Diving Deeper

By Taylor Mock, GEMM Lab intern

Greetings, all!

My name is Taylor Mock. Since February I have been volunteering in the GEMM Lab and am ecstatic to make my online debut as part of the team!

For many years, I had a shallow relationship with Hatfield Marine Science Center. As a Newport native, I would spend mornings and evenings glancing over at the Hatfield buildings while driving over the bridge to and from school. I was always intrigued. Sure, I would hear snippets of research from my peers about what projects their parents were involved in, but the inner workings of the complex mystified me.

Toward the end of my Freshman year in 2012 at Westmont College in Santa Barbara, California, my mom asked me what my summer plans were. I replied with the typical “I don’t know… Get a job?” She insisted that instead of a job I think about getting an internship; experience that will last more than a summer. I inquired through a family friend (because every person in this little community is woven together some way or another) if any internships or volunteer opportunities were available at Hatfield. She pointed me in the direction of the Environmental Protection Agency and thus began my Hatfield volunteering saga. I worked that summer, and the next, at the EPA under the direction of Ted DeWitt and Jody Stecher on denitrification studies in estuarine marshes. That summer provided me a glorious front row seat to field research and a greater understanding of my potential as a person and as a scientist. Now, this experience was marvelous, but I knew shortly after starting that my heart was elsewhere.

It was during my study abroad semester in Belize as part of my internship at the Toledo Institute for Development and Environment (TIDE) that I realized I wanted to work with marine macroorganisms. At TIDE, I engaged in radio telemetry conservation efforts tracking Hicatee (Dermatemys mawii) aquatic turtles. We would spend days on a small boat floating through canals and setting nets in hopes of capturing individuals of this small population to outfit them with radio tracking devices. These would be later used to track foraging, mating, and travel patterns in the region. It was an amazing time, to say the least. I remember waking up on my 21st birthday from my camping hammock and staring up at the lush rainforest above my head with a warm breeze across my face, followed by spending the day in the presence of these glorious creatures. It was heaven. I returned to Westmont the following term and took a Marine Mammal Eco-Physiology course and absolutely fell in love with Cetacea. Yes, I had always been captivated by this clade of beings (and truthfully when I was eight years old had a book on “How to Become a Marine Mammal Trainer”), but this was deeper. Of course, pinnipeds and otters and polar bears and manatees were enjoyable to learn about. There was something about the Cetacea though and how they migrated up and down the coast (just like me!) that I really connected with. My time learning about these animals created an intimate understanding of another group of species that developed into a rich, indescribable empathetic connection. I had to take a couple years away from scholastics and away from biology for health and wellness reasons. One day, though, a couple years after graduating and returning to Newport I rekindled with Jody from the EPA. He asked me if I would like to volunteer under Leigh Torres in the Marine Mammal Institute at HMSC. I do not think I could have possibly said no. I have been enjoying my time in the GEMM Lab ever since!

Though I am available to help anyone with any task they need, the work I do mostly centers around photogrammetry.

Using photogrammetry skills to measure gray whales in the GEMM Lab.

Photogrammetry, essentially, means geo-spatially measuring objects using photographs. What that looks like for me is taking an aerial photograph (extracted from overhead drone video footage) of a whale, running the image through a computer program called “Matlab”, taking a series of measurements from the whale (e.g., tip of the mandible to the notch of the fluke, distance between each tip of the fluke, and several measurements across the midsection of the whale). Several images of individuals are processed in order to find an average set of measurements for each whale.

Final result of the photogrammetry method on a gray whale

You might be wondering, “How can one measure the distance accurately from just a photograph?” I am glad you asked! The drones are outfitted with a barometer to measure the atmospheric pressure and, in turn, altitude. The changing altitudes are recorded in a separate program that is run simultaneously with the video footage. Thus, we have the altitudinal measurements for every millisecond of the drone’s flight. To monitor the accuracy and functionality of the barometer, calibrations are completed upon deployment and retrieval of each drone flight. To calibrate: the initial takeoff height is measured, a board of known length is thrown into the water, the drone will then rise or lower slowly above the board between 10 and 40 m, photographs of the board are then taken from varying altitudes, and are processed in Matlab.

During my time in the GEMM Lab, I have had the pleasure of completing photogrammetry assignments for both Leila on the Oregon Coast gray whale and for Dawn on the New Zealand blue whale projects. These ladies, and the other members of the GEMM Lab, have been so patient and gracious in educating me on the workings of Matlab and the video processing systems. It is a distinct honor working with them and to delight in the astounding nature of these creatures together. Each day I am struck in sheer awe of how beautiful and powerful these whales truly are. Their graceful presence and movement through the water rivals even the most skillful dancer.

Over the last 6 years, I am delighted to say that my relationship with Hatfield has become much deeper. The people and the experiences I have encountered during my time here, especially in the GEMM Lab, have been nothing short of incredible. I am sincerely grateful for this continued opportunity. It fills my soul with joy to engage in work that contributes to the well being of the ocean and its inhabitants.

Thank you, Leigh and all of the GEMM Lab members. I hope to continue volunteering with you for as long as you will have me.

Where are the eggs? Studying red-legged kittiwakes in a time of change

By Rachael Orben, Research Associate, Department of Fisheries and Wildlife, Oregon State University

In late May, I returned to St. George Island, Alaska to study the foraging ecology of red-legged kittiwakes using a mix of high-tech biologging tags, physiology measurements, and observations.  The study was designed to identify differences in behavior and physiology between birds that reproduce successfully and birds that don’t and then to see how this might carry over to the winter season (and vice versa).  Things didn’t go as planned.

A red-legged kittiwake on St. George Island. Photo C. Kroeger

This was my fourth spring on the island, and like prior seasons we arrived in late May, when birds should be building nests. However, unlike previous seasons, red-legged kittiwake’s didn’t look like they had done much nest building. I was accompanied by Abram Fleishman, a superstar MS student from San Jose State who is studying the winter spatial ecology of red-legged kittiwakes in relationship to mercury concentration in their feathers.

We immediately set about recapturing birds that had carried geolocation data loggers over the winter. We wanted to catch as many as possible before eggs were laid, so that their blood samples would represent the pre-lay period. The weather was wonderful, so it wasn’t until three weeks after we arrived that we had our first day-off. It was at about this time that I finally lost my optimism and realized the majority of red-legged kittiwakes were not going to lay eggs. By late June kittiwakes are usually incubating eggs. We only saw a handful of eggs and very few of these were being incubated. Most birds didn’t even build nests, or if they did, the nest was dismantled by other birds when the nest building pair didn’t stick around to guard their pile of mud and moss.

Nests in 2016. Laying success was also low in 2016, but even if birds didn’t lay many pairs built nests.
Same cliff (and birds) without nests in 2017.

When I designed the study, I thought collecting enough data to answer my questions about successful versus failed breeders would be hard, since failed breeders would be challenging to work with and red-legged kittiwakes typically have high breeding success, meaning that sample sizes of failed breeders would be small. Instead our three seasons occurred with progressively worse breeding success and we will now have to shift the focus of our analysis to see if we can find differences between birds that laid eggs and birds that didn’t, if we have the sample sizes! With ~80% laying success in the 9 years preceding the beginning of our study in 2015, this is something I would never have expected! The egg laying failure of 2017 is unprecedented in the productivity monitoring time series collected by the Alaska Maritime Wildlife Refuge.

Seabirds are often touted as indicator species of marine health (Cairns 1988, Piatt et al 2007), and while there are always caveats and additional questions to be answered, seabirds are reliant on the ocean for food and observing their behavior and condition tells us something about how easy (or hard) it is for them to find food.

So, what do I think the red-legged kittiwakes told us this year? I think they were squawking loud and clear, that they were not able to find myctophid fishes within their foraging range to the south and west of the Pribilofs. Myctophids are small fatty mesopelagic bioluminescent fish that come to the surface at night where red-legged kittiwakes catch them.

Besides just observing the laying failure, we were able to GPS track a few birds, collect a few diet samples, catch birds for blood and feather samples, and resight banded individuals. It is these pieces of information that I will be analyzing in the coming months to try to understand why some individuals were able to lay eggs during our study years, while most were not.  These years should also help us understand what capacity red-legged kittiwakes have to cope (or not) with changes in prey availability. However, after three years, I still don’t know what a ‘good’ year looks like for red-legged kittiwakes. Fingers crossed next season is finally a decent year for this sentinel seabird of the pelagic Bering Sea.

Pre-lay foraging trips of red-legged kittiwakes in 2015.
Pre-lay foraging trips of red-legged kittiwake in 2017. Two birds were heading north when their GPS loggers stopped recording.

You can read more about our red-legged kittiwake research in a series of blog posts written for the Seabird Youth Network, a partnership between the Pribilof School District, the Aleut Community of St. Paul Island, the City of St. Paul, Tanadgusix Corporation, the St. George Traditional Council, St. George Island Institute, the Alaska Maritime National Wildlife Refuge, and the wider scientific community. The network creates opportunities for youth to learn about seabirds with the aim of building local capacity for the collection of long-term seabird monitoring data on the Pribilof Islands.

New and old methods in our gray whale field season 2017

By Leila Lemos, Ph.D. Student, Department of Fisheries and Wildlife, OSU

On June 6th the GEMM Lab officially started the second year of fieldwork of our “Noise Physiology” Project with gray whales along the Oregon coast. To date, we have spent 14 days at sea (12 around the Newport area and 2 in Port Orford, our control area), with a total of 32:31 hrs of effort. In 29 whale sightings of approximately 40 whales we have been able to collect 6 fecal samples for hormonal analysis, to fly the drone 17 times over the whales, to deploy a GoPro 6 times for qualitative prey analysis, and to deploy a light trap 2 times for quantitative prey analysis. While this sounds good, we have only just begun, with our field season extending into October. The graph below displays the sightings and data collection by area.

Figure 1: Sightings and data collection by area and month.

We have added a couple new components to our project this year. First, we are now using a “the light trap”, as mentioned above, to capture zooplankton prey of gray whales. The light trap (Figure 2), designed by our collaborator of Kim Bernard (OSU, College of Earth, Ocean, and Atmospheric Sciences). The light trap is composed of a water jug with a cut-out cone entrance where prey might enter the jug after being attracted by the chem lights we put in the jug. The jug is weighted down to maintain position, but swivels off the drop line by its own floats; and it’s all connected to a surface float.

Figure 2: Components of the light trap.
Source: Leila Lemos

The light trap is left overnight and recovered in the next day. Trapped prey are sieved (Figure 4), stored in properly labeled jars or Ziploc bags, and kept frozen until analysis (Figure 5 and 6) including species identification, community analysis, and caloric content.

Figure 3: Todd Chandler, our research technician, preparing the light trap to be deployed in Port Orford.
Source: Leila Lemos
Figure 4: Collected preys with our light trap being sieved for storage on June 27th.
Source: Dawn Barlow
Figure 5: Kim Bernard proud of the zooplankton sample collected in Newport on June 26th.
Source: Dawn Barlow
Figure 6: Our GEMM Lab intern Alyssa holds the prey sample collected in July 1st.
Source: Leigh Torres

The second component we have added this year is the fixed-location hydrophone (Figure 7) to record acoustic noise data over the entire summer season. Last year we used a temporarily deployed “drifting hydrophone” that only recorded noise data punctually. Because of the fixed hydrophone, this year we will be able to compare our hormone data with a wider range of acoustic data, and improve our analyses.

Figure 7: Joe Haxel, our acoustician, checking the hydrophone in July 14th that was previously deployed in Newport at the beginning of the summer season.
Source: Leila Lemos

We also made our first trip down to Port Orford, our control area, to intensively collect data over only two days (July 5th and 6th). Since Port Orford is a smaller city with reduced vessel traffic, we want to evaluate if whales observed in this area show a reduced stress response when compared to the whales that inhabit the area around Newport and Depoe Bay, where vessel traffic is higher. However, we were not able to collect any fecal sample during this trip to Port Orford, so more trips south to come!

Figure 8: Sharon Nieukirk, our acoustician, Leigh Torres, and Todd Chandler checking on RV Ruby before being lifted into the water at the port of Port Orford on July 5th.
Source: Leila Lemos
Figure 9: Our mascots Pepper and Avery didn’t get to go out in the boat with us, but they enjoyed our trip to Port Orford so much that they couldn’t stay awake on the way back to Newport.
Source: Leila Lemos and Leigh Torres

The other components we used last year such as photo identification (Figure 10), fecal samples (Figures 11 and 12), drones, and GoPros are still being put to use this year. If you want to know more about our Noise Physiology project, check here.

Figure 10: Me in our boat platform waiting for whales to appear to photograph them in July 13th.
Source: Joe Haxel
Figure 11: Joe Haxel collecting a fecal sample in Newport in July 13th.
Source: Leila Lemos
Figure 12: Fecal sample collected in Newport on July 13th.
Source: Leila Lemos

We are progressively spotting more gray whales along the Oregon coast and we will continue our field efforts and data collection until October. So, for now enjoy some photos taken during the last couple of months. Until next time!

Figure 13: Gray whale’s fluke just south of the Yaquina Lighthouse, in Newport, on July 13th.
Source: Leila Lemos
Figure 14: Gray whale breaching just north of the Yaquina Lighthouse, in Newport, on July 9th.
Source: Leila Lemos
Figure 15: Gray whale breaching in Newport, on June 6th.
Source: Leigh Torres

Life in the lab: notes from a lab meeting

By Florence Sullivan, MSc, Oregon State University

One of my favorite parts about working as a member of the GEMM lab is our monthly lab meeting. It’s a chance for everyone to share exciting news or updates about their research, discuss recent advances in our field, and of course, make the schedule for who is in charge of writing the blog each week!  Our fearless leader, Leigh, usually also has an exercise for us to complete. These have varied from writing and editing abstracts for conferences, conducting mock interviews of each other, reading and discussing relevant papers, R coding exercises, and other useful skills. Our most recent meeting featured an exciting announcement, as well as a really interesting discussion of the latest International Whaling Commission (IWC) reports of the scientific committee (SC) that I felt might be interesting to share with our readers.

First, the good news – Six GEMM lab members submitted abstracts to the 2017 Society of Marine Mammalogy Conference, and all six were accepted for either a speed talk or an oral presentation! We are very proud and excited to present our research and support each other at the conference in October.

And now, a little science history:

The IWC was originally formed as a management body, to regulate the global catch of great whales. However, it never had much legal power to enforce its edicts, and was largely ineffective in its task.  By 1986 whale populations had been decimated to such low numbers by commercial whaling efforts that a worldwide moratorium on harvest was imposed. The SC of the IWC meets on an annual basis, and is made up of leading experts in the field who give advice and recommendations to the commission.  If you are interested in seeing reports from over the years, follow this link to the IWC Archive.  The reports presented by the various sub committees of the Scientific Committee are dense, packed full of interesting information, but also contain lots of procedural minutiae.  Therefore, for this lab meeting, each of us took one of the 2017 Annexes, and summarized it for the group.

Alyssa and Dawn reviewed Annex J: Report of the working group on non-deliberate human induced mortality of cetaceans.  The report shared new data about scarring rates of bowhead whales in the Bering Sea, notably, that 2.4% of the population will acquire a new scar each year, and that by the time an individual is 25 years old, it has a 40% chance of being scarred from a human derived interaction. The study noted that advances in drone technology may be an effective tool to assess scarring rates in whale populations, but emphasized that it is important to examine stranded carcasses to ground truth the rates we are able to capture from aerial and boat based photography.  The discussion then turned to the section about ship strikes, where we learned that in a comparison of fresh scars on humpback whales, and rates of voluntarily reported ship strikes, collisions were vastly under reported. Here it was noted that injuries that did not cause visible trauma could still be lethal to cetaceans, and that even moderate speed collisions can cause non-immediate lethal injury.

Leila walked us through Annex K: Report of the standing working group on environmental concerns. This subcommittee was the first one formed by the SC, and their report touched on issues such as bioaccumulation of heavy metals in whales, global oil spill emergency response training, harmful algal blooms (HABs), marine debris, diseases of concern, strandings and related mortality, noise, climate change, loss of arctic sea ice, and models of cetacean reaction to these impacts.

A few notes of particular interest:

-PCBs and other toxins are known to accumulate in killer whales, but this report discussed high levels of lead and cadmium in gray whales, leading to the question of what might be the source – sediment deposits? Fish?

-Lots of research has been done on the outfall of HABs involving domoic acid; now there is a need for research on other types of HABs

-A website has been created to increase surveillance, diagnosis and risk management of cetacean diseases, and is currently being refined: https://cdoc.iwc.int

-Changing climate is prompting distribution shifts in a number of species, putting animals at risk of interactions with shipping lanes, and increasing contact with invasive species.

-Models of cetacean bioenergetics have found that being entangled has energy costs equivalent to migration or pregnancy. Another model found that naval noise increased the metabolic rate of individuals by 30%. Models are becoming more and more accurate and complex every year, and each new one helps provide a framework to begin to assess cumulative impacts of human-cetacean interactions.

To wrap things up, I gave a brief overview of Annex N: Report of the subcommittee on whalewatching. This report gave quick updates on a number of different whale watching research projects around the world:

-Humpback whales in Hawaii change their swim speed and dive time when they encounter vessels.

-Endangered humpbacks in the Arabian Sea may need management intervention because there have been minimal advances in standards and attitudes by whale watching outfits or recreational boaters in Oman.

-Increased interactions and close encounters may be eroding the protective social barriers between bottlenose dolphins and the public.  The committee emphasizes that cetacean habituation to humans is a serious conservation cause of concern.

After research updates, the document then details a review from the working group on swim-with-whale operations. They emphasize the need for a global database, and note that the Convention on Migratory Species and the World Cetacean Alliance are both conducting reviews of this section of the whale watching industry and that a collaboration could be beneficial. Finally, this committee often gives feedback to ongoing projects and local management efforts, but is not convinced that their recommendations are being put into practice.

As one reads this litany of issues that face cetaceans in the modern world, it can be quite disheartening. However, reports like these keep researchers up to date on the current state of knowledge, areas of concern, and questions that need answering.  They help us set our priorities and determine which piece of the puzzle we are capable of tackling.  For more on some of the projects that our lab has under taken to help tackle these issues, check out Leila’s work on stress in gray whales, Dawn’s work looking at blue whales in New Zealand, Solene’s work on humpback habitat selection, or my work on vessel interactions. Individually, it’s easy to feel small, but when you look through the archives of the IWC, and realize how far we’ve come from extractive management to active conservation, you realize that every little project adds to those before it, and together, we can make a difference.

 

 

 

Migrating to higher latitudes

By Leila Lemos, Ph.D. Student, Department of Fisheries and Wildlife, OSU

On September 10th of 2015 I was catching an airplane to start a whole new phase of my life in Oregon, United States. Many thoughts, many doubts, many fears, many expectations, and one big dream that was about to come true: I was finally going to United States to work with whales.

I am from Rio de Janeiro, Brazil, a big city known for pretty beaches, tropical weather and restless nights. Thus, to arrive in a really small city on the countryside that usually rains for about six months a year was the opposite of what I was always used to. Trying to understand another language and culture differences was also not an easy step.

In addition, taking my first classes was a big challenge. It was hard to understand everything that was being said, but recording and listening to the classes afterwards definitely was what helped me the most. Also, my first meetings and discussions where I needed to explain my thoughts in another language was difficult, but when I look back and I can now see how much I have improved and it is gratifying to know that all of my efforts were worth it.

Feeling welcome was essential to start overcoming all of the difficulties. My advisor Leigh and my lab mates (Florence, Amanda, Rachael, Erin, Dawn and Courtney) always created a friendly atmosphere and I started being more confident over time. I also had amazing and understanding teachers who were patient and helped me along the way. My first roommates Jane and Angie, from US, and the students and teachers from Crossroads (an English group that I attend) made me practice English every day and I started feeling more comfortable about speaking (and also thinking) in English, and they became my “Oregon family” together with new friends I made from different nationalities. Also important were my family and friends back in Brazil that never stopped encouraging and supporting me.

Figure 1: GEMM-Lab, from left to right, starting at the top: Leigh Torres, me, Erin, Amanda, Dawn, Rachael, our interns from 2016 season (Catherine, Cat and Kelli), and Florence.

 

Figure 2: Practicing English at Crossroads.

 

The weather and seasons here are also very different from Brazil. We don’t have cold weather or snow, and we don’t see all of the changes that happen here from season to season. The first season I saw was the fall. Seeing all of the fall colors in the trees for the first time was magical and I can already say that fall is my favorite season here. The winter was a bit cruel for me, not because of the cold or eventually the snow, but because of the rain. There is a saying in my city that “people from Rio de Janeiro do not like gray days” and it is true: my mood changes with weather. However, I did travel a bit around Oregon during winter and got to enjoy the snow, and how fun is to slide in the snow, make snow angels and throw snowballs. The spring starts bringing sunny days after cold months and endless rain. Also all of the flowers around the Corvallis campus are so pretty and colorful. Finally the summer is hot, and in some days it can almost be as hot as Rio de Janeiro. However, I spend summer days in the coast, where the temperature is mild. For me, summer days are synonymous with fieldwork, since gray whales are migrating northbound and becoming resident along the Oregon coast to feed, and this is right when the fun begins!

Figure 3: Different seasons in Oregon: (A) Trees during the fall in Corvallis, (B) Winter in Crater Lake, (C) Spring at OSU campus: my office at Hovland Building, and (D) fieldwork in Port Orford during the summer.

 

I finally saw my first gray whale in July of 2016 and got to dive into all of the methodologies we wanted to apply in this project. I learned how to photograph whales for photo-identification, how to take important notes, how to collect fecal samples for hormonal analysis, and how to fly with a drone for the photogrammetry method.

Figure 4: Learning how to fly with a drone over gray whales.
Source: Florence Sullivan

 

I had to digest a lot of information while trying to equilibrate in the boat and to not get seasick. However, it was so pleasurable to see how my field skills were getting better over time and how close I was to the Pacific marine fauna.

During my master’s degree I worked on toxicology in dolphins, which means working with dead carcasses. I remember telling myself all of the time that I wanted to do something different for my PhD – that I would be involved in a project with live animals. I am very glad I could accomplish that goal. Gray whales, sea lions, seals and a variety of marine birds are just some examples of the great diversity the Pacific Ocean has to offer and I am totally amazed.

Figure 5: Great diversity of the Oregon coast. Source: GEMMLab (Leila Lemos, Leigh Torres and Florence Sullivan)

After months of fieldwork it was time to return to the land and start learning how to work with all of the data we collected. We have amazing collaborators working with us and I have had wonderful opportunities to learn from all of them about the different methods we are applying in our project.

Figure 6: Learning the hormonal analysis technique at the Seattle Aquarium.

 

Thus, after one year and a half in Oregon I can already say that I feel home. The experience as an international student is not easy, but that’s what makes it such a valuable and gratifying experience. It has been a great journey, and I hope to continue to see improvements over time and keep learning throughout this amazing project studying gray whales.

 

How Unmanned Aircraft Systems (UAS, aka “drones”) are being applied in conservation research

By Leila Lemos, Ph.D. Student, Department of Fisheries and Wildlife, OSU

 

Unmanned Aircraft Systems (UAS), also known as “drones”, have been increasingly used in many diverse areas. Concerning field research, the use of drones has brought about reduced errors, increased safety and survey efforts, among other benefits, as described in a previous blog post of mine.

Several study groups around the world have been applying this new technology to a great variety of research applications, aiding in the conservation of certain areas and their respective fauna and flora. Examples of these studies include forest monitoring and tree cover analyses, .

Using drones for forest monitoring and tree cover analyses allows for many applications, such as biodiversity and tree height monitoring, forest classification and inventory, and plant disease and detection. The Ugalla Primate Project, for example, performed an interesting study on tree coverage mapping in western Tanzania (Figure 1).

Figure 1: Tree coverage analyses in Tanzania.
Source: Conservation Drones, 2016.

 

The access to this data (not possible before from the ground) and the acquired knowledge on tree density and structure were important to better understand how wild primates exploit a mosaic landscape. Here is a video about this project:

 

Forest restoration activities can also be monitored by drones. Rainforests around the world have been depleted through deforestation, partly to open up space for agriculture. To meet conservation goals, large areas are being restored to rainforests today (Elsevier 2015). It is important to monitor the success of the forest regeneration and to ensure that the inspected area is being replenished with the right vegetation. Since inspection events can be costly, labor intensive and time consuming, drones can facilitate these procedures, making the monitoring process more feasible.

Zahawi et al. (2015) conducted an interesting study in Costa Rica, being able to keep up with the success of the forest regeneration. They were also able to spot many fruit-eating birds important for forest regeneration (eg. mountain thrush, black guan and sooty-capped bush tanager). Researchers concluded that the automation of the process lead to equally accurate results.

Drones can also be used to inspect areas for illegal logging and habitat destruction. Conservationists have struggled to identify illegal activities, and the use of drones can accelerate the identification process of these activities and help to monitor their spread and ensure that they do not intersect with protected areas.

The Amazon Basin Conservation Association Los Amigos conservancy concession (LACC) has been monitoring 145,000 hectars of the local conservation area. Illegal gold mining and logging activities were identified (Figure 2) and drones have aided in tracking the spread of these activities and the progress of reforestation efforts.

Figure 2: Identification of illegal activities in the Amazon Basin.
Source: NPR, 2015.

 

Another remarkable project was held in Mexico, in one of the most important sites for monarch butterflies in the country: the Monarch Butterfly Biosphere Reserve. Around 10 hectars of vital trees were cut down in the reserve during 2013-2015, and a great decrease of the monarch population was perceived. The reserve did not allow researchers to enter in the area for inspection due to safety concerns. Therefore, drones were used and were able to reveal the illegal logging activity (Figure 3).

Figure 3: Identification of illegal logging at the Monarch Butterfly Biosphere Reserve, Mexico.
Source: Take Part, 2016.

 

Regarding the use of drones for mapping vulnerable areas, this new technology can be used to map potential exposed areas to avoid catastrophes. Concerning responses to fires or other natural disasters, drones can fly immediately, while planes and helicopters require a certain time. The drone material also allows for operating successfully under challenging conditions such as rain, snow and high temperatures, as in the case of fires. Data can be assessed in real time, with no need to have firefighters or other personnel at a dangerous location anymore. Drones can now fulfill this role. Examples of drone applications in this regard are the detection, monitoring and support for catastrophes such as landslides, tsunamis, ship collisions, volcanic eruptions, nuclear accidents, fire scenes, flooding, storms and hurricanes, and rescue of people and wildlife at risk. In addition, the use of a thermal image camera can better assist in rescue operations.

Researchers from the Universidad Politécnica de Madrid (UPM) are developing a system to detect forest fires by using a color index (Cruz et al. 2016). This index is based on vegetation classification techniques that have been adapted to detect different tonalities for flames and smoke (Figure 4). This new technique would result in more cost-effective outcomes than conventional systems (eg. helicopters, satellites) and in reaching inaccessible locations.

Figure 4: Fire detection with Forest Fire Detection Index (FFDI) in different scenes.
Source: UPM, 2016.

 

Marine debris detection by drones is another great functionality. The right localization and the extent of the problem can be detected through drone footage, and action plans for clean-ups can be developed.

A research conducted by the Duke University Marine Lab has been detecting marine debris on beaches around the world. They indicate that marine debris impacts water quality, and harms wildlife (eg. whales, sea birds, seals and sea turtles) that might confuse floating plastic with food. You can read a bit more about their research and its importance for conservation ends here.

Drones are also being extensively used for wildlife monitoring. Through drone footage, researchers around the world have been able to detect and map wildlife and habitat use, estimate densities and evaluate population status, detect rare behaviors, combat poaching, among others. One of the main benefits of using a drone instead of using helicopters or airplanes, or having researchers in the area, is the lower disturbance it may cause on wildlife.

A research team from Monash University is using drones for seabird monitoring in remote islands in northwestern Australia (Figure 5). After some tests, researchers were able to detect which altitude (~75 meters) the drone would not cause any disturbances to the birds. Results achieved by projects like this should be used in the future for approaching the species safely.

Figure 5: Photograph taken by a drone of a crested tern colony on a remote island in Australia.
Source: Conservation Drones, 2014.

 

Drones are also being used to combat elephant and rhino poaching in Africa. They are being implemented to predict, trace, track and catch suspects of poaching. The aim is to reduce the number of animals being killed for the detusking and dehorning practices and the illegal trade. You can read more about this theme here. The drone application on combating one of these illegal practices is also shown here in this video.

As if the innovation of this device alone was not enough, drones are also being used to load other tools. A good example is the collection of whale breath samples by attaching Petri dishes or sterile sponges in the basal part of the drones.

The collection of lung samples allows many health-monitoring applications, such as the analysis of virus and bacteria loads, DNA, hormones, and the detection of environmental toxins in their organisms. This non-invasive physiological tool, known as “Snotbot”, allows sampling collection without approaching closely the individuals and with minimal or no disturbance of the animals. The following video better describes about this amazing project:

It is inspiring to look at all of these wonderful applications of drones in conservation research. Our GEMM Lab team is already applying this great tool in the field and is hoping to support the conservation of wildlife.

 

 

References

Conservation Drones. 2014. Conservation Drones for Seabird Monitoring. Available at: https://conservationdrones.org/2014/05/05/conservation-drones-for-seabird-monitoring/

Conservation Drones. 2016. Tree cover analyses in Tanzania in collaboration with Envirodrone. Available at: https://conservationdrones.org/2016/09/17/tree-cover-analyses-in-tanzania-in-collaboration-with-envirodrone/

Cruz H, Eckert M, Meneses J and Martínez JF. 2016. Efficient Forest Fire Detection Index for Application in Unmanned Aerial Systems (UASs). Sensors 16(893):1-16.

Elsevier. 2015. Drones Could Make Forest Conservation Monitoring Significantly Cheaper: new study published in the Biological Conservation wins Elsevier’s Atlas award for September 2015. Available at: https://www.elsevier.com/about/press-releases/research-and-journals/drones-could-make-forest-conservation-monitoring significantly-cheaper

NPR. 2015. Eyes In The Sky: Foam Drones Keep Watch On Rain Forest Trees. Available at: http://www.npr.org/sections/goatsandsoda/2015/05/19/398765759/eyes-in-the-sky-styrofoam-drones-keep-watch-on-rainforest-trees

Take Part. 2016. Drones Uncover Illegal Logging in Critical Monarch Butterfly Reserve. Available at: http://www.takepart.com/article/2016/06/22/drones-uncover-illegal-logging-monarch-butterfly-habitat

UPM. 2016. New automatic forest fire detection system by using surveillance drones. Available at: http://www.upm.es/internacional/UPM/UPM_Channel/News/dc52fff26abf7510VgnVCM10000009c7648aRCRD

Zahawi RA, Dandois JP, Holl KD, Nadwodny D, Reid JL and Ellis EC. 2015. Using lightweight unmanned aerial vehicles to monitor tropical forest recovery. Biological Conservation 186:287–295.

 

Beyond the Rock: Using Satellite Trackers to Study the Lives of Common Murres

By Stephanie Loredo, Seabird Oceanography Lab, OSU

Photo credit: Seabird Oceanography Lab

Common murres (Uria aalgee) are the most abundant seabird on the Oregon Coast. At least half of the population in the California Current Ecosystem breeds on the Oregon Coast (half a million seabirds). This makes them ecologically important consumers of forage fish, especially during the breeding season when they use state-waters.

While they spend most of their time at sea, murres must come to shore to breed. During this time, they are highly visible by humans as they breed in large masses on rocky islands. While they are not the most agile on land, due to their short and stubby legs, they are actually amazing divers. Their short flipper-like wings help them swim, and they typically reach depths of 30-60m to catch their prey.

Aside from their underwater aviation skills, they make great parents as well. Both parents will incubate and care for their chick – murres only lay one egg a year – until they fledge; once they leave the rock, male murres take full responsibility for their chicks while the moms go on vacation (they worked hard to lay the egg so they need some time to recuperate). After the breeding season, murres leave the rock in large quantities – this is often the last time humans will see them this year in large aggregations from shore.

Despite their omnipresence and importance as a marine predator in Oregon, there is still a lot we don’t know about murres. Where do murres go when they are not breeding? Do they migrate? Where do they feed during the breeding and non-breeding period? What habitat characteristics are associated with feeding areas? By answering these questions, we increase knowledge of murre ecology in Oregon. Moreover, a more comprehensive understanding of the year-round movements of murres aids marine spatial planners take more informed actions on the current decisions regarding offshore renewable energy development. This is what I hope to achieve through my Masters research project at OSU.

Most of what is known about the offshore distribution of murres in Oregon comes from vessel observations. However, vessel data only provide snapshots in time, and not a continuous picture of area-use. Within the Seabird Oceanography Lab (SOL), we are using individual satellite tracking devices to follow the movements of murres associated with the Yaquina Head colony, which is a prominent breeding colony in Oregon located near Newport.

A common murre displaying a satellite tag prior to release.

SOL was able to track 15 common murres associated with the Yaquina Head colony in 2015 and 2016.  These tags were deployed periodically throughout the breeding period and have been successful in tracking birds for up to three months. Thus far, we have tracking data ranging from May to December (only one bird tracked during December).

Tracking data from 2015 and 2016 of murres off the Yaquina Head colony provide an interesting comparison.  In both years, murres experienced warmer ocean conditions, high Bald eagle disturbance rates, and consequently high Western gull egg predation at the colony. Some data also indicate low prey availability.  The combination of all these factors is most likely the reason for the observed reproductive failure at the colony in both years. Tracking data showed that 13 of the 15 birds tagged dispersed from the colony earlier than expected. The maps below summarize the dispersal of birds by year and by time of deployment.

 

Each map (Left: 2015, Right: 2016) illustrates all birds that dispersed from the colony and did not engage in central-place foraging (feeding trips to and from the colony). Sample size: n2015=7, n2016_spring=1, n2016_summer=3.

Most birds made a northward movement and traveled as far north as British Columbia, Canada.  Along their movement north, they used inlets and bays, but one of the most prominent areas used was the Columbia River plume. Birds used the Columbia River mouth area during the summer and fall, with the most time spent there during the summer. Dispersal from the colony was not what we expected; we expected individuals to breed on colony and engage in central-place foraging  (feeding to and from the breeding site) nearshore until mid-August when they usually leave the rock. However, we are still interested in the habitat characteristics of feeding areas and the conditions that led to movement from one feeding area to the next.

Prior to examining habitat associations of murre feeding areas, we must first determine their behavior state at each point location derived from the satellite tags.  After data cleaning and filtering out erroneous locations, we applied a behavioral analysis (Residence in Space and Time method) to determine behaviors associated with each point location. This analysis has allowed us to distinguish between intensive foraging, transiting, and extensive foraging. Extensive foraging locations can be interpreted as a set of locations that are mostly spread out in space, where murres searched for prey. On the other hand, intensive foraging locations can be interpreted as a set of locations that are very close together in space where murres likely found prey, and thus spent more time.

We are finalizing the extraction of environmental data for each point location from satellite data. Once all data are extracted, we can begin analysis for determining what environmental conditions were sought during dispersal and what types of habitats are preferred. Some of the ocean conditions that will be examined are sea surface temperate, wind, upwelling index, and primary net productivity. Some other habitat descriptors we are interested in assessing are substrate, distance to river mouth, salinity, depth, distance to the 200-m isobath, and distance to shore. For now, exploration of data indicates differences in habitat associations by behavior and between seasons.

Sample size means everything in a study like this so I am happy to say that more data is yet to come: SOL plans to deploy 15 more tags during spring and summer of 2017. I am excited to see what the additional tagged murres will do, and whether they will follow a pattern similar to those tracked in 2015 and 2016. However this time around, we will deploy tags as late in the summer/early fall as we can, in hope of acquiring some novel winter data to fill this knowledge gap. If we are successful, we may finally have a better idea of what life is like for common murres during more of the year beyond the rock.

 

Celebrating Hydrothermal Vents!

By Florence Sullivan, MSc Student OSU

40 years ago, in 1977 OSU researchers led an NSF funded expedition to the Galapagos on a hunt for suspected hydrothermal vents. From the 1960s to the mid-1970s, mounting evidence such as (1) temperature anomalies found deep in the water column, (2) conduction heat flow probes at mid ocean ridges recording temperatures much lower than expected, (3) unusual mounds found on benthic mapping surveys, and (4) frequent, small, localized earthquakes at mid oceanic ridges, had the oceanographic community suspecting the existence of deep sea hydrothermal vents. However, until the 1977 cruise, no one had conclusive evidence that they existed.  During the discovery cruise at the Galapagos rift, the PI (principle investigator), Dr. Jack Corliss from OSU, used tow-yos (a technique where you drag a CTD up and down through the water in a zig zag pattern – see gif) to pinpoint the location of the hydrothermal vent plume. The team then sent the Deep Submergence Vehicle (DSV) Alvin to investigate and returned with the first photographs and samples from a hydrothermal vent. While discovery of the vent systems helped answer many questions about chemical and heat fluxes in the deep sea, it generated so many new questions that novel fields of study were created in biology, microbiology, marine chemistry, marine geology, planetary science, astrobiology and the study of the origin of life.

 “Literally every organism that came up was something that was unknown to science up until that time. It made it terribly exciting. Anything that came [up] on that basket was a new discovery,” – Dr. Richard Lutz (Rutgers University)

In celebration of this great discovery, OSU’s College of Earth, Ocean and Atmospheric Sciences sponsored a seminar looking at the past, present, and future of hydrothermal vent sciences. Dr. Robert Collier began with a timeline of how the search for hydrothermal vents began, and a commemoration of all the excellent researchers and collaborations between institutions and agencies that made the discovery possible. He acknowledged that such collaborations are often somewhat tense in terms of who gets credit for which discovery, and that while Oregon State University was the lead of the project, it takes a team to get the work done.  Dr. Jack Corliss proudly followed up with a wonderful rambling explanation of how vent systems work, and a brief dip into his ground breaking paper, “An Hypothesis concerning the relationship between submarine hot springs and the origin of life on Earth.” Published in 1981, with co-authors Dr. John Baross and Dr. Sarah Hoffman, they postulate that the temperature and chemical gradients seen at hydrothermal vents provide pathways for the synthesis of chemical compounds, formation and evolution of ‘precells’ and eventually, the evolution of free living organisms.

Dr. Corliss, Dr. Baross, and Dr. Hoffman were the first to suggest the now popular theory of the origin of life at hydrothermal vents. (click on image to read full paper)

Because of time constraints, the podium was swiftly handed over to Dr. Bill Chadwick (NOAA PMEL/ HMSC CIMRS) who brought us forward to the present day with an exciting overview of current vent research.  He began by saying “at the beginning, we thought, ‘No one has seen one of these systems before, they must be very rare…’ Now, we have found them [hydrothermal vents] in every ocean basin – including the arctic and southern oceans. We just needed to know how to look!”  Dr. Chadwick also reminded us that even 40 years later, new discoveries are still being made. For example, on his most recent cruise aboard the R/V Falkor in December 2016, they found a sulfur chimney that was alternately releasing bubbles of gas (sulfur, CO2 or other, hard to know without sampling) or bubbles of liquid sulfur! Check out the video below:

Some of the goals for this recent cruise included mapping new areas of the Mariana back-arc, and investigating differences in the biological communities between vents in the Mariana trench region (a subduction zone) and vents in the back arc (a spreading zone) to see if geology plays a role in biological community composition.  For some very cool video footage of the expedition and the various dives performed by the brand new ROV SUBastian (because all scientists love puns), check out the Schmidt Ocean Institute youtube channel.

Dr. Chadwick showed this video to highlight results from his last cruise.

Finally, Dr. Andrew Thurber wrapped up the session with some thoughts about hydrothermal vents from the perspective of an ecosystem services model. Even after 40 years of research, there are still many unknowns about these ecosystems.  Individual vent systems are inherently unique due to their deep sea isolation. However, most explored sites have revealed metals and mineral deposits that have generated a lot of interest from commercial sea floor mining companies. Exploitation of these deposits would be an example of ecosystem “provisioning services” (products that are obtained from the ecosystem). Other examples include the biology of the vents as a source of new genetic material, and the thermal and chemical gradients as natural laboratories that could lead to breakthroughs in pharmaceutical research. Cultural services are those non-material benefits that people obtain from an ecosystem. At hydrothermal vents these include new scientific discoveries, educational uses (British children’s television show “The Octonauts,” has several episodes featuring hydrothermal vent creatures), and creative inspiration for artists and others. Dr. Thurber cautions that there are ethical questions to be answered before considering exploitation of these resources, but there is a lot of potential for commercial and non-commercial use of vent ecosystems.

Vent inspired art by Lily Simonson

As an undergraduate at the University of Washington, I spent time as a research assistant in Dr. John Baross’ astrobiology lab. We studied evolutionary pathways of hydrothermal vent viruses and bacteria to inform the search for life on exoplanets such as Jupiter’s moon Europa.  It was very fun and exciting for me to attend this seminar, hear stories from pioneers in the field, and remember the systems I worked on in undergrad.  I may have moved up the food chain a little now, but as we all work on our pieces of the puzzle, it is important for scientists to remember the interdisciplinary nature of our work, and how there is always something more to learn.

 

 

Challenges of fecal hormone analyses (Round 2): finally in Seattle!

By Leila Lemos, Ph.D. Student, Department of Fisheries and Wildlife, OSU

In a previous blog of mine, you could read about the challenges I have been facing while I am learning to analyze the hormone content in fecal samples of gray whales (Eschrichtius robustus). New challenges appeared along the way over the last month, while I was doing my training at the Seattle Aquarium (Fig. 1).

Figure 1: View of the Seattle Aquarium.

 

My training lasted a week and I am truly grateful to the energy and time our collaborators Shawn Larson (research coordinator), Amy Green and Angela Smith (laboratory technicians) contributed. They accompanied me throughout my training to ensure I would be able to conduct hormonal analysis in the future, and to handle possible problems along the way.

The first step was weighing all of the fecal samples (Fig. 2A). Subsequently, the samples were transferred to appropriate glass tubes (Figs. 2B & 2C) for the next laboratorial step.

Figure 2: Analytical processes: (A) Sample weighing; (B) Transference of the sample to a glass tube; (C) Result from the performed steps.

 

The second conducted step was the hormone extraction. The extraction began with the addition of an organic solvent, called methanol (CH3OH), to the sample tubes (Fig. 3A & 3B). Hormones leach out from the samples and dissolve in the methanol, due to their affinity for this polar solvent.

Tubes were then placed on a plate shaker (Fig. 3C) for 30 minutes, which is used to mix the substances, in order extract the hormones from the fecal samples. The next step was to place the tubes in a centrifuge (Fig. 3D) for 20 minutes. The centrifuge uses the sedimentation principle, causing denser substances or particles to settle to the bottom of the tube, while the less dense substances rise to the top.

Figure 3: Analytical processes: (A) Methanol addition; (B) Sample + methanol; (C) Plate shaker; (D) Centrifuge.

 

After this process, the two different densities were separated: the high-density particles of the feces were in the bottom of the tube, while the methanol containing the extracted hormones was at the top. The top phase (methanol + hormones) was then pipetted into a different tube (Fig. 4A). The solvent was then evaporated, by using an air dryer apparatus (Fig. 4B), with only the hormones remaining in the tube.

The third performed step was dilution. A specific amount of water, measured in correlation with sample weight and to the amount of the methanol mixed with each sample, was added to each tube (Fig. 4C). Since the hormones were concentrated in the methanol, the readings would exceed the measurement limits of the equipment (plate reader). Thus, in order to prepare the extracts for the immunoassays, different dilutions were made.

Figure 4: Analytical processes: (A) Methanol transference; (B) Methanol drying; (C) Water addition.

 

The fourth and final step was to finally conduct the assays. Each assay kit is specific to the hormone to be analyzed with specified instructions for each kit. Since we were analyzing four different hormones (cortisol, testosterone, progesterone, and triiodothyronine – T3) we followed four different processes accordingly.

First, a table was filled with the identification numbers of the samples to be analyzed in that specific kit (Fig. 5A). The kit (Fig. 5B) includes the plate reader and several solutions that are used in the process to prepare standard curves, to initiate or stop chemical reactions, among other functions.

A standard curve, also known as calibration curve, is a common procedure in laboratory analysis for determining the concentration of an element in an unknown sample. The concentration of the element is determined by comparison with a set of standard samples of known concentration.

The plate contains several wells (Fig. 5C & 5D), which are filled with the samples and/or these other solutions. When the plate is ready, (Fig.5D) it is carried to the microplate reader that measures the intensity of the color of each of the wells. The intensity of the color is inversely proportional to the concentration of the hormone in both the standards and the samples.

Figure 5: (A) Filling the assay table with the samples to be analyzed; (B) Assay kit to be used; (C) Preparation of the plate; (D) Plate ready to be read.

 

Since this is the first fecal hormone analysis being performed in gray whales, a validation process of the method is required. Two different tests (parallelism and accuracy) were performed with a pool of three different samples. Parallelism tests that the assay is measuring the antigen (hormone) of interest and also identifies the most appropriate dilution factor to be used for the samples. Accuracy tests that the assay measurement of hormone concentration corresponds to the true concentration of the sample (Brown et al. 2005).

This validation process only needs to be done once. Once good parallelism and accuracy results are obtained, and we have identified the correct dilution factor and approximate concentration of the samples, the samples are ready to be analyzed. Below you can see examples of a good parallelism test (parallel displacement; Fig. 6) and bad parallelism tests (Fig. 7) that indicate no displacement, low concentration or non-parallel displacement; and a good accuracy test (Fig. 8).

Figure 6: Example of a good parallelism test. The dark blue line indicates the standard curve; the pink line indicates a good parallelism test, showing a parallel displacement; and the ratios in black indicate the dilution factors.
Source: Brown et al. (2005)

 

Figure 7: Examples of bad parallelism tests. The dark blue line indicates the standard curve; the light blue line is an example of no displacement; the pink line is an example of low concentration of the sample; and the green line is an example of non-parallel displacement.
Source: Brown et al. (2005)

 

Figure 8: Example of a good accuracy test while analyzing hormone levels of pregnanediol glucuronide (Pdg) in elephant urine. The graph shows good linearity (R2 of 0.9986) and would allow for accurate concentration calculations.
Source: Brown et al. (2005)

 

After the validation tests returned reliable results, the samples were also analyzed. However, many complications were encountered during the assay preparations and important lessons were learned that I know will allow this work to proceed more smoothly and quickly in the future. For instance, I now know to try to buy assay kits of the same brand, and to be extremely careful while reading the manual of the process to be performed with the assay kit. With practice over the coming years, my goal is to master these assay preparations.

Now, the next step will be to analyze all of the results obtained in these analyses and start linking the multiple variables we have from each individual, such as age, sex and body condition. The results of this analysis will lead to a better understanding of how reproductive and stress hormones vary in gray whales, and also link these hormone variations to nutritional status and noise events, one of my PhD research goals.

 

Cited Literature:

Brown J, Walker S and Steinman K. 2005. Endocrine manual for reproductive assessment of domestic and non-domestic species. Smithsonian’s National Zoological Park, Conservation and Research Center, Virginia 1-69.

Simple behavior classification of tracking data with residence in space and time

By Rachael Orben PhD., Postdoctoral Scholar in the Seabird Oceanography Lab and the Geospatial Ecology and Marine Megafauna Lab 

At 2pm, Jan 3, our paper entitled “Classification of Animal Movement Behavior through Residence in Space and Time” was published. At 14:03 I clicked on the link and there it was, type-set and crisp as a newly minted Open Access scientific contribution.

So, what is this paper about? It presents a simple – yes simple – method of identifying simple behaviors states in two-dimensional animal tracking data (think latitude and longitude). Since the paper is open access you can go find the methods there. Categorizing these “dots on a map” into behaviors allows us to ask questions about how often, why, when and where simple behaviors happen. These behaviors really are simple (hopefully the somewhat grating repetitiveness of the word ‘simple’ has driven that point home by now!). We are identifying three basic, but fundamental, states:

1) transit, characterized by fast somewhat straight line movement from a to b,

2) a sedentary state characterized by relatively more time spent in an area with little distance traveled (such as resting behavior) and

3) an active state characterized by lots of time spent in an area where an animal is also moving around a lot and covering a lot of ground.

This new method, that we termed Residence in Space and Time (RST), can assist the fast-growing, sophisticated, big-data generating, conservation-orientated field of animal movement ecology. One of the first hurdles is data exploration and visualization. Modern ecologists deploy tracking devices that collect location data remotely to understand animal distribution and behavior. But at first glance tracks (like the figure below) can look like spaghetti dinner. Identifying movement behaviors can help to us see patterns in the tangles.

24 GPS tracks of grey-headed albatross incubation foraging trips; tracked from Campbell Island, New Zealand.

So how might this method work? First lets start with a track. Below is a very short foraging trip from a thick-billed murre tracked with a GPS logger during chick rearing from St. Paul Island in Alaska (see Parades et al 2015).

p1080393
A thick-billed murre (Uria lomvia), St. Paul Island, Alaska.

The track below has points every second and we can imagine the murre flying from the colony, landing on the water, and then diving (indicated by the lack of GPS position data when the bird dives below the water to forage). Then the bird flies back to the colony to feed its chick. This trip is roughly 14 minutes long.

murretrack

So I can take this track and run RST to identify three behavior states. As color-coded below, the black points indicate transit, red indicates relatively stationary behavior, and blue indicates points where the bird was flying in a less direct manner than pure transit potentially circling around before landing and moving between dives. The high resolution of the GPS data really helps us to understand how this bird was moving. Such behavior information is easily conserved in a high-resolution track like this. Though in this case the bird did a lot of transiting and only exhibited different movement behaviors in the vicinity of the two dives.

murretrack_1sec_rst-copy

Logging locations at 1 second intervals is a stretch for the battery life of these miniaturized GPS loggers (~15g), and more often than not we would like the loggers to last much longer than 14 mins. So instead of 1 second we typically have tracks with less frequent locations. To me this is akin to taking a 1 second track and then taking off my glasses and trying to see the same behaviors. Deciphering behavior states becomes a bit (or a lot) fuzzier. In the case of this murre track, when we down-sample the locations to every 10 seconds much of the resolution of this track is lost (see plot below). What happens when we run RST?

murretrack_10sec_rst-copy

As you can see some of the behavior is maintained and some of it is a bit fuzzier.

A good rule of thumb is that if a behavior happens faster than the sampling interval the logger is recording at, then the behavior is not recorded. Seems simple, but it is an important consideration when programming loggers and designing animal movement studies. For murres these quick trips to forage for their chicks are easily lost even at a 5 minute sampling interval, which is often used in seabird tracking studies where the birds are at-sea for days. Often we work with such lower resolution location data and, instead of one trip from one bird, we have many trips from many individuals. RST allows a fast way to quickly and accurately identify simple behaviors in order to help with initial data exploration efforts and for answering more complex questions such as behavior specific habitat models.

So, if you have some tracking data – of birds, marine mammals, or your dog! – you can learn how RST works (basically by summing up time and distance covered within a circle). The R code, a short guide, and example dataset are available as links at the end of the paper.

Here is the spaghetti from above (really tracks of Grey-headed albatrosses) with the behavioral states labeled using RST:  93,481 points and this behavior classification took only 14 seconds to run!  albatrosstracks_rst