HOW TO ASSESS THE HEALTH OF YOUR ABALONE
The history of each group of abalone being examined should be closely followed, particularly that information related to stocking rate, growth performance, survival rates, feeding activity, food conversion rate, grading history, parasite or pest burdens and treatment history. Any abnormalities in any of these areas should be noted when they are observed so that these data are available for future reference if required.
Basic water quality parameters such as temperature and dissolved oxygen should be regularly monitored. Other variables such as salinity, ammonia, nitrite and nitrate, CO2, water hardness, pH, and calcium levels should be monitored in recirculation systems. Ideally these records should be available for the entire history of each group of abalone being examined.
Each abalone should be examined for shell colour and growth increments Figure 1
, shell thickness and any shell erosion or deformities Figure 2
. Before removing the abalone from the water examine them closely. Look for the movements of the feeding apparatus of mudworms Figure 3
, Figure 4
or other parasites which may live within the shell.
A 2.5 meg video of mudworms can be found here
Examine the abalone closely for abnormal behaviour (e.g. crawling out of tanks which can indicate poor water quality). Detach the abalone from the tank, turn them over and observe the righting reflex. A loss of this righting reflex indicates the abalone are weak Figure 5
. Note any abnormal areas on the shell Figure 6
, unusual colouration Figure 7
or erosion of epithelia associated with bacterial infection Figure 8
. Note any other visible lesions on the data sheet
before proceeding further.
(for abalone up to 20 mm shell length. If larger call for advice)
1. Collect your abalone. Select abalone that are behaving abnormally, or are moribund or lethargic first, then collect the remainder randomly in equal numbers from the various tanks being examined. Dead abalone are of limited value and should be noted, but not processed.
2. Measure the abalones shell length (to the nearest 1 mm) Figure 9
, Figure 10
then examine its shell and foot for mudworm blisters and other gross signs of disease.
3. Use the scalpel to cut the foot muscle free from the shell. Once the viscera and foot are removed, cut the abalone longitudinally through the majority of the viscera around 1/3 from the center line cut 1, Figure 11
, and place the first longitudinal section cut side up into the cassette. Examine the inside of the shell for fungi.
4. Then cut transverse sections through the remaining viscera cuts 2 and 3, Figure 12
and place the remains into a histology cassette cut side up (Simport 492, or Simport 493 for very small specimens).
5. Close the cassette, snap it shut and place it into 10% seawater formalin (see Recipes
page), ensuring all cassettes are covered by the fixative. Ensure that the volume of fixative used is at least 5 times that of the tissues being fixed. If any grossly visible abnormalities are observed, make sure the abnormal tissues are included in the samples excised for histopathology.
6. Any remains of the diseased abalone examined can either be frozen at -20°C for short term storage, then disposed of in landfill, and/or buried or incinerated - do not put them back into the water.
(also see Recipes
1. Follow manufacturers instructions to make up thioglycollate (29.3 grams of powder to 1 litre of freshwater). Add 20 g of salt (NaCl) to each litre of thioglycollate when using freshwater.
2. Stir then sterilise thioglycollate and universal tubes in an autoclave or pressure cooker. Cool down to body temperature then add antibiotic/mycotic solution (50 ml for each 1 L of thioglycollate). Then dispense 10 ml aliquots into sterile universal tubes. Store the sterile tubes and media in the dark until use.
3. Place tissues of interest (mantle, digestive gland, gills, foot and kidney) into tube and incubate in dark at room temperature for 7 to 14 days. If you are sending the samples by mail to DigsFish Services for analysis, this is the time to send them.
4. Remove tissues from thioglycollate, place in petri dish and flood the tissues with dilute lugols iodine. Macerate the specimen to allow iodine to penetrate.
5. Examine the stained tissues using a dissecting microscope at 40 x magnification for blue or black spheres of Perkinsus
Bacteriology uses specialised techniques which should be conducted using methods recommended by the diagnostic laboratory which will undertake the sample screening. Contact DigsFish Services or the laboratory responsible for processing your samples for more information on their desired methods of sample collection.
Excise small sections (1-2 mm3) of digestive gland, gill, mantle, epipodium, kidney, gonad or any other organ of interest and place them in a 1.5 ml eppendorf tube containing 2.5% gluteraldehyde (see Recipes
page). Store them at 4°C for 24-48 hours. Rinse once in seawater filtered to 0.22 microns and send to DigsFish Services for processing.
Excise small sections (1-2 mm3) of digestive gland, gill, mantle, epipodium, kidney, gonad or any other organ of interest and place them in a 1.5 ml eppendorf tube containing 70% ethanol (see Recipes
page). Store them at 4°C for 24-48 hours, then send to DigsFish Services for processing.