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BIODEGRADATION OF MONOETHANOLAMINE, ETHYLENE

GLYCOL AND TRIETHYLENE GLYCOL IN LABORATORY

BIOREACTORS

OLE MRKLAS1,, ANGUS CHU2, STUART LUNN3 and LAURENCE R. BENTLEY1

1Department of Geology and Geophysics, University of Calgary, 2500 University Drive NW,

Calgary, AB T4N 1N4, Canada; 2Department of Civil Engineering, University of Calgary, 2500

University Drive NW, Calgary, AB T4N 1N4, Canada; 3Imperial Oil Resources, 237 Fourth Avenue

SW, Calgary, AB T2P 0H6, Canada

(*author for correspondence, e-mail: [email protected], Tel: 403.237.4289)

(Received 8 April 2003; accepted 9 June 2004)

Abstract. The release of alkanolamines and glycols into the subsurface soils poses a potential hazard

to the environment through impacted soil and groundwater. This study investigated aerobic and

anaerobic biodegradability of monoethanolamine (MEA), ethylene glycol (MEG) and triethylene

glycol (TEG). Significant levels of MEA (31 000 mg/kg), MEG (500 mg/kg) and TEG (2100 mg/kg)

were successfully aerobically biodegraded in bioreactors. The aerobic slurry experiments suggested

initial phosphate (P) limitation, as biodegradation rates increased by one order of magnitude after

phosphate addition. Anaerobic decay of MEA, MEG and TEG was unaffected by P-addition. MEA,

MEG and TEG degradation products such as acetate, ethanol and ammonium at about 75 000 mg/kg,

8100 mg/kg and 8800 mg/kg degraded completely and did not prevent aerobic biodegradation. This

study confirms proposed biodegradation pathways of MEA, MEG, TEGand their breakdown products

in natural soil and groundwater using indigenous microbes. Levels of contamination studied here aresignificantly higher than previously reported.

Keywords: biodegradation, contamination, degradationproducts, ethanolamine, glycol, groundwater,

subsurface

1. Introduction

Alkanolamines and glycols have been commercially available for decades. The

utilization of these substances includes both household and industrial applications.Glycols and alkanolamines are extensively employed in the sour natural gas in-

dustry. Sour natural gas production is a significant portion of the global energy

markets. Alberta produces the majority of Canadian natural gas with a sour natu-

ral gas portion of approximately 40% (Sorensen et al., 1999). Alkanolamines are

basic derivatives of ammonium. The optimization and modification of the sweet-

ening process has resulted in a diverse composition of alkanolamine and glycol

mixtures for specific applications (Polaseket al., 1992; Rooneyet al., 1998). Unin-

tentional introduction of ethanolamine, ethylene glycol and triethylene glycol into

Water, Air, and Soil Pollution 159:249263, 2004.C2004Kluwer Academic Publishers. Printed in the Netherlands.

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250 O. MRKLAS ET AL.

the subsurface and groundwater is a potential hazard to the environment (Fedorak

et al., 1997; Gallagher et al., 1995; Lintott et al., 1997; Sorensen et al., 1999;

Wrubleski et al., 1997). After their release, alkanolamines and glycols may migrate

into the subsurface and groundwater. The high density and high water solubility of

these species influence their fate and transport in the subsurface and in groundwa-

ter (Sorensenet al., 1999). The combination of chemical properties and structures

inherent to these species makes alkanolamines and glycols readily biodegradable.

However, the behavior of alkanolamines in concentrations exceeding 3000 mg/L in

the presence of high levels of aerobic and anaerobic breakdown products (i.e. am-

monium, acetate and ethanol) is not well understood, especially in environmental

applications. In fact, studies looking at in-situfate and transport of alkanolamines,

glycols and degradation products have been complicated by complex geological andhydrogeological properties (McVickeret al., 1997; Sorensenet al., 1999; Strong-

Gundersonet al.,1995). In addition, it was demonstrated that MEA concentrations

exceeding 1500 mg/kg inhibitedin-situbiodegradation (Sorensenet al., 1997).

This study assesses the biodegradability of MEA, MEG and TEG in labora-

tory bench scale bioreactors. Major breakdown products of MEA, MEG and TEG

biodegradation are ethanol, acetic acid and ammonium (Bradbeer, 1965; BUA,

1994; Jones and Turner, 1973; McVickeret al., 1997). Ethanol and acetic acid may

be completely degraded via the tri carboxylic acid cycle (aerobic) or by methano-

genesis (anaerobic) (Gottschalk, 1985). Ammonium degrades by nitrification and

denitrification using shift conditions(aerobic followed by anaerobic conditions) and

carbon source supplementation. The evolution of MEA, MEG and TEG and their

breakdown products ammonium, acetic acid and ethanol is discussed with respect to

the implications for in-situ aerobic and anaerobic degradation at a decommissioned

gas plant site. Indicators such as electrical conductivity and pH measurements may

assist in monitoring degradation patterns. Explaining aerobic and anaerobic degra-

dation pathways in laboratory slurry phase experiments will assist in understanding

in-situprocesses and will aid in the design of optimal degradation conditions for

remediation regimes.

2. Methods and Materials

2.1. CHEMICALS

All chemicals were ACS grade, and purchased from Sigma-Aldrich, Canada or

Fisher Scientific, Canada.

2.2. BIOREACTORS

All bench scale bioreactors utilized the same source of soil and groundwater from

the site at a concentration of 40% w/w (100 g soil + 150 g groundwater). Initial

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BIODEGRADATION OF MEA, MEG AND TEG 251

soil and groundwater sample analyses indicated that the soil samples showed am-

monium, but no MEA concentrations and groundwater samples had both ammo-

nium and MEA concentrations. The soil and groundwater samples originated from

the former process area of a decommissioned sour gas plant site from a depth of

about 4 m. This location was chosen, because previous electrical resistivity tomog-

raphy (ERT) measurements suggested high impact levels. The slurries consisted

of four aerobic (1, 2, 3, 4), two replicate aerobic-abiotic controls, two replicate

anaerobic, and two replicate anaerobic-abiotic controls. The abiotic control slur-

ries were autoclaved three-times on three consecutive days for 1 h at approximately

100 C in autoclave bags. Soil and water samples were analyzed to determine ini-

tial concentrations after autoclaving. All Erlenmeyer flasks and glass jars were

autoclaved prior to use. Aseptic sampling procedures were used for the abioticcontrols.

2.2.1. Aerobic Slurries

The aerobic bioreactors were contained in four biotic (labeled 1, 2, 3, 4) and two

abiotic open 500 mL Erlenmeyer flasks. The flasks were wrapped with tin foil

to prevent algae growth. The flasks were mounted onto a shaker table and mixed

gently at constant speed (approximately 300 rpm). The speed was adjusted to ensure

no sedimentation. The geometry of the Erlenmeyer flasks and the fill level (about

3 cm) facilitated optimal mixing and maximal slurry/air interface area for oxygen

transfer. The aerobic bioreactors were weighed during each sampling interval and

sterile distilled deionized water was added to adjust for evaporation. The aerobicabiotic reactors were fitted with a porous foam stopper, but were not compensated

for evaporation in order to preserve aseptic conditions. Reactors 3 and 4 received

about 40 mg (P)/kg phosphate addition using potassium-di-hydrogen-phosphate

(KH2PO4) on day 11. Slurries 1 and 2 received the same treatment on day 64 of the

experiment. On day 92, slurries 3 and 4 were transferred into an anaerobic glove

box and received 200 mM acetic acid. MEA, MEG, TEG and degradation product

(acetic acid, ethanol, ammonium, nitrite, nitrate) concentrations were monitored.

The aerobic bioreactors were monitored at ambient temperature of 23 C 0.7 C.

2.2.2. Anaerobic Slurries

A dry glove box with continuous nitrogen gas flow under positive pressure createdan oxygen-free environment for two biotic and two abiotic anaerobic slurries. Ap-

proximately 250 g slurry material (40% w/w soil) was contained in each 500 mL

glass jar that was wrapped with aluminium foil to prevent algae growth. The lids

were equipped with a pressure release line open to the nitrogen atmosphere. The

slurries were mounted onto magnetic stirrers and intermittently stirred at constant

speed approximately 1 h before each sampling. Anaerobic slurries received 40 mg

(P)/kg phosphate as KH2PO4on day 11. The anaerobic slurries were held at constant

temperature of 22 C 1 C.

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2.3. ANALYTICAL PROCEDURES

The slurry sampling procedure involved the removal of about 1 mL of slurry ma-

terial from the bioreactors. The samples were centrifuged at 10 000 rpm for 5

min. The supernatant was completely removed (water extract). Subsequently, the

remaining soil at a determined moisture content was extracted with 1 M potas-

sium hydroxide (KOH) solution, mixed for 1 h and centrifuged (KOH extract).

The KOH extraction procedure was found to effectively remove sorbed MEA and

ammonium left on the soil after the original soil water extraction. Water and KOH

extracts were analyzed. MEA, ammonium, sodium, magnesium and calcium were

quantified using cation exchange chromatography with suppressed conductivity de-

tection in water mode. Acetic acid, chloride, nitrite, nitrate, phosphate and sulfatewere determined by anion exchange chromatography with chemical suppressed

conductivity detection. Anions were monitored in the water extract. MEG, TEG

and ethanol determinations were carried out on water extract by ion exclusion chro-

matography with pulsed amperometric detection. Detailed analytical protocols are

described elsewhere (Mrklaset al., 2003; Mrklas, 2002). In addition, during each

sampling interval all slurries were monitored for electrical conductivity (EC), pH

and temperature. The EC values were corrected to a reference temperature of 20 C

using external calibration with EC standards at various temperatures and linear

regression.

3. Results

The analyses of the initial mixture of collected soil and groundwater samples re-

sulted in concentrations of approximately 31 000 mg/kg MEA, 500 mg/kg MEG

and 2100 mg/kg TEG (Table I). The concentrations are comparable to in-situlev-

els found on-site. Table I outlines the average product and degradation product

concentrations in the soil, groundwater and slurry samples at the beginning of the

aerobic and anaerobic slurry experiments. Additionally, initial (final) phosphate

concentrations, pH and EC were recorded (Table I). Table II summarizes the degra-

dation pathways and Tables III and IV show the biodegradation rates for the various

species.

3.1. AEROBIC BIOREACTORS

The aerobic reactors were monitored for 98 days. Duplicate reactors showed sim-

ilar behavior and results of replicates are averaged here. Results from slurries 1

and 2 were averaged (labeled +P64) and monitored without phosphate enhance-

ments for the first 64 days, and with P-enhancements (40 mg P/kg) after day 64

(Figures 1AC). The results of biorectors 3 and 4 were averaged (labeled

+P11). Biodegradation of MEA, MEG and TEG was monitored without P-addition

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TABLE I

Characterization of initial groundwater, soil and slurry concentrations

Concentration

Species Groundwater(mg/L) Soil(mg/kg) Slurry(mg/kg)

MEA 16205 463 ND 30911 437

MEG NA NDa/ND 474 32

TEG NA 490a/514 2108 279

Ammonium 3034 75 2276 281 8843 64

Acetic acid 36410 1137 ND 75139 2347

Ethanol 3938 82 ND 8127 169

Phosphate, initial (final) NA NA

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TABLE III

Average degradation rates (/S.D.) of carbon species in aerobic bioreactor sets+P64

and +P11

+P11 +P64a

Compound Aerobic (day1) Aerobic P-enhanced (day1) Aerobic (day1)

Ethanol 0.56 0.04 0.56 0.04

MEG 0.067 0.001 0.02

Acetic acid 0.018 0.006 0.47030 0.00001 0.014 0.005

MEA 0.011 0.001 0.10 0.03 0.0078 0.0001

TEG 0.0115 0.0003 0.145 0.001 0.005 0.002

aBioreactor +P64 degradation rates P-enhanced not shown due to insufficient sam-

pling frequency.

TABLE IV

Average anaerobic biodegradation rates(/S.D.)

Compound Anaerobic (day1) Degradation kinetics

Acetic acid 0.0054 0.0001 first order

Ethanol 0.0040 0.0007 first order

MEA 0.0022 0.0002 zero order

TEG 0.0023 0.0003 zero order

+P64 showed steady acetic acid decay until day 64 and significant rate increases

were observed after day 64. (Figure 1A). The abiotic controls showed steady acetic

acid concentrations.

The MEG concentrations decreased initially at higher rates in both averaged

values +P64 and +P11. Average MEG concentrations (+P64) decreased by 36%

until day 7 and then did not significantly change until phosphate addition on day

64. MEG concentrations (+P11) declined to below detection after day 15 and

after P-addition on day 11 (Figure 1B). The abiotic controls showed steady MEG

concentrations.

MEA biodegradation had zero-order biodegradation rates (Figure 1C). Priorto P-addition the average MEA biodegradation rates +P11 and +P64 were sim-

ilar (Table III). However, these rates increased by one order of magnitude after

P-addition in both reactor sets (Figure 1C). The abiotic controls showed steady

MEA concentrations. Biodegradation rates and standard deviations are presented in

Table III and followed zero-order degradation kinetics.

MEA and ammonium partitioned into soil and water fractions (Figures 1C).

The KOH extraction method resulted in nearly complete desorption of MEA

and ammonium as indicated of recovery rates of 109 and 97% (n = 3),

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A

0

20000

40000

60000

80000

100000

0 20 40 60 80 100Time [d]

C[mg/kg]

Ac +P64 Ac +P11 Ac abiotic EtOH +P64 EtOH +P11

P-addition day 11 P-addition day 64

B

0

500

1000

1500

2000

2500

0 20 40 60 80 100Time [d]

C[mg/kg]

MEG +P11 MEG +P64 TEG +P64 TEG +P11

`


C

0

10000

20000

30000

40000

0 10 20 30 40 50 60 70 80 90 100

Time [d]

C[mg/kg]F

MEA T+P11 MEA W+P11 MEA S+P11 MEA abiotic MEA T+P64


Figure 1. Average (a) acetate and ethanol concentrations, (b) MEG and TEG concentrations and (c)

MEA concentrations in soil (S), water (W) and total (T) in [mg/kg] during aerobic biodegradation

versus time in [days]. Phosphate addition occurred on day 11 (+P11) and day 64 (+P64) as indicated

by the arrows. Error bars represent one standard deviation.

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respectively. The recovery rates were calculated as detected MEA and ammo-

nium concentrations. The total concentrations [mg/kg] were calculated as the sum

of water extract concentration in [mg/kg] and the amount in the KOH extract in

[mg/kg].

TEG biodegradation rates +P11 increased by one order of magnitude after day

21 and reached below detection levels after 32 days post P-addition (Figure 1b).

Average TEG biodegradation rates +P64 were slower until the phosphate addition

on day 64. The abiotic controls showed steady TEG concentrations.

The carbon utilization in all aerobic bioreactors occurred in the order ethanol,

MEG, acetic acid, MEA and TEG. The EC values followed similar trends as

acetic acid and MEA concentrations in the biotic and abiotic reactors, until day 60

where elevated nitrite and nitrate concentrations caused an increase in EC values(Figures 1AC, 2 and 3).

3.2. NITROGEN DISTRIBUTION

The aerobic slurries were monitored for nitrogen containing species MEA, ammo-

nium, nitrite and nitrate. Figure 3 shows averaged nitrogen concentrations +P11.

The concentration changes (+P11) of nitrogen containing species observed were

divided into four phases labeled A, B, C, and D. Phase A represents pre-phosphate

enhanced conditions and spanned the first 11 days of the experiment. Increased

carbon utilization due to P-enhancement occurred in phase B and aerobic ni-

trogen transformation from ammonium to nitrite and nitrate is seen in phase C.Phase D started on day 92 after transferring the aerobic slurries 3 and 4 into

an anaerobic environment and consequently anaerobic utilization of nitrate by

denitrification.

0

4

8

12

16

20

0 20 40 60 80 100Time [d]

E

C[mS/cm]

EC +P11 EC +P64 EC abiotic


Figure 2. Electrical conductivity (T= 20) in [mS/cm] in aerobic slurries+P64,+P11 (biotic) and C1

(abiotic) versus time [days]. Phosphate addition occurred on day 11 (+P11) and on day 64 (+P64).

Error bars represent one standard deviation.

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0

100

200

300

400

500

0 20 40 60 80 100Time [d]

C(N)[mmol/kg]

NH4 T-N NH4 S-N MEA T-N NH4 W-NNO2 NO3

A B C D

+PNH3

NH4+

NO3-

N2

NO2-

Figure 3. Nitrogen species concentrations: Total ammonium-nitrogen (NH4 Total) and ammonium

nitrogen in KOH extract (NH4 Soil), total MEA nitrogen (MEA Total), total nitrite nitrogen (NO2)

and total nitrate nitrogen (NO3) concentrations in [mmol-N/kg] averaged +P11 versus time [days].

Phases A, B, C and D indicate stages of biodegradation.

Overall ammonium concentrations decreased during the entire period of the ex-

periment. Figure 3 presents ammonium concentrations sorbed onto the soil fraction

and total concentrations in +P11. Total ammonium levels rapidly decreased during

phase A and until day 15. A slower decrease was observed between day 15 and

day 58, as indicated in phase B. A sharp decline was observed during phase C

after day 58. The sorbed ammonium concentrations were constant until day 13 and

increased from 79 mmol-N/kg to 175 mmol-N/kg until day 23 during phase B of

the experiment. The MEA degraded in phase B from 102mmol-N/kg starting on

day 13 to below detection. A sharp decrease of ammonium levels was observed

in both soil and total concentrations after day 58. The total ammonium concentra-

tion in the two abiotic controls averaged 420 58 mmol-N/kg during the entire

study.

Average nitrite concentrations (+P11) increased on day 58 and reached an aver-

age maximum of 54 mmol-N/kg in phase C on day 72. Nitrate production occurred

on day 64 and reached an average maximum level of 29 mmol-N/kg on day 92. In

phase D and after day 92, slurries 3 and 4 were placed in a dry glove box under

anaerobic conditions. The addition of 200mmol/kg acetic acid supplied sufficientavailable organic carbon to initiate anaerobic denitrification. The nitrate levels de-

clined to below detection levels on day 98 in phase D of the experiment. Finally,

bioreactors 3 and 4 (+P11) indicated that MEA, MEG, TEG, acetic acid, ethanol,

ammonium, nitrate and nitrite concentrations were below detection on day 98 at

the end of the experiment. The pH values increased with declining acetic acid

and increasing ammonium. As ammonium concentrations declined, pH declined

as well (Figure 4). The abiotic controls indicated no significant pH changes during

the entire experiment.

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6.00

7.00

8.00

9.00

10.00

0 20 40 60 80 100Time [d]

pH

[-]

pH +P11 pH +P64 pH abiotic

A B C D

+P +P

Figure 4. Average aerobic pH values +P64, +P11 (biotic) and control (abiotic) versus time [days].

Error bars represent one standard deviation. Phases A, B, C and D indicate stages of biodegradation.

0

20000

40000

60000

80000

0 20 40 60 80 100Time [d]

C[mg/kg]

MEA total EtOH MEA abiotic NH4 total TEG Ac

P-addition day 11

Figure 5. Average anaerobic biotic concentrations and control (abiotic) of acetate, ethanol, MEG,

TEG and MEA [mg/kg] during biodegradation versus time [days]. Error bars represent one standard

deviation.

3.3. ANAEROBIC BIOREACTORS

The four anaerobic slurries were monitored for 98 days (Figure 5). The anaerobic

bioreactors A1 andA2 showedsimilar degradationtrends andresults were averaged.MEA, MEG and TEG degradation was monitored without phosphate addition until

day 11 and with P-addition after day 11. Table 1 presents the initial concentrations

of chemical species in the soil, groundwater and slurries.

The alkanolamine, glycols and degradation products were not completely de-

graded in the anaerobic bioreactors. The biotic bioreactors showed MEA and TEG

concentrations decreasing by 23 and 25% during the study. Ammonium concen-

trations were constant in both abiotic and biotic anaerobic slurries. MEG concen-

trations in the biotic anaerobic slurries were stable during the entire period of the

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study and similar to MEG concentrations in the abiotic reactors. The biodegrada-

tion rates for the compounds were generally one magnitude lower than the aerobic

biodegradation rates (Tables 3 and 4).

4. Discussion

The presence of MEA, MEG, TEG and their breakdown products ethanol, acetic

acid and ammonium in the collected soil and groundwater samples agrees with

previous proposed degradation pathways of alkanolamines and glycols (Bradbeer,

1965; BUA, 1994; Jones and Turner, 1973; McVicker et al., 1997). In general,

MEA, MEG and TEG biodegradation results in ethanol, acetic acid and ammoniumevolution through deamination (MEA) and dehydrogenase (glycols) (Gottschalk,

1985).

The species present (MEA, ammonium) supplied adequate nitrogen concentra-

tions and it was observed that the aerobic biodegradation increased by one order

of magnitude and was successfully completed after the addition of phosphate. The

aerobic bioreactors showed complete biodegradation of ethanol, acetic acid, MEG,

MEA and TEG within 21 days after phosphate addition. The 3000 mg/L MEA

concentrations found in this study degraded completely and did not inhibit aero-

bic biodegradation. These biodegradation rates agree with recent literature (BUA,

1994; Philip, 1991; Sorensen et al., 1997). Therefore, it was concluded that the

slurries were initially operated under phosphate-limited conditions during the first

phase of the experiment. These results suggest that aerobic field conditions with

adequate phosphate supplies are optimal for MEA, MEG and TEG biodegradation.

Significant changes in anaerobic biodegradation rates were not observed with the

addition of phosphate. The anaerobic biodegradation rates of ethanol, acetic acid,

MEA and TEG were constant even after phosphate addition during the entire period

of the experiment. Acetic acid, however, seemed to be the preferential substrate in

the anaerobic reactors, as the degradation rate constants were the highest. MEG

concentrations did not change during the experiment. Therefore, anaerobic degra-

dation of ethanol, MEA, MEG and TEG may be inhibited by present acetic acid and

ammonium concentrations. In fact, it was previously shown that acetic acid concen-

trations of approximately 2000 mg/L significantly inhibited methane production in

anaerobic environments (Mawsonet al., 1991). Furthermore, it was reported thatanaerobic methane production was inhibited due to high ammonium/ammonia con-

centrations (Gallertet al., 1998). Thus, anaerobic decay of ethanol, MEA and TEG

was much slower than aerobic degradation, possibly attributed to inhibition and

was not complete at the end of the 98 day experiment.

MEA has been described as an ammonium derivate with comparable proper-

ties (Bollmeier, 1991; Dow, 1980). Ammonium can be sorbed onto clay minerals

(Lumbanraja and Evangelou, 1990; Olsen et al., 1999). MEA distribution between

water and soil played a major role in recent laboratory and field investigations

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(Mrklaset al., 2001; Mrklas 2002; Sorensen et al.,1999). This study presents for

the first time the detailed distribution of both ammonium and MEA in water and

soil fractions in slurry reactors during enhanced aerobic and anaerobic biodegrada-

tion. Earlier studies determined MEA and ammonium distribution coefficients of

about 1 L/kg in the same soil samples employed here (Mrklas, 2002). These distri-

bution coefficients suggested that sorption may have limited biodegradation. The

results presented here show that MEA biodegraded faster in the water phase than

MEA sorbed on the soil phase. Thus available or free MEA concentrations in the

water phase were limited by sorption during enhanced biodegradation. Bioavail-

ability of substrate plays a major role in biodegradation of contaminants (Alexander,

1999). Cation exchange and/or sorption fix the substrate and may inhibit microbial

utilization of the sorbed substrate (Nielsen et al., 1996). Therefore, the distribu-tion of substrate in a soil-water mixture determined the availability of substrate

in the water phase and, thus influences the bulk biodegradation rates. Such limi-

tation occurred during the enhanced aerobic degradation of MEA, as MEA con-

centrations in the water phase (available) declined faster than in the soil phase

(sorbed).

In addition, it was demonstrated that a decrease of total MEA concentration due

to biodegradation resulted in increased ammonium sorbed onto the soil fraction

(Figure 3). The biodegradation of alkanolamines occurred partly under ammonium

evolution (Bradbeer, 1965; Jones and Turner, 1973). As MEA molecules desorbed

from the soil phase and partitioned into the water phase the produced ammonium

sorbed onto the available cation exchange sites. The overall ammonium concen-

tration, however, declined as the pH increased to a maximum of 8.9 and ammonia

volatilization became a major release pathway. The pH increase was a direct result

of acetic acid biodegradation (Figures 1a and 4). Ammonium (NH+4) and ammo-

nia (NH3) partitioning is mainly influenced by pH changes (Canter, 1997; Larsen

et al., 2001). The ammonium equilibrium in the slurries consisted of (1) ammonium

ions in the water phase and (2) soil phase, (3) ammonia gas dissolved in the water

phase and (4) ammonia gas in the vapor phase. As the pH increased the portion of

dissolved ammonia gas (NH3) increased in relation to the dissolved ammonium and

consequently resulted in an increase of ammonia gas in the vapor phase escaping

the open bioreactors. At a pH of 8.9 about 30% of the ammonium concentration

is gaseous ammonia and it partitions into the atmosphere following Henrys law

(Larsenet al., 2001). A maximum pH of 8.9 was observed during the P-enhancedphase of the experiment in reactors 3 and 4 and therefore, ammonia volatilization

appeared to be the major mechanism for ammonium loss. Ammonia release will

not be as large a factor in highly buffered field sites, but may play a major role

in areas with high pH values. In general, results demonstrated that it is necessary

to understand the roles of sorption of both MEA and ammonium during MEA

biodegradation.

Nitrification and denitrification were demonstrated using sequential aerobic and

anaerobic conditions in +P11 (Figure 3). The production of nitrite (NO2) before

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nitrate (NO3 ) during the aerobic phase suggested the inhibition of Nitrobacter

species that are capable of nitrate production utilizing nitrite. Anthonisen et al.

(1976) demonstrated that dissolved ammonia and nitric acid concentrations in-

hibited and delayed nitrate evolution in slurry experiments. The formation of the

intermediate hydroxylamine occurs during nitrification and before the evolution

of nitrite (Brock and Madigan, 1991; Gottschalk, 1985). Consequently, we inter-

pret the observed patterns of nitrite and nitrate evolution as being due to inhibi-

tion caused by dissolved ammonia and some loss of nitrogen to hydroxylamine

formation.

Both aerobic and anaerobic abiotic controls demonstrated that chemical trans-

formation of MEA, MEG and TEG was insignificant.

5. Conclusions

This biodegradation study of MEA, MEG, TEG and their degradation products

employed natural soil,groundwater and indigenousmicrobes. Phosphatelimitations

played a significant role in aerobic biodegradation at the concentrations observed

here. This suggests that impacted areas in field scenarios may show elevated levels

for long periods until aerobic conditions and phosphate addition occur.

The KOH method enabled complete MEA and ammonium extraction from the

soil fraction due to increased pH (MEA desorption) and excess potassium con-

centration (ammonium cation exchange). This suggests that MEA sorption may

not entirely be contributed to cation exchange. The implications for the field arethat groundwater sample analyses only determine a portion of the total ammonium

and MEA concentrations in the subsurface. The degradation patterns indicated that

carbon species degraded aerobically in the order ethanol, MEG, acetic acid, MEA

followed by TEG. Therefore, TEG will degrade after the other species in this series

of contaminants and may persist in the environment the longest. The significant

acetic acid concentrations seen here at neutral pH values were buffered by the

system. At sites with low buffering capacity increasing acetic acid concentrations

caused by biodegradation may require carbonate/bicarbonate addition in order to

create neutral pH levels and optimize biodegradation.

Nitrification and denitrification were observed by sequentially changing aerobic

to anaerobic conditions, supplementing with acetate and utilizing indigenous mi-

croorganisms. Shifting aerobic and anaerobic conditions in the field, i.e. seasonal

changes or site heterogeneity (saturated/unsaturated zones) may allow for natural

in-situnitrification and denitrification.

Acknowledgement

This study was supported by the Natural Sciences and Engineering Research Coun-

cil of Canada (NSERC) and Imperial Oil Resources.

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