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Development of a harpacticoid copepod bioassay: Selection of speciesand relative sensitivity to zinc, atrazine and phenanthrene

Tristan J. Stringer a,b, Chris N. Glover a, Vaughan Keesing c, Grant L. Northcott d, Louis A. Tremblay e,n

a School of Biological Sciences, University of Canterbury, Private Bag 4800, Christchurch 8140, New Zealandb Landcare Research, PO Box 40, Lincoln 7640, New Zealandc Boffa Miskell Ltd., PO Box 110, Christchurch 8140, New Zealandd Plant and Food Research, Private Bag 3230, Hamilton 3240, New Zealande Cawthron Institute, Private Bag 2, Nelson 7042, New Zealand

a r t i c l e i n f o

Article history:

Received 15 December 2011

Received in revised form

2 April 2012

Accepted 3 April 2012Available online 21 April 2012

Keywords:

Copepod

Estuarine

Pollution

Toxicity

New Zealand

a b s t r a c t

Worldwide, estuaries are under increasing pressure from numerous contaminants. This study aimed to

identify a suitable marine harpacticoid copepod species for toxicity testing of New Zealand estuaries.

Multiple aspects were considered for species selection and included: a broad regional distribution, ease

of culture, reproductive rate under laboratory conditions, sexual dimorphism, and sensitivity to

contaminants. Five species were evaluated and two (Robertsonia propinqua and Quinquelaophonte sp.)

were able to be cultured. The relative sensitivity of these copepods to three reference toxicants was

assessed by determining the medial lethal values following a 96 h exposure (96 h LC50) to these

toxicants in the aquatic phase. LC50 values for zinc, phenanthrene, and atrazine respectively were 2.0,

0.89, and 7.58 mg/L in R. propinqua and 0.64, 0.75, and 20.8 mg/L in Quinquelaophonte sp. After

evaluating all factors involved in choosing a bioassay species for New Zealand, Quinquelaophonte sp.

was selected as the most suitable bioassay species.

& 2012 Elsevier Inc. All rights reserved.

1. Introduction

Estuaries are productive ecosystems often impacted by humanactivity (Lotze et al., 2006), and the resulting degradation of theseenvironments has significant impacts on biodiversity and theecosystem services they provide (Costanza et al., 1997). Estuariesare unique in the fact that they are affected by marine, fresh-water, and terrestrial pollution sources including agriculturalrunoff, storm water, forestry, mining, land conversion, roadrunoff, sewage, and harbor activities (Carpenter et al., 1998).Consequently, estuaries are a major sink where toxicants mayaccumulate. Pollutants are often trapped within estuaries due tothe settling of particulates, partitioning from the water columninto sediments, and binding to floating particulate and organicmatter. The high ionic strengths of saline estuarine waters canpromote the ‘‘salting out’’ of hydrophobic organic chemicals fromthe water into sediment (Brunk et al., 1997; Chapman and Wang,2001; Droppo et al., 2001).

Monitoring the impacts of anthropogenic activities on estu-aries is crucial to prevent environmental and economic lossesresulting from the collapse or contamination of the ecosystem(Chapman and Wang, 2001; Lotze et al., 2006). Laboratory-basedbioassays are powerful tools for assessing and monitoring eco-system health, especially those that use key species naturallyinhabiting estuarine sediments and which expose these animalsto environmentally-realistic conditions (Chapman and Wang, 2001).In general, marine and estuarine harpacticoid copepods are advanta-geous for this purpose owing to their short life cycles (�28 day), highreproductive rates, and their ease of culture in the lab. These proper-ties are ideal for assessing sub-lethal effects of impaired reproduction,growth, and development (Kovatch et al., 1999; Chandler and Green,2001; Bejarano et al., 2004; Greenstein et al., 2008). Harpacticoidshave close association with contaminated sediments, which makesthem very susceptible to chronic exposure to pollutants (Coull andChandler, 1992; DiPinto and Coull, 1997). Several species of harpacti-coid copepods are currently being used in sediment bioassays aroundthe world, including Amphiascus tenuiremis, Microarthridon littorale,Nitocra spinipes, Tigriopus brevicornis, Tisbe battagliai (Forget et al.,1998; Kovatch et al., 1999; Hagopian-Schlekat et al., 2001; Bejaranoand Chandler, 2003; Perez-Landa and Simpson, 2011). While previouswork identified Robertsonia propinqua as a suitable bioassay species(Hack et al., 2008a,b) there is no information on the sensitivity orutility of other native New Zealand harpacticoids as bioassay

Contents lists available at SciVerse ScienceDirect

journal homepage: www.elsevier.com/locate/ecoenv

Ecotoxicology and Environmental Safety

0147-6513/$ - see front matter & 2012 Elsevier Inc. All rights reserved.

http://dx.doi.org/10.1016/j.ecoenv.2012.04.008

n Corresponding author. Fax: þ64 3 546 9464.

E-mail addresses: tristanjstringer@gmail.com (T.J. Stringer),

chris.glover@canterbury.ac.nz (C.N. Glover),

grant.northcott@plantandfood.co.nz (G.L. Northcott),

louis.tremblay@cawthron.org.nz (L.A. Tremblay).

Ecotoxicology and Environmental Safety 80 (2012) 363–371

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organisms, and a bioassay using a New Zealand species of marinecopepod has not been developed and fully validated.

Multiple factors need to be considered in choosing a bioassayspecies. These factors include ease of culture, reproductive rate inlaboratory culture, ease of distinguishing males and females (forassessment of gender-based toxic impacts), and sensitivity tocontaminants. In most estuaries multiple sediment-dwellingcopepod species will exist; however, there is no guarantee thatany will meet all the essential criteria for a bioassay organism.

The aim of this study was to initially characterize and identifya suitable species of endemic harpacticoid copepod for use inestuarine bioassays. This was accomplished by: (1) characterizingthe suitability of five commonly occurring New Zealand harpacti-coid copepods on the basis of factors such as ease of culture andreproduction rate, (2) using acute aquatic 96-h LC50s to determinethe relative sensitivity of the best-qualified species to keyreference contaminants, and (3) use this information to identifythe endemic species with the greatest utility as a bioassay speciesfor assessing estuarine sediment health in New Zealand.

2. Materials and methods

2.1. Test species

Five commonly occurring New Zealand native harpacticoid copepods were

identified as potential bioassay species: Amphiascoides sp., Parastenhelia megaros-

trum, Quinquelaophonte candelabrum, Quinquelaophonte sp. and R. propinqua. These

species were selected as they have wide geographic distributions across New

Zealand and are easily distinguishable from each other based on differences in

overall body shape and defining characteristics of the species based on species

descriptions in Wells et al. (1982) and Hammond (1973). Quinquelaophonte genus

is characterized by serrated body segments and Q. candelabrum is distinguishable

from the yet unnamed Quinquelaophonte sp. by caudal rami morphology

(pipette shaped in Q. candelabrum) (John Wells, personal communication,

Quinquelaophonte sp. currently being described). P. megarostrum has the defining

characteristic of a large protruding rostrum (Wells et al., 1982), R. propinqua by the

stout body shape with the prosome being the widest point, and abundant spinules

on the abdomen (Hammond, 1973) and finally Amphiascoides sp. has similar body

shape to R. propinqua only smaller, thinner, and without the ornamentation on the

abdomen. All species were identified by Prof. John Wells at the Victoria University,

Wellington.

Amphiascoides sp., Quinquelaophonte sp. and R. propinqua were sourced from

Portobello Bay, Otago, New Zealand (451500S 1701390E). P. megarostrum and

Q. candelabrum were sourced from Okains Bay, Canterbury, New Zealand

(451430S 1731030E). Copepods were isolated from collected surface sediments by

gentle sieving and retaining copepods on a 125 mm sieve before being drawn out

of detritus and large sediments by attraction to a light source. Copepods were then

collected by means of a glass Pasteur pipette and transferred to Petri dishes where

they were identified to species level when sufficient taxonomic information was

available, under a dissecting microscope (Leica M125). Sandy, silty sediments

were also collected at both locations for use as the culture substrates. Sediments

were cleaned and sterilized based on methods of Chandler and Green (1996). The

sediments were washed with fresh water, sieved through a 125 mm sieve, and

autoclaved at 121 1C for 15 min. Each copepod species was monocultured in a

laboratory culture system modified from the recirculating seawater system

designed by Chandler (1986). The system included a flow-through plastic aqua-

rium (12 cm�18 cm) containing approximately 1 cm of the o125 mm sieved and

sterilized sediments overlaid with 5 cm of artificial seawater (ASW; Red Seas

Seasalts, 30%) under a dripping flow of 5 mL/min. Culture inoculation consisted of

4100 adult individuals and cultures were monitored every three day for survival

and reproduction (presence of females bearing egg sacs, hereafter referred to as

‘‘gravid’’ females) for approximately four weeks until it was determined that the

species was taking to culture or not.

The culture system was maintained at 20 1C and at 30% ASW, with a

photoperiod of 12 h light: 12 h dark. Ammonia, nitrite, and nitrate were always

below detection limit (determined by commercial aquarium test kits) and pH was

maintained at 8.2; water quality was monitored twice a week. The copepods were

fed 40 mL of a concentrated mixed suspended algal diet of Chaetoceros muelleri,

Dunaliella tertiolecta and Isochrysis galbana (1:1:1) twice a week (Chandler and

Green, 1996). Algal stocks were cultured in F2 medium, which was sub-cultured

every 7–10 day (Jeffrey and LeRoi, 1997).

For the species that were able to be cultured, the culture density was

estimated by counting adults in each of four replicate core samples (0.78 cm2)

taken from random locations within the individual culture aquaria. Only adults

were counted, as this life stage is used in toxicity tests. Total adults, the proportion

of gravid females and reproductive rate (number of eggs per female�proportion

of gravid females) were then calculated.

2.2. 96 h Acute toxicity tests

Three reference contaminants possessing a diverse range of structures and

modes of action were chosen and included a heavy metal (zinc), a polyaromatic

hydrocarbon (PAH; phenanthrene) and a pesticide (atrazine). These contaminants

also provide the advantage of being commonly used as reference chemicals in a

range of ecotoxicity test procedures, thereby providing opportunity to compare

the performance of the copepod assay against other organisms/species used in

standardized toxicity assays. Aquatic 96 h LC50 tests (median lethal concentration

after 96 h exposure to waterborne toxicant) were used as a rapid and technically

more simplistic proxy to sediment exposure. In addition, sediment toxicity can be

heavily influenced by pore-water exposure, meaning that aquatic toxicity tests

have some relevance to sediment-dwelling organisms (Simpson, 2005; Simpson

and King, 2005; Hutchins et al., 2009).

Acute toxicity tests were performed with two candidate species, R. propinqua

and Quinquelaophonte sp., as these two species performed the best in laboratory

cultures. All of the LC50 tests used laboratory-cultured copepods, described above,

and followed the methods outlined by Green et al. (1993) with modifications as

noted below.

For each test, there were five geometrically-increasing concentrations (1.5 or

2� concentration increases) of the chemical with four replicates for each

treatment and control. Three of the replicates were used for biological enumera-

tion at the termination of the test; the fourth was used for water quality and

chemical analysis. Carrier controls were performed when a carrier was necessary

for spiking atrazine and phenanthrene in the treatment groups (see below). Test

vessels were acid washed (min. 24-h ten percent nitric acid soak) and triple

acetone rinsed 50 mL borosilicate-glass Erlenmeyer flasks (LabServ), and con-

tained 30 mL of aerated 30% ASW, and 30 adult copepods (fifteen males, fifteen

non-gravid females) that were loaded at test initiation via pipette. The test vessels

were loosely covered to prevent evaporation. The tests were static (no water

exchange) and run for a period of 96 h. The test vessels were incubated in an

environmental chamber at 2071 1C, and under a 12 h light: 12 h dark cycle.

Dissolved oxygen, salinity, and pH were measured with a multi-meter (WTW

Multi 350i) at initiation and termination of the test.

2.3. Toxicant concentrations

Target toxicant concentrations were achieved by spiking ASW with aliquots of

stock solutions. For zinc, a stock solution of ZnSO4 �7 H2O (BDH cat #10299

AnalaR) was prepared in deionized water (Millipore, 0.22 mm filtered), while

dimethyl sulfoxide (DMSO, BDH cat #103234L AnalaR) and acetone (Mallinkrodt

Nanograde) were used as organic solvent carriers for technical grade atrazine

(a certified standard of atrazine (498 percent purity) obtained from Dr. Ehren-

storfer GmbH) and phenanthrene (Sigma cat #77470, 97 percent), respectively.

The proportion of carrier solution in each test vessel was kept as low as possible to

minimize the introduction of artefacts into the experiments. The final proportions

of carrier solvent in the phenanthrene and atrazine test vessels were 0.1/1000

(v/v) and 0.5/1000 (v/v) respectively (ISO 14669:1999). Carrier-solvent-only

controls were included in each schedule of testing to assess the toxic effect

arising from exposure to the carrier solvents. There were solubility limits for the

phenanthrene and atrazine of 1.1 mg/L and 33 mg/L, respectively, and the upper

concentrations were deliberately kept below solubility limits in these tests to

prevent precipitation.

Water samples (10 mL) were collected at the initiation and termination of

each test to confirm the concentration of reference compound. The concentration

of zinc in water test samples was determined by diluting samples 20� with ten

percent nitric acid and analyzing the prepared solutions by Inductively Coupled

Plasma—Mass Spectrometry (ICP—MS Agilent 7500 cx). QA/QC procedures incor-

porated into the schedule of analysis included the analysis of replicate samples

and one spiked sample within each set of twenty samples. Recovery of the zinc

spikes were 108712.1 percent (n¼4) and the relative standard deviations (RSDs)

for the mean concentration of duplicate treatment samples ranged from 0.95 to

1.03 percent.

Each atrazine and phenanthrene test solution was spiked with simazine and

anthracene-d10 (CDN Isotopes) as a surrogate spike compound before extraction.

The concentration of simazine added to the aqueous atrazine solutions varied for

the different treatments concentrations: 0.05 mg/L for control treatments, 1 mg/L

for 2 mg/L treatments, 2 mg/L for 4 mg/L treatments, 4 mg/L for 8 mg/L treat-

ments, 8 mg/L for 16 mg/L treatments, and 16 mg/L for 33 mg/L treatments.

Anthracene-d10, prepared as an acetone solution, was added to the aqueous

phenanthrene solutions to provide a final concentration of 50 mg/L. Aqueous water

samples of phenanthrene and atrazine were stabilized by the addition of 2 mL and

5 mL of dichloromethane, respectively, and stored under refrigeration prior to

extraction and analysis. Phenanthrene, atrazine and their respective surrogates

were extracted from the aqueous test solutions by liquid–liquid extraction using

T.J. Stringer et al. / Ecotoxicology and Environmental Safety 80 (2012) 363–371364

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dichloromethane, and analyzed by high resolution gas chromatography with mass

spectrometric detection for phenanthrene and electron capture (ECD) and

nitrogen–phosphorous detector (NPD) for atrazine. The 95 percent confidence

intervals for the recovery of anthracene-d10 and simazine were 10173 (n¼82)

and 10974 (n¼84) respectively. The reproducibility of analyses was excellent

with RSDs for the mean concentration of triplicate treatment solutions ranging

from 0.7 percent to 5.1 percent for phenanthrene and from 4 percent to 22 percent

for atrazine, across the range of treatment concentrations. The excellent surrogate

spike recovery and reproducibility data demonstrate the robustness of the sample

analysis methods and the concentrations of atrazine and phenanthrene deter-

mined in the aqueous test solutions.

2.4. Toxicological and statistical analysis

At test completion the numbers of surviving individuals were enumerated and

the actual concentrations of the contaminants were used to generate dose

response curves and estimate LC10, LC20, and LC50 values. The proportion of

surviving individuals was regressed against log concentration using binomial

regression. LC10, LC20, and LC50 values along with 95 percent confidence intervals

were calculated using the FIELLER procedure (a generalized linear model with a

binomial distribution and a logit link function) in the GenStat Statistical Package

(GenStat Committee, 2009) for each species (males and females combined as well

as gender-specific). Differences in species responses to concentration were tested

using a parallel slope model of the log proportion dead vs. log concentration and

examining the intercepts and the slopes. Slope and intercept estimates were

divided by the standard error of the differences and tested by t-test. Significant

difference was established at the value of Po0.05. Analyses were calculated using

the GenStat Statistical Package. Significant (Po0.05) differences in culture

density, egg number, and reproductive rate were determined using two-tailed

t-tests (in the GraphPad Prism five statistical package).

3. Results

3.1. Performance of all species in culture

Of the five species identified as potential bioassay species onlyR. propinqua and Quinquelaophonte sp. were able to reach den-sities in the lab cultures sufficient to support toxicity testing. Theother three species, Amphiascoides sp., P. megarostrum andQ. candelabrum, did not survive well in culture and populationsdied out after 3–4 weeks. Quinquelaophonte sp. produced thehighest average density of 88.5743.1 (standard deviation) a-dults/cm2, and R. propinqua had an average density of17.277.6 adults/cm2. The average egg number for each gravidfemale was similar at 15.672.5 eggs/female for R. propinqua and16.672.4 eggs/female for Quinquelaophonte sp. The percentage of

gravid females in cultures of R. propinqua was 11.6 percent andfor Quinquelaophonte sp. was 26.9 percent. Based on egg numberper female and the percentage of gravid females the calculatedreproduction rates were 1.81 and 4.47 for R. propinqua andQuinquelaophonte sp. respectively. As with all members of theorder Harpacticoida, both R. propinqua and Quinquelaophonte sp.are sexually dimorphic, and males are easily distinguished fromfemales by the enlargement of the fourth segment of the anten-nules as well as males being smaller in size, especially for R.

propinqua (see Fig. 1). These obvious morphological differencesbetween sexes are crucial for ease in distinguishing males andfemales, as specific numbers of each are required in toxicitytesting and for calculating sex-specific survival.

3.2. Toxicity results

In all tests water quality parameters were maintained withinoptimal ranges (salinity 30%, 47 mg/L DO, pH 8.0–8.5), and nitrate,nitrite, and ammonia concentrations were always below the methoddetection limits of 2.5, 0.05, and 0.25 ppm respectively (RedSeas

Marine Lab). Dissolved contaminant concentrations decreased dur-ing the tests, presumably due to combined processes of degradation,volatilisation (organic reference compounds), and absorption to thesurface of the glass test vessels. The latter is particularly importantin the case of zinc (Batley and Gardner, 1977) but is also relevant forphenanthrene. To account for the decrease in concentrations, theaverages of the initial and final concentrations were used as theexposure concentrations. Reference toxicant concentrations werereduced by an average of 13.1 percent, 11.9 percent and 9.2 percentfor zinc, phenanthrene and atrazine, respectively. In all tests themortality of test animals in control treatments was less than tenpercent, in accordance with ASTM (2008) recommendations.

Toxicity data (pooled data for both sexes) for both species areshown in Table 1 and dose response curves are shown in Fig. 2.LC50 values for R. propinqua were 2.0 (1.56–2.25) mg/L forzinc, 0.89 (0.85–0.95) mg/L for phenanthrene, and 7.58(6.12–9.39) mg/l for atrazine. The corresponding LC50 values forQuinquelaophonte sp. were 0.64 (0.51–0.82), 0.75 (0.59–1.37), and20.8 (17.9–25.2) mg/L for zinc, phenanthrene, and atrazinerespectively. Despite the fact that 100 percent mortality was notachieved in all tests, sufficient data were available for thecalculation of LC50 values.

Fig. 1. Dimorphism observed in male and female. (A) Robertsonia propinqua and (B) Quinquelaophonte sp. harpacticoid copepods (Leica M125 microscope).

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3.3. Comparative toxicity

Some significant differences were observed between the sensitiv-ities to the reference compounds exhibited by R. propinqua andQuinquelaophonte sp. While both copepods exhibited no statisticallysignificant difference in sensitivity to the PAH phenanthrene (Fig. 2B)

this was not the case for the heavy metal zinc and triazine herbicideatrazine. R. propinqua and Quinquelaophonte sp. exhibited significantlydifferent mortality responses to zinc (differences in intercept (diffintercept)¼3.865, SEdiff¼0.468, z¼8.26, Po0.001), with R. propin-

qua having an LC50 3.1 times higher than that of Quinquelaophonte sp.(Fig. 2A). R. propinqua and Quinquelaophonte sp. also exhibited distinctmortality responses following atrazine exposure as demonstrated byvalues for both intercept (diff intercept¼3.513, SEdiff¼0.718,z¼4.50, Po0.001) and slope (diff slope¼4.94, SEdiff¼0.199,z¼2.48, P¼0.02), corresponding with Quinquelaophonte sp. havingan LC50 2.7 times higher than that of R. propinqua (Fig. 2C).

3.4. Gender-specific toxicity

R. propinqua exhibited no gender-specific differences insensitivity to any of the reference compounds but there weresignificant differences between male and female responses ofQuinquelaophonte sp. to zinc and atrazine. For zinc, males weresignificantly more sensitive than females as demonstrated bydifferences in slope (diff. intercept¼3.126, SEdiff¼0.570, z¼5.49,Po0.001) and with resulting LC50 values of 0.38 (0.32–0.46) mg/Lfor males and 1.1 (0.81–1.47) mg/L for females. Males of Quin-

quelaophonte sp. also exhibited increased sensitivity to atrazinecompared with females as demonstrated by differences in slope(diff slope¼1.126, SEdiff¼0.441, z¼0.255, P¼0.02) and calcu-lated LC50 values of 13.8 (10.8–16.2) mg/L for males and 44.0(29.2–126) mg/L for females (Fig. 3B). The sensitivity of Quinque-

laophonte sp. to phenanthrene was independent of sex (males:0.64 (0.51–1.21) mg/L, females 0.79 (0.63–1.65) mg/L).

4. Discussion

There is currently no copepod bioassay validated for the assess-ment of marine sediment toxicity in New Zealand. Preliminary

Table 1Ninety-six hour acute lethal concentrations (LCx) values (mg/L) for the two

harpacticoid copepod species Robertsonia propinqua and Quinquelaophonte sp.

LC10 (95 percent CI) LC20 (95 percent CI) LC50 (95 percent CI)

R. propinquaZinc 0.86 (0.37–1.27) 1.16 (0.62–1.62) 2.0 (1.56–2.25)

Phenanthrene 0.43 (0.38–0.47) 0.53 (0.49–0.57) 0.89 (0.85–0.95)

Atrazine 1.4 (1.18–1.68) 2.65 (2.30–2.98) 7.58 (6.21–9.39)

Quinquelaophonte sp.

Zinc 0.24 (0.15–0.32) 0.34 (0.24–0.44) 0.64 (0.51–0.82)

Phenanthrene 0.28 (0.17–0.36) 0.41 (0.31–0.49) 0.75 (0.59–1.37)

Atrazine 7.11 (6.29–7.88) 10.6 (9.70–11.4) 20.8 (17.9–25.2)

Fig. 2. Dose–response curves for Quinquelaophonte sp. and R. propinqua to the

three reference contaminants. (A) Zinc, (B) phenanthrene, and (C) atrazine. LC50

values and 95 percent confidence intervals are also given. Phenanthrene and

atrazine both have solubility limits and are denoted by a dashed line.

Fig. 3. Sex specific dose response curves for Quinquelaophonte sp. to (A) zinc and

(B) atrazine. Dashed line denotes solubility limit for atrazine (B).

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studies have previously identified R. propinqua as a potentially usefulbioassay species (Hack et al., 2008a,b). However, the suitability ofother harpacticoid copepods has not been assessed. The ideal bioassayspecies is one that is easy to culture, has a high reproductive rate inlaboratory culture, has easily distinguishable sexes, is sensitive tocontaminants and is representative of species found in the wild.

4.1. Survivorship in laboratory cultures

Five species were assessed as potential bioassay organisms:P. megarostrum, Amphiascoides sp., R. propinqua, Q. candelabrum andQuinquelaophonte sp. Of these only R. propinqua and Quinquelao-

phonte sp. could successfully be cultured in the laboratory. Bothspecies have been associated with a wide spectrum of habitatconditions. The Quinquelaophonte genus is often found in detritusrich sediments (Hicks and Coull, 1983; Walker-Smith, 2004), and R.

propinqua is in muddy coastal marsh sediments (Soyer, 1970; Coullet al., 1979). Previous research by Stringer et al. (unpublished) hasshown that these two species have a preference for fine sandysediments in New Zealand. Failure of a given species to establishitself in culture may be related to changes in sediment character-istics owing to their processing and sterilization. Alternatively,copepods may have been unable to withstand the stress ofcollection and transportation. Laboratory culture, rather thanreliance on field-collected organisms, is essential for toxicitytesting as it allows for assessment on naive individuals and ensuresa more homogeneous genetic composition, which minimizesvariability and provides consistency within and between tests.The differences in sensitivity of field-collected and laboratory-reared animals have not been well studied. Some research showsthat seasonal and dietary influences can cause variability insensitivity (Sosnowski et al., 1979; Hedtke et al., 1986), promotingthe concept that controlled laboratory conditions are required forconsistency in toxicity assessments. Based on the findings ofculture survivorship, R. propinqua and Quinquelaophonte sp. wereassessed for their suitability as sediment toxicity organisms based

on the remaining criteria—sensitivity to contaminants, reproduc-tion rate, culture density, distribution and ecology.

4.2. Zinc toxicity

Zinc is one of the most problematic metals in estuaries aroundthe world and in New Zealand (Lotze et al., 2006; MfE, 1997,2007), and enters the environment through storm water, urbanrunoff and wastewater. In New Zealand zinc has been found inconcentrations above the regulatory guidelines in many areasreceiving urban stormwater inputs (e.g. the Tamaki Estuary inAuckland) (Abrahim and Parker, 2002).

Zinc proved to be the most toxic of the contaminants toQuinquelaophonte sp. and the second most toxic to R. propinqua,with LC50 values of 0.64 and 2.00 mg/L respectively. These valuesare well above the current marine water quality guideline forAustralia and New Zealand of 0.015 mg/L (95 percent speciesprotection, ANZECC, 2000). A comparison of the sensitivities ofthe two tested copepods to other estuarine and marine species isgiven in Table 2. Quinquelaophonte sp. is more sensitive to zincrelative to other evaluated copepod species. Other estuarinespecies have 96 h LC50 values ranging from 0.032 mg/L for theestuarine crab Scylla serrata to as high as 11.2 mg/L for themollusc Velesunio ambigua. Tolerance of R. propinqua to zinc wassignificantly higher than that of Quinquelaophonte sp., and issimilar to the estimated 96 h LC50 of 2.7 mg/L for adults and1.7 mg/L for nauplii (Hack et al., 2008b).

Zinc is known to have a range of impacts that contribute toacute toxicity. It has been shown to inhibit digestion and feeding inDaphnia magna and to interfere with exoskeleton maintenance andmolting in crustaceans (Poynton et al., 2007). This latter effect islikely due to the potential for zinc to share and inhibit calciumuptake pathways in crustacean epithelia (Ahearn et al., 1994).Copepods have defense mechanisms that help them regulateinternal metal concentrations and bioreactivity. One of these ismetallothionein, a protein that binds and detoxifies metals. Incopepods, as in other animals, the production of metallothioneins

Table 2Acute toxicity (96 h LC50) data for estuarine species to zinc.

Genus Species Life stage 96 h LC50 (mg/L) Reference

Copepod (Calanoid) Acartia clause Adult 0.96 Taylor (1981)

Acartia tonsa Adult 0.4 Taylor (1981)

Eurytemora affinis Adult 6.0 Taylor (1981)

Copepod (Harpacticoid) Amphiascus tenuiremis Adult 0.37 (pore-water) Hagopian-Schlekat et al. (2001)

Robertsonia propinqua Adult 2.0 (1.56–2.25) Present studyAdult 2.7a Hack et al. (2008a)

Nauplii 1.7a Hack et al. (2008a)

Tigriopus brevicornis Adult 0.715 Barka et al. (2001)

Tigriopus japonicas Adult 3.2 Taylor (1981)

Quinquelaophonte sp. Adult 0.64 (0.51–0.82) Present study

Amphipod Chaetocorophium cf. lucasi Juvenile 1.13 King et al. (2006)

Echinogammars olivii Adult 1.30 Bat et al. (1999)

Melita awa Juvenile 0.71 King et al. (2006)

Melita matilda Juvenile 0.65 King et al. (2006)

Melita plumulosa Juvenile 0.64 King et al. (2006)

Shrimp Palaemon elegans Adult 12.3 Bat et al. (1999)

Crab Pagurus longicarpus Larvae 0.6 Taylor (1981)

Paragrapsus quadridentatus Adult 14.0 Taylor (1981)

Larvae 1.3

Scylla serrata Adult 0.032 Narayanan et al. (1997)

Bivalve Dreissena polymorpha Adult 0.054 ANZECC (2000)

Mysella anomala Adult 4.40 King et al. (2004)

Soletellina alba Adult 2.90 King et al. (2004)

Velesunio ambigua Adult 11.2 ANZECC (2000)

a Range finder data, not definitive LC50.

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is induced in response to metal exposure (Barka et al., 2001;Poynton et al., 2007). The efficacy of metallothionein to detoxifytoxic metals is hypothesized to vary between species (Hook andFisher, 2002). Differential binding capacity of metallothioneinsmay be an explanation for the large variation in sensitivities ofestuarine species. This cellular mechanism may also explain theliterature data showing that copepods can have increased toleranceto metals when exposed over long time-frames (Moraitou-Apostolopoulou, 1978; Gardestrom et al., 2008), as metallothioneininduction is likely to promote survivorship. Critically, the impactsof cellular defense mechanisms such as metallothionein show howlife history affects sensitivity, and support the use of lab-rearedrather than field-collected individuals.

Another factor affecting sensitivity of copepods to zinc is thethickness of the exoskeleton. Juvenile copepods are more sensi-tive to metals than adults as a result of having a thinner bodycasing, which facilitates metal absorption (Hagopian-Schlekatet al., 2001). Although only adults were used in this study,thickness of exoskeleton may vary between species, potentiallyeffecting sensitivity. Zinc is also known to reduce chitinaseactivity and thus influence exoskeleton structural integrity(Poynton et al., 2007). These two mechanisms suggest theremay be a strong link between zinc toxicity and exoskeletonintegrity. Distinct physiological mechanisms associated withexoskeleton maintenance and development could be a contribut-ing factor explaining the differences in sensitivity to zinc betweenR. propinqua and Quinquelaophonte sp.

4.3. Phenanthrene toxicity

PAHs are some of the most common contaminants in aquaticsystems. They are also among the most toxic, displaying potentmutagenic and carcinogenic effects (Kennish, 1992). PAHs are avery diverse group of compounds structurally characterized by anumber of fused aromatic rings, with the majority detected inestuaries containing 2–5 aromatic rings. PAHs are most commonlysourced from combustion processes (Lima et al., 2005). Phenan-threne was selected as a reference PAH due to its relatively highsolubility and high toxicity and owing to its use in previous toxicitystudies with copepods and other estuarine organisms.

On exposure to waterborne phenanthrene, R. propinqua andQuinquelaophonte sp. exhibited 96 h LC50 values of 0.89 and0.75 mg/L respectively, a difference that was not statisticallysignificant. These two species of harpacticoid copepods areslightly less sensitive to phenanthrene than other copepod andestuarine species previously studied, with sensitivities varyingfrom 0.360 mg/L for the shrimp Palaemonetes pugio (60 h expo-sure) to 0.522 mg/L for the cyclopoid copepod Oithona davisae

(48 h exposure) (Table 3).Non-polar narcosis is often considered the major overt toxic

effect of PAH compounds. Non-polar narcosis can be defined asnon-specific disruption of the proper functioning of the cellmembrane, which leads to behaviors such as inhibited locomotor

activity and severely or totally inhibited capacity to respond tooutside stimuli (Ren, 2002). Bioaccumulation of PAHs can be veryrapid, reaching steady state in body tissues in less than 12 h inharpacticoid copepods (Lotufo, 1998a). Body lipid content is thebest predictor of PAH accumulation, which in turn is the bestindicator of toxicity (Lotufo, 1998b). The lipid contents ofR. propinqua and Quinquelaophonte sp. are unknown, althoughthe similarity of the species’ LC50 values suggests they may besimilar or that they have similar uptake rates.

4.4. Atrazine toxicity

Atrazine (2-chloro-4-ethylamino-6-isopropyl-amino-s-tirazine) isone of the most widely and intensively used herbicides in the world.It is applied in over 80 countries worldwide, and in the US alone, over35 million kilograms is added to agricultural land each year (Bejaranoand Chandler, 2003).

Both R. propinqua and Quinquelaophonte sp. were significantlymore tolerant to atrazine than other previously tested copepodsand estuarine species (Table 4), providing 96 h LC50 values of7.58 mg/L and 20.8 mg/L, respectively. The LC50 values of otherestuarine invertebrate species to atrazine range from as little as0.094 mg/L for the calanoid copepod Acartia tonsa, to 9.0 mg/L forthe shrimp P. pugio. In estuaries and coastal areas in the south-eastern US, environmental water concentrations of atrazine havebeen found to range from 90 ng/L to 62.5 mg/L (Bejarano andChandler, 2003). Atrazine concentrations in New Zealand estu-aries are not well characterized, but they are unlikely to exceedthe concentrations measured in south-eastern US waters. Conse-quently, acute effects of atrazine are unlikely to be observed forthese two species in the New Zealand environment.

In vertebrates, atrazine is an endocrine disrupting compound,with several modes of action suggested. It has been shown toincrease aromatase activity in humans (Sanderson et al., 2002); todisrupt the neuroendocrine control of ovarian function in femalerats (Cooper et al., 2007); to increase plasma testosterone in maleAtlantic salmon (Moore and Waring, 1998); and to induce her-maphroditism in male American leopard frogs at concentrations aslow as 0.1 mg/L (Hayes et al., 2003). As there is little known aboutinvertebrate endocrine systems it is unknown if atrazine effectssimilar pathways in copepods (Bejarano and Chandler, 2003).Further research to elucidate atrazine’s mode of action and howit affects acute and chronic toxicity is required. Such knowledgewould allow interpretation of the differential sensitivity to the twocopepod species evaluated in the present investigation.

4.5. Differences in species sensitivity

There were significant differences in the sensitivity of responseto the reference toxicants between the copepod species tested inthis study, and further differences in the sensitivity of responsereported for copepods in other studies. Some of these differencesin sensitivity may be attributed to the physical size of the tested

Table 3Acute toxicity data for estuarine species to phenanthrene.

Genus Species Life stage LC50 (mg/L) Reference

Copepod (Calanoid) Acartia tonsa Adult 0.422a Bellas and Thor (2007)

Copepod (Cyclopoid) Oithona davisae Adult 0.522a Barata et al. (2005)

Copepod (Harpacticoid) Robertsonia propinqua Adult 0.89 (0.85–0.95)b Present studyQuinquelaophonte sp. Adult 0.75 (0.59–1.37)b Present study

Shrimp Palaemonetes pugio Adult 0.360c Unger et al. (2007)

a 48 h LC50.b 96 h LC50.c 60 h LC50.

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species. Smaller species such as A. tenuiremis (0.25–0.44 mm)(Chandler and Green, 1996), generally exhibit higher sensitivity totoxicants than larger copepods such as R. propinqua (0.7–0.9 mm)and Quinquelaophonte sp. (0.6–0.8 mm). The larger surface area tovolume ratio of smaller animals means they are more susceptibleto exchanges with the environment, including the uptake oftoxicants. Furthermore, smaller animals have a reduced ‘‘buffer-ing capacity’’ to offset the impacts of bioaccumulated chemicals.It is noteworthy that the sizes of the two New Zealand speciestested in the present study were very similar, and is thus unlikelyto explain the differences between these two copepods in termsof their sensitivity to the reference toxicants.

In addition to size, there are other factors that can account fordifferences in the sensitivity of test species to toxicants. Thephysiology of different species may allow for one to be moretolerant to toxicants than the other. This may occur via processescausing an organism to metabolize a contaminant more effi-ciently or influence uptake and elimination of the contaminant,affecting bioaccumulation, and thus toxic effect. These differenceswill be contaminant-specific and will depend on the specificmode of action of a contaminant (Escher and Hermens, 2002;McClellan-Green et al., 2007). In this study Quinquelaophonte sp.was more sensitive to zinc than R. propinqua; however, theopposite was observed for atrazine. This exemplifies how dissim-ilar contaminants can affect species in different ways. In addition,there was a significant difference in the gender-associated toxi-city of Quinquelaophonte sp. to zinc and atrazine, which may bedue to the differences between the body size of males andfemales, uptake and elimination, or physiological differences(McClellan-Green et al., 2007). Interestingly, the same observa-tion was not seen in R. propinqua despite the size differencebetween genders in this species, suggesting that factors otherthan size difference are affecting gender-associated sensitivity.There is little information on the mechanisms that may beproducing these marked differences in sensitivity. Consequently,caution should be exhibited when choosing a bioassay speciesbased solely on its sensitivity to a single toxicant.

4.6. Bioassay species selection

Based on the results of acute waterborne toxicity tests therewas no strong toxicological argument favouring the selection ofeither R. propinqua or Quinquelaophonte sp. as the bioassayorganism-of-choice. Consequently, other considerations regardinglife history and ecology were considered to select the species bestsuited for use in a bioassay for New Zealand.

Both species reproduced well in culture with Quinquelaophonte

sp. having a 2.5 times greater reproductive rate than R. propinqua

(4.47 eggs/adult for Quinquelaophonte sp. and 1.81 eggs/adult for

R. propinqua). In addition, Quinquelaophonte sp. reached higherdensities during laboratory culture, producing an average densityof 88.5 adults/cm2 compared with 17.2 adults/cm2 for R. propin-

qua, providing an increased number of individuals for use inbioassays. Both species are sexually dimorphic, making it easy todistinguish males and females. Unfortunately, there is limitedecological and distribution information about these species,particularly for Quinquelaophonte sp. as it has only been recentlydiscovered and is currently being described (John Wells, personalcommunication). R. propinqua is usually found in tropical totemperate silty sands. However, there is evidence of an associa-tion with algae in shallow lagoons and it has often been found inplankton tows in the water column (Wells et al., 1982). Thepresence of R. propinqua in the water column is a potential issuewhen considering the use of this species for a sediment bioassay.It suggests this species could potentially avoid contaminatedsediments and produce misleading information regarding thetoxicity of sediment-borne toxicants. In contrast, observations ofQuinquelaophonte sp. suggest it is rarely seen swimming in thewater column and is predominantly a benthic species (Stringer,personal observations). This indicates Quinquelaophonte sp. couldbe better suited for use in sediment bioassays.

When considering the attributes required of an organism touse in sediment bioassays it is clear that Quinquelaophonte sp. isthe superior harpacticoid copepod species of those assessed inthis study (see Table 5). This is based on the observations that it issensitive to contaminants; it reproduces well in culture andreaches high densities; has a short life cycle; and its toxicresponses are likely to truly reflect sediment toxicity.

5. Conclusions

A bioassay species needs to be representative of the environ-ment under investigation, it should be among the most sensitiveof all organisms in that environment to concentrations of tox-icants that cause biological harm, and it needs to be amenable tolaboratory culture. This study showed that the harpacticoidcopepod Quinquelaophonte sp. is the best suited copepod for usein a bioassay to study the effects of toxicants in New Zealandestuaries. Quinquelaophonte sp. was chosen based on a detailedinvestigation of critical bioassay characteristics, including theorganism’s sensitivity to three representative reference toxicants.This study identified a potential bioassay species and also showedthat there can be significant differences in toxicological sensitivitybetween related animals, inhabiting similar habitats, and ofsimilar size. This shows the importance of conducting robustassessments of potential bioassay organisms prior to selection.Critically, this study has provided toxicant sensitivity data for a

Table 4Acute toxicity (96 h LC50) data for estuarine species to atrazine.

Genus Species Life stage 96 h LC50 (mg/L) Reference

Copepod (Calanoid) Acartia tonsa Adult 0.094 Ward and Ballantine (1985)

Copepod (Harpacticoid) Robertsonia propinqua Adult 7.58 (6.12–9.39) Present studyAdult 31.8a Hack et al. (2008a)

Nauplius 7.5a

Tigriopus brevicornis Adult 0.124 Forget et al. (1998)

Quinquelaophonte sp. Adult 20.8 (17.9–25.2) Present study

Shrimp Palaemonetes pugio Adult 9.0 Ward and Ballantine (1985)

Penaeus duorarum Adult 6.9 Ward and Ballantine (1985)

Crab Uca pugilator Adult 428 Eisler (1989)

Spot Leiostomus xanthurus Adult 8.5 Ward and Ballantine (1985)

a Range finder data, not definitive LC50.

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New Zealand estuarine invertebrate species that has potential tobe applied to the development of sediment and water qualityguidelines. Complementary evaluations examining the responsesof Quinquelaophonte sp. to chronic, sediment-borne exposures toreference contaminants are underway.

Acknowledgments

This work was supported by the New Zealand Ministry forScience and Innovation and the University of Canterbury DoctoralScholarship (TJS). We thank Professor John Wells for his invalu-able contribution to this study. We also thank Sally Gaw and RobStainthorpe (University of Canterbury) for assistance withmetal analysis; Guy Forrester (Landcare Research) for advice onstatistics; Katherine Trought (Landcare Research) for technicalassistance; and Christine Bezar (Landcare Research) for editorialcomments.

Appendix A. Supplementary materials

Supplementary data associated with this article can be foundin the online version at doi:10.1016/j.ecoenv.2012.04.008.

References

Abrahim, G., Parker, R., 2002. Heavy-metal contaminants in Tamaki Estuary:impact of city development and growth, Auckland, New Zealand. Environ.Geol. 42, 883–890.

Ahearn, G.A., Zhuang, Z., Duerr, J., Pennington, V., 1994. Role of the invertebrateelectrogenic 2Naþ/1Hþ antiporter in monovalent and divalent-cation trans-port. J. Exp. Biol. 196, 319–335.

ANZECC, 2000. Australian and New Zealand Guidelines for Fresh and MarineWaters. Australia and New Zealand Environment and Conservation Council,Melbourne.

ASTM, 2008. Standard Test Method for Measuring the Toxicity of Sediment-Associated Contaminants with Estuarine and Marine Invertebrates. ASTME1367-03(2008).

Barata, C., Calbet, A., Saiz, E., Ortiz, L., Bayona, J.M., 2005. Predicting single andmixture toxicity of petrogenic polycyclic aromatic hydrocarbons to thecopepod Oithona davisae. Environ. Toxicol. Chem. 24, 2992–2999.

Barka, S., Pavillon, J.F., Amiard, J.C., 2001. Influence of different essential and non-essential metals on MTLP levels in the copepod Tigriopus brevicornis. Comp.Biochem. Physiol. Part C: Toxicol. Pharmacol. 128, 479–493.

Bat, L., Gundogdu, A., Sezgin, M., Gonlugur, G., Akbukut, M., 1999. Acute toxicity ofzinc, copper and lead to three species of marine organisms from the SinopPeninsula, Black Sea. Turk. J. Biol. 23, 537–544.

Batley, G.E., Gardner, D., 1977. Sampling and storage of natural-waters for trace-metal analysis. Water Res. 11, 745–756.

Bejarano, A.C., Chandler, G.T., 2003. Reproductive and developmental effects ofatrazine on the estuarine meiobenthic copepod Amphiascus tenuiremis.Environ. Toxicol. Chem. 22, 3009–3016.

Bejarano, A.C., Maruya, K.A., Chandler, G.T., 2004. Toxicity assessment of sedi-ments associated with various land-uses in coastal South Carolina, USA, usinga meiobenthic copepod bioassay. Mar. Pollut. Bull. 49, 23–32.

Bellas, J., Thor, P., 2007. Effects of selected PAHs on reproduction and survival ofthe calanoid copepod Acartia tonsa. Ecotoxicology 16, 465–474.

Brunk, B.K., Jirka, G.H., Lion, L.W., 1997. Effects of salinity changes and theformation of dissolved organic matter coatings on the sorption of phenan-threne: implications for pollutant trapping in estuaries. Environ. Sci. Technol.31, 119–125.

Carpenter, S.R., Caraco, N.F., Correll, D.L., Howarth, R.W., Sharpley, A.N., Smith,V.H., 1998. Nonpoint pollution of surface waters with phosphorus andnitrogen. Ecol. Appl. 8, 559–568.

Chandler, G.T., 1986. High-density culture of meiobenthic harpacticoid copepodswithin a muddy sediment substrate. Can. J. Fish. Aquat. Sci. 43, 53–59.

Chandler, G.T., Green, A.S., 1996. A 14 day harpacticoid copepod reproductivebioassay for laboratory and field contaminated muddy sediments. In: Ostan-der, G. (Ed.), Techniques in Aquatic Toxicology. CRC Press, Boca Raton, FL,pp. 23–39.

Chandler, G.T., Green, A.S., 2001. Developmental stage-specific life-cycle bioassayfor assessment of sediment-associated toxicant effects on benthic copepodproduction. Environ. Toxicol. Chem. 20, 171–178.

Chapman, P.M., Wang, F.Y., 2001. Assessing sediment contamination in estuaries.Environ. Toxicol. Chem. 20, 3–22.

Cooper, R.L., Laws, S.C., Das, P.C., Narotsky, M.G., Goldman, J.M., Lee Tyrey, E.,Stoker, T.E., 2007. Atrazine and reproductive function: mode and mechanismof action studies. Birth Defects Res. Part B 80, 98–112.

Costanza, R., dArge, R., deGroot, R., Farber, S., Grasso, M., Hannon, B., Limburg, K.,Naeem, S., Oneill, R.V., Paruelo, J., Raskin, R.G., Sutton, P., vandenBelt, M., 1997.The value of the world’s ecosystem services and natural capital. Nature 387,253–260.

Coull, B.C., Bel,l, S.S., Savory, A.M., Dudley, B.W., 1979. Zonation of meiobenthiccopepods in a Southeastern United States salt marsh. Estuar. Coast. Mar. Sci. 9,181–188.

Coull, B.C., Chandler, G.T., 1992. Pollution and meiofauna—field, labratory, andmesocosm studies. Oceanogr. Mar. Biol. 30, 191–271.

DiPinto, L.M., Coull, B.C., 1997. Trophic transfer of sediment-associated poly-chlorinated biphenyls from meiobenthos to bottom-feeding fish. Environ. Sci.Technol. 16, 2568–2575.

Droppo, I.G., Lau, Y.L., Mitchell, C., 2001. The effect of depositional history oncontaminated bed sediment stability. Sci. Total Environ. 266, 7–13.

Eisler, R., 1989. Atrazine Hazards to Fish, Wildlife, and Invertebrates: A SynopticReview. U. S. Fish and Wildlife Service Biological Report.

Escher, B.I., Hermens, J.L.M., 2002. Modes of action in ecotoxicology: their role inbody burdens, species sensitivity, QSARs, and mixture effects. Environ. Sci.Technol. 36, 4201–4217.

Forget, J., Pavillon, J.F., Menasria, M.R., Bocquene, G., 1998. Mortality and LC50

values for several stages of the marine copepod Tigriopus brevicornis (Muller)exposed to the metals arsenic and cadmium and the pesticides atrazine,carbofuran, dichlorvos, and malathion. Ecotoxicol. Environ. Saf. 40, 239–244.

Gardestrom, J., Dahl, U., Kotsalainen, O., Maxson, A., Elfwing, T., Grahn, M.,Bengtsson, B.E., Breitholtz, M., 2008. Evidence of population genetic effectsof long-term exposure to contaminated sediments—a multi-endpoint studywith copepods. Aquat. Toxicol. 86, 426–436.

GenStat Committee, 2009. GenStat Reference Manual, Release 12—Part 3, Proce-dure LibraryPL20. VSN International, Oxford,.

Green, A.S., Chandler, G.T., Blood, E.R., 1993. Aqueous-phase, pore-water andsediment-phase cadmium—toxicity relationships for a meiobenthic copepod.Environ. Toxicol. Chem. 12, 1497–1506.

Greenstein, D., Bay, S., Anderson, B., Chandler, G.T., Farrar, J.D., Keppler, C., Phillips,B., Ringwood, A., Young, D., 2008. Comparison of methods for evaluating acuteand chronic toxicity in marine sediments. Environ. Toxicol. Chem. 27, 933–944.

Hack, L.A., Tremblay, L.A., Wratten, S.D., Forrester, G., Keesing, V., 2008a. Zincsulfate and atrazine toxicity to the marine harpacticoid copepod Robertsoniapropinqua. N. Z. J. Mar. Freshw. Res. 42, 93–98.

Hack, L.A., Tremblay, L.A., Wratten, S.D., Forrester, G., Keesing, V., 2008b. Toxicityof estuarine sediments using a full life-cycle bioassay with the marinecopepod Robertsonia propinqua. Ecotoxicol. Environ. Saf. 70, 469–474.

Hagopian-Schlekat, T., Chandler, G.T., Shaw, T.J., 2001. Acute toxicity of fivesediment-associated metals, individually and in a mixture, to the estuarinemeiobenthic harpacticoid copepod Amphiascus tenuiremis. Mar. Environ. Res.51, 247–264.

Hicks, G.R.F., Coull, B.C., 1983. The ecology of marine meiobenthic harpacticoidcopepods. Oceanogr. Mar. Biol. 21, 67–175.

Hammond, R., 1973. The Australian species of Robertsonia (Crustacea, Harpacti-coida), with a revised key to the genus. Rec. Aust. Mus. 28, 421.

Hayes, T., Haston, K., Tsui, M., Hoang, A., Haeffele, C., Vonk, A., 2003. Atrazine-induced hermaphroditism at 0.1 ppb in American leopard frogs (Rana pipiens):laboratory and field evidence. Environ. Health Perspect. 111, 568–575.

Hedtke, S.F., West, C.W., Allen, K.N., Norberg-King, T.J., Mount, D.I., 1986. Toxicityof pentachlorophenol to aquatic organisms under naturally varying andcontrolled environmental conditions. Environ. Toxicol. Chem. 5, 531–542.

Table 5Comparison of bioassay traits between R. propinqua and Quinquelaophonte sp. [(–) denotes no difference, (1) denotes the better suited species and (0) denotes the lesser

suited species].

Geo. dist. Life history Ease of culture Life-cycle Repro. rate Sexually dimorphic Sensitivity Total

Zinc Phenanthrene Atrazine

R. propinqua – 0 – 0 0 – 0 – 1 1

Quinquelaophonte sp. – 1 – 1 1 – 1 – 0 4

T.J. Stringer et al. / Ecotoxicology and Environmental Safety 80 (2012) 363–371370

Author's personal copy

Hook, S.E., Fisher, N.S., 2002. Relating the reproductive toxicity of five ingestedmetals in calanoid copepods with sulfur affinity. Mar. Environ. Res. 53,161–174.

Hutchins, C.M., Teasdale, P.R., Lee, S.Y., Simpson, S.L., 2009. Influence of sedimentmetal spiking procedures on copper bioavalibility and toxicity in the estuarinebivalve Indoaustriella lamprelli. Environ. Toxicol. Chem. 28, 1885–1892.

ISO (International Organization for Standardization), 1999. Water Quality—

Determination of Acute Lethal Toxicity to Marine Copepods (Copepoda, Crustacea).ISO 14669:1999.

Jeffrey, S.W., LeRoi, J.M., 1997. Simple procedures for growing SCOR referencemicroalgal cultures. In: Jeffrey, S.W., Mantoura, R.F.C., Wright, S.W. (Eds.),Phytoplankton Pigments in Oceanography; Monographs on OceanographicMethodology 10. UNESCO, France, pp. 181–205.

Kennish, M.J., 1992. Polynuclear aromatic hydrocarbons. In: Ecology of Estuaries:Anthroprogenic Effects. CRC Press, Bocca Raton, pp. 133–181.

King, C.K., Dowse, M.C., Simpson, S.L., Jolley, D.F., 2004. An assessment of fiveAustralian polychaetes and bivalves for use in whole-sediment toxicity tests:toxicity and accumulation of copper and zinc from water and sediment. Arch.Environ. Contam. Toxicol. 43, 314–323.

King, C.K., Gale, S.A., Hyne, R.V., Stauber, J.L., Simpson, S.L., Hickey, C.W., 2006.Sensitivities of Australian and New Zealand amphipods to copper and zinc inwaters and metal-spiked sediments. Chemosphere 63, 1466–1476.

Kovatch, C.E., Chandler, G.T., Coull, B.C., 1999. Utility of a full life-cycle copepodbioassay approach for assessment of sediment-associated contaminant mix-tures. Mar. Pollut. Bull. 38, 692–701.

Lima, A.L.C., Farrington, J.W., Reddy, C.M., 2005. Combustion-derived polycyclicaromatic hydrocarbons in the environment—a review. Env. Forens. 6,109–131.

Lotufo, G.R., 1998a. Bioaccumulation of sediment-associated fluoranthene inbenthic copepods: uptake, elimination and biotransformation. Aquat. Toxicol.44, 1–15.

Lotufo, G.R., 1998b. Lethal and sublethal toxicity of sediment-associated fluor-anthene to benthic copepods: application of the critical-body-residueapproach. Aquat. Toxicol. 44, 17–30.

Lotze, H.K., Lenihan, H.S., Bourque, B.J., Bradbury, R.H., Cooke, R.G., Kay, M.C.,Kidwell, S.M., Kirby, M.X., Peterson, C.H., Jackson, J.B.C., 2006. Depletion,degradation, and recovery potential of estuaries and coastal seas. Science312, 1806–1809.

McClellan-Green, P., Romano, J., Oberdorster, E., 2007. Does gender really matter incontaminant exposure? A case study using invertebrate models. Environ. Res.104, 183–191.

MfE, 1997. The State of New Zealand’s Environment. Ministry for the Environment,Wellington, New Zealand.

MfE, 2007. Environment New Zealand 2007. Ministry for the Environment, Well-ington, New Zealand.

Moore, A., Waring, C.P., 1998. Mechanistic effects of a triazine pesticide on reproductiveendocrine function in mature male Atlantic salmon (Salmo salar L.) parr. Pestic.Biochem. Physiol 62, 41–50.

Moraitou-Apostolopoulou, M., 1978. Acute toxicity of copper to a copepod. Mar.Pollut. Bull. 9, 278–280.

Narayanan, K., Lyla, P., Khan, S., 1997. Pattern of accumulation of heavy metals(mercury, cadmium and zinc) in the mud crab Scylla serrata. J. Ecotoxicol.Environ. Monit. 7, 191–195.

Perez-Landa, V., Simpson, S.L., 2011. A short life-cycle test with the epibenthiccopepod Nitocra spinipes for sediment toxicity assesment. Environ. Toxicol.Chem. 30, 1430–1439.

Poynton, H.C., Varshavsky, J.R., Chang, B., Cavigiolio, G., Chan, S., Holman, P.S.,Loguinov, A.V., Bauer, D.J., Komachi, K., Theil, E.C., Perkins, E.J., Hughes, O.,Vulpe, C.D., 2007. Daphnia magna ecotoxicogenomics provides mechanisticinsights into metal toxicity. Environ. Sci. Technol. 41, 1044–1050.

Ren, S., 2002. Predicting three narcosis mechanisms of aquatic toxicity. Toxicol.Lett. 133, 127–139.

Sanderson, J.T., Boerma, J., Lansbergen, G.W.A., van den Berg, M., 2002. Inductionand inhibition of aromatase (CYP19) activity by various classes of pesticides inH295R human adrenocortical carcinoma cells. Toxicol. Appl. Pharmacol. 182,44–54.

Simpson, S.L., 2005. Exposure-effect model for calculating copper effect concen-trations in Sediments with varying copper binding properties: a synthesis.Environ. Sci. Technol. 39, 7089–7096.

Simpson, S.L., King, C.K., 2005. Exposure-pathway models explain causality inwhole-sediment toxicity tests. Environ. Sci. Technol. 39, 837–843.

Sosnowski, S.L., Germond, D.J., Gentile, J.H., 1979. Effect of nutrition on theresponse of field populations of the calanoid copepod Acartia tonsa. WaterRes. 13, 449–452.

Soyer, J., 1970. Benthic bionomy of continental-shelf of French Catalan Coast. 3.Populations of harpacticoid copepods (Crustacea). Vie et Milieu SerieB—Oceanographie 21, 337–512.

Taylor, D., 1981. In: Williams, B.H.R. (Ed.), A Summary of the Data on the Toxicityof Various Materials to Aquatic Life. Imperial Chemical Industries Limited,Brixham, UK.

Unger, M.A., Newman, M.C., Vadas, G.G., 2007. Predicting survival of grass shrimp(Palaemonetes pugio) during ethylnaphthalene, dimethylnaphthalene, andphenanthrene exposures differing in concentration and duration. Environ.Toxicol. Chem. 26, 528–534.

Ward, G.S., Ballantine, L., 1985. Acute and chronic toxicity of atrazine to estuarinefauna. Estuaries 8, 22–27.

Walker-Smith, G.K., 2004. A new species of Quinquelaophonte (Crustacea: Cope-poda: Harpacticoida: Laophontidae) from Port Phillip Bay, Victoria, Australia.Mem. Mus. Vict. 61, 217–227.

Wells, J.B.J., Hicks, G.R.F., Coull, B.C., 1982. Common harpacticoid copepods fromNew Zealand harbours and estuaries. N. Z. J. Zool. 9, 151–184.

T.J. Stringer et al. / Ecotoxicology and Environmental Safety 80 (2012) 363–371 371