Artemia: Experimental Procedure 2016

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).

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. 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 (~36 - 48 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 10m = 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. You will need to visually monitor hatching to determine the best time to harvest.

13. Place a black plastic sheet around and over the hatching vessel, leaving a small hole, 50-70mm diameter at mid-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.

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.

 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 with 2 = high degree of swarming, 1 = small degree of swarming and 0 = random distribution (no swarming). Record the data.  

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

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 any equipment that can be recycled in the reply paid envelope and post back.  

26. Enter all your data (nauplii location in the petri dish, activity level and apparent survival % for each sample time) into the "Artemia Investigation Web Application".  

27. Using the class dataset provided graph:

a. Nauplii apparent mean survival % (Y-axis) against time (X-axis) for each salinity. Fit the best trendline possible to each treatment scatterplot (include equations and R2 values where possible) [hint: 2 point polynomials and exponential models probably will give the line of best fit]

b. Mean survival % (Y-axis) x mean activity score (X-axis) for all treatments combined. Fit a trendline (include the equation and R2 value).

c. Mean survival % (Y-axis) x mean distribution score (X-axis) for all treatments combined. Fit a trendline (include the equation and R2 value).

d. Mean survival % at 18 hours (Y-axis) versus salinity (X-axis). Add a best fit trend-line, equation and R2 value, and standard error of each mean value.

e. Mean vigour score at 18 hours (Y-axis) versus salinity (X-axis). Add a best fit trend-line, equation and R2 value, and standard error of each mean value.

f. Mean distribution score at 18 hours (Y-axis) versus salinity (X-axis). Add a best fit trend-line, equation and R2 value, and standard error of each mean value.

28. For the 18 hour % survival data perform a one-way analysis of variance (ANOVA) with post-hoc Tukey comparisons to determine any differences between salinities. 

29. For the 18 hour vigour score data perform a one-way analysis of variance (ANOVA) with post-hoc Tukey comparisons to determine any differences between salinities. 

30. For the 18 hour distribution score data perform a one-way analysis of variance (ANOVA) with post-hoc Tukey comparisons to determine any differences between salinities.   

31. Tabulate the ANOVA data to illustrate mean values and significance comparisons for each analysis into a single table. 

32. Write up a scientific style report according to the "Research Investigation Report Requirements" file and submit to the unit assessment folder.

Artemia: Research Investigation Report

Essential Report Components

Title:

The title should convey the key idea of your report or what the report will cover. Below your title you should include your name, student number, affiliations (Deakin University and the course you are enrolled in) and the date.

Abstract: 

A concise, informative overview of the research. In approximately 150 - 250 words you should briefly outline why the research was done, what the main findings were, what conclusions you came to and the implications in terms of Animal Physiology.

Introduction 

You should convey a general statement about the broad scientific context in which this research is most relevant. This should include some background information and reference to previous work to help illustrate the importance of this research. It should provide the experimental aim and objectives or hypothesis tested. In the context of the experiment you conducted, the metabolic response to extreme salinities is the focal point. Finally, you should identify the scope of the report (i.e. outline what the reader can expect to find in the report).

Materials and Methods

In this section you must describe how the experiment was performed and the conditions under which it was performed in such a way that it could be repeated by someone else. You must use photographs to help illustrate the procedure and they must be logically ordered within the text body. You will need to describe the techniques used to collect, collate, and analyse the observations that you made. DO NOT repeat verbatim the design schedule but rather summarise the essential information and include any deviations or improvisations that you made. REMEMBER that someone reading your description should feel confident to be able to repeat the experiment using the methods you describe.

Results

This section presents your observations and data from the experiment. You do not include the raw data here but rather collated class results in the form of tables, graphs and statistical summaries. The figures and tables should be 'stand-alone' , that is, they should be clearly labelled with all the information that the reader requires to interpret the results. Each figure/table should have a number and a title appropriately located. You do not go into the details of statistical analysis (if you think this is necessary add it as an appendix) but rather present the findings of this analysis in an acceptable form. You must not discuss the results but rather just present or describe them in this section.

Discussion

In this section you should refer back to your original aim and discuss whether or not it has been achieved. It is not a re-statement of the results but rather an interpretation thereof, i.e. it should answer the question what do the results mean? This is done by referring to the results and determining if they support or contradict the initial proposal/hypothesis. Once you have described and discussed the class data you might compare and contrast them with the results of other researchers experimenting in the same field, if available. You should recognise the limitations of this study and identify any areas where further research should be undertaken. Make sure the content is presented in a logical manner, that the depth and breadth of discussion is appropriate to your level of expertise and that you demonstrate evidence of original thought based on the evidence you have collected. In your discussion you must attempt to explain differences between salinities from metabolic/physiological perspectives; particularly in relation to energetics, osmoregulation and respiration (which are all key modules in this unit), including reference to relevant morphological features, and make comparison to other organisms where possible.

Assignment link -

https://www.dropbox.com/s/xu7siesomwbweq2/Artemia%20Research%20Assignment.rar?dl=0