Field work vs. lab work

During my first two years in Chile I worked primarily in the field. This year however, with the exception of one quick week in the field, I’ve been doing all my work in the laboratory. The field and the lab both have their advantages and disadvantages, pluses and minuses, ups and downs. My experience working in both environments has caused me not to pick a favorite, but rather to conclude that the strongest research approach contains both lab and field components. In this post, I’ve identified five major research concerns; experimental control, ecological relevance, animal maintenance, expenses, and working conditions. For each research concern, I’ve written a short discussion comparing and contrasting the relative benefits of lab and field approaches (note: this post is mostly geared towards those that work with vertebrate animals).

Experimental control

There are two types of things you want to control in an experiment; variables and sampling opportunities, and the lab clearly wins in both categories. In the lab, most variables can be controlled and sampling opportunities are only limited by experimental design or physiological limitations. In the field, however, there is only so much control researchers can assert over certain variables. For example, weather is pretty much uncontrollable. Sure, you can put a heating pad in a nest box or you can water an experimental plot during a drought, but that’s pretty insignificant in comparison to a temperature and humidity-controlled animal room. And as for sampling opportunities, oftentimes field researchers are subject to the whims of their animals.

                During my field research there were several variables I wish I could have controlled. For example, because degu pups are born in underground burrows and do not come aboveground until three weeks of age, there was no way for me to measure or sample the pups during this time. Additionally, because female degus live in social groups and raise their offspring in communal burrows, there was no way for me to determine which pups belonged to which mother within a social group. Using genetic information could have solved this problem, but the genetic tools for degus are still being developed.

                For my lab experiment, these problems don’t exist. I know the exact day my pups were born, who their mother is, and how much they weighed at birth. I can now weigh and measure the pups and mothers as frequently as I need to, and I can also observe the behavior of my degus by videotaping them whenever I want. This was another thing I couldn’t do in the field- because degus live underground, there was no way for me to see if my experimental treatment was affecting the level or quality of maternal care. In conclusion, the lab is a better environment for experiments that require a lot of controlled variables or specific sampling points. This comes at a cost, however, which brings me to my next research concern:

Ecological relevance

                As soon as you bring an animal into an artificial environment you alter its behavior and physiology. This can be exemplified by the struggles of captive breeding programs for endangered animals; over the years researchers have been able to determine necessary stimuli conducive to breeding, but there still are difficulties even with the best-studied species. In the lab, we know that degus will allonurse (meaning, a mother will nurse pups other than her own), but is this true in the field? Do degus only allonurse in the lab because they have plenty of food? Or maybe because of space limitations, it’s too difficult to keep litters of pups separate? Until we can put cameras in real degu burrows, we can only assume that degus allonurse in the wild.

                The point is that animals may behave differently in the lab because there may be significant factors that we, as humans, are not aware of. Animals may perceive or be more sensitive to certain wavelengths of light (unlike us, degus can see in the UV spectrum), odors, or noises. What may seem innocuous to a human may be stimulating or frightening to another animal. A combination of several factors ultimately creates an environment that is totally different and separate from an animal’s natural living conditions.

                However, there are ways to make laboratory experiments more ecologically-relevant. Some labs, for example, house their animals in outdoor enclosures. This limits the control of things like temperature and weather, and it may also be harder to sample or observe animals in such a large space, but it does allow animals to experience more natural conditions and cues. Experiments can also be more ecologically-relevant by using stimuli that animals would typically experience in the wild. For example, if you want to study the stress response of an animal, you could use more “natural” stressors such as cold, rain, or unpredictable food availability rather than something artificial like playing a loud radio for 30 minutes. Lab and field experiments aren’t necessarily separate things, and it’s up to researchers to decide where their experimental design will fit best on the continuum between experimental control and ecological relevance.

Animal maintenance

                Currently, I’ve been spending a lot of my time cleaning degu cages. I don’t mind doing animal husbandry, but at times I do feel a little under pressure because I know that I’m the sole person responsible for the health and wellbeing of my animals. It’s a responsibility I take seriously, so I make sure to give my degus the best care possible. The level of responsibility is different in the field, however. Besides checking traps often and giving degus an extra handful of oats while they’re waiting to be returned to their burrows, I can pretty much let the degus take care of themselves. So in terms of animal maintenance, the field usually wins, especially because animal care costs can be rather high, which brings me to my next point:

Expenses

To successfully carry out experiments, researchers need to pay for equipment, supplies, and field or lab assistance. If the experiment will take place far from home, researchers may also need to pay for travel, housing, and other various costs such as permits, shipping expenses, etc. Both lab and field experiments can be expensive- it really depends on your experimental design and whether you already have some of the necessary equipment.

For me, it was the really the international component that made my experiments expensive. Both my lab and field experiments required me to pay for plane tickets and housing costs, which is something a researcher wouldn’t normally have to worry about. As for lab and field-specific expenses, they ended up being about the same; for my field experiment I had to rent a truck and for my lab experiment I needed to pay for animal care during the nine months I was back in the U.S. Equipment costs were more expensive for the lab experiment (terrariums, video cameras, etc.) but only because I had access to pre-existing equipment for my field experiment (animal traps, radiocollars, etc.). So, in the end, neither the lab nor the field was more expensive than the other, and the total cost of an experiment is really determined by experimental design.

Working conditions

                I hate getting up early. Maybe not as much as some people, but when the alarm clock goes off and it’s still pitch-black outside, my inner voice grumbles a steady stream of obscenities as I get out of my warm, cozy bed. Now, you might think that getting up early is a burden that only field researchers carry, but that’s not always true. For my current lab experiment, I’ve been getting up around 6am every day for two reasons: 1) in order to be consistent with my field experiment, I’m trying to collect blood samples before 9am and 2) the Santiago metro becomes a living hell during rush hour, and in order to keep my sanity I have to catch the subway or bus before 7am. While I did have to wake up even earlier when I was doing field work (4:30am, woo-hoo!), I would allow myself to take off one day per week. For my current lab experiment, though, there’s something I have to do every, single, freaking morning. So when it comes to early mornings and total work hours, neither the lab nor the field wins.

                Fieldwork can sometimes be miserable. It can be freezing cold or baking hot, or the dew from the grass can make your boots and pant legs soaking wet. Opening 150+ traps in the dark is no fun, either, nor is hauling traps up and down a big hill (or small mountain, it depends on your perspective). In all truthfulness, my fieldwork is relatively tame (one of my friends works in a very hot, humid forest where she spends several hours a day tramping up and down steep hills, all while looking out for venomous snakes) so I really shouldn’t complain. But, nevertheless, I’ve had my share of trials and tribulations. One time I had to go to the hospital because of a really bad hand rash. One time an opilion (a very large, creepy arachnid- see photo) crawled up my pant leg. I’ve also fallen down hills, tripped over rocks, and had serious wardrobe malfunctions (i.e. ripping open the seat of my pants).

But I would never give up field work for anything. I love being outside all day, and my time in the field has allowed me to witness and experience so many cool things. I’ve seen moustached turcas (see picture) carrying grubs in their beak, running to and from their nests. I’ve seen an iguana eat an akodont (a small, mousey-like rodent) and an eagle knock a caracara (a small bird of prey) out of the sky. One of the coolest things I saw was a wasp trying to drag a paralyzed tarantula through one of my degu traps. These wasps are called “tarantula hawks” and spend their days zooming over the ground, searching for tarantulas. When they find a tarantula, they deliver a powerful string which doesn’t kill the tarantula but effectively paralyzes it. The tarantula is then dragged to the wasp’s nest where it is eaten by the wasp’s larvae after they hatch. 

So while maybe the lab is more comfortable, the field is definitely more fun. And when stuff doesn’t work out or when you make an experimental mistake, if you’re in the field you can reason that “Well, at least I got to spend the day outside.” (this quote is attributed to Dr. Sara O’Brien). Wherever I end up after grad school, I’m sure I’ll be doing field research!

The opilion.

My favorite bird- the moustached turca!

The tarantula and the tarantula hawk.

Research is hard.

It’s a simple statement, but I can guarantee that every professor, post-doc, and graduate student would agree that research, while rewarding and ultimately worthwhile, is nevertheless a difficult thing to do. Every type of biologist faces their own unique set of challenges: the biological modeler may struggle endlessly with programming code, the molecular biologist may spend months troubleshooting an experiment, and the ecologist may watch helplessly as a severe storm decimates their study species. Catastrophes befall all scientists, but there are ways to prevent and mitigate the effects of bad luck. Here are a couple of challenges I faced during my research and how I dealt (or wish I dealt) with them:

Rule #1) You can never be too prepared.

Before my first field season in Chile I spent a lot of time thinking about how I was going to collect my data, and I ended up writing several drafts of equipment lists. In the end, I brought down some equipment that ended up being unnecessary (for example, I brought 10 radio trap transmitters, these nifty gadgets send out a signal when the trap door shuts. While it was great in theory, it didn’t work well in the field because birds would get caught in the traps and cause false alarms, and also 10 transmitters wasn’t very helpful when I was  generally using 100 traps at a time). But, I also ended up bringing some equipment that really saved the day, like a hand-crank centrifuge that I was able to use when we stayed in a cabin without electricity in Parque Nacional Fray Jorge. The conclusion is that it’s best to bring more things than you think you’ll need (as long as you stay within your luggage limit).

But being prepared is more than just bringing the right equipment- it’s also about reading everything you can and communicating with your collaborators and other, more experienced researchers. Grant writing was actually a great way to prepare for this- by thoroughly reading the literature and getting feedback from my mentors, I was able to iron out some of the more theoretical aspects of my projects.

Rule #2) Don’t put all your eggs in one basket

There’s always the chance that an experiment will completely and utterly fail, so it’s a good idea to have multiple projects so you’ll be able to come home with something. During my first field season in Chile, it became apparent that my main project would require another year of field work, which meant that I would have to secure more funding. In addition to this uncertainty, my main project was also ambitious and risky. So, to make sure that my first field season (which was five months long) wasn’t wasted, I also pursued two, additional projects. In the end, I completed one of these side projects during my first field season and was able to publish a manuscript, so even if things hadn’t worked out with my main project, I would have at least gotten something out of my time in Chile.

Rule #3) Make contingency plans

No matter how much you prepare and no matter how familiar you are with your study system, key events may still not go according to plan. For example, this year I’m doing a laboratory experiment that examines the effect of stress on the quality and quantity of maternal care. The easiest way to do this experiment would have been to catch pregnant degus in the field and bring them into the lab where they could then give birth. However, transference to captivity is stressful, and I didn’t want pre-natal stress (stress while the mothers are still pregnant) to affect the pups. I could have also caught pregnant females very close to parturition, so then there wouldn’t be much time for the mother’s stress to affect the pups in utero, but then the disadvantage would be that females would still be adjusting to captivity after they gave birth and maybe this would affect maternal care.  So, I instead caught females a year before the experiment and then mated them in the lab. However, I knew that my degus might have a low fertility rate, so I made a plan in case this happened (another thing would have been to collect more animals than I needed, but this wasn’t possible logistically because of space limits. Also, it would have cost a lot more money to buy more terrariums, food, and animal care. And it also has ethical problems- you don’t want to use an unnecessary number of animals).

 

So what happened? Unfortunately, only 40% of my degus got pregnant, so I went according to my contingency plan and trapped some very pregnant degus in the field and brought them into the lab. While this wasn’t the ideal situation, I will at least be able to compare my lab-mated degus with the field-mated degus. In the end, I think it will work out, and the important thing is that I planned for this and made sure that I had the time and resources to trap these additional degus. And because the current experiment isn’t totally ideal, I’ve decided to collect some additional data to take advantage of the new group of degus (some of these wild degus, due to individual differences, will be “more stressed” than others, so I’m measuring the mother’s CORT levels to see if they correspond with the amount of maternal care they give to their pups).

Rule #4) Be realistic

Don’t try to take on more than you can manage, and if things get too busy or crazy, figure out if there’s some project aspect you can drop, delay, or share with a collaborator. I’ve had chances to collect additional data or even add projects to my current research, but I’ve sometimes passed up these opportunities so I could spend more time writing papers and grants, sleeping (this is actually really important because I had to safely drive a truck at 5:00am almost every day last season), and having fun. Before I go to Chile my advisor always says “Be safe, work hard, and make sure to have some fun,” so I try to do those three things in that order.

Rule #5) Be flexible

As I explained in Rule #3, not everything goes to plan. But oftentimes an experiment can be salvaged by altering data collection methods or changing the number of treatment groups or controlled variables. Working in the field means you can’t control everything, and this is especially true when you work with wild animals because they can leave your study area, get eaten, or refuse to be recaptured. Being a successful field researcher means you have to know when to make compromises- if you compromise too much your data may lose a lot of significance, but if you refuse to compromise you may end up with no usable data at all.

Rule #6) Always, always, always back up your data.

I’ve always strictly adhered to this rule, and it really saved my bacon when my computer completely and utterly died two days after arriving in Chile this year (screw you, Apple). Luckily, I had just backed up my data on my portable, external hard drive so I was able to easily move my files to my new laptop (I now have a Lenovo which I really like except for the Spanish keyboard). When I’m collecting data, I back up my computer at least once a week, and I also try to make sure that my data is stored in at least two separate places. I’ll also occasionally email my data spreadsheets to my advisor and collaborators, just in case. And finally, I make sure to record all my data in a lab or field notebook, and I always try to enter these data into my computer within a day of collection. For field notebooks, I highly recommend Rite-in-the-Rain notebooks which are tough, waterproof, and are used by field biologists everywhere.

Rule #7) Accept defeat gracefully and with humor

This may be the hardest rule to follow, but if you can manage to keep a positive outlook and not get overly stressed when everything goes to hell, then you’ll be a happier, healthier researcher. Every researcher, at some time in their career, will have a project that will totally fail. The best thing to do is to salvage what you can or go to one of your back-up plans, but if all else fails, then it’s best to just move on and plan for future experiments. I’ve only spent a few years in academia, but from what I have seen so far, I think that the most successful researchers are those that are resilient and optimistic. To be a successful researcher you have to accept that failures will occur, because if you give up after one or two failures, then you’ll never get a grant funded or a paper accepted.

While these seven rules are not exhaustive, they do provide a good, general philosophy for any scientist working in the field. If any readers have some additional advice, please feel free to send me your comments and I’ll post them on this blog!  

Chile, 2013!

Welcome back! This is my third and final trip to Chile, and at the end of this October I should (cross your fingers) be finished collecting data for my thesis. While I was primarily working in the field in 2011 and 2012, this year I will be spending most of my time in the laboratory. Here’s a quick synopsis of my past and current research:

As I’ve mentioned before, I’m primarily interested in the development of an animal’s hormonal stress response. The main stress hormone in degus (and humans) is cortisol (CORT), which is regulated by the hypothalamic-pituitary-adrenal (HPA) axis. Previous work on mice has shown that experiences after birth (termed “post-natal effects”) can influence the development of the HPA-axis. Mouse pups that do not receive enough licking and grooming from their mothers will end up with hyperactive stress responses, which means that they will secrete high levels of CORT in response to stressors and will have a harder time bringing CORT levels back to normal after a stressor has passed. Because stressed mothers lick and groom their pups at a lower frequency, this means that maternal stress can affect the development of the offspring stress response.

One of my main research goals is to see if stressed mothers produce pups with altered stress responses. The other main goal of my research is to determine whether the degu’s social structure can help buffer the negative effects of post-natal stress. Because degu females live in social groups and will care for other female’s pups (this reproductive strategy is called “plural breeding with communal care”), if one mother in the social group is stressed, then other females within the group may be able to compensate for the lack of maternal care.

In 2011 and 2012, I did a manipulative field experiment where I implanted certain degu mothers with CORT pellets after they gave birth. In some social groups all the mothers were implanted with CORT, while in other groups 0% or 50% of the mothers received CORT implants. After the pups emerged three weeks later, I trapped them and took blood samples to determine their stress responses.

However, one thing that I was not able to do in the field was to confirm that the CORT pellets decreased the amount or quality of maternal care. Because degus spend the first few weeks of their lives underground in burrows, it’s impossible to observe mother-pup interactions during this important time. So, what I’m doing now is basically replicating my field experiment in the lab so I can also observe the behavior of the mothers and pups.

Currently, I have 32 pregnant female degus in my collaborator’s animal rooms at the Universidad Católica de Chile. My females are housed in pairs in acrylic terrariums, and as soon as they give birth I will implant certain females with CORT and start filming them with a home security camera system. I’ll be watching the videotapes later and will score behaviors such as time spent at the nest, how often the pups nurse, and the rate of licking and grooming. When the pups are about three weeks of age, I’ll take blood samples so I can compare their stress responses to those pups that I caught in the field.

So that’s a basic overview of what I’m doing this year, and stay tuned for more posts! I’ll be writing about the importance of flexibility and contingency plans in scientific research, the comparison between doing lab and field research, and the different theories about why the HPA-axis is plastic during development.

Early environments, parental care, and the importance of touch

The conditions that we experience early in life can have profound effects on our future physiology. For example, during the winter of 1944 the Netherlands suffered a famine because the Germans blockaded food and fuel from reaching the country. Mothers who were pregnant during this time gave birth to babies that had thrifty metabolisms, which unfortunately led to a high rate of diabetes, obesity, and other problems for these "Dutch Hunger Winter" children. Because the prenatal environment was nutritionally lacking, it made sense to shift towards a thrifty metabolism, but when the environment changed a thrifty metabolism phenotype proved to be detrimental. But what the Dutch Hunger Winter children proved is that our physiology is plastic during early life, and that conditions during this time will influence our future physiology.

In addition to affecting metabolism, early life nutritional states can also influence other physiological parameters. Zebra finch chicks grow at a slower rate if their parents are inattentative and do not feed them often enough. If brood sizes are enlarged (which causes chicks to face increase competition for food), barn swallow chicks will have decreased immune responsivness as adults and male collared flycatchers will face decreased, future reproductive output. While these examples all show that adequate nutrition is important for young animals, they also show that the quality of parental care can play a large role in physiological development.

Touch is another component of parental care that has been shown to profoundly affect post-natal development in mammals. This was shown in a very crude experiment by Emperor Frederick the II of Sicily (reign 1220-1230). Frederick wanted to know what language was the "natural" language of humans, so he devised an experiment where he had some infants placed in isolated rooms where they would be unable to hear any human speech. Maids were hired to clean and nurse the babies, but the maids were not allowed to spend any more time than necessary with the infants in case they accidentally talked in the infants presence. Frederick believed that when the infants reached an age where they could talk, they would reveal the "natural" human language (most likely, he thought, Latin or Greek). What happened instead was that all the infants languished and died. And so we now know that if babies are deprived of touch, if they are denied love and affection, they will most surely die very young. (Here's a happier research example on the importance of touch; premature infants are often placed in fancy incubators because they need special respiratory equipment, protection from infectious diseases, etc. But even with this special care, many infants still fail to thrive or require longer recovery times than expected. However, by gently stroking an infant's legs and arms for just a few, short periods everyday, researchers found that growth rates increased by 50%!)

Researchers have also shown that touch is very important for the development of young rats. Rat pups that are not licked and groomed enough during the first week of life will develop hyperactive stress responses, meaning that these pups will have high corticosterone (CORT) levels in response to stressors. Additionally, these pups will have poor negative feedback, which means that it will take them longer to decrease their CORT levels after responding to a stressor. Overall, pups from low-licking and grooming mothers will be exposed to more total CORT during their lifetimes, which may make them more susceptible to stress-related pathologies. Which leads us to this question; how does low-licking and grooming cause pups to develop hyperactive stress responses?

A hyperactive stress response is characterized by high stress-induced CORT levels and poor negative feedback. Poor negative feedback is often due to low levels of CORT receptors (think about it this way; during a stressor an animal has a surge in CORT, but after the stressor passes they need to decrease CORT production. Parts of the brain know when to shut down CORT production based on how many CORT receptors are bound. So, if you have fewer receptors, it takes longer for enough CORT receptors to be bound so the brain "knows" when to shut off CORT production).

OK, so pups with hyperreactive stress responses must have lower levels of CORT receptors, and this is exactly what Liu et al. (1997) found. However, CORT receptors are not permanent fixtures- they are constantly being recycled all throughout the body, so how can an event early in life cause a permanent change in receptor expression levels? The answer is at the level of the gene; basically, conditions early in life can affect how accessible the CORT receptor gene is to transcription. Weaver et al. (1997) found that the promoter region of the CORT receptor gene was highly methylated in the hippocampus. DNA methlyation occurs when a cytosine is converted into 5-methlycytosine, and this conversion makes it more difficult for transcription factors to bind to the promoter region, thus inhibiting transcription.

Another interesting thing about pups from low-licking and grooming mothers is that they will grow up to be low-licking and grooming mothers themselves. An elegant experiment by Francis et al. (1999) showed that the low-licking and grooming trait is inherited through behavioral processes, not genetic processes. Basically, Francis et al. (1999) took pups from low-licking and grooming mothers and switched them with pups from high-licking and grooming mothers (this is called a "cross-fostering study"). When the pups grew up, their own licking and grooming frequency was determined by the licking and grooming behavior of their foster mother, not their birth mother. The low-licking and grooming trait is thought to pass behaviorally because it's been shown that increased levels of CRF (corticotropin releasing factor, an important stress hormone that stimulates release of ACTH which then causes increased secretion of CORT) inhibits maternal behavior.

For further reading: As always, I highly recommend the book "Why Zebras Don't Get Ulcers" by Robert Sapolsky (I shamelessly borrowed his examples of Emperor Frederick II, premature babies, and the Dutch Hunger Winter children). Some great, foundational papers on maternal effects on the development of the mammalian stress response include; Liu, D. et al. 1997. Maternal care, hippocampal glucocorticoid receptors, and hypothalamic-pituitary-adrenal responses to stress. Science 277:1659-1662. Francis, D. et al. (1999). Nongenomic transmission across generations of maternal behavior and stress responses in the rat. Science 286: 1155-1158. Weaver et al. Epigenetic programming by maternal behavior. Nature Neuroscience 7:847-854. 

Degus eat money; all about grant proposals

(The first half of this post is a general introduction to what grants are and how I applied for them. The second half of this post is advice for students writing grant proposals.)

Scientific research requires thoughtful preparation, careful execution, and painstaking analysis. It takes a lot of time to design a good experiment, carry it out, analyze the data, and then finally communicate the results. But what many people don't realize is that scientists have to spend additional time and effort to just get the money to do their research. This money is typically in the form of grants, which can be from the government (like the National Science Foundation, National Institutes of Health, Department of Energy, etc) or independent organizations (scientific societies, conservation groups, etc).

Typically, the Principal Investigator (the head of the lab) is the person in charge of securing research funds, while the graduate students, post-doctorates (post-docs are researchers who already have their PhDs), and technicians are in charge of carrying out the research. If a graduate student or post-doc wants to pursue a research project that's unrelated to their adviser's current grants, then they usually have to find their own research funds. That's what happened in my case, so by working on other scientist's grants and applying for my own funds, I've been able to carry out my research in Chile.

During my first field season in Chile, my collaborator from the University of Tennessee at Chattanooga (Dr. Loren Hayes) funded my research through an International Research Experience for Students (IRES) grant from the National Science Foundation (NSF). After my first field season in Chile I knew that I wanted to return, so I started applying for other grants (the IRES grant can only fund each student one time).

The first big step was finding grants; through a series of internet searches I found about 11 grants that seemed likely to fund my research project. The first place I looked at was my own university, Tufts! The Tufts Graduate School has several grants-in-aid of research that they award every semester. While small (max. of $700), these grants are a great springboard and have a high funding rate. The Tufts Graduate School also has a list of grants on their website (http://gradstudy.tufts.edu/researchteaching/opportunitiesoutsidetufts/dissertation.htm), which helped me find a few more funding opportunities. 

My adviser and thesis committee also recommended a few grants to me, like the NSF Doctoral Dissertation Improvement Grant, the American Mammalogist Society grant-in-aid of research, and the Sigma Xi grant-in-aid of research. And by chatting with other students in my department, I found out about grants available from Graduate Women in Science and the American Philosophical Society. After more internet searching, I also found grants from the Explorer's Club, the National Geographic Society, the Animal Behavior Society, and the Society of Comparative and Integrative Biology (the reason I'm listing all of these grants is so that it might be useful for other students in my field).

Then, the next big step was actually writing the grants. Writing the first grant was the most difficult, and then it got a lot easier from there since I could reuse sections from my first grant. Here's an outline for a basic grant:

1) Introduction- The first few sentences should discuss the unique phenomenon that's related to your project. You want something that will immediately interest your audience and make them agree that your project is going to help explore an important scientific question. The introduction should then continue, with each sentence logically building on the last, until you find a good place to introduce your study system. And then the introduction should continue on, narrowing and narrowing until you finally get to your hypothesis, and then don't be afraid to state your hypothesis in bold letters, italics, or whichever way you think it will best stand out!

2) Methods- This section should be brief and to the point, avoid going into too much detail! You want to convince your audience that what you're doing is feasible, but you also don't want to spend a lot of time describing your techniques. Don't forget to include the important facts like place of work, time frame, and number of animals. Try to present your steps in chronological order- this makes the section more logical and easy to read.

3) Expected Results- State what you think you're going to find, don't be afraid to use different fonts to make parts of this section stand out. Depending on the length of the grant, this can also be a good section to discuss the broader impacts of your study. How will this study make a novel contribution to your field of research? Are you going to do any outreach activities? Are you going to mentor or collaborate with other people?

4) References- Grants usually have word or space limits and you almost always want to write more than you're allowed. So, for once, avoid parenthetical citations and opt for superscript numbers. You can also save space by limiting the number of citations you use- the reviewers want to know that your statements are backed up, but they're more interested in what you're planning to do in your project.

Here are some general tips for writing grants;

-Ask for your letters of recommendation well in advance.
-Double-check that you've followed ALL grant guidelines! Many organizations will toss out a grant if you forget a section, use the wrong format, etc.
-Give yourself a lot of time to write your first grant, then the next grants will take much less time.
-Ask other students in your department if you can read their funded grant proposals. This was probably the most helpful thing for me, I learned a lot by reading other people's grants!
-Ask your peers to read your grants over, it's good to get a first round of editing before you send your grant to your adviser.
-Try to submit your grant at least one day before the deadline. Sometimes there are problems with the website because of high traffic and you'll experience technical problems, don't cut it close!
-Spend a lot of time on your grant budget. There's usually no space limit for your budget justification, so go ahead and lay out every detail if you can. I usually make a list of costs, and then I write a few paragraphs explaining how I calculated each cost.
-Start writing grants early in your career, it's best to start when you're an undergraduate! The more practice you have, the better. Many universities have undergraduate research grants, and 1st/2nd year graduate students should definitely apply for the NSF graduate research fellowship.
-Because your first grants will most likely not be funded, apply BEFORE you really need research funds so you can get feedback and improve your grant writing skills.
-That being said, don't let grant writing get in the way of your research, try to write grants only when you need the money or when you have the time to write grants.
-Don't be discouraged by rejections. There's a lot of rejection in science, and the best scientists are those that don't brood over failed attempts and instead jump back up and try again. Be persistent. 

I hope this post was informative and helpful. Grant writing isn't the most exciting thing to write about, but I spend quite a bit of time writing grants and it's an important part of scientific research, so I felt that the topic was appropriate for my blog.

Measuring Stress Hormones in Wild Animals; Different Techniques and Their Comparative Advantages and Disadvantages

Measuring circulating levels of stress hormones (cortisol, corticosterone, etc.) in wild animals can provide valuable information for many types of studies. For example, conservation biologists may want to determine if high levels of snowmobile traffic are associated with increased cortisol levels in elk. Behavioral ecologists may want to know if white-faced capuchins (a type of monkey from Central/South America) have lower levels of cortisol if they come from a troop with a high level of social support. And physiological ecologists may be interested in whether Galapagos marine iguanas have a corticosterone rhythm driven by photoperiod or tidal cycle (*see bottom of page for more details).

In order to measure levels of corticosterone or cortisol (CORT), researchers need to obtain blood, feces, urine, saliva, feathers, or hair from their animals. Here’s some information about each method and their advantages and disadvantages:

Blood

This is the most common way to measure CORT concentrations. Because hormones circulate through the blood, this method provides researchers with the most functional measure of CORT. Since CORT has a very high concentration in the blood (compared to other hormones), blood samples do not need to be very big (for degus, 30uL of whole blood is usually enough to determine baseline CORT). The downside to using blood samples is that within 3 minutes of encountering a stressor, an animal’s CORT levels start to increase. So, if a researcher wants to determine baseline levels of CORT, they must get a blood sample within 3 minutes of capture.

Another downside of using blood samples is that each sample is only a snapshot of an animal’s stress response. In order to get a more integrated picture of an animal’s stress profile, researchers usually take a series of blood samples over the course of 1-2 hours. That way, they can see the rise from baseline, the peak of the CORT response, and then the gradual decrease caused by negative feedback.

After obtaining a blood sample, the samples must be kept chilled until they can be spun in a centrifuge (you want to spin within 24 hours of collection). Once the samples are spun, the plasma or serum (the clear layer on the top) needs to be drawn off and saved for analysis. The plasma or serum must be frozen at -20˚C and can be stored for several months. These requirements are fairly reasonable, but sometimes field sites are far away from cities. Last year I worked in a national park that was pretty remote, so I had to manually spin my samples with a hand-crank centrifuge and store my samples in a cooler full of ice. Luckily, steroid hormones like CORT are fairly sturdy, so lengthy processing times aren’t a big deal.

Finally, to process the plasma samples, researchers need to run a competitive binding assay called a radioimmunoassay (RIA). Basically, known amounts of radiolabelled CORT and CORT-antibody are added to the sample. The radiolabelled CORT and normal CORT from the sample will compete to bind with the antibody. The unbound radiolabelled CORT is then washed off, and the radioactivity is measured. The more CORT you have in your sample, the more it will bind with the antibody and displace the radiolabelled CORT. Therefore, the higher the CORT in the sample, the lower the radioactivity. RIAs are fairly straightforward but require some special equipment, so they usually have to be run in laboratories that frequently measure hormone concentrations (like my lab!). The enzyme immunoassay (EIA) is another way to measure hormone concentrations and requires less equipment, but is generally more expensive to run than a RIA.

As an additional step, researchers may also want to measure levels of corticosteroid binding globulins (CBG). CORT binds to CBG in the bloodstream, and it’s thought that only unbound CORT is “free” and able to bind to cell glucocorticoid receptors. Therefore, the functional measure of CORT is the “free” CORT, so some researchers also measure CBG levels to determine the relative amounts of unbound CORT. However, it’s still under debate as to whether only unbound CORT can bind to glucocorticoid receptors, and it has also been pointed out that without CBG, CORT would be unable to circulate through the whole bloodstream and reach all the cells in the body.

My blood samples are collected in little, glass microhematocrit tubes. I put them in plastic tubes with clay in the bottom so the bottoms of the microhematocrit tubes are plugged. I then put three plastic tubes in a conical tube which are stored in a cooler full of cold packs.

Loading the centrifuge with my samples.

After spinning the blood sample, it separates into hematocrit (the bottom red stuff) and plasma (the top, clear stuff). The plasma is what has the hormone, and that's what I draw off with a glass Hamilton syringe (shown on the table).

I pipette the plasma into a plastic eppendorf tube which then goes into a -20C freezer.

Saliva

            CORT can pass from the blood to the saliva, and salivary samples are actually a common way to measure cortisol levels in humans. While captive animals can be trained to lick, chew, or drool on something, obtaining sufficient salivary samples from wild animals in non-invasive manner is most likely impossible. However, it usually takes 20 minutes after a stressor for CORT levels to increase in the salvia, so taking salivary samples could give researchers more time to collect a sample after capture.

Feces/Urine

Another way to assess CORT levels is to measure the metabolic products of CORT in the feces. However, because this method measures the metabolites of CORT, it doesn’t have quite the same functional value that a blood sample does. There are several steps involved in breakdown of CORT, and CORT metabolite formation can be affected by sex, season, metabolic rate, and diet. Therefore, experimental power is lost when using fecal samples for CORT analysis because animals cannot be compared between different seasons, locations, or sexes.

The main advantage of using fecal samples, however, is that they’re fairly non-invasive. Researchers can follow individuals and collect feces, or in the case of small mammals, check traps after a certain period of time and collect any feces from the bottom of the cage. Using feces to measure CORT levels is also great for repeated sampling, since frequent blood sampling and other more, invasive measures can change an animal’s stress response.

Using fecal samples to determine stress hormone levels makes a lot of sense if you’re working with an animal that’s endangered or difficult to capture (like an arboreal monkey). Another advantage of using feces is that they represent an integrated measure of CORT exposure; if you know how long it takes for an animal to form a fecal pellet, then you can estimate their total CORT exposure over a period of several hours.

Collecting feces is usually pretty easy, but investigators should always try to obtain fresh samples from a known individual. If this isn’t possible because the study animal is hard to find or lives in an aquatic environment, then dog trackers can be used. The Wasser Lab at the University of Washington (my alma mater!) uses canine trackers to find feces from whales, tigers, wolves, and other, elusive endangered animals (http://conservationbiology.net/conservation-canines/#scat).

 Storing fecal samples can be tricky; temperature, storage liquid, autoclaving, and storage time can all affect fecal glucocorticoid metabolite (FGM) concentrations. Sample mass can also affect FGM concentrations, as very small samples have disproportionally higher FGM levels. Getting an adequate sample mass can be difficult when studying small animals like songbirds, mice, etc., so many studies have to combine several fecal samples in their assays.

After homogenizing the fecal samples, FGM concentrations can be measured via RIA or EIA. Another downside of using fecal samples is that the CORT-antibody may not bind with all the different CORT metabolites. Therefore, researchers usually need to do a validation experiment to determine that increased CORT levels in the blood correspond to increased FGM concentrations in the feces. One advantage of using feces, though, is that by measuring levels of metabolites the researcher is essentially measuring “free” CORT, so there’s no need to measure levels of CBG.

Like feces, urine also contains glucocorticoid metabolites (and some unmetabolized CORT). In the field, however, it’s very difficult to collect urine samples in a non-invasive manner, so urine collection for CORT analysis is rarely used outside of the laboratory.

Feathers/Hair

             Believe it or not, you can actually measure CORT in feathers. During molt (feather growth), each individual feather is vascularized, and CORT from the blood can be deposited in the feather. After the feather stops growing, the blood supply is cut of and, supposedly, no further CORT can be deposited in the feather. Therefore, feathers provide a good, integrative measure of CORT during the period of molt.

            Plucking a feather is easy to do and there’s no time limit, unlike blood sampling. Using feathers can also be non-invasive if the researcher obtains naturally molted feathers. And another big advantage of using feathers to determine CORT levels is that feathers from dead birds can also be assayed (like museum specimens!). Also, feathers do not need to frozen or stored in any specific manner.

            The big downside to using feathers for CORT analysis is that we’re still not too sure what form of CORT we’re actually measuring. Different antibodies have been found to have different levels of success during feather CORT analysis, which has made researchers unsure whether they’re measuring CORT, CORT metabolites, or other molecules similar in structure to CORT.

Another downside to using feathers is that, like feces, there’s a sample mass bias where smaller feather samples have disproportionally higher CORT. Getting a large enough sample can be difficult because researchers don’t want to compromise a bird’s flying ability, and there’s also the problem that different feathers are grown at different times, so pooling certain feathers together may not be appropriate. Feather mass requirements can also prevent researchers from examining portions of the feather (sections near the end of the feather are older, and thus represent a different time period than sections near the bottom), but a recent study found that different feather sections didn’t correspond to the CORT concentrations in the blood at the time of their growth, anyway. Feather CORT can be measured via RIA, but the preparation is a real pain (you have to mince the feather into little, tiny bits).

            CORT can also be deposited in hair during follicle growth. One advantage of hair samples is that they can represent CORT exposure over a very long period of time. The downside to hair samples is that it’s hard to collect a large enough sample in a non-invasive manner. And, like feathers, there are still a lot of questions about what the antibody is actually binding to in the assay. Another disadvantage to using hair is that the sample could be contaminated with other, CORT-containing secretions, like saliva or sweat.

            So, I’ve tried to give a basic, if somewhat extensive overview of the different ways to measure CORT in wild animals. After I was almost done writing this blog, I came across a great review paper that includes everything I mentioned and more: Sheriff, M.J. et al. (2011) Measuring stress in wildlife: techniques for quantifying glucocorticoids. Oecologia 166:869–887. Also, here’s an excellent review on fecal CORT: Goyman, W. (2012) On the use of non-invasive hormone research in uncontrolled, natural environments: the problem with sex, diet, metabolic rate and the individual. Methods in Ecology and Evolution 3, 757–765. And finally, my favorite feather CORT paper: Lattin, C.R. et al. (2011) Elevated corticosterone in feathers correlates with corticosterone-induced decreased feather quality: a validation study. Journal of Avian Biology 42, 247-252.

*(Photoperiod vs. tidal rhythms: most animals have daily stress hormone rhythms that are related to foraging opportunities. For humans, the morning is an optimal time to forage, so our cortisol levels often peak right before we wake up, and these increased cortisol concentrations help us go out, run around, and collect some food. For marine iguanas, corticosterone concentrations are influenced by time of day AND tidal cycles. This is because marine iguanas need to eat algae in the inter-tidal zone and the algae are most available during low tide, but because iguanas are ectotherms and need to bask in the sun before and after swimming in the inter-tidal zone, foraging is only possible during the day. However, this relationship still needs be teased apart, since the observed CORT peak during low tides could also be influenced by food intake.)

Radiocollars

Most of my degus have given birth! Which means that things are busy in the field and I have been spending lots of time implanting cortisol pellets and collecting baseline blood samples. After I finish implanting my cortisol pellets, I will need to put radiocollars on my female degus so I can determine social group composition. While you can usually tell which degus are in which social group by seeing what burrow they're trapped at everyday, this isn't totally reliable and the best way to determine social group composition is to see who spends the night with each other in the same burrow.

The radiocollars I have been using have good signal strength and long-lasting batteries, but the collar itself irritates the degu's necks and I usually have to remove the radiocollar after a few days. So, in order to protect my degus necks, I have been remodeling some of my radiocollars. My talented friend Cecilia figured out how to do this, here's the step-by-step process:

Original radiocollar- the thin wire digs into the degu's neck, and the edges from the crimp can cut the skin.

To improve the radiocollars, we decided to replace the wires with wide, brass bands. So, for the first step, we cut strips from a sheet of brass.

Next, we marked where we wanted to put the holes. We first used a small hammer and chisel to start the holes and then an electric drill to finish.

We then smoothed the edges of the holes with a sander attachment.

Next, we soldered a small screw to the end of the brass band and then sanded down the soldered area.

We had to remove part of the transmitter to reduce weight and create a better fit. The original transmitter is on the left and the altered transmitter is on the right.

Using superglue and Poxilina (a type of cement-putty), we attached the brass band to the transmitter.

And finally, we put heat-shrink-tubing over the transmitter and lower parts of the band.

This actually took quite a bit of time, but I always enjoy the opportunity to make and alter my research equipment. You never know what sort of new skills you'll learn when you go to graduate school....