Chapter
3
ECOSYSTEM FUNCTION, CELL MICROCYCLING AND THE STRUCTURE OF
TRANSIENT BI O FILMS.
David
Graham,
M.
PATERSON1,
Rupert
PERKINS1,
Mireille
CONSAL
VEY1,
J.C. UNDERWOOD2
Gatty Marine" Laboratory.
University
of St Andrews
John Tabor Laboratories,
University
of Essex
dpl@St-andrews.ac.uk
1.
INTRODUCTION
The understanding of the complexity of benthic biofilms has progressed
rapidly in recent years, as technological advances have allowed analysis of
biofilms on more relevant temporal and spatial scales. However, many
different forms of biofilms exist and few have been extensively studied.
Biofilms form on any submerged or intermittently submerged surface and
most follow a known progression of development (or succession) towards an
equilibrium state related to environmental conditions. A particularly
dynamic sub-section of benthic biofilms are transient films that selfassembleand disperse in response to environmental conditions at the surface
of intertidal cohesive or mixed sediment systems (Paterson and Hagerthey
2001). These biofilms are comprised of motile autotrophic cells, given the
collective term microphytobenthos, and are largely comprised of diatom,
euglenid flagellates and cyanobacteria. A primary characteristic of these
organisms is their ability to migrate through sediments in response to
environmental cues and their inherent biological rhythms (Admiraal 1984).
Transient biofilms have a variable life-span related to the differential
pressures of tidal exposure, light conditions, grazing and hydrodynamic
forcing. These biofilms are formed from a matrix of cells a~d associated
extracellular polymeric substances(BPS) that many microphytobenthic cells
extrude through locomotion (Edgar and Pickett-Heaps, 1984). It is arguable
that in terms of temporal and spatial variability, transient biofilms are a
particularly complex group. Transient biofilms can form rapidly (within 20
min) (Paterson et al. 1998) and usually disperse shortly before or after the
tidal exposure period ends. At their most basic they may comprise a single
layer of cells (Paterson et al. 1998) or develop into a coherent matrix of cells
and polymeric substances,sometimes several mm in depth. The formation of
more sturdy biofilms may lead to microbial mat formation and the system
loses its transient nature, being retained over several tidal cycles, or even
become semi-permanent on a seasonal basis. An example of these semipermanent biofilms is the "bubble mat" system described in literature
(Yallop et al. 1994).
The ecological importance of transient biofilms has also been reassessed
in recent years. Although relatively thin, the films are widespread and
centres of very active primary production and biogeochemical activity
(Underwood and Kromkamp, 1999). In modern terms, the ecological role of
microphytobenthic biofilms can be seen in terms of the "ecosystem services"
they pr9vide. Transient biofilms form the basis of the food chain for many
coastal species; mediate the flux of nutrients across the sediment/water
interface (Nedwell et al. 1999) and are also "ecosystem engineers"
enhancing the resistance of the sediment to erosion (Paterson 1997, Black et
al. 2002).
Major advances in the understanding of the structure and function of
biofilm systems have been made since developments in micro-analytical
technology have allowed the examination of biofilm on a scale relevant to
the structure and processes, that occur within the film itself. These
developments have included the use of micro-sensor systems for oxygen
(Revsbech et al. 1980), nutrients and light (Consalvey, 2002) with the
analysis of biofilm properties at a 100 ~m scale (Taylor and Paterson, 1998;
Kelly et al. 200 I ). In addition, biofilms have now been viewed using a
variety of techniques, which enhance our conceptual understanding of their
structure, (Richards and Turner, 1984, Paterson, 1995, Defarge et al. 1996),
temporal plasticity (Consalvey 2002, Sauer et al. 2002,) and physiological
activity (Oxborough et al. 2000; Perkins et al. 2002). A potentially valuable
emerging technique is the ability to visualise and measure the photophysiological activity of individual cells within the biofilm. These
investigations have already demonstrated cellular responsesunder high light
conditions (Perkins et al., 2000).
There have been many attempts to define biofilms, usually based on their
taxonomy or structure (Neu 1994). Biofilms are often represented as a
matrix of relatively stable components, cell consortia, ~PS, .~nd space
arranged in three-dimensions. The forth dimension, time, is generally
ECOSYSTEM FUNCTION AND STRUCTURE OF BIOFILMS
49
considered to vary as successional processes occurring over a scale of days,
weeks and years. In permanent biofilms, the primary role of extracellular
polymeric material is anchorage. In contrast, EPS production provides the
means of locomotion to one of the most important groups of organisms
forming transient biofilms, the diatoms (Edgar and Pickett-Heap 1984). The
locomotive ability of microphytobenthos represents a possible evolutionary
strategy to maximise resource availability (light, space and nutrients) in
depositional environments. Cells rapidly congregate at the sediment surface
after the retreat of the tides. The traditional view was that cells accumulated
at the sediment surface and the relative role and turnover of cells within the
mat system was not considered. More limited locomotion is to be expected
where a biofilm forms in relation to a surface when attachment is a critical
ability. In transient biofilms, however, locomotion is the primary
characteristic. Evidence is accumulating, that locomotion of cells within the
microphytobenthic biofilm provides a mechanism whereby cells may
improve their individual "fitness" and contribute to the overall "efficiency"
of the biofilm by the sub-cycling of cells and consortia within the film on a
scale of minute to hours (Kromkamp et al. 1998, Perkins et al. 2002). This
paper presents evidence in support of the paradigm that the ecosystem
functioning of biofilm assemblages is more than the sum of the individual
parts (species or consortia). Light availability is a driving force. for
photosynthetic microbes, and migratory patterns are an adaptation to
maximising light utilisation. The temporal variability in biofilm structure,
light climate and photosynthetic activity is shown by electron microscopy,
light microscopy, light micro-profiling
and chlorophyll fluorescence
determination of cell and biofilm photophysiology.
2.
MATERIALS
AND METHODS
Sediment biofilms were collected from a variety of sites around the coast
of EuTope and the sources are indicated on figure captions. Artificial
biofilms were grown in laboratory culture using glass beads and a tidal
system with controlled temperature and day/night cycle (after Paterson,
1990).
50
3.
PATERSON. PERKlNS. CONSALVEY: UNDERWOOD
LOW TEMPERATURE
MICROSCOPY
SCANNING ELECTRON
Samples were obtained for low-temperature scanning electron
microscopy (L TSEM) by removal/excision of a 2 xI cm strip of biofilm
supported on a rigid metal foil planchette. The foil and biofilm was quench
frozen in liquid nitrogen ( -196°C) and then stored, under liquid nitrogen,
until viewed in the chamber of a specially-adapted scanning electron
microscope (JOEL 35CF SEM fitted with Oxford Instruments CT 1500B).
Samples were viewed while still frozen ( -180 OC) and surface water was
removed by heat etching (partial freeze drying) following the procedure
given in Paterson (1995).
4.
LIGHT MICROPROFILING
The light attenuation co-efficient of biofilms from 5 sites were
determined. 3 sites on the River Colne, Colchester, Essex, UK (51°50.2' N,
0°59.5' E) were investigated; a. euglenid- dominated site at Hythe and
diatom-dominated sites at Arlesford Creek and Point Clear. Sediment
biofilms from the Eden Estuary, U.K. (56°22'N, 02°51'W) and Sarilhos
Pequenos on the Tagus Estuary (38° N, 9° W) were also examined. Light
profiles through the sediment biofilms were made using fibre-optic
microsensors, constructed after Lassen et al. (1992). The fibre optic sensor
was inserted into the surface-sediment of a randomly selected sediment core,
at a 45° angle to reduce shading and moved down at 100 J.lmintervals using
a micromanipulator. The scalar irradiance at each depth was determined as a
percentage of surface incident light calculated from the associated drop in
voltage. The light extinction co-efficient (k) was calculated using the BeerLambert law:
Ez = Eo. exp( -kz)
(I)
Where Ez is the light level at depth z, Eo is the surface incident light level
and k is the attenuation coefficient.
ECOSYSTEM FUNCTION AND STRUCTURE OF BIOFILMS
5.
FLUORESCENCE MEASUREMENTS
ELECTRON TRANSPORT RATE
51
OF
Intact sediment cores were collected from Arlesford Creek and the Hythe
on the River Colne, Colchester, Essex, UK, using 7.5 cm diameter cores in
July 1999. Whereas the Arlesford Creek site is situated midway along the
estuarine salinity gradient and supports a diatom dominated biofilm, the
Hythe is at the head of the estuary and has lower salinity but higher nutrient
concentrations, and supports a biofilm comprised largely of euglenids and
cyanobacteria (Underwood et al. 1998; Perkins et al. 2002). Chlorophyll a
fluorescence images were obtained from surface scrapes (depth 1 cm)
carefully removed from the surface of the sediment cores and transferred to a
high-resolution.fluorescence imaging system (Oxborough and'Baker, 1997;
Oxborough et al. 2000; Perkiris et al. 2002). A detailed description of this
system is given in Oxborough and Baker {1997). Values of the quantum
efficiency of linear electron transport at PSII (F q'IFm1 from isolated cells
were calculated as the sum of individual pixel values for this parameter
(Oxborough and Baker, 1997; Oxborough et al. 2000).
Photosynthetic electron transport rate (ETR) versus light response curVes
were made on three replicate cuitures of Navicula salinarum, (Ehr.)
Reimann et Lewin. grown at a PPFD of 180 !.tmol m-2 s-I on a 14/ 10 h light
/ dark cycle at 18 °C in a growth cabinet using f/2 media (Guillard and
Ryther, 1962), salinity 20 and with an added antibiqtic cocktail to inhibit
bacteriafgrowth (Smith and Underwood, 1998). Data were compared to that
for intact diatom-dominated biofilms collected from Alresford Creek in July
1999. Chlorophyll fluorescence was measured using a Xenon PAM
fluorometer (Walz, GmbH, Effeltrich, Germany), the satur~~in~pulses used
for measurements of Fm' were at a PPFD of 8,600 !.tmol m s for 300 ms
and the non-actinic measuring beam frequency was set at 4 Hz. Chlorophyll
fluorescence was defined using a 680 nm bandpass filter (Coherent,
Watford, England) (Perkins et al., 2002).
ETR was calculated as the product of Fq 'IF m" photosynthetic photon flux
density (PPFD) and the specific light absorption coefficient for chlorophyll a
(a*), following Sakshauget al. (1997):
ETR=~.~.a*
F' m
a * was d~termined using an integrating
quantum
sensor. Biofilms
2
sphere and a cosine torrected
(2)
Skye
were sampled using the lens tissue technique
52
PATERSON. PERKlNS. CONSALVEY; UNDERWOOD
(Baton and Moss, 1966). Cells were re-suspended in f/2 media in replidishes. This was serially diluted to construct a standard curve of absorption
versus chlorophyll a concentration (over a range 50 to 800 ~g 1-1),from
which a* could be determined. Cultures of N. salinarum were also serially
diluted and then treated in the same manner as samples from intact biofilms.
Maximum BTR (BTRmax)and light utilisation coefficient (a) were calculated
by iteration (Long and Hiillgren, 1993), from light response curves (BTR vs.
PPFDs ofO to 830 umol m-2S-I),
6.
RESULTS
Qualitative variation in natural biofilm
structure was clearly
demonstrated by low-temperature scanning electron microscopy (Fig. 1-2).
Natural biofilms varied from isolated cells, to confluent unicellular layers
and to more extensive biofilms with several layers of cells (Fig. 1).
Comparative analysis of non-transient biofilms showed greater development
of the acellular polymeric matrix between the grains of sediment (Fig. 2a-b).
The development of the extensive films seemed to be restricted to noncohesive sediments and could be recreated in the laboratory (Fig 2d). An
extensive transient biofilm development on cohesive sediment was found
from an intertidal drainage channel from the Severn estuary which had an
unusual alignment of nitzschiod and naviculoid cells forming a matrix 500
I-Lmin extent (Fig le-f). Artificial biofilms were created which closely
resembled the structure of natural films ( compare Figs land 2d). Transient
biofilms were normally formed as a thin biogenic layer on the surface of the
bed (Fig. 2d) which disappeared as cells dispersed among the sediment
particles on downward migration. In laboratory cultures, migration ceased
after 10.:14 d and the surface layer became semi-permanent, and up to 600
I-Lmin depth after 14 d (Fig 2d). In one unusual case, natural biofilm
development was found to be discontinuous and related to the surface
topography of the sample (Fig 2e-f). In this area, diatoms were found only in
the hollows of a very uneven surface. This unusual formation was found on
the upper shore of a heavily bioturbated area on intertidal of the Severn
estuarY.
ECOSYSTEM FUNCTION AND STRUCTURE OF BIOFILMS
53
Figure I: Low-temperature scanning electron micrographs of transient biofilms. a). Mixed
assemblage of upper intertidal system dominated by cyanobacteria but still showing active
migration patterns. b). Close association between cyanobacteria and diatoms. c). Eugleniddominated biofilm.
d). Cross section of diatom-dominated biofilm showing characteristic
depth of several cells. E). Extensive transient biofilm of tidal channel (Severn estuary). F)
Detail of unusual uDriL!htDosition ofNitzschiod cells. All bar markers 100 um.
54
Figure
PATERSON. PERK/NS. CONSALVEY; UNDERWOOD
2: Low temperature scanning electron micrographs of permanent and cultured
biofilms. a).Thick cyanobacterial biofilm on the surface of non-cohesive (sandy) sediments.
b). Detail of organic matrix of thick permanent biofilm. c). Diatom biofilm cultures on a glass
bead substratum. D). Detail of organic matrix developing in glass bead culture. e). Biofilm on
the surface of heavily bioturbated sediments. Biofilm, dominated by diatoms, was found only
in the hollows. F). Detail of surface hollow filled with diatoms. All bar markers 100 ~m.
7.
LIGHT PROFILES AND BIOFILM
TYPE
The transmission of light through different biofilms was examined and
the light extinction coefficient (k) calculated. k was determined to
significantly vary between biofilm types (F4.12= 17.88, p < ~.OO5).(Table1).
Of the biofilms measured in this study, those dominated by euglenids (Hythe
ECOSYSTEM FUNCTION AND STRUCTURE OF BIOFILMS
55
site) were found to have the highest light attenuation co-efficients.
Significant differences between diatom-dominated sites were also observed.
The Arlesford Creek biofilm had the highest light attenuation coefficient
compared to the other diatom-dominated sites. Using these data light,
attenuation profiles oyer sediment depth were constructed (Fig. 3). Cells in
biofilms from the Hythe and Arlesford Creek would have experienced
almost complete darkness at a depth of 300 f.lm, whereas cells at the other
sites (diatom-dominated) would still have experienced > 4% of the incident
PPFD.
Hythe
,-
21.173
15.717
Arlesford
Creek
9.83
0.6279
Eden
9.61
1.1816
Tagus
11.14
1.6433
Point
8.
Clear
PHOTOPHYSIOLOGICAL
BIOFILMS AND CELLS
RESPONSES OF
Images obtained from the high resolution imaging system (Fig 4-6) are of
chlorophyll a fluorescence exposures (at F " just prior to application of the
saturating pulse) obtained at an actinic light level of 220 flmol PPFD,m-2 S-I.
The different community structures of the microphytobe~thic biofilms
sampled from the Hythe (Fig 4 a-b) and Arlesford Creek (Plate Fig 4c) are
clearly visible. The Hythe biofilm was comprised largely of euglenids (Fig
4a) with patches of dense cyanobacterial filaments (Fig. 4b). In contrast,
biofilms at Arlesford Creek were dominated by diatoms, principally
naviculoids and larger cells such as Pleurosigma sp., Gyrosigma sp., and
Nitzschia dubia. This variability in community structure has important
consequences when measuring variables such as. primary production and
electron transport, which vary greatly between taxa and hence biofilms. Such
variability is highlighted further by the variation in PSII quantum efficiency
(F q'IF m') shown (Fig 4c ). Individual cells showed different values of
Fq'IFm', despite being imaged at the same PPFD. Values of 0.152 and 0.315
were obtained for two cells of the same species,Nitzschia dubia. .
'jfi
PATERSON, PERKlNS, CONSALVEr; UNDERWOOD
The diatom-dominated biofilm obtained from Arlesford Creek varied in
community structure over a diel period (Fig 5). Selected fluorescence images
(imaged at 220 ~mol PPFD m-2 S-I as above) of the same biofilm (incubated
in a tidal tank system, see Perkins et al. 2002) obtained at 07:00, 13:00,
15:00 and 20:00 h demonstrated variability in biofilm appearanceover time.
Cells of Nitzschia dubia were present over the diel period, whilst smaller
naviculoids were most abundant at 07:00 h. Cells of Pleurosigma spp. were
noted primarily at 13:00 and 20:00 h, with Gyrosigma spp. particularly
apparent at 20:00 h. At least 10 images were taken at each time to ensure
those shown were representative of the biofilm overall.
The transient nature of a microphytobenthic biofilm in response to
changes in ambient light level (PPFD) was demonstrated (Fig. 6). Exactly
the same surface area of biofilm was exposed to incremental increases in
actinic light during a light response curve under the imaging system. Initially
the biofilm 'Y'fl:scomprised of a mixture of small naviculoid cells, with some
larger diatoms and euglenids (Fig. 6a, 180 ~mol m-2 S-I). However at a
higher PPFD of 1150 ~mol m-2 S-I, the surface of the biofilm was dominated
by eugl.enids(Fig. 6b). The cellular composition is clearly shown to vary in
responseto light climate.
The properties of the biofilm therefore varied depending on surface cell
type, light climate and local conditions. The performance of cells in culture
and those within a biofim matrix may also vary .This was investigated using
a Xenon PAM fluorometer (Walz, Germany). The light response curve for a
culture of Navicula salinarum grown at 180 ~mol m-2 S-I was compared to
that for an intact diatom-dominated biofilm obtained from Arlesford (Fig. 7).
The curve for N. salinarum shows a higher initial slope, corresponding to a
(0.29 :t. 0.0065 compared to 0.17 :t. 0.0014 ~mol e- m-2 ~g Chi a-1 ~mol
photons-1for N. salinarum and the intact biofilm respectively), but saturated
at a lowerETR, corresponding to ETRmax(4.5 :t. 0.57 compared to 6.1 :t. 0.42
~mol e- ~g ChI a-1 S-I for N. salinarum and the intact biofilm respectively),
compared to the biofiltn obtained from the field site (Fig. 7). This suggests a
comparatively shade-adapted photophysiology for the cultured cells
co:mpared to a light-adapted natural biofilm, with the former grown at
comparatively lower light levels than experienced by the natural biofilm in
situ in Julv 1999.
ECOSYSTEM FUNCTION AND STRUCTURE OF BIOFILMS
"7
s
'-'~
~
0Q)
0
Figure 3: Light attenuation profiles of 5 different biofilms. Hythe (euglenoid) = square;
Arlesford (:reek (diatom) = circle, Point Clear (diatom) = up triangle, Eden (diatom) = down
triangle and Tagus (diatom) = diamond.
Figure 4: Chlorophyll
a fluorescence images (pi,
imaged at 220 j.tmol PPFD m-2 S-I)
microphytobenthic biofilms dominated by euglenoids (A), cyanobacteria (8) and diatoms (C).
PSII quantum efficiencies (p q'IF m') are shown for individual diatom cells of a Pleurosigma
sp. and two Nitzschia
dubia cells (C). Plates (A) and (8) are of intact biofilms obtained from
the Hvthe and Plate (C) from Arlesford Creek in July 1999.
58
PATERSON, PERKINS, CONSALVEY; UNDERWOOD
Figure 5: Chlorophyll a fluorescence images (F', imaged at 220 I!mol PPFD m-2 S-I) of a
diatom dominated biofilm obtained from Arlesford Creek in July 1999. Images were taken at
(A) 07:00, (B) 13:00, (C) 17:00 and (D) 20:0Q h, from sediment cores incubated in a tidal tank
system. At least' 10 images were taken at each time to ensure those shown were representative
of the biofilm at each time.
~~
,~
Figure 6; Chlorophyll a fluorescence images (F') imaged at (A) 180 and (B) 1150 I!mol
PPFD m-2 S-I for a euglenoid dominated biofilm
collected from the Hythe in July 1999.
Images were taken within 8 minutes of e!lch other. This figure formed part of Fig. 3 in Perkins
et al., 2002.
59
ECOSYSTEM FUNCTION AND STRUCTURE OF BIOFILMS
7
N. salinarum
Diatom biofilm
.
6
0
)
0
0
0
0
0
5
0
4
)
)
r
,
.
3
~
.
.
.0
2
.
1
n
-i
O
='
,
0
.
0
.
.
0
.
0( 0
200
!
)
400
PPFD
800
600
(~mol
m-2
1000
S-1)
Figure 7: Light response curves of electrof! transport rate (ETR) versus light level (PPFD) for
a culture of Navicula salinarum grown at 180 I!molPPFD m-2 S-I (n=3, mean :t: s.e., closed
symbols) and a diatom-dominated biofilmfrom
Arlesford Creek collected in July 1999 (open
symbols)
9.
D ISCUSSI ON
Q1
Biofilm
structure
The structure of several types of intact microphytobenthic biofilms were
examined using L TSEM. Vari~tion in the structure, depth and coverage of
the biofilm was evident and, as expected, this variation led to differences in
light penetration through the biofilm, confirmed by light microsenor
measurements.However, it was also evident that some biofilms were highly
heterogeneousand that their formation was related to the surface micro relief
of the sediments (Fig 2e-f). This has rarely been documented and the
mechanisms behind the formation of these unusual structures are not known.
However, it is clear that the properties of the film will vary pn a s,calerelated
to the surface relief, in this case on a 50-200 ~m scale'. The region of
sediment where these biofilms were found had been highly bioturbated but
6()
PATERSON. PERKlNS. CONSALVEY. UNDERWOOD
was relatively dry at the time of sampling (during good weather). High light
and dry conditions may lead to biofilms developing in the shaded and
protected hollows of the uneven surface but this hypothesis would need
further investigation.
9.2
Light penetration
The assemblage type affected the penetration of light into the biofilm,
with light being attenuated most rapidly in euglenid biofilms (k = 21) at a
level more than 2 fold greater than the diatomaceous mat from the Eden
Estuary (k = 9). However, the depth of light penetration also varied between
diatomaceous biofilms. The variation in light penetration between different
biofilms dictates the depth to which cells remain photosynthetically active.
Greater k values will limit the photosynthetically active biomass (PAB) to
the surface .layers of the sediment and lead to a shallower zone of primary
productivity. Not only does the depth of light penetration have implications
regarding the overall primary productivity of microphytobenthic systems but
it also. influences the photophysiological state of the cells. It has been
suggested that cells are able to alter their position within the sediments
optimising light level to avoid photoinhibition/damage therefore resulting in
a maximal rate of productivity for the whole biofilm (Kromkamp et al. 1998,
Perkins et al. 2002).
9.3
Photophysiological responses of biofilms and cells
High resolution fluorescence imaging revealed a sequential change in
diatom taxa at the sediment surface; with a shift towards larger cells at the
end of the day. Furthermore short-term changes in taxa at the sediment
surface were also observed during a light curve; a shift occurring from
diatoms at low light levels to euglenid cells at high light. This demonstrates
the plasticity of microphytobenthic biofilms and the ability of cells to
respond rapidly to short-term environmental changes in light level. Vertical
migrations are widely described for a diverse range of microphytobenthic
taxa, the ubiquity of these behaviours suggeststhat these actions must confer
some evolutionary survival advantage (Cohn and Disparti 1994; Kingston
1999). Kingston ( 1999) experimentally verified the theory that upwards
migration maximises photosynthetic capacity and downward migration
minimises photo inhibition (through exposures to high light) in Euglena
proxima. Photoinhibition has rarely been recorded in benthic communities
(Blanchard and Cariou-LeGall 1994) and may be attributed to vertical
migration. The metabolic cost of a behavioural versus a ph.otophysiological
ECOSYSTEM FUNCTION AND STRUCTURE OF BIOFILMS
61
responseto high light is unknown and both strategies are likely to be used to
complement each other (e.g. Perkins et al., 2002).
A whole suite of factors is involved in the migratory response of diatoms
and other microphytobenthos, but the ultimate driving force remains
unknown. Due care must be taken with sampling to accommodate any
inherent changes in biomass based upon rhythmic migration as well as subcycling. The micio-spatial variability in biomass distribution over the
surface of the sediment may sometimes be significant. A biofilm may be
viewed as more than the sum of its parts, with surface cells shading and
perhaps benefiting the cells underneath, but themselves migrating down
when necessary,allowing other cells to take their place.
CONCLUSION
10.
The functional role of. biofilms is recognised as significant in the
structuring of coastal ecosystems (Paterson and Hagerthey, 2001). However,
is becomiJlg clear that the capabilities of individual cells cannot be used to
determine the limits of biofilm function. Evolution has resulted in a system
of dynamic turnover which maximise the functional capacities of the biofilm
and maximise the extraction of resources from the environment. More
research is now required to examine the mechanisms that allow a biofilm to
be more than the sum of its parts.
REFERENCES
Admiraal, W .1984. The ecology of estuarine sediment-inhabiting diatoms. In: Prog. Phycol.
Res. volume 3, (eds, Round. F.E. & Chapman. D.J.). Biopress. pp. 269-322.
Black, K.S., Tolhurst, T.J., Hagerthey, S.E., and Paterson, D.M. 2002. Working with Natural
Cohesive Sediments. JHydr.Eng.128(1):
Blanchard,
G. F. and v.
Cariou-Le
Gall.
1-7.
1994. Photosynthetic
characteristics
of
microphytobenthos on Marennes-Oleron bay, France: Preliminary results. J. Exp.
Bioi. Ecol. 182: 1-14.
Cohn, S. A. and N. C. Disparti. 1994. Environmental factors influencing diatom cell motility.
.I: Phycol. 30: 818-828.
Consalvey, M. 2002. The structure and function ofmicrophytobenthic
biofilms. Ph.D. Thesis.
University of St Andrews.
Defarge, C., Trichet, J., Jaunet, A., Robert, M., Tribble, J. and Sansone, F.J. 1996. Texture of
microbial sediments revealed by cryo-scanning electron microscopy. J. Sed.. Res. 66:
91S-947
"
'.
62
PATERSON. PERKINS. CONSALVEY; UNDERWOOD
Eaton, J.W. & Moss, B. 1966. The estimation of numbers and pigment content in epipelic
algal populations. Limnol. Oceanog. 11: 584-595.
Edgar, L.A. & Pickett-Heaps, J.D. 1984. Diatom locomotion. In: Frog. Fhycol. Res. 3: 47-88,
Biopress, Bristol, UK.
Guillard, R.R.L. and Ryther, J.H. 1962. Studies of marine planktonic diatoms. I. Cyclotella
nana Hustedt, and Detonula confervacea (Cleve) Gran. Can. J. Mar. Bioi. 27: 191-196
Honeywill,
C., Paterson, D.M and Hagerthey, S.E. Determination
of microphytobenthic
biomass using pulse amplitude modulated minimum fluorescence. Eur J. of Fhycol.
(In press).
Kelly, J., Honeywill, C. and Paterson, D.M. 2001. Microscale analysis of chlorophyll-a in
cohesive, intertidal sediments: the implications for microphytobenthos distribution..
Mar. Bioi. and Ecol. 81: 151-162.
Kingston, M.~B. 1999. Effect of light on vertical migration and photosynthesis of Euglena
proxima (Euglenophyta). .I: Fhycology 35: 245-253.
Kromkamp, J.~,Barranguet, C and.Peene, J. 1998. Determination of microphytobenthos PSII
quantum efficiency and photosynthetic activity by means of variable chlorophyll
fluorescence. Mar. Ecol. Frog. Ser. 162: 45-55.
Lassen, G. Ploug, H. & Jorgensen, B.B. 1992. A fibre-optic scalar irradiance microsensor:
Application for spectral light measurements in sediments. FEMS Microbial. Ecol. 86:
247-254.
Long, S.P. and Hiillgren, J-E. 1993. Measurement of CO2 assimilation by plants in the field
and laboratory. In: DO Hall, J MO Scurlock HR,Bothar-Nordenkamps
RC, Leegood,
SPLong (eds). Photosynthesis and production in a Changing Environment, afield and
laboratory manual. Chapman and Hall, St. Ives, UK
Nedwell, D. B., Jickells T. D., Trimmer, M. and R. Sanders. 1999. Nutrients in Estuaries. Ed.
D. B. Nedwell andD. G. Raffaelli. Adv. Ecol. Res: Estuaries. 29: 43-92.
Neu, T.R. 1994. Biofilms and microbial mats. In: Biostabilisation
of sediments. (Krumbein,
W.E., Paterson, D.M. and Stal, L.J., eds), BIS, Carl von Ossietzky Univ. Oldenburg,
9-15.
Oxborough, K. and Baker, N.R. 1997. An instrument capable of imaging chlorophyll
a
fluorescence from intact leaves at very low irradiance and at cellular and subcellular
levels of organization. Plant Cell Environ. 20: 1473-1483.
Oxborough, K.,Hanlon, A.R.M., Underwood, G.J.C. and Baker N.R. 2000. In vivo estimation
of the photosystem II photoch~mical efficiency of individual microphytobenthic cells
using high-resolution imaging of chlorophyll a fluorescence. Limnol 9ceanogr. 45:
1420-1425.
Paterson, D.M. 1.990. The influence of epipelic diatoms on the erodibility
of an artificial
sediment. In: proceedings of the lOth International Symposium on Living and Fossil
Diatoms. (Simola, H. ed) Joensuu, 1988. Koenigstein, 345-355.
Paterson, D.M. 1995. The biogenic structure of early sediment fabric visualised by lowtemperature scanning electron microscopy. .I: Geol. Soc. 152: 131-~40.
.
63
ECOSYSTEM FuNcTION AND STRUCTURE OF BIOFILMS
Paterson, D.M. and Hagerthey,
ecosystems: Biology
S.E. 2001. Microphytobenthos
in contrasting
coastal
and dynamics. In Ecological Comparisons of Sedimentary
shores (K.Reise, ed), Ecological stud!es, 151: 105-125
Paterson, D.M. Yates, M.G., Wiltshire,
K.H., McGrorty,
S. Miles, A. Eastwood, J.E.A.,
Blackburn, J. & Davidson, I. 1998. Microbiological
mediation of spectral reflectance
from intertidal cohesive sediments. Limnol. Oceanog. 43(6): 1207-1221.
Paterson, D.M., 1997. Biological mediation of sediment erodibility:
ecology and physical
dynamics. In: Cohesive Sediments (Burt, N., Parker, R. and Watts, J. eds), Wiley and
Sons, pp. 215-229.
Perkins, R.G., Oxborough, K., Hanlon, A.R.M., Underwood, J.C. and Baker, N.R. 2002. Can
chlorophyll
fluorescence be used to estimate the rate of photosynthetic electron
transport within microphytobenthic biofilms? Mar. Ecol. Prog. Ser. 228: 47- 56.
Revsbech, N. P., S0rensen, J., Blackburn, T. H., Lomholt, J. P. 1980. Distribution of oxygen
in marine sediments measured with microelectrodes. Limnol. Oceanogr. 25(4): 03411.
Richards. R.S. and Turner, R.J. 1984. A comparative study of techniques for the examination
ofbiofilms by scanning electron microscopy. Water Res. 18(6): 767- 773.
Sakshaug, E., Bricaud, A., Dandonneau, Y., Falkowski, P.G., Kiefer, D.A., Legendre, L.
.
Morel, A., Parslow, J. and Takahashi, M. 1997. Parameters of photosynthesis:
definitions, theory and interpretation ofresults. .I: Plankton Res. 19: 1637-1670.
Sauer, J., Wenderoth, K., Maier, U.G. and Rhiel, E. 2002. Effects of salinity, light and time on
the vertical migration of diatom assemblages.Diatom Res. 17(1), 189-203.
Taylor IS&
Paterson DM
1998. Microspatial variation in carbohydrate concentrations with
depth in the upper millimetres of intertidal cohesive sediments. Est. Coast. Shelf Sci.
46: 359-370
Underwood, G.J.C., Phillips, J. and Saunders, K. 1998. Distribution
of estuarine benthic
diatom species along salinity and nutrient gradients. Eur.l: Phycol. 33: 173-183
Underwood, G. J. C. & Kromkamp,
J. 1999. Primary production by phytoplankton
&
microphytobenthos in estuaries. Adv. in Ecol. Res: Estuaries. D. B. Nedwell & D. G.
Raffaelli. Acedemic Press, 29: 93-153.
Underwood, G.J.C. & Smith, D.J. 1998. In situ measurement of exoploymerproduction
by
intertidal epipelic diatom dominated biofilms in the Humber estuary. In: Sedimentary
Processes in the Intertidal Zone. (Black, K. S., Paterson, D. M., & Cramp, A. eds).
Geological Society, London, Special publication. 139: 125-134.
Yallop, M.L. de Winder, B., Paterson, D.M. & Stal, L.J. 1994. Comparative structure primary
production and biogenic stabilisation of cohesive and non-cohesivemarine sediments
inhabited by microphytobenthos._Est. Coastal & ShelfSci.. 39: 565-582.