stykylbak, sticklebanke, sticklebanck, stickle bag(ge),
stit(t)le bag(ge), stittle-back, stickle-back, stickleback
This page describes our methods for working with threespine
stickleback.
The three-spined stickleback (Gasterosteus aculeatus complex) is a
small fish common in many temperate coastal marine and fresh waters of
the northern hemisphere. Niko Tinbergen’s studies of the behavior of
this fish were important to the development of the field of ethology. It
has a fully sequenced genome and has become a model vertebrate for
studies in the fields of animal behavior, evolution, genetics, and
toxicology.
The photo above by N. Bedford shows a male of the benthic species
from Paxton Lake, Texada Island, BC. He is guarding a family of
stickleback fry that hatched a few days previously.
The threespine stickleback species complex reaches the height of its
diversity in lakes and streams of British Columbia. The populations
occurring here include some of the youngest species of organisms on
earth. Most water bodies contain only a single stickleback species, but
pairs of species have evolved independently in a few small lakes that
are less than 12000 years old. The system has wonderful properties that
allow us to address very basic questions concerning the roles of
resources, species interactions, phenotypic plasticity, sexual selection
and other factors in the evolution of diversity. Distinct populations
and species produce viable and fertile hybrids, making it possible to
investigate the genetic basis of species differences.
Research facilities
In addition to field sites, we conduct research on stickleback in an
indoor aquatic facility and outdoor experimental ponds.
Wet lab facilities
Our wet lab consists of two air-conditioned rooms in the InSEAS
aquatic facility (one with stand-alone aquariums and the other with a
flow-though aquarium system), plus controlled-temperature environment
chambers. These facilities contain hundreds of aquariums that currently
mainly house stickleback.
Experimental ponds
This facility includes 20 ponds, each 25 m x 15 m. An aerial view is
seen below. Click the image below to see a larger view in Google
Maps.
Each pond contains a shallow littoral area at one end and a 6 m deep
end. The littoral area contains a layer of sand and limestone gravel
extracted from surface mines near Paxton Lake, Texada Island. The ponds
have been seeded with plants and invertebrates from Paxton Lake, an
11-ha lake containing a benthic-limnetic stickleback species pair. Apart
from their construction and initialization, the ponds are unmanipulated
environments. We use them to carry out experiments on adaptation,
natural selection, and genetic mapping of natural variation in behavior
and other traits.
Mesocosms (cattle tanks)
The experimental pond facility includes 80 plastic rubbermaid
barrels, each 2 m in diameter, for experimental studies of aquatic
organisms.
Old experimental ponds
This outdoor facility is no more and has been replaced by the new
experimental ponds. It too was located on the South Campus of the
University of British Columbia.
The site contained 13 ponds. Each pond was 23 x 23 m2 with
a bottom that sloped gradually from 0 m at the edges to 3 m deep in the
center. The ponds were constructed in 1991 and seeded with plants and
invertebrates from Paxton Lake, an 11-ha lake containing a benthic and
limnetic stickleback species pair. The ponds were lined with
polyethylene overlaid with 0.25 m of sand, and were bordered with
limestone extracted from surface mines near Paxton Lake. Apart from
their construction, initialization, and use in prior experiments, the
ponds were unmanipulated environments.
How to raise stickleback
This section describes the basic procedures we use to make crosses
and raise stickleback.
Do not use water obtained from copper pipes, as the
copper ions kill stickleback.
Set up the aquarium in a cool room. 17 degrees C is perfect for our
local populations. Above 20 degrees is dangerous.
Standalone aquariums - McPhail Room
We use 100 L aquaria to keep and raise stickleback. The fish grow
well under these conditions provided densities are kept low. Standalone
tanks are stacked on shelves in the McPhail room of the InSEAS aquatic
facility. The greatest challenge is getting the nitrogen cycle going in
a new standalone tank.
All wild fish go into the McPhail Room – do not bring wild fish into
the Flow-through Room.
Initial setup
-
Dechlorinated tap water is fine. Add extra conditioning, such as Prime
or Amquel, as backup dechlorination in case of a spike in chlorine in
the water supply. Add baking soda if pH is below 7 (Vancouver water is
sometimes acidic).
-
To aid fish health, add 100-200 g of synthetic sea salt (e.g., Deep
Ocean Synthetic Sea Salts) to each 100 L aquarium. Add even more, 500 g,
if you are raising crosses between freshwater and Little Campbell River
marines (LCM), which tend to develop poorly at low salinity. This will
bring salt concentration up to about 5 ppt.
-
Add bicarbonate of soda to raise the pH to desired levels (see next
section for information on adjusting pH).
-
Add crushed coral or limestone to the tank. This keeps the pH high and
the water hard, which is best for minimizing disease. Many but not all
of our study populations live in hard water lakes.
-
One airstone per 100 L aquarium is fine (you don’t need to add one if
you are also using a sponge filter). Keep the flow light when the fish
have just hatched, because they are not strong swimmers.
-
Use sponge filters for babies and add power filters for juveniles and
adults. For small juveniles, put a sponge on the end of the intake pipe
to prevent sucking up fish. Use angel hair or double up the sponges in
the filter box for extra surface area and filtration of fine
particulates. The foam inserts of the power filters last a long time but
need periodic cleaning. Squeeze the sponge filters every two weeks or so
to keep the sponges from plugging up.
Start the nitrogen cycle
This is the greatest challenge when starting up a tank for fish. The
filters provide a surface for the bacteria that break down ammonia to
nitrite and then to nitrate. Ammonia and nitrite are toxic, whereas
nitrate is relatively harmless. However, it takes at least a month for
the bacteria to reach sufficient numbers to handle all the waste
produced by the fish.
To get the nitrogen cycle up and running:
-
Set up new tanks at least one month before introducing your valuable
fish.
-
While you wait, keep a goldfish, sculpin, or a very small number of
non-essential stickleback in the tank to help get the nitrogen cycle
going before you replace them with your valuable fish one month later.
-
Another technique, while you wait, is to add ammonia daily directly to a
fish-less tank and keep testing the water until nitrites drop to zero.
Your tank is ready to receive important fish.
-
If you don’t have the luxury of a one-month advance, but must throw your
fish straight into a new, sterile tank, then you will need to change the
water frequently to keep waste levels down for weeks while the nitrogen
cycle gets going. Add a filter sponge taken from a older tank whose
nitrogen cycle is functioning (beware of communicating disease). Try
replacing 1/3 to 1/2 of the water every two days for the first two weeks
at least. Add Amquel or other conditioner to help remove nitrite as well
as ammonia (not all products will do both). Make arrangements for water
changes on weekends if you are not around to do it yourself, especially
long weekends.
Manage the nitrogen cycle
-
Keep fish densities low, even after the nitrogen cycle has started to
work. A 100 L tank can handle about 15 adult fish when everything is
going well (maybe even 20 if you are not dealing with large benthic or
marine fish). If you go higher, be prepared for the higher risk. Go
lower if the fish are irreplaceable.
-
Remove dead fish immediately. Take special care on weekends, if you are
alone in the lab. You might be able to avert disaster this way.
-
Do not overfeed fish. If the fish food that drops to the bottom of the
tank isn’t eaten within a few minutes, siphon the excess away.
-
Test ammonia and nitrites regularly. See the next section below.
-
Apply regular water changes (see below).
-
If you are raising fish from eggs, you might end up with as many as 200
juveniles from one clutch in a single aquarium. This is unsustainable.
Families should be divided among multiple tanks as soon as the young
fish are large enough to tolerate a move.
-
Make sure your filters are working properly. Restart them immediately if
there has been a power outage.
Adjust ammonia and pH
These instructions are from the standard operating procedures,
SOP for adjusting ammonia and
the SOP for adjusting pH.
Ammonia. Test ammonia (NH3) using manufacturer’s instruction.
Ammonia levels should be at 0 ppm (mg/L) any level above zero can harm
fish. Ammonia builds up due to the breakdown of organic waste and the
nitrogen cycle and can be toxic. The presence of ammonia indicates
possible overfeeding, too many fish, or inadequate biological
filtration. If the ammonia test reveals ammonia levels above 0 ppm,
follow the instructions below:
-
Scrub off excess algae with a brush and perform a 25% water change.
-
Ensure tank is seeded/developed with nitrogenous bacteria to break down
ammonia.
-
Ensure that the filters are circulating water at a sufficiently high
rate.
-
If the tank has only a foam filter, add a higher-flow (box) filter.
-
Lower the density of fish in the tank to decrease the amount of ammonia
produced.
-
Test the water again after 24 hours.
pH. Test pH using manufacturer’s instruction. Stickleback from
most populations prefer pH between 7.4 - 7.6 (stickleback from limestone
lakes, such as Paxton, Priest, and Cranby on Texada Island, can tolerate
pH up to 8.3). If pH falls below the preferred range,
gradually
increase pH using one or more of the following methods.
-
Add limestone or crushed coral to the aquarium. The carbonate in these
compounds will react with and remove excess H+ ions in the water.
-
Crushed coral and limestone are advantageous as the release of carbonate
is governed by the surrounding water chemistry. As a result, crushed
coral and limestone will raise the pH to a maximum of 7.8 regardless of
the amount added. Crushed coral and limestone will develop a hard
surface layer and should be scrubbed once every 3 months to ensure
reactivity.
-
Add baking soda to the aquarium (gradually).
-
The direct addition of baking soda can raise the pH very quickly and it
is possible to overshoot the target pH. This can cause pH shock in the
fish. It is best to gradually add baking soda by dissolving 1 teaspoon
of baking soda in 1 cup of water for every 5 gallons of water in the
tank. Most of our tanks are 30 gallon. Add ¼ of the baking soda-water
mixture to the tank, wait 30 minutes or longer and add another ¼.
Continue this procedure until the mixture is used up. Wait 24 hours and
test the pH again.
-
Aeration can raise the pH slightly.
-
Aerate the water by adding an air stone to the fish tank or increasing
the flow of air. Excess CO2 dissolved in the water decreases pH (makes
it more acidic). As the tank water is exposed to atmospheric air through
aeration, the tendency of the dissolved gases to approach equilibrium
will transfer oxygen into the water and carbon dioxide out of the water,
resulting in a higher pH. This method will result in only a slight
change in pH. Use an alternate method if you require a larger magnitude
change.
-
Use a commercial alkaline buffer and follow manufacturer’s instructions.
Regular water changes
A functioning nitrogen cycle in an aquarium will convert toxic ammonia
and nitrites to nitrates, which are less harmful to fish. The way to
remove nitrates is via regular water changes. The following is from the
standard operating procedure
SOP
for recommended frequencies and amounts of water changes to standalone
tanks.
-
Once your tank is fully cycled (tests reveal zero nitrites and zero
ammonia), tank water needs to be changed on a routine basis.
-
The frequency and volume of water change depends primarily on the amount
of ammonia being produced; this varies with fish density and feeding
schedule (over feeding raises ammonia).
-
A 25% water change should be carried out every 2-3 weeks on healthy
tanks (low fish density, no detectable ammonia). If 25% water changes
cause too much stress, decrease the volume of water being replaced to
10% and increase water change frequency.
-
To minimize the stress caused by water changes, use aged water (same
temperature, adjusted pH levels).
-
Increase the volume or frequency of water changes when tanks show spikes
in ammonia and continue to test ammonia levels daily until the tank
stabilizes at zero.
-
To achieve targets, it may be necessary to lower fish density and the
amount of feed.
Flow-through system
The Flow Room is for lab-raised fish only, to minimize disease
transmission. Do not bring fish from the wild into this room, or dip
nets and other equipment from the McPhail room to minimize the chances
of contamination.
Initial setup of tanks
Water quality is managed centrally, so there’s little setup or
maintenance needed at individual tanks.
-
Choose empty aquaria and add your fish.
-
Look over each tank and ensure that water is flowing out of the tap and
the water level is not too high.
-
If the water level is high, lower the flow from the tap and rinse out
the sponge over the outflow pipe as it may be clogged.
Manage water quality
-
Check room temperature and record it in the daily monitoring
spreadsheet.
-
Every 2 weeks, clean the end of the pH probe on the pH doser in the back
corner of the room. Use a soft toothbrush.
-
Record the pH reading on the pH doser in the back corner of the room. pH
should be 7.4. If pH is below 7.2, clean the pH probe to ensure an
accurate reading (see above step).
-
The doser is pre-set to adjust the system pH to 7.4 by adding soda ash
from the barrel. When the pH reading stabilizes, top up the barrel by
adding dense soda ash and dechlorinated water. The amount of soda ash
added does not need to be precise.
-
To change the dosing parameters on the system, refer to the
Pinpoint
pH Controller User’s Guide.
-
Check the water temperature and O2 saturation using the PointFour
monitoring system in Room 1208A. Press the “home” button (house icon).
Our recirculation system is identified as SYSTEM B3. Measures of water
temperature and oxygen saturation should appear on the screen. Record
these values in the record book.
-
Water temperature is currently set to 17 C. The system will allow the
water temperature to fluctuate ±l degree before setting off a
temperature alarm.
-
If the temperature is outside the desired range, check that the heat
exchange pumps are functioning (you should be able to feel the machines
vibrating if you touch them). They are the two machines against the
right wall as you enter the room. They are labeled “Heat exchange pumps
for system B3”. The screen should read “17” degrees. If it the reading
is above or below 17, contact Patrick Tamkee and ask him to adjust the
“top up” water to the system in the Biosciences basement. Turning the
top up water ON will lower the water temperature in the system, as cool
water from the UBC well source will be introduced into the system.
Turning OFF the top up will allow the heat exchange pumps to maintain a
water temperature of 17 degrees. If there are any other problems with
water temperature, contact Patrick Tamkee right away.
Adjust water flow
-
The main valve that controls the water pressure is located in the back
right corner of the control room (see below).
This valve determines how much water gets diverted back to the filter
pump and into the pipes connected to the fish tanks. In an emergency
(i.e water overflowing), adjust this valve first.
-
There are additional ball valves located above each rack in the main
Flow Room that control how much water feeds each row of tanks.
BE CAREFUL when adjusting these valves. Turning off all the taps at the
tank level will cause a build up of water pressure that could lead to
cracked pipes or leaks between each joint. Always adjust the main valve
in the support room before any major adjustments at the tank level.
Make crosses
Obtaining eggs from females
A female ready to spawn can be identified by her abdomen shape at the
cloaca. If ready to lay eggs, one can almost see the first egg; at this
point her abdomen at the cloaca is sharply angled, almost like the
corner of a box. The eggs should come after gently squeezing her body
above and forward of the egg mass and, while maintaining pressure,
sliding your fingers posteriorly. The eggs, when they appear, will stick
to one another in a clump. If they dis-aggregate then the female was not
ready and you should throw the eggs out. Keep the eggs covered with
water, but keep them near the surface for oxygen.
Testes
We haven’t developed a method to extract sperm without killing the
male. Extract testes from the male after giving him an overdose of
anaesthetic. If you do not actually require separate crosses, you can
fertilize several clutches with the testes from one male. Male sperm
will keep in Hanks solution for a few days (see Storing sperm, below).
Shred the testes with tweezers and stir around the egg mass .
Remove the testes after a few minutes otherwise they might decay and
attract fungus. It is worthwhile to check that the eggs are fertilized
and begin development (the easiest way to tell is under a low-power
microscope—look for the separation of outer and inner membranes).
Raise crosses
Raise eggs
We use either of two approaches to raise the eggs to hatching. The
egg-tank method involves raising eggs for the first 7 days in small 5
gallon fish tanks and then transferring the eggs to standard 100 L tanks
just before they hatch. The 7 day period assumes that the rooms are at
17 degrees. If the room is warmer they may hatch sooner. Egg-tanks
should be cleaned out and made fresh every 2 weeks. The big-tank method
involves raising eggs entirely in the 100 L tanks.
In either case, place the eggs in a yoghurt cup having mesh screen on
the bottom. Suspend the cup from the side of the aquarium so that the
eggs are well submerged. Door screen is best, because the fry drop
through after hatching (another reason to make sure to transfer eggs
from egg-tanks to 100 L tanks before hatching begins). Put an airstone
nearby to provide oxygen and maintain a current around the eggs. Avoid
fine streams of bubbles directly underneath the eggs because air may get
trapped under in the egg mass, bringing it to the surface, where the
eggs may dry out.
The water in the tank should be dechlorinated. Adjust the pH to 7
(using baking soda or pH adjuster). Add 100-200 g of synthetic sea salt
(e.g., Deep Ocean Synthetic Sea Salts) per 100 L water. Add at least 500
g/100 L if you are raising crosses between freshwater and Little
Campbell River marines (LCM), which tend to develop poorly at low
salinity. Also add Methylene Blue solution to the tanks to help reduce
fungus attack. Mix the powder thoroughly with water in a falcon tube and
then add a few drops to the tank so that the water color is a pale blue
(don’t add the powder directly to the tank). The 100 L tanks should be
set up at least a day in advance of the egg transfer, so the filters
have time to clean the water and help dissolve/mix the salt and baking
soda. Use only a sponge filter in the tank. Make sure power
filters are shut off. Use only sponge filters until young are
large enough to handle the current and suction of a power filter.
Tend the eggs daily and remove clots of fungus that appear. Be
careful not to tear nearby eggs. You will never manage to be free of
fungus, but plenty of aeration and the methylene blue will help to avoid
the worst. Our experience is that if fungus is out of control in a
clutch of eggs only a couple of days after fertilization, then the
clutch probably wasn’t fertilized after all.
Raise young fry
When the young hatch they will sink to the bottom and stay there for
a couple of days. Then they will swim up and gulp some air from the
surface to establish neutral buoyancy. Make sure that the sponge filters
and airstones are not bubbling too vigorously at this time (keep the
power filters off until the babies are older). After a few days you will
start to see the tiny babies hanging out together off the bottom,
especially in the corners of the aquarium.
Optional: Start adding a squirt of infusoria to tanks the day they
hatch. You may also add a few drops of pet-shop liquid food for
egg-layers. They don’t eat the stuff; rather, they eat the paramecium
and possibly some bacteria. Continue adding infusoria for about 5 days
after hatch, by which time all the fry will be free-swimming.
One day after hatching, start feeding them microworms, if available,
and small quantities of first-instar brine-shrimp nauplii. Add enough
brine shrimp so that after they have fed their bellies are orange and
swelled. It is best to feed twice daily until they are a few weeks old,
but once daily will also work if you don’t mind the slower growth.
When the young reach about 2 cm, start feeding them frozen
bloodworms. Continue to feed them a small amount of brine shrimp nauplii
as well, even as adults.
Troubleshooting
Nitrites are high
Usually a problem in recently cleaned tanks, or overstocked tanks.
Monitor N levels monthly and note level on tank. When a tank has
detectable nitrites:
-
change 1/2 of the water immediately
-
add 5-15 ml of Prime (more directions on bottle of Prime)
-
add another filter
-
move some fish to another tank
-
recheck N next day, continue above steps until problem solved
Continue to replace 1/3 of the water in the tank daily until nitrites
are undetectable .
Filters not working/water not flowing properly
Check whether it’s a single filter or a bunch in one area. If single:
-
check if it’s plugged in (make sure it’s not a new baby tank and
therefore supposed to be unplugged)
-
check power bar is working
-
jiggle motor fan with finger to get it working again (sometimes they
jam)
-
check u-tube and downspout are attached properly near the motor and are
sucking up water (sometimes the filter is on, but no water is cycling –
especially important to check after cleaning filters)
-
check that the downspout isn’t blocked (plant material/dead fish/mesh
clogged with dirt)
-
check for broken parts and discard broken bits, re-assemble filter with
new part (there’s boxes of filter parts in one of the rooms, including
motors, bodies, fans etc)
-
follow the power to where it doesn’t work (the power bar has probably
gotten wet or there’s a short somewhere)
-
Jump start all filters that don’t automatically restart themselves, but
do it after you’ve sorted the problem out or you’ll be restarting things
all day.
Rooms Are Warm
This needs to be fixed immediately or fish will die.
-
If Patrick is around, tell him about the problem. If he’s not around,
phone plant ops ‘trouble calls (ext
2-2173)’ immediately, and tell them it is an animal care
emergency.
Preserve sperm for up to 2 weeks
Materials needed
-
49.5mL of Ginsburg’s ringer solution (a buffer solution - recipe is
below)
-
0.25mL of Gibco antibiotic/antimycotic (Invitrogen cat# 15240-096, 100x
concentration),
-
0.25mL of Gentamycin sulfate hydrate, (Invitrogen cat# 15750-060, 10ml,
50mg/ml liquid)
-
1 falcon tube.
Preparation of the storage solution
-
measure out 49.5mL of Ginsburg’s ringer solution and pour into a falcon
tube.
-
pipette 0.25mL of both the Gibco and Gentamycin solutions into the
Ginsburg’s solution in the falcon tube.
-
close cap and shake well in order to thoroughly mix solution
Sperm storage
-
after removing the testes from the male stickleback place each testis
(or portion of testis) in it’s own Eppendorf tube.
-
pipette in enough Ginsburg’s solution to fill half the tube.
-
make sure that each testis (or portion of testis) is completely
submerged in the solution (it should sink to the bottom).
-
store in fridge
-
replace with fresh Ginsburg’s solution every 7-10 days for storage up to
6 weeks.
Recipe for Ginsburg’s ringer solution
-
To 900 mL of ddH2O, Add: 6.6 g NaCl 0.25g KCl 0.3 g CaCl2
-
Add 0.2g NaHCO3 last
-
Mix well, and bring up to 1 L
-
Autoclave
Cryopreserve sperm
The following is derived from the
Medaka
book, which is based on
Aoki et al
(1997)
Contributed by Tom Howes: “The main modifications I made are that I
use 200 ul of medium to mince the testes (instead of 60) and store
aliquots in 5 ul microcapillary tubes (Drummond microcaps) instead of 10
ul. That gives plenty of aliquots that can be stored inside several 1.8
mL cryotubes (Nunc #377267).”
Freezing stickleback sperm
-
Remove the lids from eight 14 mL conical tubes and embed them in dry ice
in a styrofoam container (Can use a hammer to tamp them down into the
dry ice bed).
-
In a regular ice bucket, start chilling eight labeled 1.8 mL cryotubes,
and a 1.5 mL tube with the lid open and resting on the ice. Start
thawing an aliquot of Fetal Bovine Serum (heat inactivated and cleared).
-
When ready to dissect out the testes, prepare the freezing medium fresh
(180 ul FBS + 20 ul dimethylformamide) and place it in the cap of the
1.5 mL tube on ice. Place the dissected testes in the medium and mince
thoroughly. Remove any large tissue fragments.
-
Use some of the fresh sperm prep if needed (5 ul is plenty). Draw the
rest into individual 5 ul microcapillary tubes (the Drummond microcaps
come with their own squeeze bulb that you can use to create a small
amount of negative pressure). Distribute the microcap tubes to the 1.8
mL cryotubes on ice. Put the lids on the cryotubes, making sure the
capillary tubes go inside the threading of the lid so they don’t break.
-
Put the cryotubes inside the 14 mL conical tubes embedded in dry ice and
leave them there for 20 min. Then dump the cryotubes into liquid
nitrogen and move them to storage in a liquid nitrogen- cooled freezer.
Thawing an aliquot
Warm ~100 ul of Hank’s buffered saline solution to 30 C. Take out a
cryotube and keep it in liquid nitrogen until ready. Move the cryotube
onto dry ice and remove one microcap tube without thawing the others
(use forceps chilled on dry ice). Hold the microcap tube between the
fingers, and it will quickly thaw enough that you can use the
squeezebulb to eject the contents into the Hank’s solution. Move the
cryotube back to liquid nitrogen. Mix the thawed sperm solution briefly
and use for fertilization.
Prepare live food
This section provides information on preparing live food to feed
stickleback.
Brine Shrimp culture
These instructions are from the
SOP for brine shrimp
Materials needed
-
Hatching cone
-
Brine shrimp eggs
-
Clean, dechlorinated saltwater
-
Air pump and tubing
-
Warm water bath, such as a 3/4-filled aquarium, containing water heated
to 27-29 degrees C OR environment chamber heated to 25-29 degrees C
-
Aquarium heaters for water bath
-
A lamp giving off bright light, day and night
-
Scrub pad to clean hatching cone
Procedure for hatching brine shrimp eggs
-
Scrub the hatching cone to remove any bacterial slime from the sides.
-
Fill the cone with salt water, between 10-12 ppt (Note: this low
salinity is ideal for hatching brine shrimp, but a higher concentration,
28-32 ppt, is necessary to keep the shrimp alive over 24h, as they will
die at lower concentrations). Use a higher salt concentration when
rotating cultures. Adjust salinity by adding salt or water, as
necessary.
-
Insert the air line tubing into the cone and make sure the straw at the
end of the air line is inserted into the hole at the bottom of the cone.
The water should be bubbling vigorously.
-
Place the bright light above the jar and keep on always.
-
Eggs should hatch within 24 hr.
Separate the brine shrimp from the eggs
-
When hatching is complete remove the air line from the cone and leave
alone for about 15 minutes. The empty shells, if present, will float to
the top. Unhatched eggs will sink to the bottom. The live brine shrimp
(bright orange) will be hovering in the middle.
-
You may have to repeat the egg separation step a few times. It is
important that we separate as many eggs as possible as they will block
the digestive systems of the baby fish, which will cause mortality.
Microworm culture
Materials needed
-
microworm starter culture
-
a predetermined number of small Ziploc-style plastic containers
(anything with a smooth inner surface)
-
white bread (continual supply)
-
Fleishman’s dry bakers yeast (continual supply)
-
a wooden chopstick, toothpick or other suitable feeding utensil
Procedure
-
Poke minute holes (up to eight) in each of the Ziploc container lids
-
Take a small handful of white bread and moisten it thoroughly. It is
important not to soak the bread, nor to leave it too dry. The end
consistency should be similar to store-bought hummus dip
-
Add a portion of the microworm starter culture
-
Mix both the bread and culture well adding water or more bread as needed
to maintain an optimal consistency. The mixture should only take up
1/8th to 1/4th of the total volume of the container
-
Add enough yeast to just cover half of the surface of the mixture. Do
not concentrate the yeast in a single area, but spread it out over the
surface
-
Adjust consistency of mixture if needed
-
Place the lid on and the culture in a warm area that is not too dry (see
image below)
Feeding
-
Allow microworms to build up a layer around the inside surface of the
container (figure 2 above)
-
Scrape along the inside surface with the chopstick or toothpick in order
to pick up the equivalent of 2 rice grains worth of microworms (figure 3
above)
-
Dip chopstick/worm combo into the tank containing the fish to be fed
-
Dry off chopstick and repeat as necessary
Paramecium culture
Start one as soon as you start making crosses. The easy but less
reliable method is to place a pile of hay in a spare aquarium and keep
it warmer than room temperature. After a couple of weeks, stir the
contents and scoop some of the liquid into a petri dish. With a
microscope you should see Paramecium swimming around.
A more reliable but slightly more labor-intensive method, provided by
Joey Courchesne, is as follows. It makes 1L of paramecium culture.
Materials needed
-
2L flask
-
2.5g cerophyl powder (wheat grass powder)
-
3/4 g of dibase sodium phosphate powder
-
klebsiella starter culture
-
large jar (<2L)
-
a hotplate
Procedure
-
Fill flask with 1L of distilled water
-
Measure out 2.5g of wheat grass powder and 3/4 of a gram of sodium
phosphate (Sigma Aldrich cat# S-0876) and add to water
-
Bring mixture to a boil
-
Allow mixture to cool to room temperature
-
[optional step] Filter cooled mixture through cotton or nitex to filter
out wheat grass particles
-
[optional step] Autoclave cooled and filtered mixture
-
Inoculate mixture with klebsiella medium (cut off a chunk of the
bacteria rich agar and add straight to the mixture; the piece of agar
should be about the size of your thumbnail)
-
Incubate inoculated mixture overnight (8-12 hours) @ 37 degrees. The
high temperature is necessary for proper bacterial bloom
-
Pour mix into the large jar and add paramecium culture; or add mix
directly to existing paramecium culture for continued rapid growth
-
Allow paramecium population to expand for 1-3 days
-
Ripe culture should have a paramecium population visible as a “cloud” in
the jar. Sufficient population size can be confirmed with a dissection
microscope where paramecium presence should be very strongly evident.
-
Replenish culture with new food (steps 1-8) every 1-4 weeks as needed
Plan a field trip
These notes are to help plan a field trip. Send me a note if you have
modifications!
What to bring to the field
Equipment for shipping fish or eggs
-
Coolers
-
Oxygen tank
-
Battery-operated pumps
-
Batteries
-
Plastic garbage bags to line coolers so they don’t leak
-
Plastic bags for holding fish or eggs
-
Elastic bands
-
Duct tape
-
Falcon tubes & stand
Equipment for preserving fish
-
Jars
-
MS-222 (buffer with baking soda)
-
Ethanol
-
Formalin
-
Rite-in-the-rain paper
-
Pencil
Gear for catching fish
-
Traps
-
Rope
-
Dipnets
-
Pails
-
Cheese
-
Flagging tape
-
Rite-in-the-rain notebook
Gear for making crosses and keeping eggs
-
Petri dishes (10+)
-
Falcon tubes
-
Dissecting instruments (2-3 sets of tweezers, scissors)
-
Bleach or alcohol
-
Egg cups (yoghurt containers with screens)
-
Dechlorination stuff (e.g., Prime)
Gear for keeping fish
-
Air pumps
-
Tubing and air stones
-
Power filters
-
Dechlorination stuff (e.g., Prime)
-
Frozen fish food (for captives).
Clothes
-
Rain gear: jacket, pants, hood
-
Rubber boots
-
Waders
-
Life Jackets
Boat gear
-
Spare oars
-
Rope (required)
-
Bailer (required)
-
Life jackets (required)
Camping gear
-
Tent
-
Tarps
-
Sleeping bags
-
Stove
-
Cooking utensils
-
Matches
Miscellaneous
-
First Aid Kit (should be one in the lab truck)
-
Money
-
Wet suit
-
Rope
-
Binoculars
Disinfect traps
To prevent the spread of invasive species and disease organisms we
are now required to clean and sterilize all sampling equipment (traps,
seines, dip nets, boats, boots, etc.) prior to moving gear between
lakes. At minimum we are required to soak gear in a 2% solution of
household bleach for 1 minute. I would go longer, maybe 15 mintes. Rinse
with local water.
Materials
-
Garbage pail or equivalent container (30 gallons or larger)
-
Bleach
-
Measuring cup
Procedure
-
Fill garbage pail with water/chlorine mix to make the disinfectant. Mix
about 80 ml of bleach per one gallon (4L) of water. Higher concentration
is good.
-
Place traps in disinfectant for at least 1 minute (15 minutes
recommended)
-
Following disinfection, rinse traps and dry before storage
-
Disinfect the floats and strings in the same way
Notes
-
Make sure you remove any debris from the traps before you bring them
back
-
If you’re not sure that your traps are clean, it won’t hurt to
re-disinfect them
The Great Cheese Debate
When trapping sticklebacks, should a lump of orange cheddar cheese be
added in hopes of improving the catch? This is not recommended if you
are hoping to examine gut contents because some sticklebacks actually
eat the stuff. In other circumstances, however, is cheese recommended?
The debate has raged for years in the Schluter lab. McPhail swore by the
cheesy approach, Schluter also recommends it, but his lab members are
exceedingly skeptical. The question was put to the test for the first
time on June 5, 2003, by Nathan Millar and others. The test was carried
out in Klein Lake, which has a rather low density of sticklebacks.
Minnow traps were placed along the shore where they sat overnight. The
results are given below. Each number refers to the count in a single
trap (20 traps total).
|
Cheese
|
No cheese
|
|
14
|
0
|
|
12
|
3
|
|
0
|
4
|
|
3
|
3
|
|
12
|
1
|
|
23
|
1
|
|
0
|
3
|
|
22
|
4
|
|
0
|
1
|
|
|
0
|
|
|
13
|
Mean
|
9.6
|
3.0
|
**one of the traps under the no cheese columns had cheese in it, at
least initially. A two-sample \(t\)-test after a square-root (+0.5)
transformation gave \(t = 1.7386\),
\(df = 18\), \(P = 0.0992\). An approximate Wilcoxon test
(Mann-Whitney U-test) gave \(Z =
0.9628\), \(P = 0.3357\). Thus,
the results are inconclusive. Each camp can continue to hold on to
private beliefs, with the assurance that no data (yet!) will prove them
wrong.
Transport/ship stickleback
Transport live fish
Keep fish cool in pails or coolers by the lake. Use portable aerators
and air stones to keep the water bubbling. Avoid crowding, and exchange
the water frequently using lake water to maximize water quality. Keep
the water cool, as heat or oxygen stress combined with the stress of
transport will cause mortality.
Use sturdy styrofoam coolers to transport the fish. You will need a
cooler about 14 inches high if you use soda bottles rather than plastic
bags to contain the fish (make sure they are thoroughly cleaned and
rinsed to remove all odors). Other items you will need include:
-
full oxygen tank, tubing and airstone
-
clean robust plastic bags (e.g., buy at Noah’s Pet Ark) and rubber
bands, or
-
clean 2-litre soda bottles with caps
-
freezer packs or ice
-
marking pen
-
duct tape
Wrap the freezer packs or ice bag in towels or newspaper so that ice
and cold packs are not in contact with the water.
Remember that air is more limiting to the fish than water, so use
only about 20% of the container for water. Pack the fish into coolers
just before traveling back to the lab. Add a few drops of Prime or
Amquel to condition the water and remove ammonia. Add about 5 fish per
bottle or bag. Bubble oxygen into the water for a minute and then seal
bag or bottle tightly. Place bottles vertically into cooler. No more
than 40 fish in total per cooler is recommended.
When you get to the lab, open each fish bag and let it float in the
aquarium it’s going into. Wait for about 15 minutes so the water in the
bag is the same temperature as the fish tank. Every 15 minutes add 1/2
the original water volume of the bag from the aquarium. Do this at least
2 times so that the fish also gets used to the new water parameters. You
may have to dump some water out when the bag starts to fill up with
water.
Before dumping fish into tank, pour out the fish and water over the
sink into a large net. Make sure that you don’t pour any of the
lake/stream water into the fish tank as this may carry parasites. Then
dump fish into the tank.
Transport eggs/embryos
Transport eggs in falcon tubes placed in sturdy styrofoam coolers to
keep them cool and provided with sufficient oxygen. It is easier to
transport eggs that were fertilized only 0-2 days previously, because
they require less oxygen than older embryos. Other items you will need
include:
-
full oxygen tank, tubing and airstone
-
falcon tubes
-
stand or other device to hold falcon tubes upright
-
freezer packs or ice
-
marking pen
-
duct tape
Wrap the freezer packs or ice bag in towels or newspaper so that the
tubes containing the eggs are not in contact (contact will chill the
eggs and cause mortality). Place the wrapped pack or ice along the
bottom or on one side of the cooler.
Make sure the eggs are rinsed clean and that testes and other tissues
have been removed. Place one clutch of eggs into a single falcon tube
half-filled with clean lake water (preferable) or dechlorinated tap
water. Place the airstone into the tube and bubble oxygen into the water
for a minute. Remove the airstone and then cap it tightly. Place bottles
vertically into cooler (best if you have a stand for this purpose).
Secure the tubes so that they do not slide or tip during transport.
Ship live fish or eggs by air
Ship on Monday to avoid a weekend disaster if the shipment is
delayed by customs, weather, or airport issues.
If you are shipping internationally, specimens might get held up
at customs. If this happens, contact UBC’s customs broker,
Livingston, to
facilitate (link has contact information). UBC’s account number with
Livingston is 328578. Also give them a speed chart for any
charges.
If you are shipping internationally, make sure the receiver has
sent you the appropriate importation documents to include. Also print
and fill out THREE copies of the
Commercial Invoice
form. Finally, Ask Dolph to print and sign THREE copies of the
letter of declaration. An example letter is available
here. Leave
the box in Stores.
Check our oxygen tanks a few days before shipping and fill if
empty. See link on this page for instructions on how to fill the oxygen
tank.
Starve fish for one or two (preferable) days before
shipping.
Pack fish and eggs for shipment following the instructions above
for transporting fish and eggs from field to lab. Adding one teaspoon of
activate charcoal per bag of fish is recommended. Double-bag fish, and
DO NOT over-inflate bags or they will burst in transit.
Give yourself at least an hour preparation time for EACH cooler
of fish being sent (assuming that you’ve prepared other items ahead of
time: plastic bags, freezer packs, labeled boxes for coolers, full
oxygen tank). You’ll need more time if you’re sending more than one type
of fish and need to organize them while you’re packing.
Fedex Instructions
Obtain a waybill form from Zoology Stores the day before and fill
out. Call FedEx and confirm an approximate pickup time.
Air Cargo Instructions
Book the fish onto a flight 48 hours before. Look up the flight
number ahead of time.
The Air Canada booking number is 1-800-387-4865. Use the UBC account
number to book (145121), but bring a credit card to pay for the shipment
when you drop off the fish. When you book, the airline will want to know
the number, size and weight of the packages, and your name and address.
They might want to know the name and address of the recipient at this
time. They will give you a tracking/waybill number at the time of
booking. The recipient will need this information 48 hours prior to
arrival or their Customs may not release the package in time to save the
fish.
Air Canada Cargo is at 4900 Miller Road at the Vancouver
International Airport. Bring the fish to the cargo office the night
before the flight, or in the morning at least 3 1/2 hours prior to
flight time. Customs will want to view the fish before the cooler and
boxes are taped, so keep them untaped until then. Cargo might want the
cooler in a box or wrapped in sturdy paper (ask). They will not accept a
plain styrofoam cooler for fear of puncturing it during transport. Put
stickers on the box to indicate which end is up, so that the contents
aren’t inverted during transport. Make sure the statements “Live Fish”
and “Do not refrigerate” are clearly written on the box, so that they
expedite and so that they do NOT store the parcel in their 3°C cold room
before transport. Tell them that room temperature is fine.
Bring the following information and documentation to the airport.
-
Government-issued photo ID, with your name and address (e.g., drivers
license).
-
The name, address, and phone number of the person receiving the fish
(remind the recipient to bring government-issued photo ID to the airport
when they pick up the fish).
-
A declaration of importation for the US Fish and Wildlife Service. Get
the recipient to fill this out and fax to you before you go to the
airport.
-
Two copies of a letter from Dolph to US Agriculture, listing the
contents of the container.
-
Copies of the permit under which the fish were collected. You may not
need them, but the cargo people sometimes want to include documents that
the fish were collected legally.
The specific instructions above assume you are shipping to the USA.
Modify accordingly if shipping to another country.
Ship dead specimens
These instructions apply when shipping to the USA. Modify accordingly
for other countries.
We use DHL to send fish specimens, tissue, and DNA. The procedure is
similar as for an ordinary package, but with a bit more
documentation.
-
Drain all alcohol or other preservative from the specimens and pack them
into a sealed box, such as a tupperware container. In the case of fish
specimens, line the box with damp towels to prevent them drying out.
-
Print two copies of a letter from Dolph to US Agriculture, listing the
contents of the container.
-
Print and fill out TWO copies of the
Commercial Invoice
form.
-
Go to Stores and ask about how to ship.
-
To track a package, go to the
DHL
web site.
Ship stuff on dry ice
These instructions apply when shipping to the USA. Modify accordingly
for other countries.
We ONLY use FedEx, because the other couriers have restrictions on
shipping dry ice. Leave enough time in the day for you to pack up the
box and have FedEx come and pick it up from Botany/Zoology Stores.
Ship only on Monday to avoid a weekend disaster if shipment is
delayed by customs, weather, or airport issues.
If you are shipping internationally, specimens might get held up
at customs. If this happens, contact UBC’s customs broker,
Livingston, to
facilitate (link has contact information). UBC’s account number with
Livingston is 328578. Also give them a speed chart for any
charges.
Get a cardboard box and a styrofoam box that sits neatly inside
with little free space (fill any free space with scrunched up newsprint
to minimize jostling of samples). To find a matching styrofoam and
cardboard box look around labs that frequently buy enzymes, such as NAPS
or the Qaigen stores on the 3rd floor of the North wing of the
Biological Sciences building.
Place the appropriate labels on the outside of the cardboard box.
The appropriate labels include:
-
The shipper’s address and contact info. This is us.
-
The receiver’s address and contact info.
-
At least one (2 or 3 is best) dry ice warning stickers (see image
below). Please make sure that you do not write on the stickers as they
MUST not be written on. The best place to put the stickers is in the
middle of the shipping box.
The stickers may be
purchased from Botany/Zoology Stores. They are located just above the
biohazard waste disposal cards next to Scott’s office. Or, download and
print the warning
label and tape it onto the box. Just make sure that it’s a large and
clear copy of the warning label.
-
Write the code UN1845 on the box
next to the dry ice warning label (see image above). This tells the
shipper that it’s dry ice.
-
Write the total weight of the dry ice used in the shipment next to the
dry ice warning sticker (see image above). This MUST be written in
kilograms and NOT in pounds.
Get your dry ice (instructions on getting dry ice below). I’d
suggest getting at least 1 kg of dry ice but no more than 3
kgs.
Obtain a FedEx EXPANDED SERVICE INTERNATIONAL AIR WAYBILL
from Stores and fill it out as shown
here in an example.
Please DON’T get the regular INTERNATIONAL AIR WAYBILL as it won’t have
the section needed for a dry ice shipment. Our FedEx customer account
number is 359397682.
Print and fill out THREE copies of the
Commercial Invoice
form.
Ask Dolph to print and sign THREE copies of the letter of
declaration. An example letter is available
here.
Place the samples into the dry ice and close the styrofoam box
securely. Tape the cardboard box shut.
Tape a plastic window envelope to the top of the box. Combine
each of the three copies of the Commercial Invoice form and signed
letter of declaration and place them with the EXPANDED SERVICE
INTERNATIONAL AIR WAYBILL into the window envelope. Example is seen
here.
Leave the box at our shipping/receiving office. Call FedEx and
tell them the package is ready for pickup.
Get dry ice
If you really want to be thrifty, you can stop by NAPS or any
enzyme-selling lab and ask if they have any free dry ice. Otherwise
you’ll have to buy it.
To purchase, make sure you know the speed code to pay for the dry
ice. If you don’t know it, please ask Dolph.
Obtain a styrofoam box with lid. Bring it to
Chemistry Stores (corner of
Main Mall and University Blvd), located across the street and just north
of the Biological Sciences building. Once you enter the Chemistry
building, directly in front on the right is the stairs which you take
down to the bottom floor. Walk East down the hallway. You’ll walk along
a wall of lockers and will soon see the entrance of the Chemistry
stores.
You must first weigh your box before placing the dry ice inside (I
measure it in kilograms). Ask the attendant for the keys for the dry ice
room (the dry ice room is located next to the stairs you came down). Get
your ice, and weigh the box again and figure out the total weight of dry
ice taken. Record the amount in the dry ice book in Chem Stores.
Note that Chem stores closes for lunch between 11:30am and
12:30pm.
Fill the oxygen tank
-
Call Praxair 604-527-0710 to order oxygen. Our customer no. is HU839.
They will ask for a credit card number so please make sure you have it
before calling.
-
Tell them you want medical grade O2 in small bottle size and
tell them the quantity.
-
Let them know if you have any empty tanks for pickup. The delivery guy
drops off full tanks and picks up empty ones. Drop off the empty
O2 cylinders outside, next to Zoology Stores, next to the
large gas cylinders. Tear off all the old tags.
-
It usually takes 2-3 days before they arrive. They will drop off the
filled cylinders in the same area when you dropped off the empty tanks.
-
If you need to speak to our sales rep for Praxair, his name is Alan
Sechinko and he can be reached at 604 619 6420.
Tag stickleback
Elastomer tagging
This section explains how to tag threespine stickleback safely and
effectively using elastomer.
Materials
-
MS-222 (Tricaine Methane Sulfonate)
-
Sodium bicarbonate (baking soda)
-
Spatula
-
1000 ml glass beaker
-
pH test kit
-
Holding tank and recovery tank
-
Air supply and air stone
-
Tagging kit including syringe and elastomer (e.g., Northwest Marine
Technology)
Procedure
-
Fast fish for 12-24 hours prior to tagging. This will reduce fecal
contamination and risk of regurgitation.
-
Maintain adequate oxygenation via air stone in both holding and recovery
tanks.
-
Use water from the original source tank for transport, anesthetic, and
recovery. If using another water source, closely duplicate the water
quality parameters (e.g. temperature, pH, hardness, etc.).
-
Maintain water temperature at the species’ normal optimum during both
the anesthesia and recovery period.
-
Make a desired volume of MS-222 solution in the beaker (concentration of
50-75mg/L) and buffer the solution to pH of 7-7.5 with sodium
bicarbonate. Wear protective clothing, gloves, and goggles when handling
MS-222 powder. If possible, work inside a fume hood to prepare solution.
-
Anesthetize the fish in the MS-222 solution. Before tagging, make sure
that the fish has slow respiration and heart rate, and is nonresponsive
to stimuli. Observe the movement of the operculum as it opens and closes
to assess rate. If respirations become extremely slow or stop, place
fish in the anesthetic-free recovery water until respirations resume.
-
To inject a tag, insert the syringe needle into the marking location
(~1cm or less and just under the skin). Slowly withdraw the needle as
the material is injected, so that a long narrow mark is created. It is
important that the tag created is fully contained within the target
tissue.
-
Once tagged, place fish into recovery tank to recover.
Additional notes
-
To begin, anesthetize a few fish and follow them through full recovery
to ensure drug dosages and techniques are safe and provide sufficient
anesthesia.
-
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.
RFID tagging
This section explains how to insert an RFID tag into a three-spined
stickleback safely and effectively.
Materials
-
Melafix (fish medication)
-
AmQuel or other water conditioner/detoxifier
-
Methylene blue and tea tree oil
-
MS-222 (Tricaine Methane Sulfonate)
-
Sodium bicarbonate (baking soda)
-
RFID tags, 8 mm (should be 30 in a packet).
-
MK-7 Implanter
-
Surgical blade
-
1000 ml glass beaker
-
pH test kit
Procedure
-
Set up a 100-L aquarium that is completely cycled. Add about 4 mL
Melafix, 2 mL of AmQuel, about half a cup of kiln-dried salt, and
several drops of methylene blue and tea tree oil.
-
Wear protective clothing, gloves, and goggles when handling MS-222
powder. If possible, work inside a fume hood to prepare solution. Mix a
desired volume of MS-222 solution in the beaker at a concentration of
solution of 0.3g/L. Buffer the solution to pH of 7-7.5 with sodium
bicarbonate (will require about 1-2 times the amount of MS-222 added).
-
Set up multiple containers of about 1 L to hold up to 3 fish immediate
after tagging. Fill containers with dechlorinated water of room
temperature (e.g., use the water we have in the tubs for water changes).
Add a drop of tea tree oil.
-
Wipe down work surface with 95% ethanol, and fill a small dish up with
95% ethanol. Put two clean paper towels on the work surface. Put a sharp
pair of scissors, the needle portion of the MK-7, tweezers, and all tags
to be placed in fish into the ethanol-filled dish. After several
minutes, remove objects and place them on one paper towel to air dry.
-
Place a single fish in the MS-222 solution until it reaches until fish
reaches stage four of anesthesia (total loss of muscle tone and
equlibrium; slow but regular opercular rate; loss of spinal reflexes).
This will take about 1 minute for a stickleback of 5-6 cm. Stir the
MS222 solution to check the fish’s equlibrium. The rate at which the
pectoral fins beat is a good indication of muscle tone.
-
While fish is being anesthetized, put on a clean pair of latex gloves.
Use tweezers to load the glass RFID tag into the MK-7 implanter, being
careful to get the tag all the way in.
-
Gently remove the fish from the MS-222 solution using a large pair of
tweezers and place on the second clean towel. Take the clean, air-dried
scissors and make a small incision on the side of the fish, just under
the lateral line and about a centimeter behind the pectoral fin. If the
fish has lateral plates, the incision may need to be lower.
-
Insert the loaded MK-7 implanter into the incision with the needle
opening facing you, and with the point aimed toward the tail. Continue
until 3/4 of the needle opening is within the incision. Keep the
implanter as close to the dermis as possible as you insert. Then, rotate
the implanter 180 degrees so that the needle opening is facing the fish.
Push the plunger to insert the RFID tag. Once the tag is partially in
the fish, pull out the implanter as you push the plunger further to
minimize unnecessary penetration. Place tagged fish in the prepared 1-L
container.
-
Put scissors and tweezers back into the ethanol, as well as a fresh
needle. Replace paper towels and latex gloves.
-
Repeat steps 4-9 until all fish are tagged.
-
Gently transfer all fish to the prepared 100-L aquarium, while wearing a
fresh pair of gloves
-
After 24 hours, add another dose of Melafix to the aquarium water. After
3 days, replace 25% of the water in the tank.
-
Feed the newly tagged fish juvenile food for three days. Then switch to
regular adult food.
Euthanasia
This section explains how to euthanize threespine stickleback safely
and effectively.
Materials
- MS-222 (Tricaine Methane Sulfonate) (check expiry date)
- Sodium bicarbonate (baking soda)
- Forceps or long tweezers
- Spatula
- 1000 ml glass beaker
- pH test kit
Procedure
- Wear protective clothing, gloves, and goggles when handling MS-222
powder. If possible, work inside a fume hood to prepare solution.
- Make a desired volume of MS-222 solution in the beaker
(concentration of 0.5g/L). Buffer the solution to pH of 7-7.5 with
sodium bicarbonate (0.5-1 g/L).
- Place fish in the solution and wait at least 15 minutes, to ensure
that death is achieved. Verify before disposing or preserving the
carcass by monitoring the absence of respiratory or opercular movement
for at least 3 minutes.
- Use tweezers to remove the fish from the solution.
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, stain, measure
This page describes how to euthanize, stain, and measure fish.
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.
Stain fish
Stain fish preserved in ethanol with alizarin red
Warning: the procedures below will destroy DNA of the
sample.
-
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.
-
Formalin step: Transfer rinsed fish into 10% formalin for 48 hours.
Always use a fume hood when pouring formalin.
-
Rinsing (2): Pour out the formalin and rinse the fish in water for 24
hours.
-
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.
-
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.
-
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.
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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.
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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.
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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.
Measuring shape using landmarks
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To measure shape of stained fish, use the high-quality Nikon camera,
which has a lens with a flat field. Camera settings are
here
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Most of the landmarks we use were described in the paper by Albert et al
(2008). An illustration of the landmarks is
here.
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These landmarks were based on turn on those used by Walker (1997),
illustrated here.
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Caldecott & Adams (1998) used the following
skull
landmarks
Measuring lipids
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).
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Dry the fish in a drying oven and then weigh.
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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%).
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Tip contents into labeled Falcon tube. Add 8mL methanol and 8mL
chloroform.
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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.
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Top up to 25mL with chloroform and allow the sample to settle.
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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.)
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Boil off organic layer with a hot plate set to 70 deg C. Allow beaker to
cool to room temperature.
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Weigh beaker containing lipids.
© 2009-2025 Dolph Schluter