Methods of Culturing and Performing Toxicity Tests with the Australian cladoceran Ceriodaphnia dubia

Transcription

1 Methods of Culturing and Performing Toxicity Tests with the Australian cladoceran Ceriodaphnia dubia Cheryl Orr and Sharyn Foster CSIRO Land and Water Griffith September 1997 Technical Report 20/97

2 Table of Contents Page Number 1 Introduction 1 2 Washing containers 2 3 Methods of Culturing Mass Culture Culture Media Preparation DMW MQ S Supply Culture Food - Algae Stock Solutions Algae Media Plates Flasks Concentrating Algae Culture Food - YCT Feeding Transferring Culture Trays Initiating a new Culture Record Keeping 13 4 Methods for Conducting Acute Tests 13 5 Methods for Conducting Chronic Tests 14 6 Reference Toxicant Tests 18 7 Acknowledgments 18 8 References 19 i

3 1 Introduction This technical document details the methods used for culturing the cladoceran Ceriodaphia dubia, as well as its use in both acute and chronic toxicity tests at CSIRO Land and Water, Griffith. The methods used are adaptations of those of the USEPA (1991). The use of aquatic organisms to indicate toxicity is becoming more widespread because these organisms are often more sensitive to toxicants in water than chemical means of detection, and as such they give a biological perspective on possible pollution. Culturing of water fleas serves to provide a source of organisms for use in both acute and chronic toxicity tests. Ceriodaphnia dubia swim with an erratic jerking motion for a period of time and then remain still, hanging motionless in the water. They are filter feeders feeding on bacteria, detritus and algae. The life cycle consists of four distinct periods: egg, juvenile, adolescent and adult. The life span is highly variable and depends largely on temperature. Average life spans are 30 days at 25 o C and 50 days at 20 o C (USEPA, 1991). Reproduction typically begins with a clutch of eggs being released into the brood chamber. The eggs hatch in the brood chamber, and the juveniles, which are similar in form to the adult, are released when the female moults. Moulting involves the casting off of the exoskeleton or carapace. The time required to produce a first brood varies from 3 to 5 days and appears to be dependent on body size and environmental conditions. The growth rate of the organism is greatest during its juvenile stages with the body possible of doubling in size with each moult. Growth occurs immediately after each moult while the new carapace is elastic. 1

4 Following the juvenile stages comes a single adolescent instar. It is during this instar that the first clutch of eggs reaches full development in the ovary. Generally instar duration increases with age, but environmental conditions are also a factor. Four events take place in a matter of minutes at the end of each adult instar. They are: the release of young from the brood chamber to the outside, moulting, increase in size and the release of a new clutch of eggs into the brood chamber. The number of young per brood is highly variable and depends on body size, food availability and environmental conditions (USEPA, 1991). The production of males has been reported as resulting from environmental factors including low water temperatures, high population densities and a decrease in food availability (USEPA, 1991). Our data, from four years of culturing C. dubia under constant conditions, suggests that the production of males is unpredictable and can occur in any brood number (Anderson-Carnahan et al, 1995). Males (see back cover) can be distinguished from females (see front cover) by their slightly smaller size and lack of a brood chamber. Females grow to a length of approximately 1 mm (Anderson-Carnahan et al, 1995), with males achieving a length of approximately 0.8 mm. The original supply of organisms were provided by the Centre for Ecotoxicolgy (CET), Gore Hill, Sydney. This is an Australian cladoceran species and was collected from the Gibbon Pond at the Taronga Park Zoo, Sydney. 2 Washing Containers The following protocol should be followed for washing all glass and plasticware used for culturing and conducting acute and chronic tests with Ceriodaphnia dubia. Plastic cups used for housing test organisms are not washed, but used new and then discarded. 2

5 Washing procedure: a) Rinse in tap water b) Soak in detergent (scrub if needed) for at least a couple of hours, or overnight c) Soak in acid (10% HCl) for at least 1 hour, or overnight d) Rinse in Distilled water 3 times e) Rinse in Methanol if for pesticides f) Final rinse in MilliQ water. 3 Methods of Culturing 3.1 Mass Culture Mass cultures are used as a backup reservoir of organisms which are used to provide organisms to initiate cultures. Mass cultures are maintained in 1L beakers with the water to be used for culturing (see section 3.2 below) and a source of food (see sections 3.3 and 3.4 below). Culture water and food are replaced on a weekly basis providing an environment for large numbers of organisms to be produced unchecked. 3.2 Culture Media A volume of 15 ml of culture media is used for individual cultures. This volume is placed in a 60 ml plastic lily cup (Lily, N.Z). Each batch of water is analysed for ph, conductivity, Ca 2+ and Mg DMW MQ S Water The culture media or water used for culturing is prepared using a dilution of Perrier (Vergeze, France) mineral water and MilliQ water. This water source is used for its consistent quality over time and ease of preparation. To replicate the hardness of local waters, a soft water is used which requires the culture media to be a 10% dilution of Perrier. A volume of 20L is prepared in a plastic carboy by combining 2L of Perrier with 18L of MilliQ water. This water is aerated overnight prior to its use. This water is referred to as DMW MQ S (Diluted Mineral Water MilliQ Soft). 3

6 3.2.2 Supply Water As an alternative to DMW MQ S, we have access to local irrigation water which we collect from the main supply canal. This water is collected by lowering a stainless steel bucket attached to a rope into the overspill section of the canal. The water is filtered through a 125 µm mesh filter to exclude food and predators, into a 20L plastic carboy. We have had no success in culturing C. dubia in either local tap water or rain water. The presence of contaminants and a lack of nutrients are probably the reason for this. 3.3 Culture Food - Algae We use two species of green algae, Ankistrodesmus sp. and Raphidocellis subcapitata (formerly Selenastrum sp), as culture food, as these species are both easy to grow, and their cell size is suitable for ingestion by C. dubia. Whilst other species may be suitable for this purpose, these green algae appear to provide sufficient nourishment, in conjunction with YCT, for our test organism. Both species are cultured separately due to their differing growth rates and were supplied by CET Stock Solutions The following solutions are prepared in order to culture these algal species: Solution A 6.08 g MgCl.6H 2 O 2.20 g CaCl 2.2 H 2 O g NaNO 3 Combine these compounds and dilute to 500 ml with MilliQ water. Solution B 7.35 g MgSO 4.7 H 2 O dilute to 500 ml with MilliQ water. Solution C g K 2 HPO 4 dilute to 500 ml with MilliQ water. 4

7 Solution D 7.5 g NaHCO 3 dilute to 500 ml with MilliQ water. Solution E First prepare solutions ε1-ε5 as follows ε1 164 mg ZnCl diluted to 100 ml with MilliQ water ε mg CoCl 2.6H 2 0 diluted to 100 ml with MilliQ water. ε mg Na 2 MoO 4.2H 2 O diluted to 1L with MilliQ water. ε4a 60 mg CuCl 2.2H 2 O diluted to 1L with MilliQ water. ε4b 1 ml of ε4a is diluted to 10 ml with MilliQ water. ε mg Na 2 SeO 4 is diluted to 100 ml with MilliQ water. 1 ml of ε1, ε2, ε3, ε4b and ε5 to 92.8 mg H 2 BO mg MnCl 4.H mg FeCl 6.H 2 O 150 mg Na 2 EDTA this is diluted to 500 ml with MilliQ water Algal Media The algal culture solution is prepared by adding 1 ml of each of the stock solutions A, B, C, D and E to an erlenmeyer flask and diluting to 1L with MilliQ water. A bung of cotton wool is placed in the opening of the flask which is then autoclaved (Labec, Australia) to achieve sterilisation. All steps involving the transfer of algae are performed in a sterile cabinet. The cabinet (Biological Safety Cabinet Class II) is irradiated with UV light for at least 20 minutes prior to cabinet use, and the cabinet is wiped with 70% ethanol before and after use Plates Algae are cultured on agar plates to provide cells for addition to liquid media. Media plates are prepared from a 3% solution of agar diluted in algal culture media. This solution is autoclaved (Bench top Jaymac autoclave) for 20 minutes and then cooled to 5

8 about o C. It is poured, in a sterile cabinet, into plates and allowed to solidify with the lid only partially covering the plate. Upon cooling parafilm is placed around the plate to seal the base and lid together. Once prepared, plates are stored in a refrigerator until required. New culture plates are prepared from existing ones containing adequate cell growth. Inoculation involves flaming a loop to ensure it is sterile, and upon cooling scraping cells from an existing plate and redispersing them on a fresh culture plate. Inoculated plates are resealed with parafilm and placed in a controlled environment at 25 o C and a light intensity of 85 µmol m -2 s Flasks Cultures for use in cladoceran feeding are prepared in liquid culture media (section 3.3.2). A scrape from a plate culture is added to this liquid which is placed in an environmental chamber under the same conditions as previously described. This flask is mixed daily. After days the cultures should be ready to centrifuge (section 3.3.5) Concentrating Algae 50 ml centrifuge tubes are used to spin the algae with the aim of concentrating the cells to 3 x 10 7 cells ml -1 (USEPA, 1991). Both species are spun at 4,600 revolutions per minute for 10 minutes using a Heraeus Sepatech megafuge 1.0 (swing out rotor 3360, RCF3673g). The supernatant is decanted and stored, and the procedure is repeated until all has been spun. The concentrated algal cells are thoroughly mixed in a small amount of supernatant and pooled in a 50 ml measuring cylinder. The volume is recorded (for example 47 ml) and then a small magnetic mixer is added to stir the solution. 20 µl of algae is then diluted to 1 ml in a volumetric flask. After mixing this diluted solution is dispensed under the cover slip of a haemocytometer (see Figure 1) using a pasteur pipette. Individual algal cells are counted until a value of around 100 is achieved. The number of small squares required to achieve this value will vary depending upon initial inoculation size and culture performance. The number of squares is recorded. This is repeated until 6 counts have been made, with three on 6

9 each side of the haemocytometer. The mean and standard deviation are calculated. For example: 6 x 5 squares are counted to achieve the following values of cells 120, 123, 125, 121, 118, 119. The mean value is 121 and the standard deviation is 2.4. The value determined for the standard deviation should be no greater then 10 so extra squares may need to be counted if a higher value is obtained. From this mean value the following calculation is used to determine the number of cells present per ml: 121 (cell number) x 16 (total no. of small haemocytometer squares used for calibration) /5 (no. of small squares counted) x 3.968* (haemocytometer factor) x 50 (dilution) x 1000 (total cell volume) = 7.7 x 10 7 cells/ml * - The ratio of volume counted to the volume in which the units are expressed (in this case, 1 ml). From this value you need to determine what volume you need to dilute the algal concentrate to, to get a value of 3 x 10 7 cell/ml. This is found by dividing the known cells/ml by 3 x 10 7 and multiplying this by the volume of concentrate. eg. 7.7 x 10 7 cells/ml x 47 ml 3 x 10 7 cells/ml = 120 ml This volume is made up by the addition of some of the stored supernatant. Equal volumes of each concentrated algal species are combined and this solution is transferred to a labelled glass bottle and stored in the refrigerator until required. We have found that algal cultures can be stored under refrigeration for up to 1 month. 7

10 Figure 1. Diagram of a haemocytometer (Improved Neubauer - Weber England). The area pictured appears on both sides. 8

11 3.4 Culture Food - YCT YCT is the name given to a bacterial food which combines Yeast, Cerophyll (a U.S. brand name of alfalfa) and digested Trout pellets. This substance is prepared as follows: Five grams of trout pellets and 1L of Milli Q water are combined in a 1L measuring cylinder. This mixture is continuously stirred and aerated from the bottom of the container for 7 days. The digest is made up to 1L and placed in a refrigerator for 1 hour to settle prior to mixing with the other solutions. On day 6 of the digestion, 5 g of Cerophyll (dried powdered cereal leaves) is added to 1 L of MilliQ water and mixed using a magnetic stirrer overnight. This solution is settled for 1 hour prior to mixing with the other solutions. On day 7, 5 g of dry yeast is added to 1 L of MilliQ water and mixed immediately on a magnetic stirrer plate. Solutions are then mixed in a ratio of 1:1:1 after filtration through a fine Nitex 110 mesh. This mixture is placed into small containers and frozen until required. Trout pellets were provided by NSW Fisheries at Narrandera. Yeast (Fleischmann s) and dried powdered cereal leaves (Cerophyll substitute) were purchased from a local health food shop. 3.5 Feeding Cultures are fed daily to maintain the organisms in optimum condition. Individual cultures are fed at a rate of 100 µl YCT and 100 µl algae mix per 15 ml of culture media. Food should be at room temperature and thoroughly mixed prior to dispensing using a multi pipetter (eppendorf). 3.6 Transferring Individual cladocerans are transferred to fresh culture media every second day (days designated T on Figure 2). Transfers and feeding should be done at approximately the same time each day (eg. 9 am m 2 h). This is achieved using a shortened glass 9

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13 pasteur pipette approximately 4mm bore. Organisms are slowly drawn into the pipette with a small volume of media. When releasing the organism, the tip of the pipette remains under the media surface. Liquid can be poured from vessel to vessel containing the organism without fear of damage. 3.7 Culture Trays Cultures are housed in styrofoam trays with chip board backing. The chip board gives the tray stability, and the styrofoam provides a cheap, light weight, insulated material that is easy to work with. Holes are drilled in each tray to house 30 lily cups. Figure 3 presents an example of such trays. Due to their use in combination in chronic tests (see section 4) the trays have two different numbering systems (Figure 3). 3.8 Initiating a new culture New cultures are initiated from 3 rd or subsequent brood organisms that are < 24 h old in an existing culture. The performance of the adult providing the young should be examined when choosing organisms. As a guide, good brood sizes are: 1 st 3-6 young 2 nd 6-10 young 3 rd > 8 young Young from the same parent are set up in a culture board vertical row (Figure 1) requiring 5 young from each parent. This requires the young from 6 adults to be used to fill a culture. The source of young used in each new culture is recorded. Cultures are covered with a sheet of glass to prevent dust entering and to reduce evaporation. Glass sheets are wiped with methanol and dried prior to use. They are placed in an environmental chamber (Labec, Australia) with the following settings: Temperature 25 o C Light intensity 8-12 µmol m -2 s -1 Photoperiod 16:8 L:D The page number of the culture book is used to identify each culture and is recorded on the glass cover that has been placed on each tray. 11

14 Figure 3. Example of the trays used to house Ceriodaphnia dubia cultures and tests. Trays are made from styrofoam with a backing of chip board. TOP BOTTOM 12

15 3.9 Record Keeping Information describing the heritage of each culture is accurately recorded in the culture book. Figure 2 is an example of the information recorded for each culture. On a daily basis the culture is checked for survival of the adult organisms and any birth of young. The importance of good record keeping cannot be over emphasised. This information is recorded using the following symbols (see Figure 2 as an example): adult is alive adult is alive with alive young adult is alive with dead young adult is dead adult is dead and brood is alive adult is dead and brood is dead male female eggs Any other relevant observations should also be recorded. 4 Methods for Conducting Acute Tests Acute tests are performed over a 24 or 48 h period. These tests provide EC50 information on toxicants. Usually 5 concentrations of toxicant and a control source is used in these tests. 5 replicates of each concentration are used and these cups are randomly distributed in a tray using a template. The template lists the numbers 1 through 6 randomly along each of 5 rows. Test concentrations are then allocated a number, and distributed accordingly. Test volume is 15 ml and organisms are not fed during an EC50 test. For a single test, 150 < 24h old neonates of a 3 rd or subsequent brood are required. These neonates are pooled together from cultures. 5 neonates are placed in each cup making sure that the pipette is rinsed between each addition to 13

16 avoid possible carry over of toxicant. A glass cover is placed on the top and the tray is placed in an environmental chamber at the same conditions as previously described (section 3.8). Figure 4 presents an example of the information recorded at the beginning of each test. The number of organisms alive and dead at both 24 and 48 hours is also recorded. These figures are tallied to express the number of deaths per 25 organisms. This information is entered into an EC50 computer program which determines the EC50 value as well as the upper and lower confidence intervals of this estimate. This software uses the trimmed Spearman-Karber method for the determination of the EC50, and was provided to us by the USEPA. 5 Methods for Conducting Chronic Tests Chronic tests are performed over a 7 to 8 day period to determine possible sublethal toxic effects to organisms over part of their lifespan. Chronic tests require 60 < 24h old neonates with 6 neonates provided from each of 10 adults. 5 test concentrations and a control are used with 10 replicates of each concentration which are randomly arranged in 2 trays using a template. 15 ml of solution is used in each 60 ml cup and the organisms are fed 100 µl YCT and 100 µl of algal mix daily. At the same time each day, organisms are transferred to fresh solutions. The young from each adult fill all the cups in a horizontal row. This is known as blocking by parent, and ensures that the young from the same adult are exposed to each treatment. Trays are covered with a glass sheet and placed in an environmental chamber. Figure 5 presents the table used to record the performance of the test, with the same symbols being used to describe what is observed. Figure 6 presents the form which is filled out to provide the other information required about the test. The test continues until 60% of surviving control organisms have produced at least three broods of young. To be a valid test, at completion time <20% of control organisms will have died. At test completion the survival and number of young produced is summarised as presented in Figure 7. This information is entered into the computer program TOXSTAT to determine by ANOVA (Analysis of Variance) at which concentration no effect was observed (NOEL, No Observed Effect Level) and the lowest concentration at which an effect was observed (LOEL, Lowest Observed Effect Level). The TOXSTAT package was provided by the USEPA. 14

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18 Figure 5. An example of a chronic test data sheet detailing the performance of each individual in the test a a a a a a 2 a a a a a a 3 a a a a a a 4 a a5 a5 a4 a4 a 5 a4 a9 a2 a10 a9 a6 6 a a a a a a9 7 a4 a12 a10 a14 a12 a 8 a a15 a13 a15 a16 a a a a a a a 2 a a a a a a 3 a a a a a a 4 a4 a4 a4 a3 a2 a4 5 a9 a7 a8 a7 a6 a11 6 a a9 a a a a 7 a15 a a10 a11 a7 a13 8 a a12 a14 a16 a6 a a a a a a a 2 a a a a a a 3 a a a a a a 4 a3 a4 a5 a4 a3 a 5 a8 a10 a9 a7 a8 a5 6 a a a a a a6 7 a12 a11 a14 a10 a11 a1 8 a13 a12 a16 a13 a14 a a a a a a a 2 a a a a a a 3 a a a a2 a4 a 4 a4 a3 a9 a a a 5 a9 a7 a13 a10 a10 a3 6 a a12 a a14 a13 a7 7 a11 a a13 a a a 8 a13 a13 a a14 a12 a a a a a a a 2 a a a a a a 3 a a a a a a 4 a3 a4 a4 a a2 a4 5 a9 a8 a9 a a9 a8 6 a a a a a a9 7 a10 a9 a15 a4 a11 a 8 a7 a16 a15 a5 a11 x a a a x a a 2 a a a a a 3 a a a a a 4 a4 a4 a4 a a4 5 a8 a8 a9 x6 a5 6 a a a a6 7 a10 a13 a14 a 8 a10 a12 a16 a a a a a a a 2 a a a a a a 3 a a a a a a 4 a4 a a4 a4 a4 a4 5 a7 a6 a8 a8 a9 a8 6 a a a a a a 7 a14 a8 a13 a10 a7 a7 8 a13 a9 a11 a12 a8 a a a a a x a 2 a a a a a 3 a a a a a 4 a a4 a4 a4 a5 5 a4 a8 a9 a7 a9 6 a11 a a a a 7 a a11 a14 a11 a14 8 a9 a14 a9 a11 a a a a a a a 2 a a a a a a 3 a a a a a a 4 a4 a4 a4 a a4 a 5 a a6 a6 a a9 a 6 a9 a a a a a6 7 a9 a13 a7 a7 a12 a9 8 a a x10 a8 a10 a a a a a a a 2 a a a a a a 3 a a a a a a4 4 a3 a4 a3 a5 a1 a4 5 a a a a9 x4 a10 6 a9 a9 a6 a a13 7 a7 a9 a9 a12 a 8 a a a a12 a7 16

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20 Figure 7. An example of a summary of results from a Chronic test. The number of young produced by each test organism during the test is recorded as well as the number of test organisms alive at the end of the test. Conc (mg/l) No. alive/ Reference Toxicant Tests On a monthly basis an acute and a chronic test should be performed with a reference toxicant to determine whether the sensitivity of the organism is changing. These tests are performed using the chemical sodium dodecyl sulphate (SDS). Acute and chronic tests are set up as described above and a running tally of results is maintained known as a control chart. Methods for constructing a control chart are described in Anderson- Carnahan, (1994). The concentrations of SDS we use in these tests is the same each month and is as follows: 0.0, 1.25, 2.5, 5.0, 10.0, and 20.0 mg/l. 7 Acknowledgements This work was initiated by visiting US Scientist Linda Anderson-Carnahan from the USEPA. Her contribution is gratefully acknowledged, as is the technical assistance of Praba Pathmananthan, Denise DePaoli, Sue Korth, Rhonda Smith, Gillian Napier and Geoff McCorkelle. The original supply of Ceriodaphnia dubia from the Centre for Environmental Toxicology, Sydney is gratefully acknowledged. Help with the preparation of this document and ongoing support from Kath Bowmer, Wolfgang Korth and Martin Thomas is greatly appreciated. The assistance of Jane Roberts and 18

21 Peter Fairweather with proof reading is also acknowledged. For providing us with photographs of C. dubia, we acknowledge Russ Shiel (MDFRC) and Gillian Napier. 8 References Anderson Carnahan, L. (1994) Development of Methods for Culturing and Conducting Aquatic Toxicity Tests with Australian Cladoceran Moina australiensis. CSIRO Division of Water Resources Seeking Solutions Water Resources Series No.13. Anderson-Carnahan, L., Foster, S., Thomas, M., Korth, W., and Bowmer, K.H. (1995) Selection of a suitable cladoceran species for toxicity testing in turbid water. Australian Journal of Ecology. 20, USEPA Methods for Measuring the Acute Toxicity of Effluents and Receiving Waters to Freshwater and Marine Organisms (forth edition). C.I. Weber, ed. Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency, Cincinnati, Ohio. EPA / 600 / 4-90 /

22 COVER Front: A female Ceriodaphnia dubia. Actual size, approximately 1 mm. Photograph by Russ Shiel. Back: A male Ceriodaphnia dubia. Actual size, approximately 0.8 mm. Photograph by Gillian Napier.