Artemia Investigation Report

Artemia: Experimental Procedure 2017

Allow a period of approximately five days to complete this experiment at home.

1. Make a cyst hatching vessel that has a cylindroconical profile. This can easily be done

using a 1.25L water bottle and chopping the top off (keep the water).

2. Make 1 litre of seawater strength (30g/L) solution and add to the hatching vessel. If

natural clean seawater is available then use it in preference. If not then dissolve the

supplied sea salt in water (non-carbonated bottled, distilled or rain water) thoroughly

before using.

3. Place the hatching vessel in a warm location and allow at least an hour for the water to

reach about 28oC (a little warmer than normal room temperature). If you live in a cold

house you may need to purchase a heat-mat or an immersion heater in order for the

experiment to work. You probably won’t get any appreciable hatching if the

temperature is <20°C.

4. Place an air diffuser into the hatching vessel and set the air flow to very low - so there is

only mixing but no frothing or bubbles accumulating on the surface.

5. Add 2g of Artemia cysts (~250,000 cysts/gram) to the hatching vessel. Mix the cysts

thoroughly to break up any clumps and evenly distribute the cysts throughout the water

column.

6. Place a bright light (40 - 100watt or equivalent if using LED’s) over but slightly to one

side of the hatching vessel (incubator) immediately after adding the cysts. Any light

generated heat will help keep the culture warm (which is essential for successful

hatching).

7. About 6 hours after adding cysts check the hatching vessel is operating correctly and

wash cysts from the edge of the vessel using a pipette containing seawater. Thereafter

monitor it several times each day to ensure the temperature is correct (28oC is optimal,

to cold and they will not hatch) and the air flow is sufficient.

8. Close to the estimated time of peak hatch (~24 - 36 hours) label 9 plastic petri dishes

from 1-9 on the bottom with a permanent marker pen. Place the petri dishes with lids

on a bench or table that has a white/cream smooth texture. Ensure they are placed

somewhere where disturbances and potential negative influences (such as

noise/vibration; cooking smells, large variation in temperature, chemicals etc.) are

minimal or absent.

9. Make up a hypersaline (300g/L) stock sea salt solution by adding 30g of sea salt to 100ml

of water (bottled-non-chlorinated-non-carbonated; or rain or distilled) in a plastic

container. Dissolve the sea salt thoroughly before using. The saturation concentration of

sea salt is about 250g/L at 20°C so there may be some residue left after dissolving.

10. To create your salinity gradient begin by aerating the hypersaline stock solution (300g/L)

and 100ml of bottled water (in a second plastic container) for approximately 30 minutes.

Using the aerated stock solution, water and a syringe create a 10mL volume of each of

the nine salinity concentrations: 0, 15, 30, 60, 100, 150, 200, 250 and 300g/L (i.e., one

for each petri dish). To determine the proportional volumes of the hypersaline stock

solution and water to mix together to create this gradient, use the formula:

Stock volume (mls) = (test salinity/300) x 10; Distilled water (mls) = 10 – stock volume (mls)

e.g. Stock solution = (30/300) x 10ml = 1ml; Water = 10 – 1 = 9mls

NB: 30g/L (seawater strength) is the control treatment

11. After adding the 10ml volumes place lids on the petri dishes to prevent evaporation and

thus salinity changes.

12. After approximately 48 hours turn off the aerator and remove it from the hatching

vessel. If the cysts are incubated at low temperatures the hatching period will be longer

and more spread out or less synchronous or if too cold they probably won’t hatch at all.

You will need to visually monitor hatching to determine the best time to harvest but at

28°C should be about 24 – 36 hours.

13. Place a black plastic sheet around and over the hatching vessel, leaving a small hole, 50-

70mm diameter a few centimetres below the water level.

14. Place a strong light so it directs light towards the hole in the black plastic sheet

surrounding the incubator and leave for 15-30 minutes. This should cause the nauplii to

congregate toward the light.

15. Remove the black plastic, then harvest the nauplii using a plastic syringe (make sure the

volume collected is <10mls) and place them directly into a 20ml plastic bottle/vial.

NOTE: nauplii must be highly concentrated prior to harvest for this to succeed; also note

that you don’t need to harvest all the nauplii but you need a sufficient sample size for

salinity testing, several thousand will be ideal. Additionally, don’t put your hand in the

hatching medium.

16. Make up the volume in the 20ml vial/plastic bottle containing harvested nauplii to 10ml

using seawater (30g/L solution).

17. While thoroughly mixing the harvested nauplii, use a pipette to add 1 drop at a time into

each of 9 petri dishes sequentially (don't forget to take the lid off!). First add to dishes 1-

9 in order then reverse the order. Add a total of 5-8 drops to each petri dish (which will

be about 0.5ml of harvested nauplii). This procedure should ensure about the same

number of nauplii are added to each petri dish and that the salinity gradient will only be

slightly altered.

18. Immediately after adding the nauplii to your salinity gradient shine a bright light at an

angle to cast a shadow of the nauplii on the surface underneath the petri dish. Use a

magnifying glass to observe the activity level of the nauplii in the petri dishes. Using a

scale of 0-10 (0 = no activity at all; 10 = extremely active) assign an activity level to each

petri dish and record the data. Also estimate % survival in each petri dish. You will need

to patiently observe the nauplii to achieve this and have good lighting. NOTE: use the

control treatment (30g/L) as a benchmark at commencement (it should be 100% survival

and activity level of 10). You should have 100% survival in all treatment salinities at

commencement, time zero. Place the petri dish lids on after you have finished.

19. Immediately place a strong light towards one end of your petri dish array. Healthy

vigorous nauplii will swarm around the edge of the petri dish towards the light. Make a

note of whether they are concentrated towards the lit region of the petri dish or

dispersed evenly/randomly throughout the petri dish each time you observe them.

Assign categorical values to the swarming behaviour (or distribution in the petri dish)

with 2 = high degree of swarming, 1 = small degree of swarming and 0 = random

distribution (no swarming). Record the data. After recording this data you can turn off

the light source, ambient light will be sufficient until the next sampling event.

20. Check the petri dishes every hour for the first 10 hours, then at 8 hour intervals for 72

hours after commencement. Make sure you turn the light back on about 5-15 minutes

before making your observations. Check the nauplii location in relation to the light

source. Then gently swirl/mix the contents of the petri dish so as to randomly distribute

them, then observe their activity level and survival %. Record your results on a datasheet

of your design.

21. At 72 hours or earlier if 100% mortality is recorded in a petri dish, place a piece of graph

paper with 1mm x 1mm grids under the petri dish and take a digital photograph of the

petri dish so nauplii can subsequently be counted. This is quite challenging and you will

need good lighting and a reasonable quality digital camera to be successful. The count

data should be recorded as part of your experimental methods.

22. After taking the first digital photo of your petri dish containing 100% dead nauplii,

download it to a computer. Open the photograph in a suitable program such as Adobe

Photoshop. Magnify the image until you can see the nauplii (it doesn’t matter if they are

a bit blurry, you just need to be able to see them sufficiently well enough to count

them).

23. Scroll across the digital image of the nauplii and using an electronic marker of some kind,

mark each nauplii as you progressively count the entire sample. Marking nauplii during

counting will prevent counting individuals more than once.

24. If there are too many nauplii in your photograph to physically count determine the total

number of nauplii added to each petri dish by using appropriate scale-up factors (for

example, divide the petri dish area into four quadrats, count 1 and multiply by four).

25. At the completion of the test period, wash all the equipment in freshwater and return it

to your home campus (i.e., where you collected it from) so it can be recycled for

subsequent use.

26. Enter all your data (nauplii distribution in the petri dish, activity level and apparent

survival % for each sample time) into the “Artemia Investigation Web Application”

located on the SLE315 home page in cloud.

27. Using the class dataset provided graph:

a. Nauplii apparent mean survival % (Y-axis) against time (X-axis) for each salinity

(all salinities on the same graph). Add a smoothed line to each treatment.

b. Mean survival % (X-axis) x mean activity score (Y-axis) for all treatments

combined. Fit a trendline (include the equation and R2 value). (n= number of data

points = number of student replicates for each time and salinity combination)

c. Mean survival % (X-axis) x mean distribution score (Y-axis) for all treatments

combined. Fit a trendline (include the equation and R2 value). (n= number of data

points = number of student replicates for each time and salinity combination)

d. Mean survival % at 18 hours (X-axis) versus salinity (Y-axis). Add a best fit trend-

line, equation and R2 value, and standard error of each mean value.

e. Mean survival % at 18 hours (X-axis) versus vigour score (Y-axis). Add a best fit

trend-line, equation and R2 value, and standard error of each mean value.

f. Mean vigour score at 18 hours (X-axis) versus salinity (Y-axis). Add a best fit

trend-line, equation and R2 value, and standard error of each mean value.

g. Mean distribution score at 18 hours (X-axis) versus salinity (Y-axis). Add a best fit

trend-line, equation and R2 value, and standard error of each mean value.

h. Construct a time (hours) to 50% survival (Y-axis) versus salinity (X-axis) (i.e., an

LC-50) graph. Smooth a line through the data. Then add a best fit trend-line,

equation and R2 value.

28. Write up a scientific style report according to the “Research Investigation Report

Requirements” file and submit to the unit assessment folder.