Rockfish lavage

My research asks the question: do predatory fishes influence benthic communities?

To get there, we first had to determine exactly what those predatory fishes were eating.  The predators include lingcod, copper and quillback rockfish, and kelp greeling.  Anne Beaudreau, a former FHL graduate student, has already done a wonderful job of describing lingcod diets.  We chose to focus on rockfish diet.

Importantly, we wanted to sample rockfish diets without killing the fish.  Excision of the stomach is a common practice for examining stomach contents of fishes, but since rockfish populations are at risk in the Salish Sea, we didn’t want to contribute to their decline.  Gastric lavage, or stomach pumping, is a viable alternative, but had not been attempted with rockfish prior to our attempts.

Anesthesia bucket filled with 100mg/L buffered MS-222; Styrofoam cradle, with two channels for different fish sizes; Spring scale, meter stick and calipers; Hand-pumped garden sprayer.

Anesthesia bucket filled with 100mg/L buffered MS-222; Styrofoam cradle, with two channels for different fish sizes; Spring scale, meter stick and calipers; Hand-pumped garden sprayer.

We caught rockfish from less than 20m depth so as to reduce the risks of barotrauma.  This was both to reduce the risk of injury to the fish, and to ensure that the expanding gas bladder didn’t press on the stomach and cause the fish to vomit their stomach contents before we could catch them.  We also used barbless hooks to keep capture injuries to a minimum.

Once on board the boat, the fish were anesthetized in 100mg/L buffered tricaine methanosulfonate.  We had previously determined that this was a sufficient dosage to knock the fish out in about 5-10 minutes, keep them under for the 5-10 minutes of handling time, and then recover quickly in clean seawater.

Measuring the gape of a copper rockfish.

Measuring the gape of a copper rockfish.

After anesthetization, the fish were measured for mass, total length, body depth, and gape height and width.  We placed the fish upside down in a foam cradle and inserted a small plastic tube through the mouth and esophagus.  The hose end of a hand-pumped garden sprayer filled with clean seawater was inserted through the tube and into the stomach.  The sprayer hose size was stepped down in size and made smooth with surgical and Tygon tubing.

Inverted rockfish with tube and hose inserted into the esophagus.

Inverted rockfish with tube and hose inserted into the esophagus.

Because the garden sprayer was hand-pumped, we were able to easily adjust the amount of water pressure coming out of the hose.  We used this pressure to flush stomach contents out onto a collection screen.  Stomach contents were preserved in ethanol for later identification, and the fish were placed in a cooler of clean seawater to recover from anesthesia.

Several copper rockfish in the recovery cooler.

Several copper rockfish in the recovery cooler.

After the fish recovered from anesthesia we needed to return them to their capture depth, but their swim bladders were still inflated.  There are several methods for recompressing rockfish (see this video from NOAA SWFSC); we used an inverted weighted basket to return the fish.

Two copper rockfish in the return basket.

Two copper rockfish in the return basket.

Sending the rockfish back down.

Sending the rockfish back down.

We initially tested three individuals in the lab to establish an anesthetic dose and to learn the lavage methods.  All three individuals were kept for several days and appeared to recover without issues.  In our field work we captured and lavaged 29 fish, without any mortality or indication of significant injuries.  In the lab testing we offered shrimp to the rockfish, but failed to monitor how many shrimp were consumed, and we did not directly observe the fish feeding on the shrimp.  It is possible that the lavage procedure damaged or irritated the throat or stomach, preventing the fish from feeding.  If we were to repeat this work, we would want to monitor fish in the lab to determine when they start feeding again.

One of the projects that lavage could be used for is to monitor individual diets through time: fish could be tagged and recaptured and resampled periodically.  Copper rockfish are known to change their diets seasonally, with fish size, and at different locations.  Resampling individual rockfish would help determine if diet preferences are individual- or population-specific.  However, this would only be possible if the lavage procedure does not significantly alter feeding behavior.

How to Measure a Current

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A clod card made of alabaster, adhered to an acrylic plate and strapped to a brick.

One of the simplest and cheapest ways to measure how much water moves by an area is to take advantage of materials that dissolve in water.  The faster water moves past the object, the more of the material will dissolve away, proportional to the exposed surface area of the object.  Materials that have been used for this purpose include plaster of paris, alabaster, and even Lifesavers.  These are typically called “clod cards” because the first ones consisted of clods of plaster attached to paper cards.

Our lab measures current flow with a combination of clod cards made of alabaster (inexpensive to produce, easy to deploy at many sites at once) with an Acoustic Doppler Velocimeter (very expensive, can only be deployed at one site at a time).

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Derek Smith building clod cards.

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Derek Smith and Cori Kane hauling a ton of bricks.

One of the disadvantages of clod cards is that they can only tell you about relative current flow.  That is, we can tell if Site A has faster flow than Site B, but not the actual average current speed at either site.  ADVs are able to measure three-dimensional water movement (“velocimeter”) by sending sound waves (“acoustic”) to bounce off of particles in the water.  When the echos from the sound waves return to the sensor, the ADV measures the doppler shift in the waves (“doppler”).  We pair clod cards with the ADV so we can approximately calibrate dissolution of the clod cards to an average current speed.

ryan installing adv

Ryan Knowles installing the ADV (Photo by Megan Cook).

The white instrument at the top right is the actual measurement probe.  Sound waves emanate from the three prongs, bounce off of particles in the water, and are received at the midpoint of the instrument head.  The probe is elevated above the bottom so that we measure free-stream flow instead of the reduced current in the boundary layer.  The large yellow canister strapped to the cement base is the battery pack so it can record for weeks at a time.  One of the calibrating clod cards is off to the left.

Light Attenuation

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Without direct illumination from a strobe, underwater photos take on green or blue hues. These rocks are covered in bright red, orange and yellow organisms, but none of those colors appear unless we bring underwater flashlights.

Shallow coral reefs are bright and colorful.  The deep abyss is pitch black.  What about in between?

Light from the sun is composed of a spectrum of different wavelengths of energy, including visible light.  When white light passes into the ocean, these different wavelengths behave differently.  Some wavelengths are absorbed by the water very rapidly, and some are able to penetrate deep into the water.  Longer wavelenghts of light (red, orange, yellow) are absorbed more rapidly than shorter wavelengths.  If you descend in to clear open-ocean water everything looks blue:

Photo: oceanexplorer.noaa.gov

But in coastal water with lots of nutrients, like along the West Coast of the US, there are tons of phytoplankton in the water.  These phytoplankton absorb blue and red light, and reflect green light (the same reason why grass is green).  This turns coastal waters green instead of blue.

The only way to combat this light attenuation is to bring our own light sources.  We carry flashlights and camera strobes to help illuminate all of the colors in the Salish Sea:

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This photo was taken at about 20m depth. Without the camera strobe, all of these colors would appear as some shade of green.

 

Last Transect Complete!

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Tim Dwyer and me through the camera framer, just after finishing the last transect dive of 2012

After a long autumn quarter, we finally finished our annual fall surveys.  This enormous body of work (95 transects in about 75 dives) could not have been accomplished without the help of a small army of volunteers and interns.

Thanks to Annie Thomson, Aaron Galloway, Derek Smith, Noel Larson, Rhoda Green, Autumn Turner, Gavin Brackett, Ryan McLaughlin, Ryan Knowles, Jackie O’Mara, Megan Cook, Pema Kitaeff and Alex Lowe.

And thanks especially to Tim Dwyer, inexhaustible dive buddy extraordinaire.

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That’s our planning whiteboard.  All those purple boxes are transects that needed doing, and the green text are the dates we completed the transects.  They’re finally all full – whew!

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Tim and me collapsed under the weight of all the equipment it takes to get the science done.

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My face seconds after finishing the last transect.

Sunset Diving

Derek Smith and I finished up a transect dive at Shady Cove just as the sun set over Friday Harbor.  Derek snapped this photo with our photoquadrat camera setup.

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This is definitely one of the benefits of trying to dive every available slack between October and January every year!

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800th Dive

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Today marks a major milestone – 800 dives since starting graduate school.  That’s 57,065 feet down (and back) and over 417 hours underwater.  I wonder if I can make it to 900 before I graduate…  Should I even be trying to make it that many?

Photoquadrats

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Epilithic organisms in a photoquadrat at Minnesota Reef, 21m depth.  You can see cup corals, several bryozoans, hydroids, sponges, encrusting algae and a creeping sea cucumber.

A lot of the work our lab does involves taking photos of quadrats underwater.  We take photos because the depth we work at (up to 27m) prevents us from spending enough time to quantify epifauna along our transects.

Our camera setup consists of an Olympus C-8080 in an underwater housing, a strobe slave, and an Ikelite strobe, all mounted on an aluminum frame.

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Photo by Megan Cook

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Photo by Megan Cook, Ryan Knowles diving

The business end of the framer measures 35x25cm.  When we take photos we rest this rectangle flat against the bottom, and use the rulers to set an accurate scale for measuring mobile fauna on the computer.  We quantify percent cover of sessile organisms by dividing the photo into 24 blocks and estimating basal and canopy cover in each block.

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