DYNAMICS OF ESTUARINE MICROPHYTOBENTHOS IN A
SHALLOW WATER SAND BOTTOM HABITAT
by
Jeffrey Garner Allison
B.S., The University of West Florida, 2000
A thesis submitted to the Department of Biology College of Arts and Sciences The University of West Florida In partial fulfillment of the requirements for the degree of Master of Science
2006 The thesis of Jeffrey Garner Allison is approved:
Joe E. Lepo, Ph.D., Committee Member Date
Jane M. Caffrey, Ph.D., Committee Member Date
Richard A. Snyder, Ph.D., Committee Chair Date
Accepted for the Department/Division:
George L. Stewart, Ph.D., Chair Date
Accepted for the College:
Jane S. Halonen, Ph.D., Dean Date
Accepted for the University:
Richard S. Podemski, Ph.D., Dean of Graduate Studies Date
ii ACKNOWLEDGMENTS
I am grateful for loving, caring, and kind family and friends through which my emotional stability, trust and support was upheld. I would like to thank my major advisor Dr. Snyder as well as my committee members, Dr. Lepo and Dr. Caffrey, for the opportunity to complete a
Master of Science in Biology degree. My thesis work would not have been possible without their guidance and lasting direction. Likewise, the support of my colleagues, Matt Wagner, Mike
McAllister, Alex Ren, Paige Cramer, Ashley Moore, Kristen Hellein, Joe Moss, Hugo Castillo,
Alan Knowles, and Melissa Ederington-Hagy enabled me to accomplish my goals and were instrumental throughout experimentation and analysis. Technical information and assistance was generously provided by Jim Hammond, Jan Macauley, Steve McLin, and Tanya Streeter. I especially appreciate the ability to use the facilities at Gulf Islands National Seashore, thanks to
Riley Hoggard and Permit #: GUIS-2005-SCI-003.
iii TABLE OF CONTENTS
Page
ACKNOWLEDGMENTS ...... iii
LIST OF TABLES...... vi
LIST OF FIGURES ...... vii
ABSTRACT...... x
CHAPTER I. INTRODUCTION...... 1 A. Benthic Microalgal Communities...... 1 B. Measuring Benthic Microalgal Production...... 4 C. Measuring Benthic Respiration...... 7 D. Benthic Microalgal Biomass...... 10 E. Benthic Nutrient Fluxes ...... 10 F. Benthic Chamber Technique...... 12 G. Measuring Phytoplankton Production...... 14 H. Microalgae in Biofilms ...... 15 I. Enzymatic Activity Assays...... 16 J. Study Area ...... 20
CHAPTER II. OBJECTIVES & HYPOTHESIS ...... 23
CHAPTER III. MATERIALS & METHODS ...... 24 A. Site Characteristics...... 24 B. Physicochemical Determinations...... 24 C. Chamber Procedures ...... 25 D. Biofilm Analysis ...... 27 E. Water Column Analysis...... 27 F. Sediment Procedures...... 28 G. Oxygen Determinations ...... 29 H. Biomass Determinations ...... 30 a. Chlorophyll a ...... 30 b. Biofilm Biomass by Optical Density ...... 31 c. Organic Content...... 31 iv d. Elemental Analysis ...... 31 I. Compensation Point Determinations ...... 32 J. Nutrient Analysis ...... 32 K. Enzyme Assays ...... 33 L. Statistical Analysis...... 37
CHAPTER IV. RESULTS ...... 38 A. Ambient Physicochemical Conditions...... 38 B. Sediment, Water Column and Biofilm Biomass...... 46 C. Benthic Chamber Results...... 56 D. Compensation Point Analysis...... 61 E. Chamber Nutrient Fluxes...... 62 F. Enzyme Activity Data...... 64
CHAPTER V. DISCUSSION...... 67 A. Conclusions...... 80
REFERENCES ...... 82
APPENDIX...... 98
v LIST OF TABLES
Table Page
1. Sampling Dates, Incubation Times, and Ambient Dissolved Oxygen Concentrations of Bottom Water at Butcherpen Cove ...... 39
2. Pearson Correlations for Benthic Chamber Production, Respiration, and Benthic Photopigments as well as Correlations with Biofilm Photopigments, Nutrients, and Physicochemical Parameters...... 52
3. Pearson Correlations for Phytoplankton Versus Biofilm Production, Respiration, and Benthic Photopigments as well as Correlations with Phytoplankton, Sediment, and Biofilm Photopigments, Nutrients, and Physicochemical Parameters...... 53
4. Pearson Correlation Matrix for Biofilm Production, Respiration, and Photopigments; Biomass; Water Column Nutrients; Esterase Activity; and Temperature ...... 54
vi LIST OF FIGURES
Figure Page
1. Location of Butcherpen Cove within the Pensacola Bay estuary in northwest Florida along the northern Gulf of Mexico coast; Approximate location of National Weather Service precipitation gauge and The University of West Florida ...... 22
2. (A) A dark benthic chamber with attached tubing for syringe sampling and current simulation device. (B) A benthic periphytometer containing acrylic sampling plates and glass slides. Image is from April 15, 2005, showing attached freshwater macroalgae. (C) A clear benthic chamber and greenhouse shade cloth. Internal stirring paddles are visible in this image...... 26
3. Total recorded rainfall (mm) in Pensacola, Florida during the experimental season of 2005 (February through August) ...... 40
4. Percent transmission of PAR (% PAR) to the benthos and bottom water salinity measures during 2005 (February through August) ...... 41
- - -2 -1 5. Mean (± 1 SE) light/dark sediment fluxes of NO3 + NO2 (µmol · m · hr ) - - and water column concentrations of NO3 + NO2 (µM) in 2005 (February through August) ...... 42
+ -2 -1 6. Mean (± 1 SE) light/dark sediment fluxes of NH4 (µmol · m · hr ) and + water column concentrations of NH4 (µM) in 2005 (February through August)...... 43
7. Mean (± 1 SE) water column concentrations of total kjeldahl nitrogen (µM) and total phosphorous (µM) in 2005 (February through August) ...... 44
-3 -2 -1 8. Mean (± 1 SE) light/dark sediment fluxes of PO4 (µmol · m · hr ) and -3 water column concentrations of PO4 (µM) in 2005 (February through August)...... 45
-2 -1 9. Mean (± 1 SE) light/dark sediment fluxes of SiO2 (µmol · m · hr ) and water column concentrations of SiO2 (µM) in 2005 (February through August)...... 45 vii 10. Sediment core total carbon (nmol/cm2), total nitrogen (nmol/cm2), and total phosphorous (µmol/cm2) during 2005 (April through August) ...... 47
11. Sediment core and biofilm carbon/nitrogen ratios during 2005 (April through August) ...... 47
12. Biofilm total carbon (µmol/cm2), total nitrogen (µmol/cm2), and total phosphorous (µmol/cm2) during 2005 (April through August) ...... 48
13. Sediment photopigments, Chl a, Chl c, and Pheo a (µg/cm2) during 2005 (March through August) ...... 49
14. Biofilm photopigments, Chl a, Chl c, and Pheo a (µg/cm2) during 2005 (April through August) ...... 50
15. Near bottom water photopigments, Chl a, Chl c, and Pheo a (mg/m2) during 2005 (March through August) ...... 50
16. Sediment and biofilm ash free dry weight (mg/cm2) during 2005 (April through August) ...... 55
17. Biofilm optical density, ash free dry weight (g/cm2), and biofilm Chl a (µg/cm2) during 2005 (April through August)...... 55
18. Benthic net and benthic microalgal gross production and benthic respiration (mg C · m-2 · hr-1) during 2005 (February through August)...... 57
19. Bottom water temperature (ºC), phytoplankton gross production (mg C · m-2 · hr-1), and phytoplanktonic net production and planktonic respiration (mg C · m-2 · hr-1) during 2005 (February through August)...... 58
20. Bottom water temperature (ºC), biofilm gross production (mg C · m-2 · hr-1), and biofilm net production and respiration (mg C · m-2 · hr-1) during 2005 (February through August) ...... 59
21. Phytoplankton and biofilm gross production (mg C · m-2 · hr-1) during 2005 (February through August) ...... 60
22. Planktonic, benthic, and biofilm production/respiration ratios during 2005 (February through August) ...... 60
23. Benthic photosynthetic efficiency (mg C · mg Chl a-1 · m-2 · hr-1) response to irradiance intensity (μE · m-2 · s-1) ...... 61
viii 24. Biofilm photosynthetic efficiency (mg C · mg Chl a-1 · m-2 · hr-1) response to irradiance intensity (μE · m-2 · s-1) ...... 62
25. Fluorescein diacetate activity (mM · m-2 · hr-1) and respiration rates of biofilms (mg C · m-2 · hr-1) during 2005 (April through August) ...... 65
26. Alkaline phosphatase activity (µM · m-2 · hr-1) of biofilms during 2005 (April through August) ...... 65
27. Denitrification (acetylene block assay) rates and nitrogen fixation (acetylene reduction assay) rates (nmol N · g DW-1 · hr-1) of biofilms grown on artificial substrata during 2005 (April through August)...... 66
28. Percentage of incident light reaching the bottom of Pensacola Bay. From Murrell (unpublished data), reprinted here with permission ...... 74
ix ABSTRACT
DYNAMICS OF ESTUARINE MICROPHYTOBENTHOS IN A SHALLOW WATER SAND BOTTOM HABITAT
Jeffrey Garner Allison
Microphytobenthos inhabiting subtidal estuarine sediments are important contributors to marine food webs and biogeochemical transformations of nutrients.
Production and respiration rates, biomass estimations, and nutrient flux rates in
Butcherpen Cove, Pensacola Bay, Florida, were investigated from February through
August 2005 for the benthos, phytoplankton, and biofilms grown on artificial substrates.
Extracellular enzyme activity was assayed on slurries of biofilm material. Production and respiration were measured by oxygen changes in light and dark incubations in situ.
Microphytobenthic production and biomass decreased following major storm events.
Water column phytoplankton and periphyton did not respond to rainfall and appeared to be controlled by temperature. Phytoplankton production and respiration increased through the sampling season and variability was low. Periphyton grown on artificial substrates did not correlate with benthic processes, but appeared to be responding to water column conditions. Bioavailability of water column nutrients did not seem to be as important to benthic production as much as benthic fluxes. The data suggest nitrogen and phosphorous nutrient co-limitations existed for both phytoplankton and benthic x -3 microalgae in Butcherpen Cove. When dissolved PO4 increased in the water column
alkaline phosphatase activity was suppressed. Esterase activity of biofilms on artificial
substrates reflected biofilm community respiration. N-fixation and denitrification rates
were low and mutually exclusive.
xi CHAPTER I
INTRODUCTION
Benthic Microalgal Communities
Benthic microalgae, microphytobenthos, periphyton, and biofilms are communities dominated by Bacillariophyceae. Benthic diatoms are tolerant of a wide range of temperature, light, and salinity regimes. Microphytobenthos may also include chlorophytes, cyanophytes, and other photoautotrophic taxa that can be locally dominant in algal mats, but are usually present as minor components of shallow water estuarine benthos. These epibenthic communities are a primary source of carbon for estuarine food webs (Kang, Kim, Lee, Lee, & Hong, 2003), have significant roles in ecosystem level nutrient cycling (Nicholson, Longmore, & Berelson, 1999), and aid in sediment stabilization (Holland, Zingmark, & Dean, 1974).
The relative contribution of benthic microalgae to ecosystem level food webs and overall productivity is considered to be high (Cahoon, 1999; Kang et al., 2003; Schmid-
Araya & Schmid, 2000). Stable isotope tracer studies have shown that benthic consumers
(deposit and suspension feeders, predators, and omnivores) are supported predominantly from microphytobenthic production (Kang et al.; Sullivan & Moncreiff, 1990). Kang et al. suggested that benthic microalgae may be more important in bays with tidal flats and intertidal zones, than in offshore waters, although benthic algae production has been
1 2
found to equal water column production on the nearshore mid-Atlantic US continental
shelf (Cahoon & Cooke, 1992).
In addition to the significance of benthic microalgae as a food source for benthic
invertebrates, microphytobenthos are also known to furnish organic carbon to the
planktonic system through suspension into the water column caused by tidal currents and
wind-induced turbulence (Brandini, da Silva, Pellizzari, Fonseca, & Fernandes, 2001).
Surface wave energy acts as a subtidal pump bringing bottom water up to mix into the
water column through bottom hydrostatic pressure fluctuations, especially in shallow
waters (Malan & McLachlan, 1991). Shallow subtidal environments also have burial of
organic matter through advective transport (Hüettel & Gust, 1992).
Acknowledgment of the importance of benthic microalgae to estuarine
ecosystems are common in the literature, yet only a few studies (Cahoon, 1999; Schreiber
& Pennock, 1995) have attempted to evaluate the contribution of microphytobenthos
production to total estuarine production. Scaling up point measurements to ecosystem
levels propagates errors from the high degree of spatial and temporal heterogeneity of the
microphytobenthos within estuaries (Underwood & Kromkamp, 1999), but the published
estimates are instructive. Cahoon, Redman, and Tronzo (1990) found that up to 80% of
the chlorophyll a (Chl a) in Onslow Bay, NC was associated with the sediment and the
benthic Chl a almost always surpassed the value of Chl a for the integrated water column
samples. In Weeks Bay, Alabama, benthic production was estimated to contribute 21%
(90 g C · m-2 · yr-1) to total system production (Schreiber & Pennock). Thom and Albright
(1990) estimated Puget Sound’s coarse grained, sandy sediments to have annual net benthic production nearly equal to the phytoplankton (676 and 649 g C · m-2 · yr-1,
3 respectively). Conversely, phytoplankton production (468 g C · m-2 · yr-1) was slightly greater than benthic microalgal production (339 g C · m-2 · yr-1) in a Gulf of Mexico seagrass habitat (Daehnick, Sullivan, & Moncreiff, 1992; Moncreiff, Sullivan, &
Daehnick, 1992).
Estimates for benthic microalgae production in shallow estuarine and intertidal zones studies have documented high productivity rates. Temperate zone studies have indicated values for annual productivity as high as 892 g C · m-2 · yr-1 and hourly production rates topping 0.8 g C · m-2 · hr-1 (Hargrave, Prouse, Phillips, & Neame, 1983).
Production rate estimations for microphytobenthos in tropical zones can be even higher with annual values as high as 3760 g C · m-2 · yr-1 (Hawkins & Lewis, 1982).
Several investigators (Sullivan & Moncreiff, 1988; Cahoon & Cooke, 1992) observed a single late winter-early spring biomass peak that was triggered by high nutrients, increasing temperature and increasing day length coincident with the spring bloom in the water column. A decrease in biomass in late spring was attributed to increased grazing pressure as opposed to decreased production. However, Sullivan and
Daiber (1975) suggested that seasonal variation in benthic biomass was solely regulated by variations in nutrient concentrations.
Shallow subtidal environments are exposed to a wide range of light intensities due to water cover (Pinckney & Zingmark, 1993), variations in turbidity (Brotas & Catarino,
1995), and cloud cover (Cahoon & Cooke, 1992). Patchy distributions, or micro- heterogeneity, of benthic microalgae biomass and production has often been reported for estuarine habitats (Admiraal, 1984) and has been attributed to variations in light and temperature. Light penetration into sediments is grain size dependent and often limited to
4 the upper 5 mm of sediment. Light scattering by sediment particles may double light availability in the surface layers beyond measures of direct incident light (Lassen, Ploug,
& Jørgensen, 1992). Benthic primary production is often reduced by cloud cover, with light reductions of an order of magnitude. In deeper waters, Cahoon and Cooke suggest that light and nutrient availability on the nearshore shelf seafloor do not substantially limit benthic primary production since macroalgae were observed beyond the North
Carolina shelf break at 55 m depth. In an earlier study by Cahoon et al. (1990), Chl a was found over the entire continental shelf bottom off of Onslow Bay, NC from depths of 15 to 41 m. A significant positive correlation between the concentrations of ATP and Chl a in these data suggested the presence of viable microphytobenthos.
Measuring Benthic Microalgal Production
Methods to measure benthic primary production have included both in situ methods (benthic chambers) and laboratory studies (core samples). Methods for analysis of production have included O2 exchange methods, pore water chemistry changes, radiocarbon tracer techniques, O2 microelectrode methods, use of an InfraRed Gas
Analyzer to measure fluxes of gaseous carbon dioxide, and mathematical extrapolations based on the relationship between measures of light intensity and benthic microalgal biomass. Benthic chambers allow measurements of known area of sediment in contact with a known volume of overlying water. The primary assumption involved in benthic chamber + O2 exchange method is that the rate of O2 release (production) or uptake
(respiration) is independent of the O2 concentration in the water overlying the sediment
5
(Hofman & de Jong, 1993). This allows determination of community production (P) to respiration (R) ratio (P:R) in addition to gross primary production (GPP) and net primary production (NPP) estimates. A major advantage of this method is that the sediment surface area enclosed by chambers can be large, minimizing the effect of micro- heterogeneity. Alteration and manipulation of sediments is minimal with chamber deployments. Incubation occurs under near ambient environmental conditions. Chamber studies also incorporate the intact benthic infauna community contributing to community respiration and nutrient fluxes across the sediment-water interface.
Despite these benefits, relatively few studies have attempted to measure benthic microalgal production in situ, mostly because of the logistical difficulties involved.
Laboratory analysis of core samples allows greater control of variables to estimate potential production, but also involves substantial disruption of the benthic boundary layer (BBL) during both the actual sample grab and the transport to the analytical laboratory. In situ incubation production and respiration are subject to variability associated with natural light regimes and ambient temperature conditions. Other methods used in estimating benthic primary production present more compromises than the O2 exchange method using benthic chambers. Benthic chambers have been used to obtain estimates of in situ benthic nutrient fluxes in the deep-sea, shallow coastal systems, and estuaries (Berelson et al., 1987; Fisher, Carlson, & Barber, 1982; Nicholson et al., 1999).
Defining the O2 gradients with O2 microelectrodes across the redox discontinuity layer (RDL) permits the calculation of O2 fluxes into or out of the sediment when both the diffusion coefficient and the porosity are known (Hofman & de Jong, 1993). O2 microelectrodes offer several advantages for estimating production including relative
6
inexpensiveness, ease of calibration, precision, accuracy, and simplicity in use. They also
can be used with exposed intertidal sediments, cores, or for direct in situ measures on submerged sediments. Because of the small area characterized, scaling up observations to large areas (e.g., estuaries) can be problematic. Microelectrodes are also very thin and easily breakable (Hofman & de Jong). O2 consumption rates cannot be acquired directly
so the O2 gradients measured by microelectrodes must be converted to fluxes using an
appropriate sediment diffusion coefficient. After darkening the sediment surface oxygen
demand can be accounted for by calculating the initial slope of oxygen decrease at 50 μm
intervals (Pinckney & Zingmark, 1993). Each depth interval is integrated over the entire
sediment profile to give a depth-integrated area (i.e., per m2) estimate of GPP. Unlike 14C
or coring methods, microelectrode techniques are nondestructive to the sediment surface
and allow for multiple measures on the same core sample or sediment patch in situ
(Pinckney & Zingmark).
In benthic chamber experiments, net O2 fluxes can be derived by transforming the
change of O2 concentration into a flux through the sediment-water interface.
Phytoplankton O2 production/consumption in the chamber water is accounted for to
isolate net benthic primary production. Net benthic primary production (mg C · m-2 · hr-1) can be calculated from dissolved oxygen (DO) concentration changes in light chambers using the formula for non-standard sample volumes (Strickland & Parsons, 1972).
7
NBPP = (([DO]t2 – [DO]t1) * V * 12)/(PQ * H * A) (1)
where:
NBPP = net benthic primary production in mg C · m-2 · hr-1
[DO]t2 = dissolved oxygen concentration in mmol/L at the end of incubation
[DO]t1 = dissolved oxygen concentration in mmol/L at the beginning of incubation
V = volume under the benthic chamber
12 = the atomic weight of carbon
PQ = photosynthetic quotient (mol O2 evolved/mol C fixed), 1.2 (Strickland & Parsons,
1972)
H = incubation time in hours
A = area under the benthic chamber.
Grant (1986) recommends an equation to convert production in oxygen units to carbon units in order to compare various studies: μmol O2 * 0.012 = mg C (assuming PQ
= 1). However, there is no uniform agreement in the literature on conversion factors,
-2 -1 -2 -1 Cahoon (1999) offers a conversion equation of: mg C · m · hr = mg O2 · m · hr *
0.32. In the current investigation, mmol O2 * 12 = mg C was used as the conversion
factor.
Measuring Benthic Respiration
Measurement of benthic respiration, as biological oxygen consumption, is
complicated by chemical oxygen demand (COD) and variation in biological oxygen
8 demand (BOD) in the sediments. Several researchers (Hargrave, 1969; Hopkinson, Day,
& Gael, 1978; Patmatmat, 1971) suggested the magnitude and temporal patterns of benthic respiration are a function of organic matter supply rates. One problem in the use of benthic chambers is that they limit the supply of labile organic matter to the sediment surface and disturb the natural flow regime of the ambient water affecting diffusion and advection processes (Hundling & Hargrave, 1973). Respiration rates of microalgae include photorespiration and may be higher in the light than dark (Cahoon & Cooke,
1992). Although respiration rates are rarely constant over diel or seasonal cycles, an assumption is made that respiration is constant in order to extrapolate measures to daily or annual rates.
Values for benthic respiration should be calculated in a similar fashion as production estimations using the dark benthic chambers and according to:
RESP = (([DO]t2 – [DO]t1) * V * 12)/(RQ * H * A) (2) where:
RESP = benthic oxygen consumption (the sum of biological respiration and chemical oxygen demand; Patmatmat, 1971) in mg C · m-2 · hr-1
RQ = respiratory quotient (mol C respired/mol O2 consumed), 1.0
(Strickland & Parsons, 1972)
Other parameters are same as NBPP.
Oxygen consumption can be converted to carbon equivalents by assuming that for every mL of O2 (44.6 μmol) consumed, approximately 0.42 mg of CO2 carbon was
9 produced (Rowe & Phoel, 1992). This also relies on the assumption that the organic matter consumed is composed equally of lipids, carbohydrates, and proteins. Patmatmat
(1971) included both BOD and COD as sediment oxygen demand (SOD).
Determining the fraction of the benthic respiration rate that is BOD can be problematic. Jørgensen (1985) found that a significant portion of oxygen consumption is due to the oxidation of sulfides. Forja and Gomez-Parra (1998) commented that a reduction in the oxygen concentration inside the chambers potentially affects the metabolism of the aerobic microorganisms and may raise the level of the RDL in the sediments. A few studies (Hopkinson et al., 1978; Hargrave, 1969) have inoculated cores with antibiotics to determine the contribution of bacterial respiration to BOD. Hopkinson et al. added formaldehyde (5%) to samples so abiotic O2 consumption could be subtracted away from the total O2 uptake. In their study, BOD accounted for 51% of total O2 consumption. COD was observed by Hopkinson et al. to be much less a factor with in situ benthic chambers due to their larger cross sectional area (twice that of the cores used) and the lower disturbance of the benthic electrochemical gradient. Increased COD may be caused by exposing reduced materials during core sampling.
Gross primary production (GPP) is calculated by correcting net production values for SOD. GPP is a better measure of microalgal activity than net production (Cahoon &
Cooke, 1992) since the measurement of respiration is not specific to the algae. For benthic microalgae, NPP has been reported as > 90% of GPP (Cahoon & Cooke; Colijn
& de Jonge, 1984; Grant, 1986; Pomeroy, 1959), so that GPP may be used as an estimate of microalgal growth with less error than measuring NPP by incorporating estimates of
SOD.
10
Benthic Microalgal Biomass
Difficulties with calculating biomass from cell abundance or counting microalgae
has led researchers to quantify benthic microalgal biomass using chlorophyll a (Chl a;
MacIntyre, Geider, & Miller, 1996). Benthic microalgal biomass was best estimated as
chlorophyll a (mg Chl a/m2) according to Whitney and Darley (1979) with a hexane
extraction to limit interference from degraded pigments common in sediment samples.
The ratio of total pheopigments to Chl a has been used as an indicator of the
physiological state of microalgal communities, with higher values indicating a stressed or
declining community while lower numbers suggest an actively growing community
(Bidigare, Frank, Zastrow, & Brooks, 1986). Production to biomass ratio (P:B), has also
been used as an indicator of condition. This ratio, also known as the photosynthetic
-2 -1 efficiency, is calculated as net production (μmol O2 · m · hr ) divided by biomass (mg
Chl a/m2) and in benthic production studies ranges from 0.1 to 0.8 (Grant, 1986;
Pinckney & Zingmark, 1993; Sullivan, & Moncreiff, 1988).
Benthic Nutrient Fluxes
Microbial breakdown of organic material in sediments results in nutrient regeneration and an outward flux to the overlying water column. At times, sediments may also take up nutrients from the water column, but net flux is generally outward to the water column (Billen, 1978; Callender & Hammond, 1982; Hopkinson, 1987). Benthic nutrient fluxes can be an important source of limiting nutrients for planktonic production
(Fisher et al., 1982). While nitrogen is often the limiting nutrient for marine systems, P-
11 limitation and co-limitations for P and N are common in estuaries (Flemer, Livingston, &
McGlynn, 1998). Murrell et al. (2002) found evidence of phosphorous limitations in
Pensacola Bay during summer months when productivity is high and allochthonous nutrient supply is low.
Several methods exist for estimating benthic nutrient fluxes. Rowe, Clifford,
Smith, and Hamilton (1975) first measured nutrient regeneration with benthic chambers and this technique has been used in a wide variety of marine environments from shallow intertidal zones (Asmus, 1986) to deep ocean depths of more than 6000 m (Smith, Laver,
& Brow, 1983). Placement of in situ diffusive samplers, modeling porewater profiles, sediment-core incubations, and the placement of benthic chambers on the sediment surface have been used in various investigations. Determining porewater profiles assumes solute transport only by molecular diffusion. Sediment-core incubations involve disruption of the natural gradients as previously noted. Diffusive samplers require long incubations and are limited by membrane type and the time to reach equilibrium
(Nicholson et al., 1999). Interception of nutrients by benthic microalgae must also be considered if the gross flux is desired. Net flux is more easily obtained. An initial nutrient sample should be taken to allow for the determination of whether the actual benthic chamber placement had any effect on the solute concentration. Rowe and Phoel (1992) calculated net nutrient flux rates according to the formula:
12
FLUX = [(Ci – Cx) * V]/(H * A) (3) where:
FLUX = benthic nutrient flux in μmol · m-2 · hr-1
Ci = initial concentration
Cx = final concentration
V = volume under the benthic chamber
H = incubation time in hours
A = area under the benthic chamber.
Benthic Chamber Technique
In order for benthic chamber experiments to properly estimate benthic primary production, one must assume that each sample taken is from homogeneous, well-mixed overlying chamber water. The stirring rate inside the chamber should mimic the ambient near bottom water flow to avoid build up of concentration gradients that would inhibit diffusive fluxes. However, Berelson et al. (1987) found that varying the chamber stirring rate did not affect nutrient fluxes. Forja and Gomez-Parra (1998) found a homogeneous nutrient distribution inside the benthic chamber if the stirring device of the chamber accurately reproduced natural current. However, Hüettel and Gust (1992) showed that excessive stirring in core samples increased advective porewater flushing.
In situ benthic chambers have been judged to give the most accurate estimate of fluxes, because chambers include the effects of diffusion and incorporate the exchange across the BBL created by bioperturbation. Bioturbation effects may include an increase
13 in the surface area of sediments, sediment mixing, biodiffusion, bioirrigation, horizontal and vertical redox gradients and even spherical gradients (Santschi, Hohener, Benoit, &
Buchholtz-ten Brink, 1990).
There is no uniformity to the design and construction of benthic chambers (Forja
& Gomez-Parra, 1998), and a wide variety of chamber features and shape (cylindrical, hemispherical, square box, semi-ellipsoid) and size have been employed. Material of construction (e.g., glass, acrylic) and degree of transparency have also been variable, as well as methods of agitation and sampling. There are also differences in methods of calculating fluxes from the measured concentration changes over time.
Diver-sampled chambers are the most popular due to cost effectiveness and data quality. Fully automated designs are more readily implemented across a wide range of conditions, but for shallow waters where small-scale heterogeneity is suspected, the replicated diver-sampled chambers may be more accurate and efficient. Diver deployment of simple benthic chambers can be a major disadvantage when considering weather, water depth, water temperature, wind speed, current velocity, water visibility, bottom time limitations, and risk to personnel. Benthic chambers cannot be used for sediments with high relief, or in conditions of strong currents.
Nicholson et al. (1999) tested the effect of sophistication of chamber design on the measurement of benthic nutrient fluxes in shallow (< 30 m) marine systems. The simple benthic chamber was much more practical for shallow water than the automated design. Advantages exist for the high priced, sophisticated chambers, such as remote deployment eliminating diver placement and retrieval of domes, and continuous automated sampling. Disadvantages associated with remote deployment include lack of
14
knowledge of how well the chamber is settled on the bottom and no accounting for
macroorganisms or burrows covered by the system.
In benthic chambers, net oxygen fluxes over the sediment-water interface are
measured and there is no distinction between gross production, consumption, and
transport of oxygen during incubation in the light chambers (Hofman & de Jong, 1993).
Extreme changes in DO within the chambers during incubation can affect the processes
being measured (Hopkinson, 1978). Metal should be avoided in construction because of
the potential for it to act as a cathode when brought in contact with reduced sediments
resulting in O2 loss (Cramer, 1989).
Measuring Phytoplankton Production
The O2 exchange method has not been widely used in recent years for phytoplankton primary production in favor of radioisotope methods (Kinney & Roman,
1998). The O2 exchange method measures net community production in light bottles and
with the addition of dark bottles, gross production rates can be estimated with the
assumption that respiration is equal in both light and dark bottles. The 14C-bicarbonate
tracer method is much more routinely employed, although Underwood and Kromkamp
(1999) point out uncertainties as to whether this method is actually measuring GPP or
NPP or something in between. Although the 14C method of determining production
values is more precise and sensitive than the Winkler oxygen titration method, the 14C
method is unable to measure respiration entirely. In applying the tracer technique to
benthic production Revsbech, Horgensen, and Brix (1981) acknowledge a major
15
difficulty in measuring the total inorganic carbon in the water at the sediment-water
interface, which is necessary to calculate the specific activity of CO2 available for uptake.
Microalgae in Biofilms
Benthic microalgae on hard surfaces are referred to as periphyton, biofilms, or aufwuchs (Wainright, 1990). Biofilms concentrate and integrate organic and inorganic constituents of the surroundings in which they develop (Jeffrey & Paul, 1986; Snyder,
Lewis, Nocker, & Lepo, 2005; Zoell, 1943). Researchers and managers are constantly searching for new and improved methods to assess water quality and ultimately predict changing conditions in the marine environment. Biofilms grown on artificial substrates have the potential to be stress indicators in marine ecosystems since they meet certain criteria (e.g., cost effectiveness and sensitivity) important in choosing and developing new methods (Mattila & Räisänen, 1998).
Microalgae are constantly mixed and transferred into the water column from the benthic boundary layer mainly through advection and bioperturbation, especially in shallow waters. These suspended microalgal can colonize surfaces in the estuary, including artificial substrates. In shallow waters of subtropical and temperate estuaries during the summer index period, temperature and light are not usually limiting and nutrient availability at the benthos often controls production and growth. The growth of microalgae on artificial substrates would indicate nutrient limitation or eutrophication from nutrients available in the water column. Artificial substrates deployed near the benthos have the potential to incorporate both water column nutrients and nutrients
16 released from the sediments. Under similar light conditions experienced by benthic algae, periphytometers could potentially reflect benthic processes.
Several experimental media have been used as artificial substrates for biofilm growth including microscope cover slips, glass slides, acrylic plates, and glass fiber
(GF/C) filters (Mattila & Räisänen, 1998). Although incubation lengths have no uniformity in the literature varying from days to weeks, previous investigations have found that seven-day incubations in a subtropical estuary maximized accumulation of microalgal biomass while limiting calcareous invertebrates (e.g., barnacles. spirorbids, bryozoans; Moss, Nocker, Lepo, & Snyder, in press; Nocker, Lepo, & Snyder, 2004;
Wagner, 2006).
Enzymatic Activity Assays
Many microbial biogeochemical processes can be monitored by enzymatic activity. Processes involved in sediment nitrogen fluxes, such as net ammonification
+ + - (NH4 release from organic matter), nitrification (oxidation of NH4 to NO3 ), nitrogen
- fixation (microbial reduction of N2 to NH3), and denitrification (reduction of NO3 to N2O and N2) can indicate the magnitude of N cycling activity, and bioavailability of N species.
Biological nitrogen fixation is confined to specialized groups of prokaryotes, both auto- and heterotrophs, which possess the enzyme nitrogenase. In unvegetated shallow coastal sediments where light is not limiting, dense populations of benthic N-fixing cyanobacteria may develop and contribute fixed N to the ecosystem (Herbert, 1999). N- fixation rates can be estimated by the acetylene reduction assay (ARA) because
17
nitrogenase catalyzes reduction of both the N≡N of N2 and the C≡C of acetylene (C2H2;
Burris, 1972). Acetylene is stoichiometrically reduced to ethylene, which can be
quantified by gas chromatography with a flame-ionizing detector.
N-fixation rates are historically low for unvegetated marine sediments ranging
from 0.002-0.65 g N · m-2 · yr-1 with the highest of these rates deriving from organically rich sediments, such as in Waccasassa Bay, a subtropical estuary in central Florida
(Brooks, Brezonik, Putman, & Keirn, 1971). Metabolizable carbon substrate is required as a source of reducing power to support heterotrophic N2 fixation and is the likely
limiting factor for heterotrophic N-fixation in unvegetated sediments (Herbert, 1999).
Another cause of low N-fixation rates is high nitrogen bioavailability in the water column
+ - and sediments. Biologically available N, such as NH4 or NO3 , inhibits both activity and
biosynthesis of nitrogenase. Because N2 fixation requires high amounts of ATP, the direct
assimilation of fixed N is energetically advantageous to the microbes (Herbert).
Denitrification is a reductive process, mediated by heterotrophic bacteria utilizing
- NO3 as a terminal electron acceptor in respiration, reducing it to either N2O or N2.
Denitrification decreases the amount of bioavailable N to primary producers by releasing
these gaseous endproducts to the atmosphere. Under eutrophic environmental conditions
with low O2 concentrations, denitrification provides a mechanism for the removal of N
from the system thereby reducing the rate of eutrophication. However, low DO can also
- lead to a reduction in nitrification which often represents a significant source of NO3 for
denitrifiers (Joye & Hollibaugh, 1995). Herbert (1999) notes that subtidal denitrification
rates are subject to significant change on a diel basis due to growth of benthic microalgae.
O2 produced during photosynthesis can diffuse into the surface sediments, lowering the
18
RDL and inhibiting dissimilatory nitrate reduction which is the first step in
denitrification.
Denitrification is commonly measured by the acetylene inhibition technique, or
acetylene block assay (ABA). This assay is based on the inhibition of nitrogen oxide
reductase by acetylene (Balderston, Sherr, & Payne, 1976). The major advantages of
ABA are simplicity, sensitivity, and low expense. The most notable drawback of ABA is
- that at the low NO3 conditions often found in marine sediments (< 10 μM) inhibition of
15 15 N2O formation is incomplete. A more accurate technique, the N-tracer technique, or N isotope pairing method, not only determines denitrification rates with no inhibitor required but simultaneously measures nitrification, nitrate ammonification, and N-
- + mineralization rates in a single experiment if NO3 and NH4 fluxes are known (Nielsen,
1992).
Alkaline phosphatase activity (APA) in marine organisms was examined by
Kuenzler (1965) and Perry (1972). The phosphatase assay uses a fluorogenic artificial
substrate, difluoromethylumbelliferyl phosphate (DiFMUP), to determine potential
catalytic capabilities. The substrate is hydrolyzed by phosphatase to produce
difluoromethylumbelliferone (DifMUF), which fluoresces blue under UV excitation.
APA is assayed by adding DiFMUP at saturating concentrations to samples and
measuring fluorescence accumulation over time. The rate of change in fluorescence,
when calibrated against standards of the pure DiFMUF fluorophore, yields a measure of
the phosphatase activity.
Phosphatase activity assays can aid in the assessing the state of phosphorous
(phosphate) bioavailability and the utilization of dissolved organic phosphorous (DOP) as
19 a source of P for bacteria and primary producers. High APA is an indication of P- limitation. High external orthophosphate concentrations result in competitive inhibition of phosphatase, so APA will be suppressed (Hernandez, Niell, & Whitton, 2002).
However, at constant high P concentrations of > 300 μg/L (9.6 μM) for several days,
Hernandez et al. (2002) found that APA may adapt and increase.
General heterotrophic activity of microbes has been commonly estimated by a variety of techniques, including the fluorescein diacetate (FDA) assay. FDA is a non- polar esterified compound that can be hydrolyzed by nonspecific intra- and extra cellular esterases (Guilbault & Kramer, 1964) and thus may indicate general heterotrophic activity of bacteria and eukaryotic cells. Its ability to assay extracellular enzyme activity makes it useful for determining the activity of biofilms, where extracellular enzymes are found throughout the polymer matrix. FDA diffuses freely into intact cells where esterases hydrolyze it to the more polar fluorescein. Accumulation of fluorescein is a function of membrane potential and intracellular activity, therefore rendering the FDA turnover a useful indicator of cell viability and activity (Battin, 1997). FDA is hydrolyzed by enzymes from bacteria, fungi, rotifers, and algae and for that reason is a potential indicator for total esterase activity of biofilms (Battin) and sediments (Battin; Meyer-
Reil, 1990). The FDA assay is sensitive enough to detect spatial patterns of biofilm hydrolytic activity and identify active biological zones in marine sediments (Battin).
Fluorescein is a widely used fluorogenic compound with excitation at a 488 nm wavelength, a high extinction coefficient, and a high quantum yield.
20
Study Area
Pensacola Bay system, at 30.4º N latitude, is a subtropical (falling between 23.5 –
40º N/S latitude), river-dominated estuary located in northwest Florida along the northeastern Gulf of Mexico coast (Figure 1). Pensacola Bay is a moderately sized (373 m2), drowned river valley estuarine system (Schroeder & Wiseman, 1999) consisting of 5 interconnected river-estuarine basins and 1 lagoonal sound: Blackwater River and Bay,
Yellow River, East River and Bay, Escambia River and Bay, Pensacola Bay, and Santa
Rosa Sound. Each individual basin is characterized by a relatively shallow shelf, a slope area, and a coastal plain similar to the adjacent systems, Perdido and Choctawhatchee
Bays (Livingston, 2003). Access to the Gulf of Mexico from Pensacola Bay is through a narrow inlet (926 m) at the southwestern terminus of the bay. Pensacola Bay salinity ranges from mesohaline (10 – 20) to nearly oceanic (> 30) in the deeper areas. Diurnal tides occur with a microtidal range of 0.5 – 1.0 m (Livingston, 1999).
The average water residence time for the entire Pensacola Bay system is about 25 days (Solis & Powell, 1999). River flow, wind and tides, and the intrusion of higher salinity Gulf of Mexico waters contribute to periodic stratification of the water column within Pensacola Bay (Livingston, 1999). Flushing of the system during spring floods could have an adverse effect on the benthos (Livingston). The drainage basin for the 3 primary rivers (Escambia, Blackwater, and Yellow Rivers) covers about 17,500 km2 extending well into Alabama. Land use is dominated by agriculture and silviculture
(Livingston). The mean depth for Pensacola Bay system is about 3 m (Olinger et al.,
1975).
21
Butcherpen Cove (Figure 1) is a shallow embayment along the southern shore of
Pensacola Bay with no direct freshwater inflow. Pensacola Bay at this location exchanges water with Escambia and East Bay. This study was located in < 1.5 m depth at
Butcherpen Cove of Pensacola Bay, FL near the coordinates 30.3703 ºN and 87.1428 ºW.
Butcherpen Cove is located directly north of Gulf Islands National Seashore (GINS)
Naval Live Oaks area, Gulf Breeze, FL. The cove sediments are dominated by coarse grained sand (quartzite) in series of troughs and bars with a wavelength of approximately
30 m.
22
University of West Florida
Rain Gauge
Butcherpen Cove
Figure 1. Location of Butcherpen Cove within the Pensacola Bay estuary in northwest
Florida along the northern Gulf of Mexico coast; Approximate location of National
Weather Service precipitation gauge and The University of West Florida. 1From
“Evidence that phosphorus limits phytoplankton growth in a Gulf of Mexico estuary:
Pensacola Bay, Florida, USA,” by M. C. Murrell, R. S. Stanley, E. M. Lores, G. T.
DiDonato, L. M. Smith, and D. A. Flemer, 2002, Bulletin of Marine Science, 70(1), p.
157. Adapted with permission of the author.
CHAPTER II
OBJECTIVES & HYPOTHESIS
The primary objective of this study was to document benthic microalgal productivity and biogeochemical fluxes across the sediment-water interface. A secondary objective was to compare microphytobenthos dynamics with autotrophic activity in the plankton and in biofilms on artificial substrates. The specific hypotheses tested during this study included:
1. The dynamics of benthic microalgal production are the same as that of
phytoplankton over the seasonal change from winter through summer.
2. The dynamics of benthic microalgal production are the same as that of
periphyton over the seasonal change from winter through summer.
3. The dynamics of phytoplankton production are the same as that of
periphyton over the seasonal change from winter through summer.
4. Benthic microalgal production is controlled by benthic flux.
5. Benthic microalgal production is controlled by water column chemistry.
6. Periphytometer responses are proportionally representative of benthic
processes.
7. P:R ratios, P:B ratios, and compensation points for microphytobenthos and
periphyton grown on artificial substrates are the same. 23 CHAPTER III
MATERIALS AND METHODS
Site Characteristics
Selection of the study site was based on accessibility, convenience, favorable dominant wind direction, lack of nearby freshwater input, and the lack of a need for boat operations. A 5 m by 10 m grid was established in a Butcherpen Cove trough between bars. Chambers were placed within the grid using a random number generator (Zar, 1996) leaving at least 1 m distance from the adjacent bar to eliminate edge effects.
Physicochemical Determinations
A HydroLab® DataSonde 4 was programmed to take water quality measures
(temperature, conductivity/salinity, turbidity, pH, % oxygen saturation, and DO) every 15 min throughout the week long incubation. The DataSonde 4 was secured in a 10.2 cm
PVC cylinder 10 cm above the bottom with holes (3.8 cm) for water flow throughout the cylinder. A handheld YSI® Model 85 (San Diego, CA) was used to record bottom water temperature, salinity, and DO at the beginning, middle, and end of incubations.
Incubations were started before solar zenith and ended before afternoon thunderstorms which are common during the summer months. Incubation times ranged from 4 hr
(summer months) to 7 hr (winter and spring months; Table 1). Light readings (μE · m-2 · 24 25
s-1) were taken every hour during the day with a Li-Cor LI-190SA radiation sensor at the
seasurface and seafloor allowing calculation of % PAR transmission.
Chamber Procedures
Light and dark benthic chambers, 3 of each, were constructed from clear acrylic
domes of 10.25 in. radius (0.26035 m), 7.5 in. height (0.1905 m), and 0.027 m3 volume
(27.044 L) covering an area of 0.213 m2 (Figure 2A and 2C). Dark chambers were covered with 6 mil thick black plastic sheeting (Film-Gard®) to block light (Figure 2A).
Stirring devices were assembled with hemispherical plastic cups transferring external
current to internal acrylic stirring paddles (Figure 2C). Domes were fitted with two
barbed hose fittings and silicone tubing (3 mm ID; 6 mm OD) to allow sample water to
be removed by a 60 mL syringe (B-D® Luer-Lok™ syringe) and to allow make-up water
to enter chambers. Make-up water dilution of chamber water was assumed to be
negligible. Dome water samples were drawn to 60 mL, then inverted underwater, and
excess water and any air bubbles were removed by adjusting the volume to 50 mL.
Syringes were then sealed with B-D® Luer-Lok™ syringe tip caps. Chamber samples
were collected in the same order for each sampling event to account for sequential
sampling time delays. Actual sampling times were recorded for each syringe.
Chambers were carefully placed on the benthos to avoid disturbance of the
sediment-water interface. Macrophytes, large polychaete burrows, tunicates, and hard
substrates were avoided. The chambers were secured to the bottom by piling surrounding
sand over each dome’s 2 in. (5.08 cm) external flange. This technique supported an
26
assumption that negligible water exchange occurred between inside and outside of the
domes during incubations. Chambers were allowed to stabilize for 10-15 min before
initial (TO) samples are gathered. Time zero (TO) dome samples were taken for dissolved
oxygen (2; DO), total nutrients (1; TN), and dissolved nutrients (1; DN). Coded sample
syringes were secured to a dive slate with rubber straps. After a 4 to 7 hr incubation
period, Time X (TX) benthic chamber samples were extracted in the same manner as TO
samples.
A B
C
Figure 2. (A) A dark benthic chamber with attached tubing for syringe sampling and current simulation device. (B) A benthic periphytometer containing acrylic sampling plates and glass slides. Image is from April 15, 2005, showing attached freshwater macroalgae. (C) A clear benthic chamber and greenhouse shade cloth. Internal stirring paddles are visible in this image.
27
Biofilm Analysis
Biofilms were grown on glass microscope slides (38.7 cm2) and ⅛ in. (0.32 cm) thick acrylic plates (360 cm2) fitted into acrylic racks lashed to a PVC frame anchored in concrete (Figure 2B). Slides and plates were deployed and incubated 10 cm above the benthos at Butcherpen Cove for the seven-days prior to benthic chamber experiments.
During the chamber stabilization period, biofilms slides were retrieved and placed in 50 mL glass shell vials underwater to avoid air bubbles. Three vials were left clear and three were covered with aluminum foil for a dark treatment. Vials were secured to a periphytometer with zip-ties for the incubation period. Additional biofilm substrates
(plates and slides) were transported on ice to the Wetlands Research Laboratory at the
University of West Florida (UWF) in acrylic racks draped with wet paper towels.
Material scraped from artificial substrates was used for determining enzyme activities
(see Chapter III, Section H), quantities of photopigments (see Chapter III, Section I), carbon and nitrogen content and ratio (C:N), total phosphorus (TP), and AFDW (%OM) in replicate analyses. Material for enzyme assays was transferred to serum bottles.
Material for elemental analyses was collected on GF/F filters with 10-15 psi of vacuum pressure. GF/F filters were stored in a Petri dish (C:N, TP, AFDW) or a foil envelope
(Chl a) at -80 ºC until analysis.
Water Column Analysis
Initial (TO) ambient water column samples were collected at the site by filling an amber 1 L polypropylene bottle with near bottom water. From this 1 L bottle, samples for
28
total nutrient analysis (TKN, TP) were collected by pouring unfiltered water into
prelabeled 125 mL polypropylene bottles containing 0.4 mL of concentrated H2SO4.
Samples for dissolved nutrient (NH3, NO3, NO2, OP, Si) analysis were filtered through 27
mm ashed (500 ºC for 1 hr) Whatman® glass fiber filters (GF/F) held in Swinnex® filter holders. Filtrate was collected in prelabeled 40 mL polypropylene bottles. Three replicate photopigment samples were filtered from the 1 L bottle onto 27 mm ashed GF/F filters and the volume filtered recorded. Filters were folded over, blotted dry, and stored in labeled foil envelopes. Samples for nutrients and chlorophyll analysis were kept on ice in the field and held in a lab freezer (-80 ºC) until analysis. Water column production and respiration were determined in 300 mL BOD bottles that were filled underwater with near bottom site water. Water was allowed to circulate within BOD bottles for about 30 s.
Bottles were then capped off underwater and incubated in a PVC BOD bottle rack in situ.
Replicate light and dark BOD bottles were incubated in parallel with benthic chambers.
BOD racks were incubated on the bottom with line and buoy tethered them. Once
incubations were complete, 3 TX light and 3 TX dark BOD bottles were collected.
Sediment Procedures
Sediment samples were randomly collected from within a 1 m by 1 m square grid
placed adjacent to the benthic chamber grid. Sediment sample locations within the grid
were determined using a random number generator (Zar, 1996). Three replicate cores
were taken for each sediment analysis (C:N, AFDW, Chl a, Chl c, Pheo a). A cut off 15
mL Falcon™ (Fisher®) centrifuge tube (1.3 cm diameter) was inserted to ~2 cm depth and
29
using two gloved fingers as stoppers a sediment sample was transferred to a prelabeled 15
mL centrifuge tube. Sediment samples were stored on ice while in the field and during
transport to UWF. In the lab, samples were centrifuged in an International Equipment
Co.® IEC Clinical Centrifuge Model CL on speed 4 for 5 min. The supernatant was
removed by aspiration and tubes were stored at -80 ºC until analysis.
Oxygen Determinations
Dissolved oxygen concentrations were determined by Winkler titration of water samples (Eaton, Clesceri, Rice, & Greenberg, 2005). 300 mL samples were fixed in the field with sequential addition of 2 mL each of MnSO4 (36.4%) and Alkaline Iodide Azide
(50% sodium hydroxide, 15% potassium iodide, 1% sodium azide; LabChem Inc®). The
resulting precipitate was allowed to settle for at least 15 min and samples were remixed
by inversion, then stored on ice until return to the lab. Syringes and vials containing 50
mL samples were fixed with 1 mL of MnSO4 and Alkaline Iodide Azide by inserting
blunt ended B-D® 16G 1½ PrecisionGlide® needles attached to a 20 mL B-D® Luer-
Lok™ syringe containing each reagent. Syringes were re-capped, mixed and stored on ice
until return to the lab. Samples removed from domes were fixed in the field within 15
min of sampling. In the lab, BOD bottles, syringes, and shell vials were acidified (2 mL
for 300 mL samples, 1 mL for 50 mL samples) with concentrated H2SO4, mixed well to
dissolve the flocculent, and titrated with a 0.0125N sodium thiosulfate (Na2S2O3) solution
(LabChem Inc®). Samples received 0.5 mL of a starch indicator solution (< 1.0% acetic
30 acid, < 0.1% starch; Fisher®) as an endpoint marker once the sample turned a pale-straw color. Winkler analyses were completed within 12 hr of sample collection.
Biomass Determinations
Chlorophyll a
Biomass was quantified by total chlorophyll (Chl a) according to the methods of
Dandonneau and Neveux (2002). For Chl a analysis, sediment samples or filters were extracted in 8-10 mL of 90% acetone in centrifuge tubes (15 mL). Ultrasonication (30 s) occurred under reduced light and samples were then extracted overnight at -20 °C.
Samples were then centrifuged in an International Equipment Co.® IEC Clinical
Centrifuge Model CL on speed 4 for 5 min and supernatant was pippetted into 2 mL microfuge tubes. Extracts were measured for their fluorescence on a Perkin Elmer LS45 luminescence spectrometer with Xenon lamp. Each scan consisted of 16 4 nm steps, beginning at excitation wavelengths of 406 nm, and emission (fluorescence) wavelengths for detection of Chl a were between 666 and 668 nm, Chl c at 630 nm, and Pheo a from
646 to 656. Pheophytin a was quantified by acidifying the stock Chl a (Sigma®) standard according to the methods of Welschmeyer (1994). Chl c was quantified using standard material extracted from the Cryptophyceae (DHI Water and Environment®). Pigment concentrations in acetone extracts were converted back to sediment area, volume filtered, or biofilm area sampled.
31
Biofilm Biomass by Optical Density
Optical density of biofilms on artificial substrates was determined by digitizing
air-dried biofilm plates on an Epson 636 scanner with an Epson EU-14 transparency unit
at a resolution of 200 dpi. Digital images were analyzed with the National Institutes of
Health image analysis software (NIH Image) to obtain the average and standard deviation
of pixel densities on a gray scale of 1 – 256.
Organic Content
Total organic content of sediment samples and biofilm filters was determined
using standard method 10300C for ash-free dry weight (AFDW; Eaton et al., 2005).
Samples were oven dried (105 °C) for 24 hr and the weight was recorded. Samples were
then combusted in a muffle furnace (500 °C) for 4 hr. Samples were then re-wet with
reagent grade H2O and dried at 105 °C for another 24 hr and weighed. The difference between weights, representing the material lost on ignition, was recorded as the organic content per area (biofilms) or dry weight (sediments).
Elemental Analysis
Organic content of sediments and biofilm material on filters was also measured by
elemental analysis of C and N content. Samples were oven dried (105 °C) for 24 hr in
aluminum weigh dishes. Sediment samples were homogenized with mortar and pestle
after drying and stored in 20 mL glass vials in the dessicator until analysis. Filters were
transferred directly to the dessicator. Subsamples (150 mg) of homogenized sediment and
whole biofilm filters were analyzed on a Thermo Finnigan® Flash EA 1112 Series
32
elemental analyzer. Acetanilide (Microlab®) was used as a standard. Aspartic acid
(Thermoquest®), acetanilide, and a soil standard (Thermo Electron Corporation® #0704)
were run as check standards every 10 samples.
Compensation Point Determinations
A 3-day sampling event, June 15-17, 2005, generated data to establish an
irradiance curve for the intact benthos and biofilms. Seven domes were deployed and
immediately covered with commercial shade cloth (Greenhouse Mega Store®) of varying density (30, 40, 50, 60, 70, 80, and 90% shade), along with black (100%) and clear
(< 1%) domes. The shade cloth was suspended above the domes on a PVC frame (1 m by
1 m square) with 0.75 m legs (Figure 1C). Each leg was anchored in the sand. Actual irradiance intensity (μE · m-2 · s-1) for each treatment was recorded by Li-Cor light meter as the independent variable. Photosynthetic efficiency (PE) was obtained by dividing measured net production by biomass (mg C · mg Chl a-1 · m-2 · hr-1) as the response
variable. After a brief acclimation period, duplicate T0 samples were collected for each
dome and fixed with Winkler reagents for DO analysis. Following an approximate 4 hr
incubation (10:00 – 14:00), TX samples were collected and fixed.
Nutrient Analysis
-3 + - - All samples for nutrient analysis (TP, TKN, SiO2, PO4 , NH4 , NO3 + NO2 )
were stored at -20 ºC until analysis by standard methods (US EPA, 1993; US EPA,
-3 + - - 1983). Detection limits for TP, TKN, SiO2, PO4 , NH4 , NO3 , and NO2 were 1 µM, 4
33
µM, 3 µM, 0.064 µM, 0.357 µM, 0.036 µM, and 0.036 µM respectively. When the
nutrient was non-detectable the detection limit was reported. Samples for total nutrients
(TP, TKN) were acid digested in a fume hood on a 50-place AI Scientific® AIM500
Programmable Digestion System with a BD50 Microprocessor Control. 20 mL of
preserved sample or 2 g of homogenized sediment was placed into 100 mL borosilicate
glass tubes for digestion at 350 ºC for 3.5 hr.
A 2-channel Bran-Luebbe® AutoAnalyzer 3 with Compact Sampler, multitest manifolds and a high sensitivity digital spectrophotometer (Bran-Luebbe® Digital
Colorimeter) was used to analyze samples. National Institute of Standards and
Technology (NIST) traceable solutions were used in the preparation of all quality control
samples and standards. For each analytical batch, one standard source was used for the
calibration standards (either LabChem® or Ricca®) and a different source was used for
calibration verification (either LabChem® or Ricca®). Matrix spikes were made from
either source, purchased at a concentration of 1000 mg/L and diluted to the required
analytical concentrations. Matrix spikes were run every 10 samples to check the recovery
of known amounts of silica, nitrogen and phosphorous. Benthic nutrient fluxes were
determined from analysis of nutrient autoanalyzer results (Equation 3).
Enzyme Assays
Activity assays were performed with biofilm slurries obtained by scraping the
periphyton covered plates with a rubber-tip squeegee into a polypropylene weigh boat.
After rinsing the plate and squeegee with 20 mL of filtered (GF/F filter at 10-15 psi) site
34
water, biofilm slurries were transferred to 140 mL labeled serum bottles using a 1½ in.
(3.81 cm), 16-gauge needle attached to a 60 mL syringe. Serum bottles were then sealed
with sterile crimp cap stoppers. Time recorded incubations (0-30 min) allowed for
enzyme activity rates to be determined. Activity rates were calculated based on biofilm
area (m-2) sampled and time (hr-1). All labeled serum bottles were incubated under low light conditions and kept on New Brunswick Scientific Co., Inc. Series 25 Shaker
(Edison, NJ) on low speed at room temperature.
Heterotrophic activity as FDA esterase activity of biofilm slurries was measured as fluorescein product formation. Fluorescein was quantified by absorbance at 490 nm using a ThermoSpectronic® Genesys20 spectrometer following methods derived from
Battin (1997). Due to light sensitivity of fluorescein, all serum bottles were wrapped in foil. A standard solution of 1.0 M fluorescein (Sigma Ltd.®, FW 376.3) was prepared by
dissolving 37.6 mg of fluorescein in 80 mL of acetone:water (1:1). This solution was
diluted 1:5 with water to prepare the working stock (200 mM). A 1:10 dilution of the
working stock was then serially diluted to determine a standard curve of absorbance
versus concentration. An FDA stock solution (2 mM) was prepared in the same manner
as fluorescein with 83.3 mg of FDA in acetone:water. The assay was initiated with the
addition of 4 mL of FDA stock solution to test suspensions. Biofilm slurry samples (1.5
mL) were removed with a glass syringe (5 cc) and filtered through a mixed cellulose ester
(MCE) membrane disposable syringe filters (0.22 µm) into a silica glass cuvette (1 cm2)
at 15, 30 and 45 min for quantification of fluorescein. Culture fluid from Bacillus cereus
grown on 1:10 tryptic soy agar (TSA) medium for more than 3 days and less than 1 week
was used for a positive control. 39 mL of this culture broth was added to a serum bottle
35
and 1.5 mL was removed for each sample. A negative control consisted of Bacillus
cereus broth (37 mL) acid-killed with 2 mL of concentrated H2SO4.
Alkaline phosphatase activity (APA) was determined by adding 10 µM DiFMUP
to biofilm slurries, and to positive and negative controls. The fluorescent product was
quantified using a Perkin Elmer® LS45 luminescence spectrometer following the methods of Perry (1972). Samples were excited at a 365 nm wavelength and fluorescent emissions were read at 450-460 nm. As with the FDA assay, this assay was light sensitive, so all containers were wrapped in foil. 3 mL of biofilm slurry was pipetted into 2 microfuge tubes and centrifuged (Eppendorf® Centrifuge 5417C) for 2 min at 10,000 rpm. The
supernatant was poured into a quartz cuvette (1 cm2) and analyzed on the
spectrofluorometer as the sample blank. Substrate was then added to the serum bottle
followed by vigorous shaking. 3 mL of this mixture was centrifuged as before and
analyzed as the TO sample. This process was repeated every 15 min for 1 hr. For positive
controls, 3 mL of culture fluid from Bacillus cereus grown on 1:10 TSA medium for more than 3 days and less than 1 week was analyzed as above. Negative controls consisted of Bacillus cereus broths (20 mL) were acid-killed with 1 mL of concentrated
H2SO4, and 3 mL samples were run in parallel with biofilms and positive controls.
An anaerobic environment was created for acetylene block assays (ABA) by
degassing sample bottles with argon in a COY® Laboratory Products, Inc., Model AAL
(Automatic Air Lock; Grass Lake, MI) hood equipped with arm sleeves. Containers were sealed with sterile serum stoppers under anaerobic conditions. Nutrients (1.5 mg each) were added to the ABA bottles in the form of solid KNO3 and glucose prior to sealing.
® N2O gas accumulation was measured with a Hewlett Packard 5890 Series II gas
36 chromatograph equipped with an electron-capture detector according to the methods of
Balderston et al. (1976). N2O standards were prepared in a helium environment by placing 4 glass beads inside a 2000 mL round bottom flask (RBF) sealed with a large stopper and silicon cement. 2 mL of pure N2O was injected into the flask to prepare a
1:1000 dilution at approximately 1 ATM. A 5-point standard curve (peak height vs. N2O concentration) was generated by injecting subsequent volumes (1.0, 0.5, 0.25, and 0.1 mL) of standard into the GC with a 1 mL tuberculin B-D® syringe with a 27 gauge needle. ABA bottles were injected with acetylene (20 mL) by syringe to approximately
10% of serum bottle volume to initiate assays. Headspace samples (1 mL) were removed from sample bottles over time (typically at 1, 4, and 24 hr). The first 0.5 mL of each sample was injected into the air, and the remaining 0.5 mL of sample was injected into the GC. Time intervals were adjusted based on initial activity. Paracoccus denitrificans
(ATTC® Number 13543) grown on 1:10 TSA medium for more than 3 days and less than one week provided culture broth for positive controls. Negative controls (Paracoccus denitrificans) were killed with the addition of 1 mL of concentrated H2SO4.
For N-fixation acetylene reduction assays (ARA), C2H4 gas was measured from headspace samples of biofilm slurry incubations by a Shimadzu® GC-17A gas chromatograph equipped with a flame-ionizing detector following the methods of Burris
(1972). Standards were prepared in the same manner as ABA stocks with 2 mL of C2H4 gas. A 5-point standard curve (peak height vs. C2H4 concentration) was generated by injecting subsequent volumes (1.0, 0.5, 0.25, and 0.1 mL) of standard into the GC. ARA bottles were injected with acetylene to approximately 10% of serum bottle volume to initiate assays. Headspace samples (1 mL) were removed from sample bottles over time
37
(typically at 1, 4, and 24 hr). The first 0.5 mL of each sample was injected into the air,
and the remaining 0.5 mL of sample was injected into the GC. Time intervals were
adjusted based on initial activity. Azotobacter vinelandii (ATTC® Number 12837) grown
on Burk’s N-free medium for more than 3 days and less than 1 week provided culture
broth that was used for positive N-fixation controls. Negative controls consisted of
Azotobacter vinelandii broth that was acid-killed with 1 mL of concentrated H2SO4.
Statistical Analysis
The multivariate data set was analyzed for normality (SPSS software v. 13.0,
SPSS Inc.®) and transformed if necessary either logarithmically (log X +1) or by taking
the natural logarithm. Levene’s homogeneity of variance tests revealed differences
between sample populations over the course of the study and transformations were
unsuccessful. Pearson nonparametric correlations were performed on all normalized
variables. Confidence intervals (95%) were determined for nutrient flux data in order to
establish if a flux was significantly different from zero. Means, standard deviations, and
standard errors were calculated in MS® Excel.
CHAPTER IV
RESULTS
Ambient Physicochemical Conditions
Experiments were conducted at Butcherpen Cove once in February and March, three times in April and May, and twice each in June, July and August (Table 1).
Experimental days were chosen in response to storm events and for days with relatively clear skies. Prevailing southerly winds resulted in minimum wave action in Butcherpen
Cove during sampling events. DO concentrations of the ambient seawater remained above 7 mg/L during winter and spring sampling, and above 5.9 mg/L for all summer incubations (Table 1).
A record monthly maximum rainfall of 621 mm was recorded for April 2005 by the National Weather Service at Pensacola Regional Airport, which is approximately 10 km from the study site. Two other significant precipitation events occurred during the sampling season as a result of tropical storm (T. S.) Arlene passing west of Pensacola on
June 11, 2005, and Hurricane Dennis (category 4) passing east of Pensacola on July 10,
2005. T. S. Arlene dropped 119 mm of rain while Hurricane Dennis contributed 97 mm.
During the study period, 1661 mm fell on Pensacola, over 600 mm above the average of
1031 mm for the same time period. The greatest 24 hr rainfall total was 354 mm on April
1, 2005 (Figure 3).
38 39
Table 1
Sampling Dates, Incubation Times, and Ambient Dissolved Oxygen Concentrations of
Bottom Water at Butcherpen Cove
Experiment Sampling Incubation Ambient DO number date time (h) (mg/L)
1 18-Feb-05 6 9.0 2 18-Mar-05 7 9.5 3 15-Apr-05 7 7.7 4 22-Apr-05 7 8.5 5 6-May-05 7 11.4 6 13-May-05 6 8.2 7 20-May-05 6 7.8 8 8-Jun-05 5 7.2 9 24-Jun-05 5 7.4 10 8-Jul-05 4 7.1 11 29-Jul-05 4 6.4 12 17-Aug-05 4 6.2 13 24-Aug-05 4 8.0
Macroalgae (Enteromorpha spp. and Ectocarpus spp.) were found in Butcherpen
Cove in April 2005 and grew on periphytometers. It is not known if this was a consequence of the high rainfall and freshwater influx to the bay, or if these species are a normal part of the spring transition in Pensacola Bay system. The heavy rains of April
2005 increased turbidity in the water column reducing the available light at the benthos.
40
Percent transmission of photosynthetically active radiation for the 1 m water depth was
reduced from 85 to 14.6 from March 18 to April 15, 2005, but recovered within three
weeks to 49.5 on May 6, 2005, and averaged 55.1 ± 7.95 for the summer index period
(Figure 4). Quantum flux at the benthos ranged from 225 to 1700 µE · m-2 · s-1 for the
entire study period. Afternoon thunderstorms, which are normal for the summer period,
also reduced light availability. On April 22, 2005, turbidity was high and afternoon cloud
shading reduced surface intensity from 1450 to 390 µE · m-2 · s-1 at 16:00 hr.
2 0 0
1 5 0 ) m m (
l l 1 0 0 a f n i a R
5 0
0 F e b M a r A p r M a y J u n J u l A u g S e p
Figure 3. Total recorded rainfall (mm) in Pensacola, Florida, during the experimental season of 2005 (February through August).
Butcherpen Cove was a mesohaline habitat with bottom water salinity ranging
from 1.3 (April 15, 2005) to 21.1 (June 24, 2005) with an average (± 1 SE) of all
measurements taken of 11.63 ± 5.53 for the study period. The average change of bottom
41
water salinity between high and low tides throughout the study period was 2.5 ± 3.6.
From April through August average salinity differences between surface and bottom
waters were 1.3 ± 3.0. Salinity was greatly reduced due to the large rainfall events of
April from 18.3 on March 18, 2005, down to 1.3 on April 15, 2005 (Figure 4). Bottom
water temperatures increased from February to June and then were relatively stable
through August ranging from 16 ºC on February 18, 2005, to 31 ºC on August 24, 2005
(Figures 20 & 21). The average change of temperature between high and low tides
throughout the study period was 2.13 ± 0.8 ºC. From April through August the average
temperature differences between surface and bottom waters was 0.42 ± 0.57 ºC. The
highest temperatures were in surface waters while greatest salinity values were recorded
in the bottom waters.
25 100 Salinity % PAR 20 80 n o i s s i
15 60 m y t s i n n i a l r a T
S 10 40 R A P
% 5 20
0 0 Feb Mar Apr May Jun Jul Aug Sep
Figure 4. Percent transmission of PAR (% PAR) to the benthos and bottom water salinity measures during 2005 (February through August).
42
- - Bottom water dissolved NO3 + NO2 concentrations averaged 0.10 ± 0.03 µM for
winter sampling, spiked in April 2005 to 3.28 µM coincident with heavy rains, tapered
off in May 2005, and averaged 0.04 ± 0.02 µM for June through August 2005 (Figure 5).
- - Storm events after April did not significantly affect NO3 +NO2 concentrations. Dissolved
+ NH4 in the bottom water was negatively affected by rainfall events with highs just before major storms of 6.98 and 6.76 µM on March 18, 2005, and June 24, 2005, respectively and lows of 0.36 µM (the detection limit) on June 8, August 17 and 24, 2005
(Figure 6). N
8 O 3 -
6 + N
4 O 2 -
2 ( ) 1
0 ) - h
·
2 40 - m
·
l 20 o m µ ( 0 x u l F
- -20 2
O Dark Domes N -40 Light Domes + - 3 O N Feb Mar Apr May Jun Jul Aug Sep
- - -2 -1 Figure 5. Mean (± 1 SE) light/dark sediment fluxes of NO3 + NO2 (µmol · m · hr ) and
- - water column concentrations of NO3 + NO2 (µM) in 2005 (February through August).
43
10 N
8 H 4
6 +
( µ
4 M )
2 0
) 150 1 -
h Dark Domes
·
2 100 Light Domes - m
·
l 50 o m µ
( 0
x u l F
-50 + 4
H -100 N
Feb Mar Apr May Jun Jul Aug Sep
+ -2 -1 Figure 6. Mean (± 1 SE) light/dark sediment fluxes of NH4 (µmol · m · hr ) and water
+ column concentrations of NH4 (µM) in 2005 (February through August).
TKN in bottom water did not follow any definable patterns, averaging 19.3 ± 7.6
µM with a high of 32 µM on June 8 and April 8, 2005. Lowest values of TKN were 7.5
µM on June 24, 2005 (Figure 7). TP was detectable in bottom water on seven of 13 experiments with a median of 1.1 µM for the first three experiments and 1.6 µM for the last four experiments. A peak of 2.6 µM was seen on August 17, 2005. TP concentrations for the middle six experiments (April 22, 2005 – July 8, 2005) were undetectable (Figure
7).
44
50 4
W . C . TK N 40 W . C . TP 3 )
30 ) (
2 (
N P K T
T 20
1 10
0 0 Feb M ar A pr M ay Jun Jul A ug Sep
Figure 7. Mean (± 1 SE) water column concentrations of total kjeldahl nitrogen (µM) and total phosphorous (µM) in 2005 (February through August).
-3 Bottom water dissolved PO4 concentrations were only detectable on seven of 13
sampling days. The first three experiments averaged 0.13 ± 0.03 µM. Midsummer peaks
occurred on June 24, 2005, of 0.18 µM and 0.11 µM on July 8, 2005, and a late summer
increase was observed in the final two experiments of 0.13 ± 0.05 µM (Figure 8).
Near bottom water dissolved SiO2 averaged 66.2 ± 7.9 µM over the entire
sampling season with a consistently increasing seasonal trend. The low of 24.3 µM was
recorded for February 18, 2005, and the high of 137.1 µM was recorded on August 24,
2005 (Figure 9).
45
0.3 P O
0.2 4 3 -
(
0.1 M )
) 4 1 - 0.0 h
·
2 Dark Dom es -
m Light Domes
2 ·
l o m µ (
0 x u l F
- 3
4 -2 O P Feb M ar A pr M ay Jun Jul A ug Sep
-3 -2 -1 Figure 8. Mean (± 1 SE) light/dark sediment fluxes of PO4 (µmol · m · hr ) and water
-3 column concentrations of PO4 (µM) in 2005 (February through August).
120 S i O 2
80 ( M
40 )
0
) 1500 1 - D ark D om es h 1000
2 Light Domes - m
500 l o
m 0 (
x -500 u l F
2 -1000 O i
S -1500 Feb M ar A pr M ay Jun Jul A ug Sep
-2 -1 Figure 9. Mean (± 1 SE) light/dark sediment fluxes of SiO2 (µmol · m · hr ) and water column concentrations of SiO2 (µM) in 2005 (February through August).
46
Sediment, Water Column and Biofilm Biomass
Sediment total carbon (TC) ranged from 38.4 to 158.3 nmol C/cm2 and total nitrogen (TN) ranged from 2.5 to 15.8 nmol N/cm2 (Figure 11). Sediment values of TC remained relatively constant all year except for two experiments. On May 13, 2005, both
TC and TN content spiked to 158.3 ± 57.4 and 15.8 ± 4.2 nmol/cm2, respectively. On
August 24, 2005, the carbon and nitrogen values were again higher at 155.6 ±16.0 and
11.7 ± 6.4 nmol/cm2, respectively (Figure 10). Sediment total phosphorous (TP) was also fairly constant ranging from 0.42 to 1.2 µmol P/cm2, and increased with the spike of C and N on August 24, 2005, but not with the increased C and N seen on May 13, 2005.
Sediments of Butcherpen Cove had C:N ratios averaging 13.96 ± 2.64 for the entire sampling season, while biofilms averaged 5.91 ± 2.02 (Figure 11).
Periphyton TC and TN did not track changes in sediment C and N, but increased throughout the sampling season with low values on June 8, 2005, of 0.01 ± 0.003 and
0.004 ± 0.002 µmol/cm2, respectively. Greatest biofilm TC and TN concentrations were seen on the final two experiments averaging 17.43 ± 0.47 and 3.85 ± 0.04 µmol/cm2, respectively (Figure 12). Biofilm TP also increased in concentration during the sampling season ranging from 0.02 – 0.17 µmol/cm2.
47
25 300 2.0
250 )
20 1.6 2 Core TN - ) ) m 2 - 2 - c
Core TC l m
200 m c o Core TP c l
l 15 1.2 m o o ( m
m s n n ( 150 u
(
o n n r e o
10 0.8 o g b h o r r 100 p a t s i C o N h
5 0.4 P 50
0 0 0.0 Apr May Jun Jul Aug Sep
Figure 10. Sediment core total carbon (nmol cm-2), total nitrogen (nmol cm-2), and total
phosphorous (µmol cm-2) during 2005 (April through August).
2 5 Biofilm C:N C o r e C :N 2 0 n e g o
r 1 5 t i N
:
n
o 1 0 b r a C 5
0 A p r M a y J u n J u l A u g S e p
Figure 11. Sediment core and biofilm carbon/nitrogen ratios during 2005 (April through
August).
48
30 5 0.25
25
4 0.20 ) 2 ) m 2 ) c 2 20 m c m s
3 0.15 l c Biofilm Carbon o s l l Biofilm Nitrogen m o o 15 (
m
m Biofilm TP s 2 0.10 ( ( u
o n n 10 r e o o g b h r o
1 r 0.05 a p t s i C 5 o N h P 0 0 0.00
Apr May Jun Jul Aug Sep
Figure 12. Biofilm total carbon (µmol/cm2), total nitrogen (µmol/cm2), and total
phosphorous (µmol cm-2) during 2005 (April through August).
On an area basis sediment Chl a content was two orders of magnitude greater than
the seven-day old biofilm content and four orders of magnitude greater than water
column values for the 1 m depth at the study site. However, the sediment Chl a had the
most variation and biofilm Chl a tended to increase through the summer. Cores averaged
17.94 ± 6.42 µg Chl a/cm2 for the entire sampling season with the highest values on April
8, 2005, of 24.57 ± 3.7 µg Chl a/cm2 and lowest values during the previous experiment
(March 18, 2005) at 7.27 ± 2.46 µg Chl a/cm2 (Figure 13), biofilms averaged 0.3 ± 0.03
µg Chl a/cm2 with 0.997 ± 0.02 (July 29, 2005) and 0.019 ± 0.003 (June 8, 2005) being the maximum and minimum estimates (Figure 14). Phytoplankton averaged 3.42 ± 0.93 mg Chl a/m2 on an areal basis (Figure 15). Pheo a values exceeded Chl a on every
49 sampling occasion for all three matrices with an average sediment ratio of Pheo a to Chl a of 1.16 ± 0.08 (Figure 13), an average biofilm ratio of Pheo a to Chl a of 1.19 ± 0.11
(Figure 14), and an average water column ratio of Pheo a to Chl a of 1.25 ± 0.13 (Figure
15).
35 Core Chl a 3.0 Core Pheo a 30 Core Chl c 2.5 ) 2 ) m
25 2 c
/ 2.0 m
c g
/
(
20 g a
(
o 1.5 C e
l h 15 h P
C & 1.0 a
l 10 h C 0.5 5
0 0.0 Mar Apr May Jun Jul Aug Sep
Figure 13. Sediment photopigments, Chl a, Chl c, and Pheo a (µg/cm2) during 2005
(March through August).
50
1.4 0.14
1.2 0.12 Biofilm Chl a ) 2 Biofilm Pheo a
m 1.0 0.10 c
Biofilm Chl c / )
2 g
0.8 0.08 m ( c
/
a
g o
0.6 0.06 e (
h c
P l
0.4 0.04 h &
C a
l
h 0.2 0.02 C
0.0 0.00
A pr M ay Jun Ju l A ug S ep
Figure 14. Biofilm photopigments, Chl a, Chl c, and Pheo a (µg/cm2) during 2005 (April
through August).
7 0.5 Water Chl a 6 Water Pheo a
) 0.4 2 Water Chl c m
/ 5
) g 2 m m
( / 0.3 4 A g
o m ( e
h C
P 3 l 0.2 h &
C A
l 2 h C 0.1 1
0 0.0 M ar A pr M ay Jun Jul A ug Sep
Figure 15. Near bottom water photopigments, Chl a, Chl c, and Pheo a (mg/m2) during
2005 (March through August).
51
In all three media, Chl c was roughly one order of magnitude less than Chl a for each corresponding sample and followed the seasonal trend of Chl a and Pheo a.
Maximum Chl c values were seen in the sediments on April 8, 2005, at 2.39 ± 0.44 µg
Chl c/cm2 (Figure 13). Sediment Chl a and Chl c significantly correlated (Table 2). Water
column Chl a and Chl c did not significantly correlate (Table 3). Water column Chl a
also correlated with sediment and biofilm Chl a (Table 3). Biofilm Chl a and Chl c had
the tightest correlation (Table 4).
AFDW determinations began on April 8, 2005, and continued through the
remainder of the sampling season. On average, sediment AFDW (15.53 ± 9.96 mg/cm2)
measures were three orders of magnitude greater than biofilms (0.23 ± 0.21 mg/cm2).
Sediment AFDW was lowest on April 22, 2005, (7.3 ± 1.1 mg/cm2) and greatest on July
29, 2005, (33.7 ± 12.1 mg/cm2). Biofilm AFDW was lowest on April 8, 2005, (0.04 ±
0.01 mg/cm2) and greatest on August 24, 2005, (0.56 ± 0.04 mg/cm2). Sediment and
biofilm total organic content were correlated with each other, and followed an increasing
trend through the year correlated with temperature (Table 4; Figure 16). Biofilm mean
optical density was 72.8 ± 66.4 for the entire sampling season with highs of 188.5 ± 2.94
on August 24, 2005, and lows on May 20, 2005, of 18.2 ± 4.74. All three biofilm biomass
estimations were significantly correlated (Table 4; Figure 17).
52
Table 2 Pearson Correlations for Benthic Chamber Production, Respiration, and Benthic Photopigments as well as Correlations with Biofilm Photopigments, Nutrients, and Physicochemical Parameters
Benthic Parameters NPP GPP RESP Chl a Chl c Pheo a Dome NPP — Dome GPP .92** — Dome Resp .60** .74** — Core Chl a - .17 - .09 - .09 — Core Chl c .07 .09 .15 .62** — Core Pheo a .01 - .06 - .13 .98** .61** — Sediment TP - .13 .20 .08 .41* .47* .42* Sediment TN - .13 .08 .29 .44* .37 .35 Biofilm Chl a .46* .51* .39 .27 .19 .27 Biofilm Chl c .46* .50* .35 .23 .20 .25 Biofilm Pheo a .48* .53* .40 .27 .20 .27 Biofilm AFDW .67** .63** .39 - .12 - .16 - .10 Biofilm TP .72** .56** .72** .26 .39 .27 Biofilm TN .64** .35 .57* .17 .14 .17 Biofilm TC .65** .38 .61** .16 .10 .15 Temperature - .02 - .12 .01 .41** - .27 .40* Salinity .35** .18 - .20 - .47** - .36* - .44** %PAR .43** .35 .08 - .22 .07 - .22 WC TKN - .22 - .27 - .29 .46* .57* .34
NH4 Flux .36* .26 .37* - .32 - .01 - .28 FDA .63** .66** .43 .24 .22 .21
Note. NPP = net production (n = 39); TN = Total Nitrogen (n = 26); GPP = gross production (n = 39); TC = Total Carbon (n = 26); Resp = respiration (n = 39); % PAR = Percent of light reaching bottom (n = 39); Chl a = Chlorophyll a (n = 39); WC = Water Column; Chl c = Chlorophyll c (n
= 39); TKN = Total Kjehldahl Nitrogen (n = 26); Pheo a = Pheophytin a (n = 39); NH4 Flux = Ammonia Flux from Sediments (n = 39); AFDW = Ash Free Dry Weight (n = 32); FDA = Esterase Activity (n = 18); TP = Total Phosphorous (n = 26). * P < 0.05; ** P < 0.01
53
Table 3 Pearson Correlations for Phytoplankton Versus Biofilm Production, Respiration, and Benthic
Photopigments as well as Correlations with Phytoplankton, Sediment, and Biofilm
Photopigments, Nutrients, and Physicochemical Parameters
Phytoplankton Parameters Wc NPP Wc GPP Wc Resp Wc Chl a Wc Chl c Wc Pheo a Wc NPP (39) — Wc GPP (39) .70** — Wc Resp (39) .38* .80** — Wc Chl a (42) .26 .63** .60** — Wc Chl c (42) - .23 - .08 .15 .29 — Wc Pheo a (42) .16 .53** .51** .94** .36* — Biofilm NPP (33) .47** .71** .68** .70** - .34 .60** Biofilm GPP (33) .46** .70** .70** .67** - .44* .61** Biofilm Resp (33) .14 .60** .49** .39* - .34 .51** Biofilm Chl a (24) - .03 .59** .72** .45* - .13 .44* Biofilm Chl c (24) - .03 .55** .67** .39 - .09 .39 Biofilm Pheo a (24) - .05 .59** .73** .47* - .13 .46* Biofilm TP (16) - .32 .53** .82** .70** - .13 .80** Biofilm TN (22) - .22 .60** .82** .56** - .29 .43* Biofilm TC (22) - .21 .59** .82** .53* - .11 .38 Biofilm OM (22) - .16 .50* .81** .35 - .29 .30 Optical Density (19) .27 .42 .68** .44 - .07 .43 Core Chl a (39) .11 .33* .23 .67** .29 .66** Core Chl c (39) - .34* - .13 - .07 .08 .42** .23 Core Pheo a (39) .15 .35* .23 .61** .25 .61** Temperature (39) .52** .75** .71** .68** - .26 .52** Wc TKN (26) - .19 - .16 .03 .41* .35 .52**
Wc NH4 (26) .15 - .20 - .16 - .55** - .05 - .62**
Wc NO3+NO2 (26) - .32 - .56** - .55** - .37 .14 - .28
Wc SiO2 (26) .17 .42* .48* .41* - .40* .39 FDA (18) - .52* .47 .85** .69** - .22 .54*
Note. Abbreviations are identical to Table 2.
* P < 0.05; ** P < 0.01; (n)
54
Table 4 Pearson Correlation Matrix for Biofilm Production, Respiration, and Photopigments; Biomass; Water Column Nutrients; Esterase Activity; and Temperature
Biofilm Biofilm Biofilm Biofilm Biofilm Biofilm Biofilm Biofilm Biofilm Biofilm Biofilm Wc Wc Wc FDA
NPP GPP Resp Chl a Chl c Pheo a TP TN TC Pixels OM Temp NO3+NO2 PO4 SiO2 (m2) NPP — GPP .98** — Resp .61** .68** — Chl a .87** .85** .69** — Chl c .86** .82** .64** .99** — Pheo a .86** .84** .72** .99** .98** — TP .90** .83** .84** .86** .82** .89** — TN .78** .79** .79** .92** .86** .94** .85** — TC .79** .77** .79** .91** .86** .93** .88** .99** — Pixels .88** .84** .59* .77** .76** .79** .92** .78** .80** — AFDW .86** .79** .77** .88** .83** .91** .93** .95** .95** .85** — Temp .75** .72** .34 .47* .39 .49* .55* .81** .71** .53* .64** —
NO3+NO2 - .38 - .44* - .47* - .45 - .37 - .49* - .69* - .71** - .64** - .56* - .67** - .54** —
PO4 .47* .47* .67** .41 .39 .44 .75** .57* .60* .74** .53* .06 - .48* —
SiO2 .62** .72** .67** .56* .47 .60* .64* .71** .63** .52* .60* .65** - .55** .14 — FDA(m2) .77** .69** .68** .79** .72** .81** .98** .84** .86** .90** .96** .70** - .76** .57* .68** —
Note. NPP = net production (n = 33); Temp = Temperature (n = 39); WC = Water Column; GPP = gross production (n = 33); TP = Total Phosphorous (n = 16); NO3+NO2 = Nitrate + Nitrite (n = 26);
Resp = respiration (n = 33); TN = Total Nitrogen (n = 22); PO4 = Orthophosphate (n = 26); Chl a = Chlorophyll a (n = 24); TC = Total Carbon (n = 22); SiO2 = Silica (n = 26); Chl c = Chlorophyll c (n = 24); OD = Optical Density (n = 19); FDA = Esterase Activity (n = 18); Pheo a = Pheophytin a (n = 24); AFDW = Ash Free Dry Weight (n = 22). * P < 0.05; ** P < 0.01
55
50 0.7
S ed A F D W 0.6 )
) Biofilm AFDW 2
2 40 m m 0.5 c c
/ /
g g m
m 0.4 ( (
30
W W D D 0.3 F F A A
t 20 m l n
0.2 i e f o m i i B d
e 0.1
S 10
0.0
0 A p r M ay Ju n Jul A u g S ep
Figure 16. Sediment and biofilm ash free dry weight (mg/cm2) during 2005 (April through August).
250 0.7 1.2
0.6 1.0 200 y t i
0.5 )
Optical Density 2 s 0.8 ) 2 n m
e AFDW m c
D c 0.4 /
Biofilm Chl a l
150 /
g a
( 0.6 g
c i µ t ( W
0.3 p D a
O l F 0.4 h
e 100 A
g 0.2 C
a m l r i m l f e
0.2 i v f o
0.1 i o A i 50 B B 0.0 0.0
0 Apr May Jun Jul Aug Sep
Figure 17. Biofilm optical density, ash free dry weight (g/cm2), and biofilm Chl a
(µg/cm2) during 2005 (April through August).
56
Benthic Chamber Results
Net benthic community production averaged 8.91 ± 8.28 mg C · m-2 · hr-1 for the entire sampling season. Maximum net production values for the benthos were recorded on July 8, 2005, averaging 21.25 ± 0.96 mg C · m-2 · hr-1 while minimum values were
observed on April 15, 2005, averaging -2.33 ± 0.39 mg C · m-2 · hr-1, coincident with high
turbidity from rainfall. Net benthic production appeared to be controlled by percent
transmission of PAR (p = .01), which increased with increasing salinity (p = .01; Table
2).
Benthic respiration did not show a dramatic response to the April rain events, but
did decrease following T. S. Arlene and Hurricane Dennis. Overall, sediment oxygen
demand was less variable than net production. Benthic respiration averaged 7.14 ± 5.99
mg C · m-2 · hr-1 for the sampling season with maximum values occurring on August 24,
2005, averaging 16.69 ± 11.62 mg C · m-2 · hr-1 while minimum respiration was seen on
May 6, 2005, averaging 0.7 ± 6.06 mg C · m-2 · hr-1 (Figure 18). A direct effect of
temperature was not observed on respiration estimates for sediments, plankton, or
biofilms, indicating that additional factors were influencing respiration rates in
Butcherpen Cove. GPP was positive at all sampling times.
57
60 ) 1 -
h 50
Benthic Resp ·
2
- Benthic NPP
m 40 Benthic GPP ·
C
g 30 m (
.
p 20 s e R 10 &
, P
P 0 N
, P
P -10 G
-20 F eb M ar A p r M ay Ju n Ju l A u g S e p
Figure 18. Benthic net and benthic microalgal gross production and benthic respiration
(mg C · m-2 · hr-1) during 2005 (February through August).
Phytoplankton net and gross production and plankton respiration were calculated
for the entire 1 m depth water column and expressed on a m2 basis. Phytoplankton net primary production in 2005 averaged 22 ± 13.3 mg C · m-2 · hr-1 for the entire sampling
season. A seasonal low of 5.6 ± 1.7 mg C · m-2 · hr-1 was recorded on February 18, 2005,
and the highest peak on August 17, 2005, of 46.6 ± 16.5 mg C · m-2 · hr-1. Plankton
respiration values averaged 29.0 ± 19.3 mg C · m-2 · hr-1 for the season, with minimum
and maximum values of 12.4 ± 3.7 (April 15, 2005) and 59.4 ± 2.9 mg C · m-2 · hr-1
(August 24, 2005), respectively (Figure 19). Phytoplankton GPP correlated highly with
temperature changes, increasing throughout the spring to late summer (Table 3; Figure
19). Plankton respiration was also correlated to temperature (Table 3).
58
140 35
) Phyto Resp 1 - 120 h Phyto NPP
· Phyto GPP 30 2
- 100 Temp m
·
80 25 ) C C
o g 60 (
e m ( 20 r 40 u P t a S
20 r E
15 e R p 0 m &
e , T P -20 10 P N
-40 , 5 P
P -60 G -80 0 Feb Mar Apr May Jun Jul Aug Sep
Figure 19. Bottom water temperature (ºC), phytoplankton gross production (mg C · m-2 ·
hr-1), and phytoplankton net production and plankton respiration (mg C · m-2 · hr-1) during
2005 (February through August).
Minimum net production values for biofilms were recorded on February 18, 2005,
(66.7 ± 24.9 mg C · m-2 · hr-1) and a maximum production of 1807.3 ± 114.1 mg C · m-2 ·
hr-1 was recorded on July 29, 2005, (Figure 20). Biofilm net production averaged 657.7 ±
107.5 mg C · m-2 · hr-1 for the entire sampling season. Biofilm respiration averaged 126.3
± 16.2 mg C · m-2 · hr-1. Biofilm GPP and temperature (Figure 20) had a slightly lower correlation than phytoplankton GPP and temperature (Figure 19) although biofilm NPP had a higher correlation to temperature (Table 4).
59
2500 35 ) 1
- Biofilm Resp h
Biofilm NPP · 2000 2 Biofilm GPP - 30
m Temp
· )
C C
1500 o
(
g
25 e r m ( u
t
P 1000 a r S e E
20 p R
m
& 500 e
, T P
P 15 N
0 , P P G -500 10 Feb Mar Apr May Jun Jul Aug Sep
Figure 20. Bottom water temperature (ºC), biofilm gross production (mg C · m-2 · hr-1), and biofilm net production and respiration (mg C · m-2 · hr-1) during 2005 (February through August).
Neither phytoplankton nor biofilm GPP was significantly related to the benthic
GPP. However, phytoplankton and biofilm GPP were significantly correlated (Table 3,
Figure 21). Sediment community P:R ratios showed net respiration on three of 13 experiments and net production on 10 of 13 (Figure 22). P:R ratios were comparable for sediment (1.75 ± 1.63) and phytoplankton (1.13 ± 0.67), while biofilm P:R ratio was much greater at 9.01 ± 8.28, reflecting the new growth on the artificial substrates.
Extreme P:R ratios (23.66 ± 0.49) were seen for biofilms on May 13, 2005. Biofilm P:R ratios only fell below 1.0 in one of 13 experiments on February 18, 2005, 0.56 ± .01
(Figure 22).
60
2500 120 )
Biofilm GPP 1 - h
Phyto GPP 100 · )
2000 1 - 2 - h
m ·
· 2
- 80 C m
1500 · g
m C
(
60 g n m o ( t
1000 k m n
l 40 i a l f o p i o t B 500 20 y h P
0 0 Feb Mar Apr May Jun Jul Aug Sep
Figure 21. Phytoplankton and biofilm gross production (mg C · m-2 · hr-1) during 2005
(February through August).
5 3 0 D o m e P :R
R P h yto P :R :
P 2 5 Biofilm P:R
n 4 o t
k 2 0 n a l R :
p 3 P o 1 5
t y m l h i f P
1 0 o i
d 2 B n a
s 5 o h
t 1 n
e 0 B
0 F eb M ar A p r M ay Ju n Ju l A u g S ep
Figure 22. Phytoplankton, benthic, and biofilm production/respiration ratios during 2005
(February through August).
61
Compensation Point Analysis
Compensation point determinations made from biomass-specific production
(photosynthetic efficiency, PE) plotted against irradiance were 394.3 µE · cm-2 · s-1 for
the benthos (Figure 23), and 12.5 µE cm-2 s-1 for the biofilms (Figure 24). Maximum PE
for the benthos (6.5 e+6 mg C · mg Chl a-1 · hr-1) was four orders of magnitude less than biofilm PE (2.5 e+10 mg C · mg Chl a-1 · hr-1) for the same time period, June 15–17, 2005.
Light saturation was not found for either benthic or biofilm microalgae.
8e+6 ) 1 - r
h 6e+6
·
1 - a
l 4e+6 h c
g m
·
2e+6 C
g m ( 0 E P
c i h t -2e+6 n e B
-4e+6 0 200 400 600 800 1000 1200 Irradiance (E · m-2 · s-1)
Figure 23. Benthic photosynthetic efficiency (mg C · mg Chl a-1 · hr-1) response to
irradiance intensity (μE · m-2 · s-1).
62
3e+10 ) 1 -
r 3e+10 h
·
1 -
a 2e+10
l h c
g 2e+10 m
·
C 1e+10 g m (
E 5e+9 P
m l i f
o 0 i B
-5e+9 0 200 400 600 800 1000 1200 Irradiance (E · m-2 · s-1)
Figure 24. Biofilm photosynthetic efficiency (mg C · mg Chl a-1 · hr-1) response to irradiance intensity (μE · m-2 · s-1).
Chamber Nutrient Fluxes
- - NO3 +NO2 average flux was greatest in the clear chambers (-1.09 ± 5.95 µmol · m-2 · hr-1), while dark domes were an order of magnitude less (-0.26 ± 3.73 µmol · m-2 ·
-1 - - hr ) with slightly less variability. Sediment uptake of NO3 + NO2 was seen when
-2 -1 - averaging all domes over the entire sampling season, -0.67 ± 4.88 µmol · m · hr . NO3
- + NO2 flux measures showed a large sediment uptake in both light and dark domes on
April 15, 2005, with the light chamber sediment uptake (-20.77 ± 29.7 µmol · m-2 · hr-1) double that of the dark chamber (-11.36 ± 55.5 µmol · m-2 · hr-1; Figure 5). In the following experiment (May 6, 2005), a small sediment release occurred in both domes
63
with the dark (5.28 ± 6.79 µmol · m-2 · hr-1) doubling that of the light domes (2.37 ± 0.97
-2 -1 - - µmol · m · hr ), the only NO3 +NO2 flux to be significantly different from zero. The
- - remainder of the sampling season revealed little to no net fluxes for NO3 +NO2 (see
Figure 5).
+ Significant non-zero NH4 fluxes appeared to be responsive to storm disturbance,
and seasonal progression. Sediment release occurred in the spring sampling, with the
exception of February 18, 2005, where sediment uptake in the dark domes was -15.8 ±
27.4 µmol · m-2 · hr-1. Near the beginning of summer (May 13, 2005) a sediment uptake
trend began in both chambers that progressed until Hurricane Dennis passed, with highest
sediment uptake in dark chambers on June 24, 2005 (-48.5 µmol · m-2 · hr-1). Sediment
release occurred in the light chambers on July 8 and July 29, 2005, of 77.58 ± 93.9 and
11.45 ± 20.0 µmol · m-2 · hf-1, respectively. The final two sampling events showed no net
+ flux in either direction (Figure 6). Overall net NH4 flux estimates were sediment releases
+ of NH4 in clear chambers from the sediments of Butcherpen Cove, 5.97 ± 28.24 µmol ·
-2 -1 + m · hr . In dark domes, NH4 was taken up by the sediments at rates of -5.71 ± 19.21
-2 -1 + µmol · m · hr for the entire season, while the overall combined flux of NH4 was 0.13
± 24.4 µmol · m-2 · hr-1 (see Figure 6).
-3 Butcherpen Cove sediments had very low but significant non-zero PO4 fluxes on
7 of 13 experiments. An average sediment release occurred in the light domes of 0.31 ±
0.8 µmol · m-2 · hr-1 for the sampling season, while dark domes exhibited a sediment
-2 -1 -3 uptake of -0.18 ± 0.53 µmol · m · hr . PO4 combined flux for the season averaged
-2 -1 -3 0.07 ± 0.71 µmol · m · hr . The midsummer peak of water column PO4 coincided with
-3 -2 an sediment release of PO4 in the light domes of 1.95 ± 0.74 and 2.13 ± 0.89 µmol · m
64
· hr-1 on June 24 and July 8, 2005, respectively. These outward fluxes were followed by
sediment uptake of -0.66 ± 1.73 µM · m-2 · hr-1 on July 29, 2005 (Figure 8).
SiO2 fluxes were not significantly different from zero for seasonal averages in
either light or dark chambers. However, SiO2 fluxes differed in light and dark chambers
at 96.7 ± 116.9 and -92.5 ± 93.6 µmol · m-2 · hr-1, respectively. The combined average showed sediment release of 2.04 ± 372.2 µmol · m-2 · hr-1. The largest flux occurred in
clear domes on August 17, 2005, with a sediment release of 1149.05 ± 296.3 µmol · m-2 ·
hr-1 (Figure 9).
Enzyme Activity Data
Fluorescein diacetate assay (FDA) esterase activity in biofilm material followed a
linear trend that was correlated to temperature, biofilm production and biomass, and
dissolved silica (Table 4). Highest FDA estimates were on the final experiment at 753.4 ±
62.7 mM · m-2 · hr-1 while lows of 15.5 ± 1.2 mM · m-2 · hr-1 occurred on April 8, 2005,
(Figure 25). FDA results tracked changes in biofilm respiration closely except for the
post-storm sampling in April (see Figure 25). Alkaline phosphatase activity (APA)
activity appeared to increase following major storm events with highs in early May and
late July of 16.4 ± 0.5 and 11.6 ± 1.4 µM · m-2 · hr-1, respectively. A season low occurred
on June 24, 2005, 0.48 µM · m-2 · hr-1 just before Hurricane Dennis while previous lows
occurred in April with a monthly average of 1.02 ± 0.07 µM · m-2 · hr-1 (Figure 26).
65
1000 350
FD A(m 2) )
300 1 -
800 Biofilm RESP h
) 2 - 1
- 250 m h
C 2
-
600 g
2 200 - m (
m
P M
150 S 400 E m (
R
A
100 m D l i F f
200 o i
50 B
0 0 Apr M ay Jun Jul Aug Sep
Figure 25. Fluorescein diacetate activity (mM · m-2 · hr-1) and respiration rates of biofilms (mg C · m-2 · hr-1) during 2005 (April through August).
2 0
Phosphatase Activity 1 6
) 1 - h
·
2 1 2 - m
·
M µ (
8 A P A
4
0 A p r M a y J u n J u l A u g S e p Figure 26. Alkaline phosphatase activity (µM · m-2 · hr-1) of periphyton during 2005
(April through August).
66
Biofilm ARA experimental incubations yielded little if any nitrogen fixation
activity for the sampling season. The final two experiments were an exception, with
average biomass specific ARA rates of 10.7 nmol · g DW-1 · hr-1 (Figure 27). Biomass
specific ABA rates declined through the year from the high of 6.35 nmol · g DW-1 · hr-1
on April 15, 2005, to 0.32 nmol · g DW-1 · hr-1 on July 29, 2005, (Figure 27). Areal
estimates of denitrification rates averaged 3.84 ± 0.68 nmol · m-2 · hr-1 for the sampling
season, similar to biomass specific activity.
2 0
Denitrification 1 - 1 5 Nitrogen Fixation h
·
1 - W D
g 1 0
·
N
l o m
n 5
0 A p r M a y J u n J u l A u g S e p
Figure 27. Denitrification (acetylene block assay) rates and nitrogen fixation (acetylene reduction assay) rates (nmol N · g DW-1 · hr-1) of biofilms grown on artificial substrata
during 2005 (April through August).
CHAPTER V
DISCUSSION
This research provides new information for both Pensacola Bay and estuaries in general. Estimates of O2 and nutrient fluxes in situ are new for Pensacola Bay. Estimates of benthic processes following major storm events in a Gulf of Mexico estuary add to our knowledge of disturbance effects on geochemical transformations, biological production, and heterotrophic activity. Simultaneous comparison of microalgal production on the benthos, on artificial substrates and in the water column provides a unique comparative data set.
The dramatic rainfall events of April 2005 resulted in changes to the salinity and light regimes of Pensacola Bay and Butcherpen Cove that directly affected the dynamics of the microphytobenthos. However, light limitations to the bottom were not as severe during the tropical storm and hurricane when compared to April rain events, despite a reduction in salinity with the freshwater pulse in all storm events. When rainfall in the vicinity increased, salinity and percent transmission of PAR coincidentally decreased.
Percent transmission of light to the benthos and salinity correlated significantly at p =
.01, r = 0.691 indicating a freshwater source for turbidity. Microphytobenthic production was drastically reduced when turbidity increased coincident with freshwater intrusion in early April 2005. This storm event interrupted what appeared to be an increasing trend of
67 68
benthic community production from winter to early spring, reducing net production from
14.74 to -2.33 mg C · m-2 · hr-1, resulting in a net heterotrophic community.
August GPP averages in this study, 23 mg C · m-2 · hr-1 are equal to averages
observed in August in Massachusetts Bay (Cahoon, Beretich, Thomas, & McDonald,
1993). After converting Pinckney and Zingmark (1993) O2 data to carbon equivalents,
production values at Butcherpen Cove were much higher all season than for the subtidal
sands adjacent to submerged seagrass in the North Inlet estuary near Georgetown, SC
where the highest values (3.3 mg C · m-2 · hr-1) were recorded in January. In the Ems-
Dollard estuary bordering the Netherlands and Germany, Colijn and de Jonge (1984)
found hourly rates for all stations averaged 37 mg C · m-2 · hr-1 with a maximum of 100
mg C · m-2 · hr-1 during summer months, considerably higher than the maximum
measured for Butcherpen Cove. Butcherpen Cove microphytobenthic GPP scaled up to
daily averages (0.3 g C · m-2 · d-1), based on actual day length multiplied by hourly rates,
were comparable to Weeks Bay, AL averages of 0.2 g C · m-2 · d-1 (Schreiber & Pennock,
1995).
Cahoon and Cooke (1992) found benthic microalgal net production was 36% of
GPP on their sandy inshore site in Onslow Bay, NC at a depth of 18 m.
Microphytobenthic net production at Butcherpen Cove was 54% of gross production.
Biofilm NPP was 83% of GPP, closer to the predicted physiological value for microalgae of 90% (Pomeroy, 1959) and reflecting the dominance of new algal growth in the biofilm communities and the absence of sediment heterotrophic and abiotic oxygen demand.
Daily benthic respiration (24 hr) at Butcherpen Cove averaged (0.2 g C · m-2 · d-1),
identical to summertime oxygen consumption rates determined in Buzzards Bay, MA
69
(Banta, Giblin, Hobbie, & Tucker, 1995), comparable to Caminada Bay, LA marsh
sediments (0.4 g C · m-2 · d-1) with temperatures similar to the current study (Hopkinson et al., 1978), and much lower than estimates in Chesapeake Bay of 0.75 g C · m-2 · d-1
(Officer, Lynch, Kemp, & Boynton, 1985) with similar temperatures to the Buzzards Bay
work.
Benthic respiration was affected by rainfall events with decreased values
following storms, although the reductions were not as dramatic as production losses.
Benthic respiration rebounded from storm disturbances faster while production continued
to decline in subsequent incubations, suggesting that heterotrophic activity was less
affected by storm disturbances. Hurricane Dennis resulted in a major reduction in
respiration estimates from the sampling season’s second highest value (13.78 ± 0.32 mg
C · m-2 · hr-1) to an order of magnitude lower (3.58 ± 2.5 mg C · m-2 · hr-1). Following this
low, benthic respiration was increasing on August 17, 2005, while benthic community
production continued to decrease.
Following the high rainfall of April, the microphytobenthic production recovered
during May (rainfall total = 1.95 in.; 4.95 cm) until early June when T. S. Arlene rains
again reduced net production to near 0 values (-0.01 ± 3.41 mg C · m-2 · hr-1) resulting in
net heterotrophy again on June 24, 2005. However, rapid recovery of the
microphytobenthos was seen following T. S. Arlene with the highest production
estimates of the sampling season recorded during the subsequent experiment, 14 days
later (July 8, 2005). April rain and T. S. Arlene equally lowered production from ~15 to
~0 mg C · m-2 · hr-1. Hurricane Dennis with less rainfall affected NPP less dramatically,
but NPP was reduced by over half from 21.25 to 8.51 mg C · m-2 · hr-1. However,
70
microphytobenthos production recovered within 1 month after April rains but took longer
(1.5 months) to recover after Hurricane Dennis, possibly reflecting wind-induced mixing
and disturbance of the benthos from the hurricane.
P:R ratios provided evidence of Butcherpen Cove’s trophic status during 2005
with photoautotrophic processes dominating the benthos during February and March
(pre-storm) and heterotrophic activity controlling post-storm processes in April.
Autotrophy exceeded respiration again in May and remained high through August with
the exception of June 8, 2005, just before T. S. Arlene. Biofilm and water column P:R
ratios provided evidence of photoautrophically dominated communities throughout the
sampling season.
In other systems, both the microphytobenthos and phytoplankton have been
shown to exhibit a late winter and early spring biomass bloom (Rabalais, 2002). Benthic
microalgae appeared to experience the initial phases of a spring bloom as production
reached near season highs in March, but this trend was truncated by turbidity from heavy
rains on April 1, 2005. Phytoplankton in Butcherpen Cove’s shallow depth (≤ 1.5 m) did not show a typical spring bloom in 2005.
Throughout the sampling season in Butcherpen Cove, phytoplankton production estimates were comparable to previous estimates from adjacent Gulf of Mexico estuaries.
Phytoplankton production estimates from Weeks Bay, AL during summer months were comparable to the current study. However, maximum values seen in the fall of 1991 were near 3 g C · m-2 · hr-1, significantly greater than any measures taken in Butcherpen Cove
(Schreiber & Pennock, 1995). The average and range of phytoplankton production
measures in Apalachicola Bay, FL (32; 4 – 75 mg C · m-2 · hr-1; Pennock et al., 1999) was
71 close to Butcherpen Cove’s range (22; 4 – 50 mg C · m-2 · hr-1) from February to August.
Maximum values occurred in summer months in both studies.
Biofilm and phytoplankton production did not show a response to April rainfall events as did the benthos, but appeared to be controlled by the temperature. Combined with a lack of correlation with benthic processes, correlation of biofilms with planktonic production showed biofilm growth responding to water column conditions rather than benthic conditions. Periphyton biomass, production, and respiration all followed the temperature trend and did not show a response to rainfall events. Periphyton biomass did not respond to salinity and turbidity changes associated with storm events as did the benthic biomass. Therefore periphytometers were not proportionately representative of the benthos. However, biomass increased on all three media following Hurricane Dennis.
Biofilms showed the least variability in production, respiration, and biomass of the three media, and the greatest production estimates. Although biofilm respiration was much less variable than production, winter and spring production and respiration measures were of similar magnitude while summertime estimations of biofilm net production were 1 order of magnitude greater than respiration. Brandini et al. (2001) found a wide range of < 0.01 to 1.4 g C · m-2 · d-1 for periphyton GPP in 1-m depth of a subtropical Brazilian estuary, similar to Butcherpen Cove’s large range over the study period. However the lowest values recorded in that study (5.6 mg C · m-2 · d-1) were 2 orders of magnitude less than the lowest values (59.7 mg C · m-2 · hr-1) from Butcherpen
Cove. At the high end of the range, these two systems were comparable.
Several methods of biomass estimation (Chl a, AFDW, and optical density) were employed for biofilms grown on artificial substrates and all exhibited high correlations
72 with one another. This indicates that optical density can be a rapid and accurate method for an estimation of periphyton biomass. Biofilm pigments (Chl a, Chl c, and Pheo a) had a close relationship with periphyton production, which was also observed by Brandini et al. (2001). Biofilm biomass was less variable than sediment microalgal biomass estimates suggesting that periphytometers are a more stable estimator of biomass response to environmental conditions, with sediment measures affected by benthic heterogeneity.
Sediment and water column microalgal biomass estimations also followed the salinity and turbidity patterns set by rainfall events. Chl a, Chl c, and Pheo a varied in a manner similar to microphytobenthic production, salinity, and % transmission with values decreasing post storm event and recovering shortly thereafter. The exception to this trend was the increase in biomass following Hurricane Dennis. All three media exhibited Pheo a : Chl a ratios that were an indicative of active growth with low values
(approximately one) for the sampling season.
The linkage between benthic production and Chl a at Butcherpen Cove was not significant despite both variables responding similarly to storm events. Masini and
McComb (2001) also found no significant relationship between microphytobenthic production and biomass in the Swan-Canning estuary, Australia. Several authors have correlated biomass and production with r values as high as 0.55 (Colijn and de Jonge,
1984; Grant, 1986; Sullivan and Moncrieff, 1988). Pickney and Zingmark (1993) attributed high correlations (r = 0.7) to advanced O2 microelectrode techniques.
Phytoplankton gross production was highly correlated to Chl a and Pheo a concentrations
(Table 3), but not related to Chl c. Biofilm Chl a and production showed the strongest relationship (85%), likely due to the new microalgal growth comprising the biofilms.
73
Biofilm samples, representing new growth of periphyton over a seven-day period,
were not coupled with a significant detrital carbon reservoir or the oxidation-reduction
processes dominating the sediment. Average biofilm C:N ratios (6) for the sampling
season were very close to the empirical value (7) determined by Redfield (1963) for algal
biomass, while sediments were double (14), consistent with a greater detrital carbon
component. Sediment TC and TN spiked in May and in late August coincident with
microphytobenthic production. TP within sediments also appeared to be related to algal
growth showing a resurgence following Hurricane Dennis and following the TC and TN
trend during the rest of the year. Biofilm TC, TN, and TP followed the seasonal
temperature trend and correlated tightly with periphyton production and biomass
indicating a relationship with a periphytic carbon reservoir.
The findings of this study have implications for the overall productivity of the
Pensacola Bay System. Murrell (unpublished data) estimated average photic depth of
Pensacola Bay based on turbidity measures for 1% light transmission, to be
approximately 5 m, meaning that 78% of the bay could potentially support
microphytobenthos production (Figure 28). However, net community metabolism was
equal to zero for the benthic community at approximately 400 µE · m-2 · s-1 irradiance,
suggesting that net benthic production would be limited to shallower parts of the bay.
Surface irradiance on a sunny day in the middle of summer can be around 2000
µE · m-2 · s-1, making the microphytobenthos community zero net production estimate
20% of surface irradiance, which would indeed limit net benthic production to mainly along the shorelines in water of < 2 m depth. Biofilm production values that do not incorporate SOD, suggest that microalgal growth would occur over a much broader area
74
10%
1
<1%
Figure 28. Percentage of incident light reaching the bottom of Pensacola Bay. 2From
Murrell (unpublished data), reprinted here with permission.
of the bay bottom, with a compensation point of 12.5 µE · m-2 · s-1 or 0.6% of summer
surface irradiance. Although saturation irradiance values were not discernable from the
experiments at Butcherpen Cove, values in the literature range from 250 to 650 µE · m-2 ·
s-1 and Brandini et al. (2001) estimated the periphyton from 1 m depth to be saturated at
320 µE · m-2 · s-1 while the benthos and biofilms at Butcherpen Cove at 1 m depth showed
no evidence of saturation ≤ 1000 µE · m-2 · s-1.
- - NO3 +NO2 concentrations in near bottom water significantly increased immediately following the major rainfall events of early April 2005 although neither
75
- - T. S. Arlene nor Hurricane Dennis significantly affected NO3 +NO2 concentrations.
- - Some of the NO3 +NO2 pulse in April was absorbed by the sediments as evident by the large uptake on April 15, 2005, followed by regeneration on the subsequent 2
- experiments. Advective pumping of NOx has been theorized to significantly control primary production on the benthos (Risgaard-Petersen, 2003; Rizzo & Christian, 1996) and may have facilitated the growth of microphytobenthos during the recovery process in
- late April. Risgaard-Petersen also found that additions of NO3 increased sediment uptake and light incubation estimates were much larger than dark incubations. DiDonato, Lores,
Murrell, Smith, and Caffrey (2006) working with stirred cores taken from deeper depths
- - in Escambia Bay found low NO3 +NO2 fluxes. Similar magnitudes were observed in
Butcherpen Cove and other Gulf of Mexico estuaries (Twilley, Cowan, Miller-Way,
Montagna, & Mortazavi, 1999). Sediments from the deeper parts of the estuary are finer grained and have a larger detrital carbon component than the sands from Butcherpen
Cove. Net uptake has been documented in previous investigations of river-dominated estuaries (Teague, Madden, & Day, 1988).
+ Irregular seasonal patterns were observed for water column NH4 concentrations,
+ though the major storm events were associated with rapid declines of NH4 in near bottom waters. April rains diminished the seasonal highs seen in March. Pre- and post-
+ Hurricane Dennis, sediments were a sink for NH4 with regeneration occurring in subsequent experiments in clear chambers. Risgaard-Petersen (2003) and Rizzo, Lackey, and Christian (1992) observed sediment release in the dark and uptake in light treatments of core samples from Neuse River estuary, North Carolina. The opposite was detected in
+ situ for Butcherpen Cove sediments with overall releases of NH4 in light domes while
76 uptake was seen in dark domes, similar to spring and summer experiments of Marinelli,
+ Jahnke, Craven, Nelson, and Eckman (1998). Marinelli et al. NH4 flux estimates from depths of 14 to 40 m off the coast of Georgia and eastern Florida were comparable to
+ Butcherpen Cove. The tight correlations found between NH4 and oxygen uptake by
Rizzo and Christian (1996) were less significant in this study (Table 2), but the
+ magnitude of NH4 uptake was consistent with the findings of DiDonato et al. (2006) at
-48 to 110 µmol · m-2 · hr-1.
According to the US EPA (1992), estuarine concentrations of TKN and TP are considered high if they exceed 1.0 and 0.1 mg/L (71 and 3.2 µM, respectively). TP estimates were close to this level on 1 sampling occasion (August 17, 2005) while TKN never approached this threshold. TKN estimates for the water column in Butcherpen
Cove during this study were minimal compared to previous studies in Pensacola Bay. The highest TKN and TP values recorded in Butcherpen Cove during this study were an order of magnitude less than the highest reported TKN and TP (600 µM and 60 µM, respectively) values for Pensacola Bay (Lewis et al., 2002). Near bottom water concentrations of TKN were highest prior to T. S. Arlene, lowest on the following experiment June 24, 2005, just before Hurricane Dennis, and increased through the remainder of the season consistent with an increase in phytoplankton biomass and production.
3- PO4 in the bottom water of Butcherpen Cove was minimal all season and similar to the findings of DiDonato et al. (2006), but low compared to highs of 2 µM observed by
Murrell et al. (2002) in upper Pensacola Bay. A small pulse (0.2 µM) was observed in the weeks after T.S. Arlene. Although fluxes were also very low in Butcherpen Cove
77
compared to previous estimates within Pensacola Bay, as well as other estuaries,
sediments exhibited a net regeneration (2 µmol · m-2 · hr-1) just prior to Hurricane Dennis
and minor uptake under light incubations subsequent to Dennis. Cowen and Boynton
(1996) considered their estimates to be low at -16.5 to 148 µmol · m-2 · d-1 in Chesapeake
Bay estuary. Rizzo et al. (1992) found no response of phosphorous fluxes to changes in
irradiance levels, observed slightly elevated fluxes under dark incubations (< 25 µmol ·
m-2 · hr-1), and suggested the benthos in Neuse River estuary was P-limited while the
phytoplankton was N-limited. Similarly to Rizzo et al., Butcherpen Cove sediments
released phosphorous from light incubations while uptake occurred in dark chambers.
The only nutrient to follow a seasonal temperature trend was SiO2 with lows in the early spring/winter and highs in late summer. Highest water column values observed in Butcherpen Cove (in August) were comparable to the highest concentrations seen by
Murrell et al. (2002), although their high values were observed in February. The current study reports water column SiO2 measurements that were double those of DiDonato et al.
(2006) at 67.6 µM, but their greatest values were in July, similar to Butcherpen Cove.
Measured SiO2 fluxes were high compared to other Pensacola Bay estimates as well as
estimates from other Gulf of Mexico estuaries (Twilley et al., 1999). However, high
values in this study were similar to a small California slough at 1500 µmol · m-2 · hr-1
(Caffrey, Harrington, & Ward, 2002). In Pensacola Bay, DiDonato et al. measured fluxes
that were highly variable and averaged roughly one order of magnitude less than the
present study. Chesapeake Bay estimates from Cowen and Boynton (1996) were much
greater ranging from -30 to 450 µmol · m-2 · d-1, only half of the current study estimates.
78
SiO2 is typically considered limiting in warm summer water columns relative to cooler
months. The opposite was observed over the shallow benthos of Butcherpen Cove.
Biofilm respiration and heterotrophic activity measured as FDA hydrolysis
exhibited a tight positive correlation during the late spring and summer index period.
Despite the uncoupling between respiration and esterase activity observed in the two
experiments following April rainfall events, the overall correlation was significant (Table
4) suggesting FDA hydrolysis may be an easy and accurate method of estimating
heterotrophic activity. Butcherpen Cove total biofilm esterase activity was several orders
of magnitude higher than the tropical, freshwater biofilms of the Njoro River, Kenya
(Mathooko, Mokaya, & Leichtfried, 2002). Estimates of esterase activity in a temperate,
unpolluted stream (0.4 mM · g-2 · hr-1; Battin, 1997) were four orders of magnitude
greater than freshwater biofilms of Kenya, but two orders of magnitude less than the
current study (51.9 mM · g-2 · hr-1). Previous estimates of total esterase activity for shallow, temperate marine sediments (Köster, Dahlke, Meyer-Reil, 2005) were similar to freshwater biofilms of Kenya. The trend of low esterase activity in winter and early spring months and high in late summer was the opposite of the pattern seen in the subtropical, shallow estuarine sediments of Brazil (Crapez, Baptista Neto, & Bispo,
2003). Crapez et al. sediment esterase activity was two orders of magnitude less than the current study’s estimations from a similar habitat.
Results of alkaline phosphatase activity in biofilms were consistent with the
-3 bioavailability of PO4 suppressing APA (McComb, Bowers, Posen, 1979). Low rates of
-3 APA were observed in the current study in April and again in late June when PO4
increased in the water column. Alkaline phosphatases are excreted from algal cells when
79
-3 inorganic PO4 limits growth (McComb et al.). Higher rates of APA were seen during
-3 periods when PO4 was minimal in the water column suggesting P-limitation in
-3 Butcherpen Cove following the pulses in water column PO4 . As previous experiments have indicated, biofilm APA rates observed in this study were inversely correlated with
-3 PO4 (p = 0.2, r = -0.42). APA rates in the current study were two orders of magnitude greater than the freshwater biofilm rates investigated in numerous Mediterranean and
Atlantic fluvial systems (Romaní & Sabater, 2000).
N2 fixation in Butcherpen Cove was consistent with previous studies of seasonal changes. N2 fixation in estuarine sediments hits maximum rates in late summer (Currin,
Joye, & Pearl, 1996; Herbert, 1999; Joye & Pearl, 1994). In Butcherpen Cove, ARA of nitrogen fixation yielded no detectable rates from April through July due to the high N availability in the water column and sediment. In August, minimal but measurable
- - + fixation occurred when there was no NO3 +NO2 or NH4 in the water column. Rates of ethylene production in biofilm assays were low, from 0 to 18.6 nmol N · g DW-1 · hr-1 but comparable to southwest Florida mangrove sediments (Pelegrí, Rivera-Monroy, &
Twilley, 1997) and transplanted marsh sediments in the Newport River estuary (Currin et
+ al.). Nitrogenase activity in August provided NH4 to the benthos when the water column concentration was depleted.
Results of the ABA in Butcherpen Cove were consistent with the bioavailability
- - of NO3 +NO2 in the water column. Denitrification appeared to be coupled to the
- - sediment uptake of NO3 +NO2 associated with the rainfall events of April and potentially
- - aided in the removal of excess NO3 +NO2 loaded into Pensacola Bay system during these storms. Although their estimates were three orders of magnitude greater than the current
80 study, Caffrey and Kemp (1990) also found that denitrification rates of bare sand in
- Chesapeake Bay were highest in spring when NO3 concentrations were elevated. The highest potential ABA measured from biofilms was an order of magnitude less than previous estimates of denitrification within Pensacola Bay ranging 31 – 132 nmol N · g-1 · hr-1 (Flemer, Lores, & Bundrick, 1998). ABA activity in biofilms occurred when salinity in Butcherpen Cove was reduced, similar to the findings of Rysgaard, Thastum,
Dalsgaard, Christensen, and Sloth (1999) in Danish estuarine sediments. Denitrification that may have been occurring deeper in the sediments was not measured in this investigation. Additionally, denitrification may have been underestimated using the ABA because acetylene inhibits nitrification which is often coupled to denitrification (An &
Joye, 2001; Risgaard-Petersen, 2003).
Conclusions
For the entire study period benthic production accounted for a significant portion
(28.8%) of the total production in 1 m of water at the studied site within Butcherpen
Cove. Production rates obtained from in situ incubations of Butcherpen Cove sediments exhibited an irregular trend related to salinity, turbidity, and irradiance conditions. Storm events tended to reduce benthic production, and respiration, with heterotrophic activity rebounding faster than benthic production. Bioavailability of water column nutrients did not seem to be related to benthic production as much as benthic fluxes. Periphytometers supporting the growth of benthic microalgae were not responsive to benthic fluxes.
Biofilm production more closely tracked phytoplankton response in the water column and
81 thus appeared to be controlled by water chemistry. Evidence of nitrogen and phosphorous nutrient co-limitations exists for Butcherpen Cove which is congruent with Flemer et al.
(1998) findings for the plankton of an adjacent estuary, Perdido Bay. N-fixation occurred when there was no nitrogen in the water column potentially supplying nitrogen to the benthos. Denitrification was coupled to the sediment uptake and water column presence
- - of NO3 +NO2 potentially removing excess nitrogen. Results are consistent with the conclusions of Murrell et al. (2002) in that Pensacola Bay and northern Gulf of Mexico estuaries are more storm event dominated rather than seasonally-driven.
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APPENDIX
Copyright Permission Letter
98 99
From: [email protected] [mailto:[email protected]] Sent: Thu 3/9/2006 7:24 PM To: Jeffrey Allison Subject: Re: permission request??
Hi Jeff, Neither of those plots are published, so the appropriate way to cite them is "M. C. Murrell, unpublished data"
Good luck in finishing up.
Mike
Michael C. Murrell, Ph.D., Research Ecologist USEPA Gulf Ecology Division, 1 Sabine Island Dr., Gulf Breeze, FL 32561 email: [email protected], voice: 850-934-2433, fax: -2401
From: Jeffrey Allison [email protected] Sent: Thu 03/09/2006 06:25 PM To: Michael Murrell/GB/USEPA/US@EPA Subject: permission request??
Hi Mike, I was hoping to use two of your figures from your GERS presentation in my thesis. One being the incident light regime of PBS and the other being a hypograph of bay area % and euphotic depth. Is this acceptable to you? If so, how would I go about reference citings? I think one of the figures has a label of Murrell et al. (2005), can you expand this?
Thanks, Jeff
Jeffrey G. Allison Graduate Research Assistant Univ. of West Florida – CEDB