decomposition of mytilus edulis: the effect on sediment nitrogen and carbon cycling

www.elsevier.com/locate/jembe

Journal of Experimental Marine Biolog

Decomposition of Mytilus edulis: The effect on sediment nitrogen

and carbon cycling

Bente A. Lomstein *, Lise Bonne Guldberg, Jesper Hansen

Department Biological Sciences - Microbiology, University of Aarhus, Building 540, Ny Munkegade, DK-8000 Aarhus C, Denmark

Received 3 June 2005; received in revised form 22 August 2005; accepted 9 September 2005

Abstract

The benthic degradation of mussel tissue (Mytilus edulis) was studied in a continuous flow-through system over a 32 day

incubation period. Sediment chambers without mussels served as controls. The inflowing artificial seawater and the outflow water

were analyzed for dissolved organic nitrogen (DON), short chain fatty acids (SCFA), dissolved inorganic nitrogen (DIN),P

CO2

and O2 during the course of incubation. Sediment profiles of particulate organic carbon (POC), particulate organic nitrogen (PON),

total hydrolyzable amino acids (THAA), pore water concentrations of DON and DIN and turnover rate of dissolved free amino

acids (DFAA) were measured at four different times during the 32 day experiment. Immediately after the addition of mussel tissue,

the chambers became completely anoxic and there was an increase in carbon oxidation and the efflux of DON, SCFA and NH4+

from the sediment+mussel layer to the overlaying water. During the first 9 days there was a net buildup of DON, and NH4+ in the

sediment followed by a net consumption of the respective N-species during the remainder of the experiment. During the course of

incubation 41% of the organic content of the added mussel tissue was released from the sediment as DON, whereas most of the

other mussel-N effluxed the sediment as NH4+. Only 8% of the added mussel-N remained by the end of the experiment. There were

indications of stimulated bacterial growth in both the mussel amended and the unamended sediments. This was measured as a net

increase in THAA, which could only be explained by net bacterial growth and/or protein synthesis. During mussel decomposition

both the estimated bacterial carbon incorporation efficiency and the C:N ratio of the substrates used by the bacteria were low. This

resulted in a low bacterial nitrogen demand. As a consequence, almost all of the nitrogen mineralized within the sediment was

released to the water column as NH4+.

D 2005 Elsevier B.V. All rights reserved.

Keywords: Bacterial growth; Benthic decomposition; Coastal environment; Mytilus edulis; Nitrogen cycling

1. Introduction

Shallow marine sediments are areas of intense bio-

geochemical activity. The sources of organic matter in

these sediments are pelagic and benthic primary and

secondary producers and terrigenious material. The

0022-0981/$ - see front matter D 2005 Elsevier B.V. All rights reserved.

doi:10.1016/j.jembe.2005.09.003

* Corresponding author. Tel.: +45 89423243; fax: +45 89422722.

E-mail address: [email protected] (B.A. Lomstein).

quality and quantity of organic matter introduced into

the sediment have been found to have an overall deter-

mining effect on benthic mineralization (e.g. Blackburn

and Henriksen, 1983; Lomstein et al., 1998; Sloth et al.,

1995; Pedersen et al., 1999; Neubauer et al., 2004). In

order to understand the controls of benthic mineraliza-

tion and hence the fraction of gross NH4+ mineralization

remaining for benthic-pelagic coupling, it is essential to

gain a detailed insight into the different flows and

processes involved in the decomposition process. How-

y and Ecology 329 (2006) 251–264

B.A. Lomstein et al. / J. Exp. Mar. Biol. Ecol. 329 (2006) 251–264252

ever, as discussed by Pedersen et al. (1999) and Neu-

bauer et al. (2004), little is known about the factors

controlling sediment processes and the exchange of

nutrients across the sediment water interface. Previous

studies of organic matter decomposition in controlled

laboratory systems have focused on the decomposition

of primary producers (Enoksson, 1993; Pedersen et al.,

1999; Neubauer et al., 2004). The latter two investiga-

tions concentrated on the decomposition of nitrogen-

poor materials in the form of seagrass leaves and roots,

whereas the first study was on the decomposition of the

diatom Skeletonema costatum. The present study builds

on and extends these former studies by investigating the

decomposition of high-quality and nitrogen rich organic

matter present in the blue mussel Mytilus edulis. The

trigger for this study was an anoxic event in Mariager

Fjord, Denmark, in August 1997, which caused mor-

tality of the entire population of the blue mussel (M.

edulis) in the inner fjord (Fallesen et al., 2000). The

extensive anoxia in 1997 was attributed to an unusual

long period with calm and warm weather that reduced

the input of oxygen from the air (Fallesen et al., 2000).

In addition a heavy rainfall in the end of July supplied

nutrients to the fjord from the surrounding catchment

area and directly by rainwater (16 ton-N within 3 days).

This input should be compared to a normal monthly

input during summer months of 80–100 ton-N month�1

(Sørensen and Wiggers, 1997). A mass bloom of the

dinoflagellate Prorocentrum minium was observed in

the beginning of August with biomasses of ~800 Ag-Cl�1 in the inner part of the fjord and biomasses up to 28

mg-C l�1 in other parts of the fjord. Decomposition of

the P. minimum bloom increased the oxygen demand in

the fjord (Fallesen et al., 2000). At some point, the

limited oxygen input could no longer meet the oxygen

demand and all mussels down to 8–10 m died. During

the period of anoxia (2 weeks) there was a rapid

recycling of NH4+ and the water column concentration

(0–10 m) of NH4+ increased to 189 AM within a few

weeks (Sørensen and Wiggers, 1997). In the inner part

of the fjord the increase in NH4+ was strongly affected

by the decomposition of M. edulis tissue.

We postulate that anoxic decomposition of mussel

tissue at the sediment surface would lead to a signifi-

cant release of DON compared to NH4+ as lysis and

hydrolysis took place close to the sediment–water in-

terface. Further, most of the NH4+ mineralized would be

liberated to the overlying water with only little NH4+

incorporation into the growing populations of hetero-

trophic bacteria. The rationale for the latter part of the

hypothesis is that it is assumed that anaerobic bacteria,

with low carbon incorporation efficiencies, dominate

the decomposition process. The low carbon incorpora-

tion efficiency combined with a low C:N ratio in the

mussels degraded would in turn lead to a low nitrogen

demand by the heterotrophic bacterial populations.

2. Materials and methods

2.1. Sampling

Sediment was collected from a M. edulis bed in

Limfjorden, Denmark, in February 1998. The water

depth was 0.5 m. Sediment was sampled in Plexiglas

cores (i.d. 6 cm). The upper 12 cm of the sediment was

sieved (1-mm mesh size) to remove macrofauna and to

homogenize the sediment. After returning to the labo-

ratory the sediment was transferred to 7 continuous

flow-through Plexiglas chambers (dimensions: length/

with/height=25/25/14 cm) and they were carefully

overlaid with a water column. The resultant sediment

volume in each chamber was 6250 cm3 and the surface

area was 625 cm2. The volume of the overlaying water

column was 2500 cm3. The chambers were kept in the

dark, at the in situ temperature in August, 1997 (20 8C)for 4 days before the experiment was initiated.

M. edulis was sampled at a water depth b1 m and

kept in seawater from the sampling location and in the

dark until the water was bubbled with nitrogen to create

anoxic conditions. The mussels were killed by exposure

to 200 AM H2S for 2 days. In order to quantify the

amount of added mussel tissue, the soft mussel tissue

was carefully removed from the shells and weighed

before it was added to the sediment surface.

2.2. Experimental set-up

The sediment chambers were placed in a dark shad-

ed water bath at 20 8C (Fig. 1). The water overlying the

sediment (reservoir water) was replaced with nitrogen-

free artificial seawater (Kester et al., 1967) with the

following modifications: the salinity was 24.6 psu,

there was no addition of vitamins, NO3� or PO4

3�.

The water overlying the sediment was stirred with a

centrally placed magnet (45 rpm). After 7 days the

chambers were connected to the water reservoir with

artificial seawater. The reservoir water was pumped

through the sediment chambers with a Watson–Marlow

peristaltic pump with a flow rate of 5 ml min�1. The

tubes from the reservoir to the chambers were gas tight

butyl rubber. The tubes were disconnected from the

chambers twice during the experiment and rinsed with

1 N HCl followed by a thorough wash with artificial

seawater to avoid bacterial growth in the system.

Fig. 1. The continuous flow-through system consisted of a reservoir with artificial seawater (A), a Watson–Marlow pump (B), a sampling port for

inflow water (C), sediment chambers (D), sampling port for outflow water (E) and a water bath (F) where the temperature was maintained at 20 8C.

B.A. Lomstein et al. / J. Exp. Mar. Biol. Ecol. 329 (2006) 251–264 253

2.3. Fluxes of solutes across the sediment–water

interface

During the first 20 days of incubation, the sediment–

water fluxes were high and variable, and this period

was considered as the preincubation period (data not

shown). Day 0 in the following refers to the day the

mussel tissue was added onto the sediment surface to

four of the 7 chambers (Mussel+). The remaining

chambers without mussel tissue (Mussel�) served as

controls. The addition of mussel tissue corresponded to

the wet weight of mussel tissue per m2 in the inner part

of Mariager Fjord (6.7 kg m�2; Sømod, 1992). The

total organic carbon (OC) and organic nitrogen (ON) in

the added mussel tissue were 19.9 mol-C m�2 and 4.0

mol-N m�2, respectively.

In- and outflow water from the chambers was col-

lected every second or third day throughout the 32 day

incubation period. The water was analyzed for O2,PCO2, dissolved organic nitrogen (DON), dissolved

free amino acids (DFAA), total hydrolyzable amino

acids (THAA), short-chained fatty acids (SCFA),

urea-N, NH4+ and NO3

�.

The sediment–water fluxes were determined using

the equation of Nishio et al. (1982):

F ¼ DC� V=A ð1Þ

where DC is the concentration change between in- and

outflow water, V is the flow rate, and A is the sediment

surface area in the chamber.

O2 was measured by the Winkler method (Strickland

and Parsons, 1972) andP

CO2 was analyzed on a flow-

injection system (Hall and Aller, 1992). O2 andP

CO2

analysis were carried out b2 h after sampling.

Samples for DON, DFAA, and THAA were filtered

through a 0.2-Am Sartoriusk filter. Samples for SCFA,

urea-N, NH4+ and NO3

� were unfiltered. All samples,

except THAA, were frozen for later analysis.

Samples for total dissolved nitrogen (TDN) were

analyzed on a modified Antek 7000 system as de-

scribed in Lomstein et al. (1998). The concentration

of TDN was estimated from a calibration curve made

on a Tris-buffer and the concentration of DON was

estimated as the difference between TDN and dissolved

inorganic nitrogen (DIN). The concentration of DFAA

was determined by high-performance liquid chromatog-

raphy (HPLC; Waters Chromatographic System) on o-

phthaldialdehyde-(OPA)-derivatized products (Lindroth

and Mopper, 1979). Immediately after sampling THAA

samples were added 12 N HCl in a 1:1 v:v ratio and the

head space in the sample container was replaced with

N2. Hydrolysis was carried out at 110 8C for 24 h. After

hydrolysis, 100 Al of the sample was dried in a vacuum

desiccator for 6 h. The sample residue was dissolved in

1 ml Milli-Q water and filtered through a 0.2-AmSatoriusk filter. The THAA content of the sample

was measured as DFAA as previously described. The

concentration of dissolved combined amino acids

(DCAA) was determined as the difference between

individual amino acids of THAA and DFAA. SCFA

was determined by high-performance liquid chromatog-

raphy (HPLC-IC Sykam) as described in Bøtte and

Jørgensen (1992). The SCFAs identified were acetate,

propionate, butyrate, isobutyrate, formate and valerate.

In addition ethanol was quantified by the same method.

NH4+, NO3

� and urea-N were measured spectropho-

tometrically using an Alfa- Laval Bran+Luebbe auto-

analyzer using the following methods: NH4+ (Bower

and Holm-Hansen, 1980), NO2�+NO3

� (Grasshoff et

al., 1983) and urea (Price and Harrison, 1987).

2.4. Sediment and mussel characteristics

At day 0 when mussel tissue was added to the

Mussel+ chambers one of the control chambers

(Mussel�) was gently removed from the continuous

flow-through system. The sediment was sectioned

into the following depth intervals: 0–1, 1–2, 2–4, 4–6

and 6–10 cm. The remaining two Mussel� chambers

were sacrificed on days 17 and 33 after mussel addition.

ig. 2. Efflux ofP

CO2 from the sediment to the overlying water (A)

nd O2 uptake (B) in the Mussel+ and Mussel� chambers. Error bars

present the standard deviation of the mean (n =2).


The four Mussel+ chambers were sacrificed on days 2,

9, 16 and 32, respectively. The overlying water and the

mussel tissue were carefully removed before the sedi-

ment was sliced into the depth intervals described

above. The following parameters were measured in all

sediment chambers: sediment specific density, water

content, particulate organic carbon (POC), particulate

organic nitrogen (PON) and THAA.

The specific density of the sediment (g cm�3) was

determined gravimetrically on triplicate 1-cm3 samples.

The porewater content was determined as the weight

loss of fresh sediment dried at 105 8C for 24 h. The

contents of POC and PON were measured on dried,

homogenized, H2SO3 treated sediment on a Carlo Erba

NA-1500 HCN analyzer. THAA was determined on 1

cm3 fresh sediment to which there was added 10 ml 6 N

HCl. Analysis was performed as described for THAA

in inflow and outflow water, with the exception that the

sample residue was dissolved in 5 ml Milli-Q water

instead of 1 ml Milli-Q water.

The M. edulis tissue was analyzed for the following

parameters: wet weight, water content and the content of

organic carbon (OC), organic nitrogen (ON) and mussel

THAA with the respective procedures described above.

2.5. Porewater DON and DIN

Porewater from the respective depth described above

was obtained by centrifugation at 1000�g for 10 min.

Samples for DON, THAA and DFAA analysis were

filtered though a 0.2-Am Sartoriusk filter. Samples for

urea-N and NH4+ were not filtered. All samples were

frozen for later analysis except THAA samples that

were preserved in 12 N HCl in a 1:1 v:v ratio. Analysis

of the respective porewater constituents was as previ-

ously described.

2.6. Turnover of porewater DFAA

The turnover of DFAA was estimated from the turn-

over of 14C-glutamate and 14C-alanine. Incubations

were performed as follows at the sediment depths pre-

viously described: (1) 10 Al 14C – [U] – amino acid

tracer was injected into 1 cm3 sediment in a N2-flushed

exetainer (glutamate 1.58 nCi ml�1, 266 nCi nmol�1;

alanine 1.49 nCi ml�1, 155 nmol�1; Amersham) and

incubated in a time course (0, 20, 45 min); (2) turnover

activity was stopped by the addition of 1 ml 2.5% w:v

NaOH; and (3) the sediment �NaOH suspension was

thoroughly mixed and frozen for later analysis. The

final concentrations of tracer were always b10% of

the respective ambient pools.

Radioactivity was counted in a Packard 2200 CATri

Carb Liquid Scintillation analyzer. The turnover rate

constants of glutamate and alanine were estimated by

the steady state model II described in Lund and Black-

burn (1989) and the turnover rate of DFAA was esti-

mated as the average turnover rate constant for alanine

and glutamate multiplied with the pool of DFAA.

3. Results

3.1. Efflux ofP

CO2 from the sediment and O2 uptake

In the Mussel+ chambers there was an increase in

the efflux ofP

CO2 from 18 mmol m�2 day�1 at day 0

to 554 mmol m�2 day�1 at day 2 (Fig. 2A). From days

2 to 13, theP

CO2 efflux fluctuated in the range of

434–626 mmol m�2 day�1 before progressively de-

creasing to 91 mmol m�2 day�1 by day 32. ThePCO2 efflux in the Mussel� controls remained rela-

tively constant at b57 mmol m�2 day�1.

All O2 in the inflow water was consumed in the

Mussel+ chambers, which explains why there was only

a minor increase in the O2 uptake during incubation

compared with theP

CO2 efflux. Hence, O2 uptake

increased during the first 5 day of incubation from 10 to

20 mmol m�2 day�1 and then remained within the

range 20–24 mmol m�2 day�1 for the remainder of

the experiment (Fig. 2B). Oxygen uptake in the

Mussel� chambers remained within the range of 6–

F

a

re


13 mmol m�2 day�1 during the 32 day incubation

period.

3.2. Efflux of SCFA, DON and DIN

The efflux of SCFA increased rapidly from non-

detectable at day 0 to a maximum of 1263 mmol-C

m�2 day�1 at day 2 (Fig. 3A). After day 15 the efflux

Fig. 3. Efflux of SCFA (A), DON (B), DCAA-N (C), DFAA-N (D), urea-

Mussel� chambers (open symbols). Outflow water for SCFA-C, DFAA-N

occasions, due to the large number of parameters analyzed for in the present

the SCFA-C efflux, 0–22% for the DFAA efflux, 6–34% for the NH4+ efflux

addition.

of SCFA-C could no longer be detected. Acetate

accounted for 36–84% of the SCFA-C efflux and was

the most important of the identified SCFAs effluxing

from the sediment. In the Mussel� controls there was

no detectable efflux of SCFA.

In the chambers amended with mussel tissue, the

DON efflux increased from non-detectable at day 0 to

516 mmol-N m�2 day�1 at day 1. After day 1 DON

N (E), NH4+ (F) and NO3

� (G) in the Mussel+ (closed symbols) and

, NH4+ and NO3

� analysis were only analyzed in replicates on a few

experiment. The coefficient of variation varied between 6 and 19% for

and it was highly variable for the NO3� efflux (9–482%) after mussel

Table 1

The content of organic carbon (OC), organic nitrogen (ON) and

THAA-N and the relative contribution of THAA-N to the ON pool

in mussel tissue after 0, 2, 9, 16 and 32 days of incubation

Day OC

(mol m�2)

ON

(mol m�2)

THAA

(mol-N m�2)

THAA/ON

(%)

0 19.9 4.0 1.9 47

2 11.6 2.2 0.6 27

9 2.8 0.4 0.2 44

16 2.0 0.3 0.1 33

32 1.6 0.3 0.1 30


efflux decreased rapidly and remained at b6 mmol-N

m�2 day�1 after day 18 (Fig. 3B). In the control

chambers there was no detectable efflux of DON.

Fig. 4. Depth profiles of POC (A), PON (C) and THAA-N (E) on day 2, 9

profiles of the respective pools from Mussel� at days 17 and 33 were not inc

the Mussel� chamber at day 0. The area integrated (P

0–10 cm) pools of

chambers. Error bars represent the standard deviation (n =2) of the mean. T

Data in Fig. 3C show that the DCAA-N efflux

increased from 47 mmol-N m�2 day�1 at day 0 to a

maximum of 121 mmol-N m�2 day�1 at day 2. After

day 5 the DCAA efflux remained b25 mmol-N m�2

day�1. In the Mussel� chambers the DCAA efflux

decreased from 47 mmol-N m�2 day�1 at day 0 to

b22 mmol-N m�2 day�1 during the remaining part of

the experiment. The efflux of DFAA increased from b1

mmol-N m�2 day�1 at day 0 to a maximum of 115

mmol-N m�2 day�1 at day 1 (Fig. 3D). After day 5 the

DFAA efflux remained b3 mmol-N m�2 day�1. The

DFAA efflux remained b0.2 mmol-N m�2 day�1

throughout the incubation period in the controls.

Urea-N efflux increased from b0.1 mmol-N m�2

day�1 to a maximum of 6 mmol-N m�2 day�1 at

, 16 and 32 in the Mussel+, and day 0 in Mussel� chambers. Depth

luded, as they could not be distinguished from the respective profiles in

POC (B), PON (D) and THAA-N (F) in the Mussel+ and Mussel�HAA was not replicated.


day 2 (Fig. 3E) in the Mussel+ chambers, whereas in

the controls the urea-N efflux remained b0.1 mmol-N

m�2 day�1. After day 5 the urea-N efflux remained

b0.7 mmol-N m�2 day�1 in the Mussel+ chambers. In

the Mussel+ chambers the efflux of NH4+ increased

from b1 mmol m�2 day�1 at day 0 to a maximum of

496 mmol m�2 day�1 at day 2 (Fig. 3F). By the end of

the experiment NH4+ efflux had declined to the levels

recorded in the controls. In the Mussel+ chambers the

efflux of NO3� decreased from 0.5 mmol m�2 day�1 at

day 0 to b0.2 mmol m�2 day�1 for the remainder of

the experiment whereas in the controls NO3� efflux was

in the range 0.3–0.9 mmol-N m�2 day�1 over the 32

day incubation period (Fig. 3G).

3.3. Mussel and sediment characteristics

Mussel OC and ON decreased with time of incuba-

tion from 19.9 and 4.0 mol m�2 at day 0, respectively,

to 1.6 and 0.3 mol m�2 at day 32, respectively (Table

1). THAA in the mussel tissue decreased from 1.9 mol-

Fig. 5. Total loss of mussel OC and sediment POC and the efflux of DOC andPCO2 in Mussel� (A right panel), loss of mussel ON and sediment PON a

PON and efflux of DON and DIN in Mussel� (B right panel).

N m�2 at day 0 to 0.1 mol-N m�2 at day 32. THAA

accounted for 27–47% of mussel ON during the time

course of the experiment (Table 1).

At all sampling times there was a decrease in the

content of POC, PON and THAA-N with sediment

depth in the Mussel+ chambers (Fig. 4A, C, E);

most of the change taking place within the upper 2

cm of the sediment. There was a decrease in surface

(0–1 cm depth) POC and PON from 1207 Amol-C

cm�3 and 154 Amol-N cm�3, respectively, at day 2

to 925 Amol-C cm�3 and 107 Amol-N cm�3, respec-

tively, at day 32 (Fig. 4A, C). The contents of POC

and PON did not vary much with depth in the

controls compared to Mussel+ chambers and were

in the range of 719–937 Amol-C cm�3 and 48–74

Amol-N cm�3, respectively, throughout the incuba-

tion period. In the Mussel amended chambers the

depth integrated (P

0–10 cm) content of POC and

PON decreased from 86 mol-C m�2 and ~7 mol-N

m�2 at day 2, respectively, to 79 mol-C m�2 and

~6 mol-N m�2 at day 32, respectively (Fig. 4B, D).

PCO2 in Mussel+ (A left panel), loss of POC and efflux of DOC and

nd the efflux of DON and DIN in Mussel+ (B left panel) and loss of


In the Mussel� controls the depth integrated (P

0–

10 cm) content of POC and PON decreased from 82

mol-C m�2 and ~6 mol-N m�2 at day 0, respec-

tively, to 80 mol-C m�2 and ~5 mol-N m�2 at day

33, respectively.

In the upper 0–1 cm sediment horizon of the Mus-

sel+ chambers THAA-N decreased from 57 Amol-N

Fig. 6. Depth profiles of DON (A), DCAA-N (C), DFAA-N (E) and NH4+

Mussel� chamber. Depth profiles of the respective pools from Mussel� c

distinguished from the respective profiles in Mussel� chamber at day 0. Are

(F) and NH4+ (H) in the Mussel+ and Mussel� chambers.

cm�3 at day 2 to 43 Amol-N cm�3 at day 16, after

which time the surface THAA-N content steadily in-

creased reaching 58 Amol-N cm�3 by day 32 (Fig. 4E).

In contrast in the control chambers the THAA-N con-

tent remained relatively constant in the range of 25–28

Amol-N cm�3 throughout the experiment. The depth

integrated (P

0–10 cm) THAA-N content remained

(G) on days 2, 9, 16 and 32 in Mussel+ chambers and on day 0 in

hamber at days 17 and 33 were not included, as they could not be

a integrated (P

0–10 cm) pools of DON (B), DCAA-N (D), DFAA-N

Fig. 7. Depth profiles of DFAA-N turnover rates on days 2, 9, 16

and 32 in Mussel+ chambers and day 0 in Mussel� chamber (A)

Depth profiles of DFAA-N turnover from Mussel� chamber at days

17 and 33 were not included, as they could not be distinguished

from the turnover rates in Mussel� chamber at day 0. Area inte

grated (P

0–10 cm) turnover rates of DFAA-N in Mussel+ and

Mussel� chambers (B).


elevated in the Mussel amended chambers (2.4–2.5

mol-N m�2) throughout the experiment compared to

the controls (2.2–2.4 mol-N m�2; Fig. 4F).

In the Mussel+ chambers the efflux ofP

CO2+

SCFA-C+THAA-C accounted for 90% of the carbon

loss from the mussel tissue and sediment during the

32 day incubation period (Fig. 5A). Similarly, over

the same time period, the efflux of DON+DIN

accounted for 95% of the nitrogen loss from the mussel

tissue and sediment (Fig. 5B). In the unamended

controls the efflux ofP

CO2+SCFA-C+THAA-C

accounted for 146% of the carbon loss from the sedi-

ment and the efflux of DON+DIN for 69% of the

observed nitrogen loss.

3.4. Porewater profiles and pool

Two days after the addition of the mussel tissue

the surface concentrations of DON, DCAA and

DFAA were all enhanced above the respective con-

centrations in the control chambers (Fig. 6A, C, E).

At day 9 and 16 the concentration of DON in the

Mussel+ chambers was elevated throughout the sed-

iment. The area integrated (P

0–10 cm) DON pool

increased from 200 mmol-N m�2 at day 2 to 297

mmol-N m�2 at day 9 after which time the DON

pool decreased to 39 mmol-N m�2 at day 32 (Fig.

6B). In the Mussel� control chambers the DON pool

decreased from 34 mmol m�2 at day 0 to 12 mmol

m�2 at day 32.

The area integrated pools (P

0–10 cm) of DCAA

and DFAA decreased from 94 and 18 mmol-N m�2 at

day 2, respectively in the Mussel amended chambers to

13 and b1 mmol-N m�2 at day 32, respectively (Fig.

6D, F) whereas in the controls the area integrated pools

of DCAA and DFAA decreased from 11 and b1 mmol-

N m�2 at day 0, respectively, to 6 and b1 mmol N

m�2 at day 32, respectively (Fig. 6D, F).

Two days after the addition of the mussel tissue the

surface concentration (0–1 cm depth) of NH4+ in-

creased to 4 Amol cm�3 compared to b0.1 Amol

cm�3 in the controls (Fig. 6G). At days 9 and 16

the concentration of NH4+ in the Mussel+ chambers

was elevated throughout the sediment. The area inte-

grated NH4+ pool (

P0–10 cm) in the Mussel+ cham-

bers increased from 111 mmol m�2 at day 2 to 226

mmol m�2 at day 9, after which time NH4+ decreased

to approximate 40 mmol m�2 by the end of the

experiment (Fig. 6H). There was a slight increase in

the area integrated NH4+ pool in the Mussel� controls

from 13 mmol m�2 at day 0 to 19 mmol m�2 at day

32 (Fig. 6H).

.

-

3.5. Sediment DFAA-N turnover

At day 2 after mussel addition there was a maximum

in the turnover rate of DFAA within the upper 1 cm of

the sediment of ~15 Amol-N cm�3 day�1 (Fig. 7A). In

comparison the turnover rate of DFAA remained b2

Amol cm�3 day�1 throughout incubation in the un-

amended controls. The area integrated (P

0–10 cm)

turnover rate of DFAA in the Mussel+ chambers de-

creased from 265 mmol-N m�2 day�1 at day 2 to 22

mmol-N m�2 day�1 at day 9, after which time the

turnover rate of DFAA remained b27 mmol-N m�2

day�1 (Fig. 7B). In the controls the area integrated

turnover rate of DFAA was always b2 mmol-N m�2

day�1 (Fig. 7B).

4. Discussion

4.1. Composition of efflux solutes

There was a rapid response in the efflux of SCFA

and DON after the addition of M. edulis tissue to the

sediment surface.


Over the time course of the Mussel+ incubation

SCFA dominated the identified DOC efflux (SCFA+D-

CAA-C+DFAA-C) accounting for 67% of the identi-

fied DOC. The most important SCFA component was

acetate, which accounted for 36–84% of the SCFA

efflux. The large efflux of fermentation products

(SCFA) during the first 8 d after mussel addition indi-

cates that respiration initially lagged behind fermenta-

tion. These results are in agreement with those of

Holmer and Kristensen (1994) who demonstrated that

high loads of organic matter saturate sulfate reduction

with substrate, which then leads to the accumulation of

SCFA in the sediment principally in the form of acetate.

A large fraction (42%) of the added mussel-N es-

caped mineralization and effluxed the sediment as

DON; only 8% of the added mussel-N remained by

the end of incubation. It is possible that part of the

DON efflux may have been due to leakage of solutes

from the mussel tissue. In accordance with this the

DON efflux was dominated by DCAA-N+DFAA-N

that accounted for 57% of the DON efflux. This should

be compared with the initial 48% contribution of

THAA-N to mussel ON. Further, as proposed by Black-

burn and Blackburn (1993) there is a high probability

that products of hydrolysis escape into the overlying

water, when hydrolysis occurs at the sediment–water

interface.

In the Mussel� controls DCAA-N was the only

identified DON component and it was used as a min-

imum estimate of the DON efflux. TDN analysis of

outflow water failed to show any DON due to analytical

problems.

It is likely that the unidentified DOM efflux in the

Mussel+ amended chambers was composed of N-free

DOM molecules together with N-containing organic

molecules. Among the N-free DOM molecules that

may have been of importance were polysaccharides,

sugars and lipids. The N-containing organic molecules

that were not identified may have been volatile amines

(methyl-, dimethyl- or trimethylamines). This conclu-

sion is based on the unpleasant smell that developed in

the Mussel+ chambers and the fact that TMAO is

present in M. edulis (King, 1988). Further, trimethyla-

mine precursors are easily converted to trimethylamine

by anaerobic microorganisms in marine sediments

(references in Sørensen and Glob, 1987).

Within a few days after the addition of mussel tissue

carbon oxidation (i.e.P

CO2 efflux) and NH4+ efflux

reached maximum activities, which were indicative of a

maximum bacterial mineralization activity. The high

NH4+ efflux of ~500 mmol m�2 day�1 was five-fold

higher than the highest reported NH4+ efflux from a

marine fish pond at a temperature similar to that used

in the present experiment (Lefebvre et al., 2001). In the

present study the NH4+ efflux during the first 3 days

after addition of mussel tissue would have resulted in

an NH4+ concentration in a 5 m water column of 180

AM. In comparison the in situ NH4+ concentration in

Mariager Fjord (water depth 0–10 m) increased to 189

AM within a few days after the water column became

anoxic (Sørensen and Wiggers, 1997).

In contrast to the stimulated NH4+ efflux after mussel

addition there was a decrease in the efflux of NO3� that

can be explained by the complete consumption of O2 in

the Mussel amended chambers. Denitrification was pos-

sibly also reduced after mussel addition since the only

source of NO3� for denitrification was that resulting

from sediment nitrification; there was no NO3� in the

inflow water to the continuous flow-through system.

The C:N ratio in mineralization products (P

CO2:

DIN)efflux was estimated from the following equation

due to the lack of information on the actual denitrifica-

tion rate:

C:Nmin¼X

CO2efflux= NHþ4 þ 2� NO�

3

� �� efflux

ð2ÞAs denitrification was based solely on NO3

� generated as

a result of sediment nitrification it can be assumed that

the upward flux (efflux) of NO3� was equal to the

downward flux of NO3� to the denitrification zone;

thus denitrification equalled the NO3� efflux. This has

previously been shown to be a valid assumption in a

seasonal study described by Jørgensen (1996). The

molar C:N ratio in the mineralization products

(P

CO2:DIN) that effluxed from the sediment in the

Mussel+ chambers during the course of the experiment

was low (3.9) and is consistent with the initial low molar

C:N ratio of 5 in the added mussel tissue. In the Mussel�controls the C:N ratio in the mineralization products that

effluxed from the sediment was 10.1 over the 32 day

incubation period and this value is within the range of

ratios that can be estimated from other studies (2–156;

Blackburn et al., 1996 and references therein, Burdige

and Zheng, 1998; Pedersen et al., 1999). As stated by

Fenchel et al. (1998) the C:N ratio in the organic matter

degraded is often inferred from theP

CO2:DIN ratio in

efflux mineralization products. This assumption is only

valid if it can be assumed that the bacterial biomass

remains in steady state.

4.2. Indications on stimulated bacterial growth in the

sediment

We postulated that the addition of mussel tissue to

the sediment surface would stimulate bacterial growth,


but that the nitrogen demand by the bacteria was low

due to an anticipated low carbon incorporation effi-

ciency of the anaerobic bacteria and a low C:N ratio

in the mussel substrate supplied. The net increase in

sediment THAA-N of 126 mmol-N m�2 in the Mus-

sel+ chambers and 71 mmol-N m�2 in the controls

during the time course of the experiment can only be

explained by an increase in bacterial biomass and/or

proteins. Net synthesis of THAA-N requires that non-

THAA-PON is mineralized to NH4+ and incorporated

into THAA-N.

The accumulation in sediment THAA-N during the

incubation period was equivalent to an increase in

bacterial cells of 0.9�108 cells cm�3 in the Mussel

amended chambers and 0.5�108 cells cm�3 in the

controls. The nitrogen content in natural aquatic bac-

terial cells was obtained from Fagerbakke et al. (1996)

and it was assumed that protein is the dominant

nitrogen containing organic molecule in bacterial

cells. In comparison, Pedersen et al. (1999) estimated

the increase in bacterial cell number, during decom-

position of eelgrass leaves, to have been 3.4�108–

4.0�108 cells cm�3. The inferred response in bacte-

rial production might have been the result of supply of

energy sources from the easily hydrolyzable mussel

tissue.

Our use of THAA as a measure of net bacterial

growth implies that there was a stimulation of bacterial

growth or protein synthesis down to a depth of ~2 cm

in the sediment. In agreement with this, Graf (1987)

showed that benthic biomass and metabolism were

stimulated down to at least 7 cm after the addition of

diatoms to the sediment surface. Benthic biomass was

measured as ATP and metabolism as heat production.

Diffusion estimates show that diffusion can easily ac-

count for high activities at a depth of 3–4 cm within 1

week. This estimate was based on the equation of

Carslaw and Jaeger (1959) and a diffusion coefficient

of 5�10�6 cm�2 s�1 (Blackburn and Blackburn,

1993).

4.3. Conceptual models of N- and C-cycling during M.

edulis decomposition

Further discussion will be related to Fig. 8A–B in

which the integrated area measured and estimated

nitrogen and carbon transformation rates (mol m�2

32 days�1) can be seen in relation to each other

during the entire incubation period in the Mussel

amended chambers. Data for Mussel� controls are

not shown as the measured and estimated rates were

very low.

Gross NH4+ mineralization (d) was estimated as

follows:

d ¼ iþ change in NHþ4 poolþ NHþ

4 efflux ð3Þ

where i is NH4+ incorporation into bacterial biomass

(i.e. net THAA-N increase).

DFAA-N turnover to NH4+ accounted for 70% of

gross NH4+ mineralization, which indicates that there

were other sources than amino acids for NH4+ produc-

tion. In comparison Lomstein et al. (1998) estimated

that DFAA turnover was responsible for 58% of gross

NH4+ mineralization in the shallow cove Knebel Vig,

Denmark. Among other sources for gross NH4+ miner-

alization is urea-N, which can account for a substantial

fraction of NH4+ production in marine sediments (e.g.

Lomstein et al., 1998; Pedersen et al., 1999).

Based on the inferred non-steady state in bacterial

biomass the carbon incorporation efficiency (E) was

estimated from the following Blackburn (1980) non-

steady-state equation:

E ¼ i= Co � Ncð Þ þ i½ � ð4Þwhere Co is carbon oxidation, i is NH4

+ incorporation

into bacterial cells and Nc is the molar N:C ratio of

bacterial cells (0.2; Fagerbakke et al., 1996). The esti-

mated average E value in the Mussel+ chambers was

0.06 (Table 2). In accordance with this Stouthamer

(1979) obtained E-values within the range of 0.05–

0.27 for pure cultures of anaerobic bacteria depending

on the organism and the substrate used. In the control

chambers the average E-value was 0.33, which may

reflect that both aerobic (with relatively high E-values)

and anaerobic bacteria (with low E-values) participated

in the degradation of organic matter.

The molar C:N ratio in the substrate degraded by the

bacteria was estimated from the Blackburn (1980)

equation:

Ns ¼ E � Nc � d=i ð5Þ

where Ns is the molar N:C ratio in the substrate de-

graded. The average C:N ratio in the substrate degraded

in the Mussel+ and Mussel� chambers was 3.7 and

10.2, respectively (Table 2). The low C:N ratio in the

substrate utilized by bacteria during mussel decompo-

sition showed that the bacteria degraded organic mate-

rial with a lower C:N ratio than their own (~5). The low

C:N ratio in the substrate utilized by the mussel degrad-

ing bacteria combined with the low carbon incorpora-

tion efficiency was the cause of the almost complete

release of NH4+ mineralized within the sediment (Fig.

8A). During the course of incubation, 95% of gross

NH4+ mineralization effluxed to the overlying water

Fig. 8. Conceptual models of total nitrogen- (A) and carbon cycling (B) in Mussel+ chambers during the entire incubation period. All rates are in

mol m�2 32 days�1. The rate of change in bother PONQ was estimated as the difference between the net change in PON minus the net change in

THAA-N during the course of incubation. The rate of change in bother POCQ was estimated in an equal manner. The rate of PON hydrolysis was

estimated as the DON efflux plus the gross NH4+ mineralization rate. The minimum rate of POC hydrolysis was estimated as the SCFA+THAA-C

efflux (used as minimum DOC efflux) plus the sum of carbon oxidation and C-incorporation into the bacterial biomass. The net changes in

dissolved pools are not included as they were always much smaller than 0.1 mol m�2 32 days�1.


column. In contrast to this, only 37% of gross NH4+

mineralization effluxed to the overlying water in the

Mussel� controls, whereas the remaining 63% was

Table 2

Summary of carbon oxidation (Co), gross NH4+ mineralization (d),

NH4+ incorporation (i) and the C:N ratio in the substrate degraded

(C:Nsub) during the entire incubation period in the Mussel+ and

Mussel� chambers, respectively

Co (mol-C

m�2 32

days�1)

d (mol-N

m�2 32

days�1)

i (mol-N

m�2 32

days�1)

E C:Nsub

(mol mol�1)

Mussel+ 10.1 2.8 0.1 0.06 3.7

Mussel� 0.7 0.1 b0.1 0.33 10.2

incorporated into the bacterial biomass (Table 2). The

cause of this difference is a combination of a high

substrate C:N ratio (10.2) in the controls compared to

Mussel amended chambers (3.7) and the fact that deg-

radation was carried out by both aerobic and anaerobic

bacteria in the Mussel� chambers, with a resultant

higher average carbon incorporation efficiency (0.33),

than in the Mussel+ chambers (0.06).

5. Conclusions

In summary our hypothesis on factors that cause

rapid recycling of nitrogen during anaerobic degrada-

tion of M. edulis has been confirmed by the results


obtained in this study. These results all point in the

same direction—the anaerobic degradation of high-

quality organic matter such as M. edulis tissue at the

sediment surface has severe ecological consequences

for the environment: most of the mussel tissue (N90%)

was degraded within 2 weeks, which was the duration

of the anoxic event in Mariager Fjord. A large fraction

of the DOM produced escaped mineralization and

effluxed to the overlying water together with most

of NH4+ produced during mineralization (Fig. 3). The

consequence of the relatively large efflux of high-

quality DOM (high content of SCFA, DCAA and

DFAA) to the overlying water column is that it even-

tually will undergo mineralization with resultant con-

sumption of oxygen in the water column. The almost

complete liberation of NH4+ mineralized within the

sediment forms the basis for a new phytoplankton

bloom, which is actually what happened in Mariager

Fjord.

Acknowledgments

We thank Rikke O. Holm for skillful technical as-

sistance. This study was financed by grants from the

Danish Natural Science Research Council, grant nos.

9901859 and 9901860 (equipment) and the Carlsberg

Foundation, grant nos. 990330 and 0203/20. [SS]

References

Blackburn, T.H., 1980. Seasonal variations in the rate of organic-N

mineralization in anoxic marine sediments. Biogeochimie de la

matiere organique a l’ınterface eau-sediment marin. Colloq. Int.

C.N.R.S. 293, 179–183.

Blackburn, T.H., Blackburn, N.D., 1993. Rates of microbial processes

in sediments. Philos. Trans. R. Soc. Lond. 344, 49–58.

Blackburn, T.H., Henriksen, K., 1983. Nitrogen cycling in different

types of sediments from Danish waters. Limnol. Oceanogr. 28,

477–493.

Blackburn, T.H., Hall, P.O.J., Hulth, S., Landen, A., 1996. Organic-N

loss by efflux and burial associated with a low efflux of inorganic

N and with nitrate assimilation in Arctic sediments (Svalbard,

Norway). Mar. Ecol. Prog. Ser. 141, 283–293.

Bøtte, H.F., Jørgensen, L., 1992. Evaluation of low-conductance

eluents for suppressed ion-exclusion chromatography. J. Chroma-

togr. 602, 27–31.

Bower, C.E., Holm-Hansen, T., 1980. A salicylate-hypochlorite meth-

od for determining ammonia in seawater. Can. J. Fish. Aquat. Sci.

37, 794–798.

Burdige, D.J., Zheng, S., 1998. The biogeochemical cycling of dis-

solved organic nitrogen in estuarine sediments. Limnol. Oceanogr.

43, 1796–1813.

Carslaw, H.S., Jaeger, J.C., 1959. Conduction of heat in solids. Trans.,

2nd ed. Oxford Univ. Press, pp. 1–510.

Enoksson, V., 1993. Nutrient recycling by coastal sediments: effects

of added algal material. Mar. Ecol. Prog. Ser. 92, 245–254.

Fagerbakke, K.M., Heldal, M., Norland, S., 1996. Content of carbon,

nitrogen, oxygen, sulfur and phosphorus in native aquatic and

cultured bacteria. Aquat. Microb. Ecol. 10, 15–27.

Fallesen, G., Andersen, F., Larsen, B., 2000. Life, death and revival

of the hypertrophic Mariager Fjord, Denmark. J. Mar. Syst. 25,

313–321.

Fenchel, T., King, G.M., Blackburn, T.H., 1998. Bacterial Biogeo-

chemistry: The Ecophysiology of Mineral Cycling, 2nd ed. Aca-

demic Press, San Diego, pp. 1–307.

Graf, G., 1987. Benthic energy flow during a simulated autumn

bloom sedimentation. Mar. Ecol. Prog. Ser. 39, 23–29.

Grasshoff, K.M., Ehrhardt, M., Kremling, K., 1983. Methods of

Seawater Analysis. Trans. Verlag Chemie.

Hall, P.O.J., Aller, R.C., 1992. Rapid, small-volume, flow injection

analysis for CO2 and NH4+ in marine and freshwaters. Limnol.

Oceanogr. 37, 1113–1119.

Holmer, M., Kristensen, E., 1994. Organic matter mineralization in an

organic-rich sediment: experimental stimulation of sulfate reduc-

tion by fish food pellets. FEMS Microbiol. Ecol. 14, 33–44.

Jørgensen, B.B., 1996. Material flux in the sediment. In: Jørgensen,

B.B., Richardson, K. (Eds.), Eutrophication in Coastal Marine

Ecosystems. Coastal and Esturarine Studies, IV Series, vol. 52.

American Geophysical Union, Washington, DC.

Kester, D.R., Duedall, I., Connors, D.N., Pytkowicz, R.M.,

1967. Preparation of artificial seawater. Limnol. Oceanogr.

12, 176–178.

King, G.M., 1988. Distribution and metabolism of Quaternary amines

in marine sediments. In: Blackburn, T.H., Sørensen, J. (Eds.),

Nitrogen Cycling in Coastal Marine Environments. John Wiley

& Sons, Chichester, pp. 143–174.

Lefebvre, S., Bacher, C., Meuret, A., Hussenot, J., 2001. Modeling

approach of nitrogen and phosphorus exchange at the sediment–

water interface of an intensive fishpond system. Aquaculture 195,

279–297.

Lindroth, P., Mopper, K., 1979. High performance liquid chromatog-

raphy determination of subpicomole amounts of amino acids by

precolumn fluorescence derivatization with o-phthaldialdehyde.

Anal. Chem. 51, 1667–1675.

Lomstein, B.Aa., Jensen, A.G.-U., Hansen, J.W., Andreassen, J.,

Hansen, L.S., Berntsen, J., Kunzendorf, H., 1998. Budgets of

sediment nitrogen and carbon cycling in the shallow water of

Knebel Vig, Denmark. Aquat. Microb. Ecol. 14, 69–80.

Lund, B.Aa., Blackburn, T.H., 1989. Urea turnover in a coastal

marine sediment measured by a 14C-urea short-term incubation.

J. Microbiol. Methods 9, 297–308.

Neubauer, A.T.A., Pedersen, A.-G.U., Finster, K., Herbert, R.A.,

Donnelly, A., Viaroli, P., Giordani, G., de Wit, R., Lomstein,

B.Aa., 2004. Benthic decomposition of Zostera marina roots: a

controlled laboratory experiment. J. Exp. Mar. Biol. Ecol. 313,

105–124.

Nishio, T.I., Koike, I., Hattori, A., 1982. Denitrification, nitrate

reduction, and oxygen consumption in coastal and estuarine sedi-

ments. Appl. Environ. Microbiol. 43, 648–653.

Pedersen, A.-G.U., Berntsen, J., Lomstein, B.Aa., 1999. The effect

of eelgrass decomposition on sediment carbon and nitrogen

cycling: a controlled laboratory experiment. Limnol. Oceanogr.

44, 1978–1992.

Price, N.M., Harrison, P.J., 1987. Comparison of methods for the

analysis of dissolved urea in seawater. Mar. Biol. 94, 307–317.

Sloth, N.P., Blackburn, H., Hansen, L.S., Risgaard-Petersen, N.,

Lomstein, B.Aa., 1995. Nitrogen cycling in sediments with dif-

ferent organic loading. Mar. Ecol. Prog. Ser. 116, 163–170.


Stouthamer, A.H., 1979. The search for correlation between theoret-

ical and experimental growth yields. Int. Rev. Biochem. 21, 1–47.

Strickland, J.D.H., Parsons, T.R., 1972. A practical handbook of sea

water analysis, 2nd ed. Bulletin-Fisheries Research Board of

Canada, vol. 167. Fisheries Research Board of Canada, Ottawa.

Sømod, B., 1992. Blamuslingernes produktion, assimilation og filtra-

tionskapacitet i Mariager Fjord. Arhus Amt. Miljøkontoret, Arhus.

(in Danish).

Sørensen, J., Glob, E., 1987. Influence of benthic fauna on trimethy-

lamine concentrations in coastal marine sediments. Mar. Ecol.

Prog. Ser. 39, 15–21.

Sørensen, H.M., Wiggers, L., 1997. Notat. Iltsvind i Mariager

Fjord 1997 - Status. 1997. Arhus Amt, Natur-og Miljøkontoret.

(in Danish).

decomposition of mytilus edulis: the effect on sediment nitrogen and carbon cycling

Documents