After long weeks of preparation and anticipation, our very own made-from-scratch “buzzbox” is nearing completion!
The “buzzbox” is an original piece of machinery developed by a colleague of Professor Meyer. The buzzbox is essentially a simple circuit capable of generating a mild electric pulse. The two ends of the circuit are bare electrode wires. When these two wires are placed in a conductive material (e.g. sea water), the circuit is completed and an electric current can travel between the wires.
Our buzzbox will be used for staining animals for live imaging. Up until now, we’ve stained the animals by soaking them in a seawater/dye solution and allowing the cells to indiscriminately absorb the dye. While this technique has been easy to do and useful for certain experiments, it doesn’t allow for any amount of precision. With the buzzbox, we can use brief electric pulses to discriminately draw the dye into the cells. Based on the positioning of the wires, we can accomplish extremely precise staining. We can stain a small region of cells, a broad surface layer of cells, or even individual cells. Utilizing the buzzbox for staining will enable us to conduct a wider variety of experiments, generate fine-tuned, crisp images, and allow us to use different types of dyes.
Of course having the buzzbox itself exciting, but I’m extra excited to announce that I built the entire buzzbox with my own two hands (with an enormous amount of help from Professor Agosta and Clark’s Physics department)! Despite having never set foot in a workshop prior to that day, I managed to assemble, connect, and solder the box’s entire plethora of wires, dials, lights, and switches. I’m very proud of myself, to say the least.
Although it doesn’t fully work yet (we’ve yet to attach the electrode wires), the core circuitry itself is completed. Switching on the toggle and seeing the little red indicator light turn on was a small but thoroughly relished victory. Here’s hoping we’ll be able to complete construction and use it very soon!
Til next week,
In a few weeks, the Meyer lab will be involved in the Eureka! Program, a local science outreach program for middle school girls. Eureka! (associated with Girls Inc.) provides young girls with the knowledge, the experience, and the role models to encourage their interest in the sciences and higher education. Even today, mathematics and science programs in high school and beyond see an overwhelming male majority. Euerka! aims to change that by showing female students that science is fun and accessible.
Over the course of five weeks, students in the Eureka! program will spend time at Clark University, Worcester Polytechnic Institute, and Becker College, participating in a variety of classes and activities. For two days, the students will be here in our own lab learning about annelids.
In preparation for the Eureka! students, I’ve been preparing a number of activities, including a simple oil pollution experiment. Pollution, especially oil pollution, is a widespread problem for a number of marine species. Delicate invertebrates, like our own beloved Capitella teleta, are particularly sensitive to this pollution. I’ve designed an experiment that will highlight the dramatic effect even small amounts of oil can have on the development of C. teleta. The girls will be given several separate dishes of C. teleta embryos. They will then treat some dishes with a “high concentration” oil solution, some with a “low concentration” oil solution, and some will be left alone as a control. They will make predictions about the effect of the oil on the embryos, leave them overnight in their various solutions, and then observe them the next morning.
Since very little literature (read: none at all) exists on C. teleta’s tolerance for oil pollution, I did some preliminary experiments to figure out what concentrations we’d use for the actual experiment. With some motorcycle oil Professor Meyer brought from home, I made several different mixes of oil and seawater. Without any idea of what concentrations to use, I arbitrarily picked a few that covered a wide range (from 1 part per million (0.001%) to 1 part to thousand (0.1%)). Ideally, I aimed to find a low concentration that would have little to no effect, and a higher concentration that would cause clear physical deformities without outright killing the embryos.
My initial batch of test dishes went very poorly. Because I apparently can’t do math at 9am, I severely miscalculated the concentration of the oil. All of my batches were 1000-fold the intended concentration (oops!!). Unsurprisingly, the highest concentrations were dead by the end of the day. However, the lower concentrations (those that survived, anyway) displayed noticeable deformities
The next day, I tried another set of batches (this time I double-checked all my math!). I was able to find two perfect concentrations of oil. 1uL of oil per 1mL of water (or 1 part per thousand, 0.1%) has only a slight effect on the embryos after 24 hours. It makes them swim a little more slowly and causes their bodies to become a little lumpy. The “high” concentration is double that, 2uL/1mL, or 2 parts per thousand (0.2%). This concentration causes severe physical deformities and reduced swimming speed. Now knowing these experiment parameters and what to look for in the embryos, I was able to design the protocol sheet (the instructions, so to speak) and the observation worksheet.
Hopefully the students will enjoy the experiment and learn a bit about the extreme harmful effects that oil pollution can have on marine creatures!
Til next week,
This past week has primarily been spent conducting further calibration experiments, this time with the equipment itself rather than the experiment parameters. We’ve noticed that the microscope we use for live imaging movies seems to have an issue with focal plane slippage. When we film movies of animals over long periods of time (often a minimum of 1.5 hours), the focal planes seem to slowly slide out of focus. By the end of the movie, almost nothing is in focus, and it’s extremely difficult to observe development over time.
Below are two images of the same animal from a single movie. The first image is one Z-slice at time zero (the beginning of the movie). The second image is the same Z-slice after about an hour and a half (the end of the movie). Notice how the second image is out of focus and the animal seems to be further away from the lens.
Since we were filming live animals suspended in water, we initially assumed this slippage was due to the animals themselves sinking in the water over time, slowly moving away from the microscope lens above them. However, this same issue occurred even when the animals were positioned flush against the glass slide, thereby making it impossible for them to sink any further.
It was therefore concluded that the issue lay with the microscope itself. Perhaps the stage (the platform on which the slides rest) was sinking over time, or the grease & clay on the slides were heating up and melting, or the lens (which shifts up and down to film the Z-stack) wasn’t realigning correctly… or, crazily enough, the air conditioning vent on the ceiling, its breeze barely noticeable to human senses, was actually pushing the stage down on a microscopic level. The possibilities were nearly endless. We needed to solve this mystery before we proceeded with further live imaging movies. Otherwise, all our experiments would continue to be mucked up.
So, obviously, we conducted some experiments with sparkly nail polish.
Sparkly nail polish has the benefit of being inanimate, so we know it won’t move on its own. It also has conveniently suspended particles in clear enamel. Perfect.
We filmed several movies with the nail polish, experimenting with the size of the Z-stack, its depth and location, the thickness of the Z-slices, the length of the movie, whether or not the air vent was covered with a box… In all of these movies, we still observed a significant shift in the focal planes over time. Observe the images below. The nail polish itself it obviously not moving, yet the sparkles get very out of focus over time.
However, on the bright side, the degree of slippage has been dramatically reduced since we’ve updated the microscope software. Previously, we’d see shifts of ~17 micrometers*, a huge shift on the microscopic scale. Now, the overall shift is less than 2 micrometers. It’s now fairly manageable rather than completely detrimental.
The images below are from another nail polish movie, after the software update. In the previous nail polish movie, there was an overall shift of 8-10 micrometers. These two images show a shift of less than 2 micrometers. While still apparent, it’s not nearly as severe.
Unfortunately, the ultimate cause of this problem remains unknown. We weren’t able to draw any solid conclusions about the cause of the focal plane slippage from the nail polish movies. None of the tested factors seemed to have any consistent, noticeable effect. Professor Meyer believes it may simply be due to background vibration caused by people walking around in the lab, an unavoidable complication.
Our microscope mystery now somewhat solved, I’ll be returning to regular experiments. This week I’ll be continuing work on Hoechst concentrations, incubation times, and permeability.
Til next week,
(* 1 micrometer is one-thousandth of a millimeter, or about 0.000039 inches. Needless to say, it’s a very small unit of length. When you’re working with embryos, however, 17 micrometers is huge.)
This last week has been a flurry of experiments. Professor Meyer’s experience with live imaging is limited, so I’ve been conducting a variety of logistical experiments to figure out ideal imaging conditions.
For instance, I’ve been trying to find the best conditions for incubating the embryos in the fluorescent Hoechst dye. The better the dye is absorbed by the cells during incubation, the brighter and clearer the fluorescent images will be. Since we’ve been experiencing difficulties with the dye fading too rapidly (making multi-hour movies impossible), I’ve also been trying to find the conditions under which the dye will last the longest. Better absorption and longer fluorescent will enable us to take longer, more informative movies.
Thus far, I’ve experimented with exposure time, incubation temperature, and cell permeability.
Early last week, I conducted a simple multivariate incubation experiment. Embryos were collected and placed in six separate dishes. Three dishes were incubated in the Hoechst solution at 16C (~61F), while the other three dishes were incubated at room temperature (21C, 70F). In each of the temperature groups, one dish was incubated in the dye for 1 hour, one dish was incubated for 2 hours, and one dish was incubated for 4 hours.
At the end of each dish’s incubation period, we took time-zero images of the embryos. This was just to compare the initial brightness of the Hoechst. There was a very noticeable difference in the initial brightness of the dye between groups. All the embryos incubated at room temperature had brighter fluorescence than the embryos incubated in the cooler refrigerator, regardless of incubation time. This would seem to suggest that higher temperatures, at least within the tested range, lead to better uptake of the dye.
Below, on the left, is a time-zero fluorescent image of an embryo incubated in Hoechst for 1 hour at 16C. On the right is a time-zero image of an embryo incubated in Hoechst for 1 hour at 21C. The bright green dots are cell nuclei where the dye has been absorbed. Note the difference in the brightness between the two images.
Longer exposure time had mixed results… The 2-hour groups were brighter than the 1-hour groups, but the 4-hour groups were about the same, if not dimmer in some cases, compared to the 2-hour groups. These results were boggling, so we repeated a simplified version of the experiment. We compared two groups: embryos were incubated in the dye at room temperature for either 1 hour or 4 hours. This time, the 4-hour group did appear to be noticeably brighter.
I also experimented with increasing the embryos’ permeability, or the ease with which molecules can pass through the cells’ membranes. Higher permeability means that more molecules (e.g. more of the dye) can more easily be absorbed into the cell. A group of embryos were briefly exposed to a solution of sodium citrate sucrose, which softens the egg envelope and increases permeability. The sodium-citrate-sucrose-treated embryos were then incubated in the Hoechst dye and imaged. Compared to a control group (only incubated in Hoechst, with no exposure to the sodium citrate sucrose), the treated embryos showed significantly brighter fluorescence. The dye also remained brighter and more visible for the duration of the multi-hour movie. Yes! Success at last!
This treatment may be the key to creating ideal dye brightness for our movies! The downside is that the treated embryos are extremely tricky to work with; their softened egg envelopes become sticky and very vulnerable to damage. I accidentally ‘retired’ an entire dish of treated embryos when I attempted to pipette them into a fresh dish. Every single embryo got stuck to the inside of the pipette tip and wouldn’t budge. When I tried to shake them loose by pumping water in and out of the pipette, the embryos all lysed (their cells burst open and the organisms were instantly killed). …Oops.
Fortunately (or unfortunately…?), despite my remarkably quick and efficient manslaughter of innocent worm babies, this blunder wasn’t a particularly novel mishap. Worm death is not an uncommon tragedy in the annelid lab. The vast majority of the worms born in the lab are euthanized in the pursuit of scientific research*. On the bright side, our lab still boasts a far lower mortality rate than the worms’ natural habitat. It isn’t perfect, but it’s a fair sight better than the salt marshes. (I’m sure the worms would tell you so themselves, but they’re too busy bumming around in cozy, smelly mud.)
Til next time,
(*As an important disclaimer, all worms are treated humanely and ethically, in living and in death.)
For the past week I’ve been working on creating live imaging movies in the lab with C. teleta embryos. The first step is soaking the embryos in a fluorescent dye (in our case, Hoechst dye). Hoechst is a nuclear dye, meaning it’s soaked up by the DNA in the cells’ nucleuses. Later on, when we look at the embryos under a fluorescent microscope, the cells’ nucleuses will appear as bright dots inside the organism.
After the embryos have soaked in the dye for a few hours, we plate them on an airtight glass slide to keep them from drying out while we film our movie. The tools of the trade include vacuum grease (to form the seal), modeling clay (to make “feet” for the coverslip), and eyelash brushes. Yes, eyelash brushes are exactly what they sound like: a tiny brush made with a single human eyelash. They’re used to rotate and position the microscopic embryos once they’re on the slide. Professor Meyer described hairloop brushes, another common tool of cellular biologists. Researchers will select a piece of their own hair (hopefully clean!), form a small loop, and secure it to the end of a narrow tool. The taut loop is then used to cut microscopic cells and organisms. I had never realized how weird and creative lab tools can be!
In any case, back to the embryos.
The embryos (now properly plated on a slide) are placed in the fluorescent microscope. The software can be programmed to take a photo of the slide at specific time intervals (in our case, every 15 minutes). Additionally, the microscope can take “Z stacks”, or multiple stacked photos in the Z plane. The Z axis, in the 3D world, is up and down. Therefore Z planes are basically slices, or cross-sections, that stack vertically (like a packet of cheese slice singles). When you take a Z stack, you end up with is a series of images that, together, create a 3D model of an organism. So instead of a single 2D image of the organism at a particular depth (cheese slice #12, from the middle of the package), you get numerous slices covering a range of depths (cheese slices #1-25).
In our case, many of our initial movies had the microscope taking a Z stack of ~20 simultaneous photos every 15 minutes. In tiny C. teleta embryos, our slices were only a few micrometers thick (very very very thin cheese slices).
We imaged the live embryos for about 2 hours. Since the organisms were still alive and developing, the resulting movies allowed us to watch real-time embryonic development, in 3D, with visible nucleuses showing the movement and division of individual cells. That’s some seriously awesome stuff for the first week.
If this sounds too good to be true, that’s because it is. The main problem with our movies is that the Hoechst dye is fading too quickly. Below are fluorescent images from the lab demonstrating this fading. Both images are of the same Z slice, in the embryo, over time. The first image, on the left, is from time zero (the beginning of the imaging movie). The image on the right is from 1.5 hours later. Note the significant decrease in dye brightness.
Often the dye is completely bleached (no longer visible) within 30 minutes to an hour. That means we can no longer observe nucleuses, ergo cell movement and division. A maximum of one hour is an extremely small time window and doesn’t allow us to film actual development. This upcoming week, I’ll be conducting a few small scale experiments with the Hoechst dye. My goal is to find ideal conditions for incubating the embryos in the dye. I’ll be trying to find a way to make the dye soak in better or last longer, allowing us to create longer, more useful live imaging movies.
I’ll be mostly experimenting with incubation temperature, exposure time, and embryo stage. Wish me luck!
Til next week!
Today marked the first day of my LEEP Pioneer project! This morning I met with Professor Meyer in her research lab here on campus in the Lasry Center for Bioscience. There I was introduced to the other students who will also be working in the annelid lab this summer. We’re all working with the organism C. teleta and studying its neural development, but we’re all researching different mechanisms and processes. One student is investigating how Notch (a signaling protein crucial to neural development) affects cell fate and patterning in C. teleta. Another student is causing the expression of certain genes that produce a mutant third eye. I myself will be working with two other students to produce live imaging movies of developing neural systems, using fluorescent dyes.
After actually setting foot in the lab today, I can’t wait to really get started. Up until now, this entire project has existed only in the abstract. I’ve filled out forms, exchanged emails, had meetings… but the project never felt real until today. Everything about the lab, from the extremely mundane (sifting bowls of mud to keep the annelids comfortable) to the extremely novel (looking at live early-stage annelid embryos under a microscope) to the extremely confusing (every single research paper I’ve been given to read) has finally become a reality. It’s still going to be a while before I can start making live imaging movies, since I need to get familiar with the equipment and the organisms first. Even so, I’m looking forward to getting back into the lab tomorrow. It’s going to be a long, busy, complicated summer, but I wouldn’t have it any other way.
(Plus, I’m even getting my own lab coat. Best job perk ever.)
Til next week!
In preparation for my LEEP project, I’ve been working my way through several relevant research papers. One such paper, written by Professer Meyer herself, explores the genetic and cellular mechanisms involved in neurogenesis in Capitella teleta. To be honest, the paper was a bit over my head. Fluorescent cell tagging, the type of work I’ll be doing, was but one of many methods used to model brain development in the annelid. Several approaches, “including Dil lineage tracing, immunohistochemistry, BrdU labeling, and gene expression analyses” (Meyer & Seaver, 2009), were used to describe C. teleta‘s early neural development. Don’t ask me to try and explain what those methods actually are; despite reading the paper, I’m just about as confused as you are! Hopefully my own experiments will be a bit simpler and easier to understand. The complexities of gene expression and chemical signalling are not my area of expertise.
From what I can understand, the details of C. teleta‘s early neural development is not yet entirely understood. This annelid’s brain develops very differently compared to vertebrates, insects, and other lophotrochozoans. That said, many analogous processes, genes, and cellular mechanisms can be found. The evolutionary significance of C. teleta’s pattern of neural development has yet to be determined. It may shed some light on the neurogenesis of other annelids and lophotrochozoans, and the evolutionary history of neurogenesis in more advanced organisms.
In humans and other vertebrae, neurogenesis occurs via a process called “neuralation”. (See image below)
The neural plate, originally located on the exterior surface of the ectoderm (cells that will later become the skin and other external tissues), folds inward and is covered over by the ectoderm. The neural plate then forms into an internal tube (aptly named the “neural tube”) which develops into the spinal cord and brain.
Neural development in invertebrates is completely different. In annelids, there is no cohesive plate of neural precursor cells, nor are there mass migrations of large groups of NPCs. Rather, individual cells ingress (move inward) from the external surface of the ectoderm, migrating one by one to the internal layers of the developing ectoderm. Once there, gene expression and chemical signalling promote neural development. This model is supported by the various research methods described above by Meyer and Seaver.
Although my experiment will be limited to fluorescent cell tagging (no complicated gene expression analysis for me, thank you very much!), it’s my hope that my research will aid in further understanding the processes by which annelids’ early brains develop. By using dyes and live cell imaging, I’ll be able to observe in real time the ingression of individual ectodermal neural precursor cells. In simpler terms: I get to watch a tiny worm’s brain form right before my very eyes. That sounds pretty darn cool to me.
Til next week,
Ladies, gentlemen, those in-between, and/or none of the above: welcome! I am proud to present my LEEP Pioneer blog. Over the course of the summer, this blog will chronicle the thrilling journey of my LEEP project from confused, blundering beginnings to slightly less confused, triumphant completion. And, hopefully, I’ll uncover some prize nuggets of knowledge along the way.
First things first, let me introduce myself.
Psychology has been my passion ever since my very first term of freshman year, and my enthusiasm and fascination has only grown over time. While the human mind is too tremendously complex to ever understand completely, it’s my desire to one day understand it well enough that I can fulfill the capital ‘G’ Goals of psychology: “prediction and influence”. Whether it be through research, consultation, counseling, or any other field of work, I want to use my passion for psychology to improve the lives of others in whatever way I can.
My love of biology quickly developed as I began to branch out into neurobiology and studies of the brain itself. Though often ignored or undervalued by the psychological community at large, the physical body is equally intricate, elegant, and relevant compared to the metaphysical “mind” it houses. In fact, the dualist body/mind distinction is fundamentally inaccurate: no activity, even just thinking, is “purely” psychological. There are always underlying biological processes at work, even if we can’t yet measure, observe, or fully understand them. Therefore, as I see it, an understanding of biology is not only helpful but necessary if one is to truly understand the human mind.
This reasoning is what motivates my double-major, and my passion for neurobiology is what inspires my LEEP Pioneer project.
For my LEEP Pioneer project, I will be working with Professor Néva Meyer here in Clark’s own biology department, researching the development of neural stem cells in the embryos of Capitella teleta. As the annelid’s central nervous system is analogous to human brains and spinal cords, insight into C. teleta’s neural development can be applied to further understanding human neural development. Research of this nature has relevance in a number of fields, including neurobiology, phylogenetics, medicine, and mental healthcare. The significance and potential utility of analogous model organisms like C. teleta cannot be overstated.
It is my hope that through this project, I will be able to contribute to the larger community of neurobiological research, thus aiding (in my own small way) in the betterment of human lives. More realistically (and more quantifiably), I have a few smaller, more personal goals. My objectives include cultivating an understanding of the neural systems of annelids, developing my research & procedural skills, and refining my ability to write & present scientific material. And, of course, I aim to thoroughly enjoy myself along the way!
Here’s to the journey ahead. Cheers.
My project doesn’t officially start til the beginning of June, but until then I’ll try to provide relevant updates and background info!
Tune in next week for a brief introduction to annelids, nervous systems, and development!