measure

This page describes how to euthanize, stain, and measure fish.

Euthanasia

This section explains how to euthanize threespine stickleback safely and effectively.

You will need the following materials:

  • MS-222 (Tricaine Methane Sulfonate)
  • Sodium bicarbonate (baking soda)
  • Forceps or long Tweezers
  • Spatula
  • 1000 ml glass beaker
  • pH test kit

Procedure:

  1. Make a desired volume of MS-222 solution in the beaker (concentration of 0.5g/L) and buffer the solution to pH of 7-7.5 with sodium bicarbonate (0.5-1g/L). Wear protective clothing, gloves, and goggles when handling MS-222 powder. If possible, work inside a fume hood to prepare solution
  2. Place fish in the solution until death is achieved. Verify the animal is dead before disposing or preserving the carcass by monitoring the absence of respiratory or opercular movement for at least 3 minutes.

Additional notes:

  • Dispose of MS-222 waste by flushing down the drain to a sanitary sewer with an excess of water.
  • If in a remote location where a sewer may not be readily available, further dilute the solution with water and dump wastes on land in a location away from water.
  • Do not discard MS-222 directly into surface water, storm water or catch basins.
  • MS-222 is a light sensitive chemical and should be stored in a dark container or in a cabinet/drawer.

Preserve fish

In 95% ethanol

The advantage of this method is preservation of DNA. However, formalin fixes the tissues best for measurement.

  • Euthanize fish with a lethal dose of MS-222 (see above).
  • Make sure that the fish are not too crowded in the jar or the water contents of the fish dilutes the ethanol.
  • Seal jar cap with parafilm to minimize evaporation.
  • Jars may need top-up every few years to compensate for evaporation.

In 10% formalin

Warning: It is difficult to extract and amplify DNA from fish preserved in formalin. It is best to set aside a sizable sample of tissue to preserve in 95% ethanol before the fish goes into formalin.

  • Euthanize fish with a lethal dose of MS-222 (see above).
  • Always use formalin a fume hood.
  • Dilute formalin to 10% corresponding to 3.7% formaldehyde (100% formalin is 37% formaldehyde).
  • Place fish in formalin solution. Make sure that the fish are not too crowded in the jar or the water contents of the fish dilutes the solution.
  • Allow the specimens to fix in the formalin for about two weeks before measuring.
  • For long term storage, seal jar cap with parafilm to minimize evaporation. Jars may need top-up every few years to compensate for evaporation.

Stain fish

Stain fish preserved in formalin with alizarin red

Warning: the procedures below will destroy DNA of the sample.

  1. Rinsing (1):
    Strain formalin into receptacle. Rinse fish in water for 24 hours. Dispose of the formalin safely.
  2. Staining with alizarin red:
    Add four pellets of KOH to 100ml of water to which you add enough alizarin red powder to turn the solution purple (“Welch’s Grape Juice” color). It should be just fine as long as the amount of KOH used is less than 0.5g per 100ml . Place the rinsed fish (pour rinse water through strainer) into this alazarin red solution. Stain for 24-48 hours (time may need to be adjusted depending on size of fish, etc.). Mix the vials of stained fish every few hours to maximize dye exposure.
  3. Rinsing (2):
    Strain KOH/alizarin red solution into receptacle. Rinse fish well with water, up to 24 hours. This will usually clear away any stain picked up by non-bony tissues. Dispose of the KOH/alizarin red solution safely.
  4. Storing:
    Store rinsed fish in 40% isopropyl alcohol. We usually get our isopropyl alcohol from the fish museum.

Fish are now ready to be measured or kept in storage. Seal the jars with parafilm to minimize evaporation. Check fluid levels in jars every year or so.

Stain fish preserved in ethanol with alizarin red

Warning: the procedures below will destroy DNA of the sample.

  1. Rinsing (1):
    The fish will need to be transferred gradually to a water-based solution, otherwise the little bodies fly apart. From 95% ethanol, transfer to 70% ethanol for 24 hrs; 50% ethanol for 24 hrs; 20% ethanol for 24 hrs; tap water 24 hrs.
  2. Formalin step:
    Transfer rinsed fish into 10% formalin for 48 hours. Always use a fume hood when pouring formalin.
  3. Rinsing (2):
    Pour out the formalin and rinse the fish in water for 24 hours.
  4. Staining with alizarin red:
    Add four pellets of KOH to 100ml of water to which you add enough alizarin red powder to turn the solution purple (“Welch’s Grape Juice” color). It should be just fine as long as the amount of KOH used is less than 0.5g per 100ml . Place the rinsed fish (pour rinse water through strainer) into this alizarin red solution. Stain for 24-48 hours (time may need to be adjusted depending on size of fish, etc.). Mix the vials of stained fish every few hours to maximize dye exposure.
  5. Rinsing (3):
    Strain KOH/alizarin red solution into receptacle. Rinse fish well with water, up to 24 hours. This will usually clear away any stain picked up by non-bony tissues. Dispose of the KOH/alizarin red solution safely.
  6. Storing:
    Store rinsed fish in 40% isopropyl alcohol. We usually get our isopropyl alcohol from the fish museum.

Fish are now ready to be measured or kept in storage. Seal the jars with parafilm to minimize evaporation. Check fluid levels in jars every year or so.

Staining armor on live adult fish using calcein

Provided by Pam Colosimo.
REAGENTS: Calcein (Sigma # C0875). Calcein is a fluorescein-iminodiacetic complex that fluoresces green when combined with calcium. Examination for fluorescence can be carried out under UV microscopy in the usual manner for green fluorescence. Use the Nikon dissecting microscope in the lab – it has a mercury bulb UV light source. The filters are the same for observing GFP.

  1. Make 10 mg/ml (100X) Calcein solution in dH20. Cover this solution with tin foil. I have been storing this at 4 C for a month and it is still fine.
  2. Place adult fish in a beaker of fish tank water. (We use .35% Instant Ocean formula in dH20.) Since calcein binds calcium phosphate, it is important not to use water that contains that salt). I put one fish in 100 ml of fish tank water, but you can probably fit up to 5 fish in there. Add 1 ml of the 100X solution.
  3. Cover beaker with foil and leave for 3-4 hours. If you are in a rush, the fish will be stained within 1 hour, but it might be difficult to see all of the plates, especially the most anterior plates. I left one fish swimming in the solution overnight and the staining looked great.

Measure fish

There is some variation in how these traits are measured, and it is best that all fish be measured by the same person to ensure consistency.

Basic measurements

Measuring shape using landmarks

  • To measure shape of stained fish, use the high-quality Nikon camera, which has a lens with a flat field. Camera settings are here
  • Most of the landmarks we use were described in the paper by Albert et al (2008). An illustration of the landmarks is here.
  • These landmarks were based on turn on those used by Walker (1997), illustrated here.
  • Caldecott & Adams (1998) used the following skull landmarks

Lipid extraction

These methods were contributed by Karl Heilbron, and are based on Post and Parkinson (2001, Ecology 82:1040–1051) and Folch et al., (1957, J. Biol. Chem. 266:497–509).

  • Dry the fish in a drying oven and then weigh.
  • Homogenize dried sample with a mortar and pestle, then transfer to the tared cap of a 50mL Falcon tube. Reweigh (some loss occurs during the homogenization process, ~10%).
  • Tip contents into labeled Falcon tube. Add 8mL methanol and 8mL chloroform.
  • Bring solution to a boil in a 61 deg C water bath (~5 minutes) (coat hanger contraptions hold tubes in the water bath). Let cool to room temperature in fume hood.
  • Top up to 25mL with chloroform and allow the sample to settle.
  • Filter solution through No. 1 Whatman filter paper into a separatory funnel. (Wash filter paper with 10mL 0.9% saline).
  • Cap and vigorously shake separatory funnel. Uncap and let sit for ~15+ minutes while phases separate. The longer you give them, the better. Drain bottom organic layer into a pre-weighed beaker. (Rinse separatory funnel between extractions using a scrubbing brush but no soap.)
  • Boil off organic layer with a hot plate set to 70 deg C. Allow beaker to cool to room temperature.
  • Weigh beaker containing lipids.