. 2) The inoculum plug is removed when extramatrical hyphae appear and the disc is added with radiolabeled compound.

J POUCH I

44



Figure 2. Exposure chamber and trap system used for testing the ability of H.crustuliniforme to metabolize organic compounds.



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PHYTOREMEDIATION OF SOIL AND WATER CONTAMINANTS

organisms (control). The plant and plant-fungus tests were run in triplicate and the control in duplicate. Specially designed exposure chambers were used to measure the metabolism of l^c-atrazine to ^ C 0 2 (Figure 2). Individual chambers holding a single pouch were comprised of a glass cylinder (30 cm tall, 90 cm diameter) with two rubber stoppers at each end. The top and bottom stoppers were fitted with glass tubes inserted through them to permit air to enter the bottom and leavefromthe top. Each chamber was connected in series by Tygon tubing with two C02 traps, and an empty, dry catch-tube. The air entering the CO2 traps (35x200 mm test-tube) was drawn through a fritted glass bubbler tube submerged in 40 ml of full-strength Carbo-Sorb a CO2 trapping agent purchased from Packard Instruments (Downers Grove, IL). The second CO2 trap was used to monitor the possible saturation of the first trap that was intended to retain all evolved CO2. A steady flow of air was pulled through the chamber/trap system by a vacuum pump. During the course of a 32 day incubation period water was added to individual pouches through a sterile needle, which was inserted through the top stopper and into a sterile pipette held in each pouch (Figure 2). Water was added as needed from a syringe fitted to an Acrodisc sterilizing filter (Fisher) attached to the needle. Control pouches possessing no plants or fungi also received water. All chambers were illuminated 14 h a day with a beam of light transmitted through fiber optics. The beam was focused on the needles of the ponderosa pine in chambers holding plants. Each chamber received approximately 500-600 pEinsteins of illumination from 17 fiber-optic fibers whose ends were positioned approximately 3 cm from the glass chamber. The bundle of 17 fibers ran to a trunk line connected to a Fiberstars Inc. (Fremont, CA) illuminator model FS-FIB-401S equipped with a Fiberstar silver halide bulb. This novel method of illumination eliminated the temperature problems normally associated with conventional lighting of small enclosed chambers. Radioactivity (cpm) evolved from each incubation chamber was determined at regular intervals by collecting two 2 ml samples from each trap (Figure 2). Prior to each sampling the absorbent volume was adjusted to the starting volume (40 ml) by adding additional Carbo-Sorb. Each 2 ml Carbo-Sorb sample was combined with 2 ml of Permafluor E+ scintillation fluid purchased from Packard Instruments. The mixed samples were left in the dark for 24 h and then the radioactivity was determined with a Beckman scintillation counter set for 10 min assays. The total cumulative amount of radiation (cpm) evolved over time was determined for each incubation chamber. At each sampling time the cpm values of the two 2 ml aliquots removed from a trap were averaged and then adjusted for background radiation by subtracting the cpm value of the two control chambers that possessed no plant or fungal tissue. The amount of radiation in each 40 ml trap at each sampling time was determined by multiplying the cpm value of the 2 ml aliquot by 20. The cumulative amount of radiation that had been released from each incubation chamber at each successive sampling time was determined by adding the sum of the cpm values of previously removed 2-ml aliquots. The percent mineralization of the starting atrazine-i^c to ^ C 0 2 at different times during the course of the experiment was determined by dividing the total acumulative amount of released as CO2 by the amount of radiation provided at the beginning of the experiment as l^C-atrazine The rate of atrazine degradation to ^C02 was calculated by dividing the pmoles of atrazine metabolized per day by the dry weight of either the root-fungus or the estimated fungus in contact with the exposure disc at the completion of the experiment. The total root-fungus weight was established by removing the paper towel and associated root-fungal tissue from individual pouches. The root and fungal tissue directly beneath the exposure disc were removed and dry weights were



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determined. The amount of fungal weight was then estimated by considering 25% of the total weight to be fungal weight. This correction was based on the 25-40% root to fungal percent ratios reported by Harley and Smith (6) and knowledge that the fungal masses observed in the pouches in this study were considered less developed than hyphae associated with plant roots grown in soil. For comparative purposes, the percent mineralization rates reported by Donnelly et al. fi) for several different ftmgi were converted to pmoles atrazine-day" 1-mg dry weight"! (Table I). This was accomplished by multiplying the reported percent mineralization rates by the total pmoles of atrazine provided and dividing these rates by days and fungal weight. Analysis of variance was used to compare the ability of plant and plant-fungus to mineralize l^C-atrazine. The radioactivity present in the last four samples collected were used for these analyses. Treating these as equivalent samples even though they were not collected on the same day was justified by virtue of the fact that the release of radioactive C02 had stopped, thereby suggesting that the substrate had been used up and all of the last four values were equally representative of the terminal state of mineralization. Results and Discussion Carbon-14 was releasedfromboth the mycorrhizae and plant-only systems throughout the course of the experiment. There was substantial variability among the replicates of both the mycorrhizae and plant systems, but in general the amount of ^C02 produced from the mycorrhizae replicates exceeded thatfromthe plant replicates throughout the experiment (Table Π). Statistical analyses of the acumulative ^C02 values at the end of the experiment showed a significant difference between the two systems. The rate of atrazine mineralization by the mycorrhizae was 2-fold faster than the rate by the plant alone (Figure 3). The enhanced rate may have been due to direct metabolism of the xenobiotic by the fungus or perhaps facilitated movement of the compound into the plant root through the fungal hyphae, followed by plant metabolism of atrazine. In either event the ectomycorrhizal system enhanced the degradation of atrazine. Previous studies with pure cultures of ectomycorrhizal fungi have shown that this group of fungi have the genetic capability to degrade xenobiotics, but the degradation of atrazine by the mycorrhizal system used in this study is the first evidence of enhanced xenobiotic degradation by roots colonized with mycorrhizae. The rate of atrazine mineralization by the plant-fungus system was compared to rates reported in the literature for pure cultures of ectomycorrhizae by converting the cpm values in Table I to pmoles atrazine-day"!-nig dry weight" 1. This conversion was made by using the specific activity of the parent material to convert the cpm of ^C02 released to pmoles of atrazine metabolized. Rates of atrazine degradation to ^C02 were calculated by dividing the pmoles of atrazine metabolized per day by the dry weight of tissue in contact with the exposure disc at the completion of the experiment. Data from the work of Donnelly et al. (1) were recalculated in a similar fashion as explained in the materials and methods. The rate of atrazine degradation calculated for Hebeloma crustuliniforme and ponderosa pine roots was 9.0x10"7 pmoles atrazine-day" 1 -mg dry weight" 1. Assuming that the fungal tissue was responsible for all of the degradation and that the fungus accounted for 25% of the total root-fungal weight (6) then it was estimated that the degradation rate of Hebeloma crustuliniforme by itself was 3.6x10"^ pmoles atrazine-day" 1-mg dry weight "1. This value is approximately 10-fold lower than values calculatedfromthe pure culture data reported for several different species of ectomycorrhizae by Donnelly et al (7). The difference in rate between the two studies involves several possible explanations. The lower rate



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Table I. Mineralization of Atrazine (pmoles atrazine-day-i-mg dry weight" 1) by four different ectomycorrhizal fungi in pure culture as compared to roots of Pinus ponderosa with colonized Hebeloma crustuliniforme. Colonized Plant Pure Culture^ Rhizopogon Gauteria Gauteria Radiigera Root and Fungus othii atragleba Fungus vulgaris crispa 3.6xl09.0xl09.9xl0" 8.3xl0" 2.3xl03.3xl0*Data takenfromDonnelly et al fij. Weight of fungus was considered to be 25% of the total weight of the colonized root (6) 2

5

5

5

5

7

6

2

c

0.21

0.1 H

Day Into Experiment Figure 3. Mineralization of atrazine (p=0.13). Error bars represent standard deviation around the mean of thefinaltwo samples pooled together.



GASKIN & FLETCHER

Table Π. Amount of radioactivity (cpm) recovered in CO2 when l^C-atrazine was provided to different systems PLANT PLANT-FUNGUS Day a Β C A B C 2 172 204 196 166 173 170 Sample 22 54 46 16 23 20 Corrected Sample 440 1080 920 320 460 400 Total Count* 440 1080 920 320 460 400 Cumulative Count 170 187 197 213 234 228 6 Sample 20 37 48 63 84 79 Corrected Sample Total Count 1260 1680 1580 400 740 960 Cumulative Count 1304 1788 1672 432 786 1000 177 182 173 209 239 265 10 Sample 27 32 23 Corrected Sample 59 89 115 Total Count 1080 1780 2300 540 640 460 Cumulative Count 1250 2056 2550 612 760 596 169 191 239 258 239 312 14 Sample 19 41 89 Corrected Sample 104 89 162 Total Count 2080 1780 3240 380 820 1780 Cumulative Count 2368 2234 3720 506 1004 1962 182 203 238 272 253 349 18 Sample 32 53 88 Corrected Sample 122 103 199 Total Count 2440 2060 3980 640 1060 1760 Cumulative Count 2936 2692 4784 804 1326 2120 181 212 243 272 230 382 22 Sample 31 62 93 122 80 232 Corrected Sample 2440 1600 4640 620 1240 1860 Total Count 3180 2438 5842 848 1612 2396 Cumulative Count 174 179 249 286 248 403 26 Sample 24 29 99 Corrected Sample 136 98 253 2720 1960 5060 480 580 1980 Total Count 3704 2958 6726 770 1076 2702 Cumulative Count 173 220 239 270 216 405 30 Sample 23 70 89 Corrected Sample 120 66 255 Total Count 2400 1320 5100 460 1400 1780 Cumulative Count 3656 2513 7272 798 1954 2700 A , B, and C represent triplicate exposure chambers ^The cpm of each sample taken. The subtraction of background radiation (average pouch cpm 150). The correction of volume by multiplying the cpm of a 2ml sample by 20, which brings the volume up to 40ml. The addition of all previous 2ml samples removed. A


b

0

1

6

a

c

d

e


160



may be a feature of species difference since unfortunately Hebeloma crustuliniforme was not a part of the Donnelly study and no comparable pure culture data on xenobiotics metabolism is available from another source. Another possibility is that ectomycorrhizal fungi on colonized roots do not metabolize xenobiotics as they do in pure culture, and the degradation observed in our study was actually plant degradation facilitated by hyphal uptake of atrazine. A third explanation is that the fungal tissue was the primary contributer to atrazine degradation, but we overestimated the amount of fungal tissue by assuming that 25% of the fungal root weight was due to the fungus. Each of these possibilities or some combination of them serve as explanations for the 120-fold difference in reported rates (Table I), and emphasizes the need for additional research. Summary The mycorrhizal system of ponderosa pine and Hebeloma crustuliniforme was shown to mineralize atrazine to a greater extent than the plant by itself. The ability of ectomycorrhizae to enhance the metabolism of atrazine when the fungus is associated with the host plant suggests that ectomycorrhizae could play an active role in remediating organic, soil pollutants. The advantage of ectomycorrhizae systems in bioremediation over that of other systems such as free-living bacteria or fungi is that degrading strains of these organisms introduced at toxic waste sites are often difficult to maintain for extended periods of time because of competition from wild types whereas ectomycorrhizae introduced along with their host plant theoretically have a survival advantage over competing organisms since the host plant provides an ecological niche for the ectomycorrhizal fungus. The general ecology and structure of ectomycorrhizae are also favorable for soil bioremediation since the long hyphae of the fungus, increase the surface area already explored by roots, and thereby greatly increase the contact between potentially bioreactive organisms and soil contaminants. Although ectomycorrhizae hold great promise in bioremediation, the slow rate of mineralization observed with our test system (ponderosa pine/Hebeloma crustuliniforme) clearly shows that the maximum potential of ectomycorrhizal systems in bioremediation can only be realized through more intensive efforts to identify those plant-fungal systems with the greatest genetic potential to degrade a broad spectrum of xenobiotic compounds. Literature Cited 1. Donnelly, P.K.; Entry A.E.; Crawford, D.L. Appl. Environ. Microbiol. 1993, 59, pp. 2642- 2647 2. Robideaux, M . L . ; Fowles, N.L.; King,R.J.; Spriggs, J.W.; Rygiewicz P.T. Abst.l2 Annual Meeting. Society of Environmental Toxicology and Chemistry, Seattle, W A ; 1991, p. 175. 3. Donnelly, P.K.; Fletcher J.S. Bull. Environ. Contam. Toxicol. 1995, 54, pp. 507-513. 4. Fortin, J.Α.; Piché, Y ; Godbout, C. Plant and Soil. 1983, 71, pp. 275-284. 5. Fortin, J.A.; Piché, Y.; LaLonde, M . Can. J. Bot. 1980, 58, pp 361-365. 6. Harley, J.L.; Smith, S.E. Mycorrhizal Symbiosis. Academic Press, London. 1983. th


The Metabolism of Exogenously Provided Atrazine by the

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