An assessment of the ecological health of Eprapah Creek FINAL Data Report submitted to Kinhill Ltd by Marine Botany University of Queensland Adrian B. Jones BSc (Hons) PhD Joelle Prange BSc (Hons) William C. Dennison BA MS PhD November 1999 Marine Botany Marine Botany THE HE UNIVERSITY NIVERSITY OF F QUEENSLAND UEENSLAND
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An assessment of the ecological health
of Eprapah Creek
FINAL
Data Report
submitted to
Kinhill Ltd
by
Marine Botany
University of Queensland
Adrian B. Jones BSc (Hons) PhD
Joelle Prange BSc (Hons)
William C. Dennison BA MS PhD
November 1999
Marine BotanyMarine Botany
TTHE HE UUNIVERSITYNIVERSITYOOF F QQUEENSLANDUEENSLAND
2
TABLE OF CONTENTS
Executive Summary 3
Introduction 4
Materials and Methods 5
Study Region 5
Water Quality Procedures 6
Sediment Nutrient Fluxes 6
Phytoplankton Bioassays 6
Plant Tissue %N and δ15N 7
Results 9
Physical Water and Sediment Quality Analyses 9
Salinity 9
Dissolved Oxygen 9
pH 9
Secchi Depth 9
Sediment Nutrient Fluxes 10
Bioindicators 11
Phytoplankton Bioassays 11
Tissue Nitrogen Content 12
δ15N Stable Isotope Ratio of Nitrogen 12
Discussion 15
Physical Water and Sediment Quality Analyses 15
Water Quality 15
Sediment Nutrient Fluxes 15
Bioindicators 16
Phytoplankton Bioassays 16
Tissue %Nitrogen Content 16
δ15N Stable Isotope Ratio of Nitrogen 17
Conclusions & Recommendations 19
References 21
3
Executive Summary
The ecological health of a small tidal creek, Eprapah Creek, flowing into Moreton Bay was
examined in Sept. 1999 using water quality analyses, phytoplankton bioassays, sewage plume
mapping using stable isotopic signatures and sediment nutrient flux measurements. Results
were compared with a previous survey conducted in April-May 1997 as part of a University
of Queensland PhD thesis (A. Jones). The ecological health of Eprapah Creek appears to be
compromised with sewage-derived nutrients. Of greater concern is the likelihood of a system
that is actively degrading, with several indications that the ability of the creek to assimilate
nutrients has been reduced since 1997. The ecological health indicators indicating degraded
conditions include high sediment nutrient efflux, low rates of sediment denitrification, high
nutrient stimulation in phytoplankton bioassays and a strong sewage signature in marine
plants. The overall recommendation regarding sewage treatment upgrades is to enhance the
nutrient removal capacity of the treatment processes.
4
Introduction
Eprapah Creek extends from Mount Cotton, through residential areas in its lower reaches,
before discharging into Moreton Bay north of Victoria Point. The creek is approximately 2-
5 m deep, 15 km in length, and has a standing body of water at low tide. It receives point
source discharge (2400 m3 d-1 containing 4.5 mg N L-1 and 8.0 mg P L-1, which equates to
10.8 kg N d-1 and 19.2 kg P d-1) (Redland Shire Council, pers. comm.) from the Victoria
Point sewage treatment plant, located ~2.6 km from the mouth. The sewage treatment plant
services approximately 14 000 people and utilises secondary (activated sludge) treatment
techniques. The plant will be upgraded to service 42 000 people by 2002. Water quality
monitoring by the Redlands Shire Council over the last few years has identified Eprapah
Creek as a waterway with consistently poor water quality (with many parameters outside
ANZECC guidelines), especially in the vicinity of the sewage treatment plant.
The primary aim of this project was to conduct a comprehensive survey of biological assays
and water column and sediment nutrient parameters to determine the ecological health prior
to a planned increase in the output of the sewage treatment plant. From these pre-upgrade
surveys a monitoring program will be developed to monitor the changes in ecological health
of the creek as a result of the increase in sewage discharge. Additionally, the results from
this study are compared with those obtained in 1997 surveys conducted by Jones (1999), to
determine changes to ecological health in Eprapah Creek over the last two years. Based on
these analyses, recommendations are made regarding the level of sewage upgrade to maintain
existing ecological health.
5
Materials and Methods
Study Region
Four sites were chosen within Eprapah Creek and one located near Coochiemudlo Island
(27.56255 ºS, 153.33009 ºE) as a reference site. An upstream site was situated towards the
tidal limit (AMTD 3.8 km) of the creek (AMTD ~3 km; 27.58205 ºS, 153.28548 ºE) which is
approximately 0.5 km upstream from the sewage treatment plant (STP). A site was situated
at the STP outlet (AMTD ~2.6 km; 27.58163 ºS, 153.28986 ºE). Additional sites were
located midway (AMTD ~1 km; 27.57781 ºS, 153.29242 ºE) between the STP and the mouth
of the creek and at the creek mouth (AMTD 0 km; 27.56512 ºS, 153.28967 ºE) (Fig. 1).
These sites were chosen to correlate with existing sites sampled by Redlands Shire Council
and the sites used by Jones (1999) in 1997. Sampling took place on the 13th September, 1999
during the ebbing tide.
Brisbane
MoretonBay
•
⊗
0 0.5 1.0
kilometres
N
Eprapah Ck
SewageTreatment
Plant
OysterPoint
VictoriaPoint
PointHalloran
Coochie-mudloIsland
Coochiemudlo Site Mouth Site
Mid Site
STP SiteUpstream Site
Figure 1 Eprapah Creek sampling sites for ecological health monitoring.
6
Water Quality Procedures
Salinity (expressed on the Practical Salinity Scale1), pH and dissolved oxygen were measured
with a Horiba U-10 water quality meter (California, U.S.A.).
Secchi depth was determined by lowering a 30 cm diameter secchi disk (black and white
alternating quarters) through the water column until it was no longer possible to distinguish
between the black and white sections.
Sediment Nutrient Fluxes
At the mid and mouth sites (sediment substrate was too rocky to core at other sites), four
replicate sediment cores (2.4 L perspex cores) were sampled to a depth of 10 cm and sealed
with a PVC endcap, trapping the overlying site water in the core to minimise sediment
mixing. Cores were transported back to the laboratory the overlying water was removed and
replaced with filtered water from a low nutrient, oceanic influenced site off North Stradbroke
Island. The cores were incubated in a water bath at room temperature with negligible
ambient light for 24 hours. Water samples were collected initially and after 1, 3, 6, 12 and 24
hours. At each sampling interval, 50 mls of water was collected from each core and filtered
through a 0.45 µm Millex GV Millipore filter unit. Samples were analysed for dissolved
nutrients, nitrogen (NH4+), nitrogen oxides (NO3
-, NO2-) and phosphorus (PO4
3-) by the
NATA accredited Queensland Health Analytical Services Laboratory in accordance with the
methods of Clesceri et al. (1989) using a Skalar autoanalyser (Norcross, Georgia, U.S.A.).
Flux rates were calculated as µmol m-2 h-1 of NH4+, NO3
-+NO2- and PO4
3-.
Phytoplankton Bioassays
Phytoplankton bioassays were conducted with ambient phytoplankton assemblages collected
from four sites in Eprapah Creek and Moreton Bay (Fig. 1). One 30 L drum of water was
collected from each site, kept cool and shaded, and returned to an outdoor incubation facility.
Four litres of water from each site was filtered through a 200 µm mesh (to screen out the
larger zooplankton grazers) into sealed transparent 6 L plastic containers and placed in
incubation tanks filled with water (2 m diameter, 0.5 m deep). Temperature was maintained
1 Practical salinity (S) is the ratio of the conductivity of a sample of seawater at 15 ºC compared to that of a
defined potassium chloride (KCl) solution. Seawater with a practical salinity of 35 will have the same
conductivity as a solution of 32.4356 g of KCL in 1 kg of water.
7
at ±2°C of the ambient water temperature by flowing water through the tanks and light levels
were maintained at 50% of incident irradiance with neutral density screening. For each site
there were six bioassay containers, each with a different nutrient treatment. Samples were
spiked to make the following concentrations: NO3- (200 µM); NH4
+ (30 µM); PO43- (20 µM);
SiO32+ (66 µM); all nutrients at those concentrations (+All); and a control (no nutrient
addition). The concentrations were chosen, as they are known to be saturating for
phytoplankton in estuarine environments. At identical daily circadian times, all bioassay
bags were gently shaken and 20 mL from each container was poured into pre-rinsed 30 mL
glass test tubes and placed in darkness for 20 minutes to allow photosystems to dark adapt.
Chlorophyll a concentrations were determined from in vivo fluorescence (indicating
phytoplankton biomass) on a Turner Designs Fluorometer. An initial measure (time = 0) was
taken on the control treatment and then for all treatments daily for 7 days.
Over the 7-day period settlement of suspended solids within samples may occur and light
availability increase above ambient levels. The response of the plankton community in the
control bioassay container gives an indication of the ambient light conditions. Light
stimulated phytoplankton bloom potential was calculated as the difference between initial
(time = 0) and maximum in vivo fluorescence values in the control water sample over the 7 d
incubation. Nutrient stimulated bloom potential was calculated as the difference between the
maximum response in the nutrient treatments and the maximum response in the control
(referred to as the stimulation factor). This stimulation factor can be used to determine the
relative importance of the different nutrient additions compared with light.
Plant Tissue %N and δ15N
Samples of seagrass (Zostera capricorni), mangrove (Avicennia marina), and macroalgae
(Catenella nipae) were collected, placed on ice and returned to the laboratory and prepared
for analysis of %N, δ15N. In the case of the seagrass and mangroves, the second youngest
leaves were chosen, and for the macroalgae a single mangrove pneumatophore covered in
macroalgae was collected for each replicate. Two replicates for each plant type were
collected at each site.
Samples were oven dried to constant weight (24 h at 60 °C), ground and two sub-samples
were oxidised in a Roboprep CN Biological Sample Converter (Europa Tracermass, Crewe,
8
U.K.). The resultant N2 was analysed by a continuous flow isotope ratio mass spectrometer
(Europa Tracermass, Crewe, U.K.). Total %N of the sample was determined, and the ratio of 15N to 14N was expressed as the relative difference between the sample and a standard (N2 in
air) using the following equation (Peterson & Fry, 1987):